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PROGRESS

IN

Nucleic Acid Research and Molecular Biology Volume

9

This Page Intentionally Left Blank

PROGRESS IN

Nucleic Acid Research and Molecular Biology edited b y

J.

N. DAVIDSON

Department of Biochemistry The University of Glasgow Glasgow, Scotland

Volume

WALDO E.

COHN

Biology Division Laboratory Oak RidgeNational Oak Ridge, Tennessee

9

7969

ACADEMIC PRESS New York and ond don

COPYRIGHT @ 1969, BY ACADEMICPRESS, INC. ALL RIGHTS RESERVED. NO PART OF THIS BOOK MAY BY PHOTOSTAT, WRITTEN

MICROFILM,

PERMISSION

FROM

BE REPRODUCED

OR ANY THE

OTHER

I N ANY FORM, MEANS,

WITHOUT

PUBLISHERS.

ACADEMIC PRESS, INC. 111 Fifth Avenue,New York,New York 10003

UnitedKingdom Edition published by ACADEMIC PRESS,INC. (LONDON) LTD. Berkeley SquareHouse,London W.l

LIBRARYOF CONQRESSCATALOQCARD NUMBER:63-15847

PRINTED I N THE UNITED STATES OF AMERICA

Listof Contributors Numbersinparentheses refer to thepageson whichtheauthorscontributions begin.

E. I.BUDOWSKY (403), Institute forChemistry of NaturalProducts, AcademyofSciences of USSR,Moscow,USSR H. FRAENKEL-CONRAT (1) , Department of Molecular Biology and Virus Laboratory and spaceSciences Laboratory, University of California, Berkeley, California D. T. KANAZIR(117) ,* Faculty ofSciences, University ofBelgradeBoris Kidrich Institute for NuclearSciences-Vintcha, Belgrade, Yugoslavia

N. K. KOCHETKOV(403), Institute of OrganicChemistry, Academyof Sciences of USSR,Moscow,USSR E. PALEEEK(31), Institute of Biophysics, Czechoslovak Academyof Science, Brno,Czechoslovakia BERNARDPULLMAN (327), University of Paris, Institut de Bwlogie Physico-Chimique, Paris, France ALBERTE PULLMAN (327), University of Paris,Institut de Biologic Physico-Chimique, Paris, France JOHN P. RICHARDSON(75), Institut de Biologie Mole culaire, Universitd de GenBve,Geneva, Switzerland TATSUYASAMEJIMA(223), Department of Chemistry, College of Science Tokyo, Japan and Engineering, Aoyama Galcuin University, B. SINGER( l ) ,Department of Molecular Biology and Virus Laboratory and SpaceSciences Laboratory, University of California, Berkeley, California P.M. B. WALKER (301), Department of Zoology, The University, Edinburgh, Scotland JEN TSIYANG (223), Cardiovascular Research Institute and Department of Biochemistry, University of California Sun Francisco Medical Center, Xan Francisco, California

*

PRESENT ADDRESS: Depnrtnirnt ofBiology, Johns Hopkins University, Baltimore, Maryland, V

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Preface In this volume of Progress in NucleicAcidResearchand Molecular Biology an attempt has been made to concentrate on articles dealing with the physicochemical aspects of nucleic acid studies, but not to the exclusion of other topics. The contributions follow our usual pattern of attempting topresent “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 persona1 expression than is normally met in review articles. T o 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. 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 or 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 again to remind readers that we wish them to write to us with their comments. Abbreviations used for nucleic acids and their derivatives are now well established by the authority of the International Union of Biochemistry and the International Union of Pure and Applied Chemistry. Those pertinent to our subject are not listed a t the beginning of each chapter, but will be found on the following pages.

J .N.D. W.E.C. Januarg, 1969

vii

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Abbreviations and Symbols All contributors to this Series are asked to use the terminology (abbreviations and symbols) as formulated by the IUPAC-IUB Commission on Biochemical Nomenclature (CBN) and approved by IUPAC and IUB, and the Editors endeavor to assure conformity. These Rules have been published in many journals ( I )and compendia ($) in four languages and are available in reprint form from the NAS-NRC Office of Biochemical Nomenclature (OBN) (31,as stated in each publication, and are therefore considered to be generally known. Those used in nucleic acid work, as set out in section 5 of the above Rules (1) and recently revised and expanded (8, 3) are given in condensed form (I-V) below for the convenience of the reader.

1. Bases, Nucleosides, Mononucleotides 1. Freebases(in tables, figures, equations, chromatograms) are abbreviated Ade, Gua, Hyp, Xan, Cyt, Thy, Oro, Ura; Pur =I any purine, Pyr = any pyrimidine, Base =any base. The prefixes S-, H,, F--, Br, Me, etc., may be used for modifications of these. (in tables, figures, or equations) are abbreviated, in the 2. Free ribonucleosides same order, Ado, Guo, Ino, Xao, Cyd, Thd, Ord, Urd (qrd), Puo, Pyd, Nuc. Modifications may be expressed as indicated in (1) above. Sugar residues may be specified by the prefixes r (optional), d (=deoxyribo), a, x, 1, etc., to these, or by two threeletter symbols, as in Are-Cyt (for aCyd) or dRib-Ade (for dAdo). 3. Mono-,di-,and triphosphates of nucleosides (5‘) are designated by NMP, NDP, NTP. The N (for “nucleoside”) may be replaced by any one of the nucleoside and 5’- are used as prefixes when necessary. The symbols given in 11-1 below. 2-, 3 -, prefix d signifies “deoxy.” [Alternatively, nucleotides may be expressed by attaching P to the abbreviations in (2)above.]

II. Oligonucleotides and Polynucleotides 1. Ribonucleoside Residues (a) Common: A, G, I, X, C, T, 0, U, 9,R, Y, N (in the order of 1-2 above). (b) Base-modified : sI or M for thioinosine = bmercaptopurine ribonucleoside ; sU or S for thiouridine; brU or B for 5-bromouridine; hU or H for 5,6-dihydrouriprefixes dine. Other modifications are similarly indicated by appropriate lower-case (in contrast to 1-1 above). (c) Sugar-modified: prefixes are d, a, x, or 1 as in 1-2 above; alternatively, by italics or boldface type (with definition) unless the entire chain is specified by an appropriate prefix. The 2’-O-methyl group is indicated by sufiz m (e.g., -Am- for 2’-O-methyladenosine, but -mA- for N-methyladenosine) . (d) Locants and multipliers, when necessary, are indicated by superscripts and subscripts, respectively, e.g., -m:A- = 6-dimethyladenosine ; -s U-or -’S- = thiouridine ; -ac‘Cm- = 2’-0-methyl-4-acetJylcytidine. ( c )When space is limited, as in two-dimensional arrays or in aligning homologous letter, the suffixes overthe sequences, the prefixes may be placed over thecapita2 phosphodiester symbol (see ref. 8, H62-63). ix

X

ABBREVIATIONS

AND SYMBOLS

2. Phosphoric Acid Residues [left side = 5’, right side = 3t (or 2’11 (a) Terminal: p; eg., pppA . . . is a polynucleotide with a 5’-triphosphate at one end. (b) Internal: hyphen (for known sequence), comma (for unknown sequence) ; unknown sequences are enclosed in parentheses. E.g., pA-G-A-C(C*,A,U)A-U-G-C p is a sequence with a (5’) phosphate at one end, a 2’:3’-cyclic phosphate at the other, and a tetranucleotide of unknown sequence in the middle. (Only codon tiplets are written without some punctuation separating the residues.)

>

3. Polarity, or Direction of Chain The symbol for the phosphodiester group (whether hyphen or comma or parenthesis, as in 2b) represents a 3’-5‘ link (i.e,, a 5’ . . . 3‘ chain) unless otherwise indicated by appropriate numbers. ‘‘Reverse polarity” (a chain proceeding from a 3 terminus at left to a 5‘ terminus a t right) may be shown by numerals or by rightto-left arrows. Polarity in any direction, as in a two-dimensional array, may be shown by appropriate rotation of the (capital) letters so that 5’ is at left, 3 at right when the letter isviewed right-side-up.

4. Synthetic Polymers The complete name or the appropriate group of symbols (see 11-1 above), enclosed in parentheses if complex, is either (a) preceded by “poly,” or (b) followed by a subscript n. The conventions of 11-2b are used to specify known or unknown (random) sequence, e.g., polyadenylate = poly A or A,,a simple homopolymer; poly (3adenylate, 2 cytidylate) = poly (A&,) or (As,C2),, a random copolymer of A and C in 3:2 proportions; poly (deoxyadenylate-deoxythymidylate)= poly d(A-T) or d(A-T),, an alternatingcopolymer of dA and dT; poly (adenylate, guanylate, cytidylate, uridylate) = poly (A,G,C,U) or (A,G,C,U)., a random assortment of A, G, C ,and U residues, proportions unspecified. The prefix copoly or oligo may replace poly, if desired. The mbscript “n” may be replaced by numerals indicating actual size.

111. Association of Polynucleotide Chains 1 .Associated (e.g., H-bonded) chains, or bases within chains, are indicated by a center dot (not a hyphen or a plus sign) separating the complete names or symbols, e.g.: poly A-poly U or (A),. (U). poly Am2 poly U or (A).*2(U), poly (dA-dC) ‘poly (dG-dT) or (dA-dC), (dG-dT) ,; also, “the adenine-thymine base pair” or “A-T base pair” in text. 2.Nonassociated chains are separated by the plus sign 2Cpoly A*poIyUl 5poly A . 2poly U poly A (II4a) or 2CA..UnI 5 An*2Un f An (114b). 3.Unspecified or unknown association is expressed by a comma (again meaning “unknown”) between the completely specified residues. Note:In all cases, each chain is completely specified in one or the other of the two systems described in 11-4 above.

-

+

ABBREVIATIONS

xi

AND SYMBOLS

IV. Natural Nucleic Acids RNA DNA mRNA; rRNA; nRNA D-RNA; cRNA tRNA

ribonucleic acid or ribonucleate deoxyribonucleic acid or deoxyribonucleate messenger RNA; ribosomal RNA; nuclear RNA “DNA-like” RNA; complementary RNA transfer (or acceptor or amino acid-accepting) RNA; replaces sRNA, which is not to be used for any purpose aminoacyl-tRNA “charged” tRNA (i.e., tRNA’s carrying aminoacyl rcsidues) ; may be abbreviated to AA-tRNA alanine tRNA or tRNA normally capable of accepting alanine, to form tRNA*’”, etc. alanyl-tRNA The same, with alanyl residue covalently attached, etc. alanyl-tRNA or Note:fMet = formylmethionyl alanyl-tRNA*’“ Isoacceptors are indicated by appropriate subscripts, e.g., tRNA;””.

V. Miscellaneous Abbreviations Pi,PPI inorganic orthophosphate, pyrophosphate RNase, DNase ribonuclease, deoxyribonuclease Others listed in Table I1 of Reference 1 may also be used without definition. No others, with or without definition, are used unless, in the opinion of the editors, they increase the ease ofreading. Enzymes I n naming enzymes, the recommendations of the IUB Commission on Enzymes, approved by IUB in 1964 (4), are followed as far as possible. At first mention, each enzyme is described by either itssystematic name or by the equation for the reaction catalyzed, followed by its EC number in parentheses. Subsequent mention may use a trivial name. Enzyme names are not to be abbreviated except when the substrate has an approved abbreviation (e.g., ATPase, but not LDH, is acceptable).

REFERENCES 1. J .Biol. Chem.241, 527 (1966) and elsewhere. 2. “Handbook of Biochemistry” (H. A. Sober, ed.), Chemical Rubber Co., Cleveland, Ohio, 1968,pp. AS9, G3-8, HlP19,H62-5. 3.In press; available, as are all CBN Rules, from the Office of Biochemical Nomenclature (W. E. Cohn, Director), Biology Division, Oak Ridge National Laboratory, Box Y, Oak Ridge, Tennessee, 37830, USA. 4. “Enzyme Nomenclature,” Elsevier Publ. Co., New York,1965. (Also in ref. a.1

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Contents LIST OF CONTRIBUTORS .

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PREFACE .

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v vii

ix xvii xxiii

The Role of Conformation in Chemical Mutagenesis

B. SINGERAND H. FRAENKEL-CONRAT

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I. Introduction . . . . . . . . . . . . . 11. Chemistry of Action of Mutagens . . . . . . . . . 111. The Reactivity and Mutability of Polynucleotides . . . . . IV. The Reactivity and Mutability of Double-Stranded Polynucleotides V. The Reactivity and Mutability of Protein-Encased Nucleic Acids . VI. Conclusions . . . . . . . . . . . . . . Addendum . . . . . . . . . . . . . . References . . . . . . . . . . . . . .

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Polarographic Techniques in Nucleic Acid Research

E. PALEEEK I, Introduction . . . . . . . . . . . . . 11. Principles of Polarography . . . . . . . . . 111. The Behavior of Low-Molecular Weight Nucleic Acid Components IV. The Behavior of Deoxyribonucleic Acids . . . . . , V. The Behavior of Polyribonucleotides . . . . . . . VI. Concluding Remarks . . . . . . . . . . . References . . . . . . . . . . . . .

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RNA Polymerase and the Control of RNA Synthesis JOHN P. RICHARDSON I. Introduction . . . . . . . . . . . . . 11. General Concepts of the Regulation of RNA Synthesis . . 111. Purity and Physical Properties of RNA Polymerase Preparations IV. The Transcription Mechanism . . . . . . . . V. Selective Transcription . . . . . . . . . . VI. The Mechanism of RNA Chain Initiation . . . . . . VII. Termination Signals . . . . . . . . . . . VIII. Inhibitors . . . . . . . . . . . . .

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CONTENTS

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111 112 116

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Radiation-Induced Alterations in the Structure of Deoxyribonucleic Acid and Their Biological Consequences

D. T. KANAZIR I . Introduction . . . . . . . . . . . . . . . I1. Evidence Favoring the Idea That DNA Is an Important Macromolecular Target for Lethal Radiation Effects in Living Systems . . . . . I11. A General Survey of the Physical and Chemical Nature of RadiationInduced Damage to DNA . . . . . . . . . . . IV . Radiation Action on DNA Replication . . . . . . . . V . Radiation Effects on DNA Transcription and on the Biosynthesis of RNA . . . . . . . . . . . . . VI. Biological Consequences Resulting from Radiation-Induced Damage to DNA . . . . . . . . . . . . . . VII . A Working Hypothesis . . . . . . . . . . . . References . . . . . . . . . . . . . . .

117 120 132 142 166 204 207 214

Optical Rotatory Dispersion and Circular Dichroism of Nucleic Acids

JEN TSIYANG AND TATSUYA SAMEJIMA I. Introduction . . . . . . . . . . I1. Optical Rotatory Dispersion and Circular Dichroism I11. Mononucleosides and Mononucleotides . . . IV. Oligonucleotides . . . . . . . . . V . Synthetic Polynucleotides . . . . . . V I . DNA and RNA . . . . . . . . . VII . Complexes of Nucleic Acids . . . . . . VIII . Visible Rotatory Dispersion . . . . . . IX . Concluding Remarks . . . . . . . . References . . . . . . . . . .

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224 225 231 238 249 262 280 289 291 295

The Specificity of Molecular Hybridization in Relation to Studies on Higher Organisms

P. M . B . WALKER I. Introduction . . . . . . . . . . . I1. Factors Affecting the Stability of the Formed Duplex .

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I11. Rates of Reassociation IV . Practical Implications of V . Conclusions . . . References . . .

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Q u a nt urn-Mec hanical I nvestiga t ions of the Electronic Structure of Nucleic Acids a n d Their Constituents

BERNARDPULLMAN AND ALBERTE PULLMAN I. Introduction . . . . . . . . . . . . . . . I1. Types of Calculation . . . . . . . . . . . . . 111. Schematic Description of the Methods of Calculation . . . . . IV. Problems Investigated . . . . . . . . . . . . V . Electronic Properties of the Purine and Pyrimidine Bases . . . . VI . Interbase Interactions . . . . . . . . . . . . VII . Problems in Radio- and Photobiology . . . . . . . . . VIII . Electronic Factors in Mutagenesis . . . . . . . . . IX. Conclusion . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . .

328 329 332 340 340 354 375 383 393 395

The Chemical Modification of Nucleic Acids

N . K . KOCHETKOVAND E . I. BUDOWSKY I . Introduction . . . . . . . . . . . . . . . 11. Possibilities of Chemical Modification in the Study of the Primary and Secondary Structures of Nuclcic Acids . . . . . . . . . 111. Importance of the Chemical Modification Methods for Studies of Nucleic Acid Function . . . . . . . . . . IV . Reactions Used for the Chemical Modification of Nucleic Acids . . V . Modification of the Uracil Nucleus with Hydroxylamine . . . . VI . Modification of the Cytosine Nucleus with Hydroxylamine . . . . References . . . . . . . . . . . . . . . AUTHORINDEX . .

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Contents of Previous Volumes Volume 1

"Primer" in DNA Polymerase Reactions

F. J. BOLLUM The Biosynthesis of Ribonucleic Acid in Animal Systems

R. M. S. SMELLIE The Role of DNA in RNA Synthesis

JERARD HURWITZAND 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

AARON BENDICHAND HERBERTS. 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 by X-Ray Scattering Techniques

VITI!ORIO LUZZATI Molecular Mechanisms of Radiation Effects

A. WACKER AUTHOR INDEX-SUBJECTINDEX

Volume 2 Nucleic Acids and Information Transfer

LIEBEF. CAVALIERI AND BARBARAH. ROSENBERG Nuclear Ribonucleic Acid

HENRY HARRIS xvii

xviii

CONTENTS

OF PREVIOUS VOLUMES

Plant Virus Nucleic Acids

ROY MARKHAM the Nucleases of Escherichia coii

I.R. LEHMAN Specificity of Chemical Mutogenesis

DAVIDR. KRIEG Column Chromatography of Oligonucleotides and Polynucleotides

MATTHYS STAEHELIN Mechanism of Action and Application of Azapyrimidines

J. SEODA The Function of the Pyrimidine Base in the Ribonuclease Reaction

HERBERT WITZEL Preparation, Fractionation, and Properties of sRNA

G. L. BROWN AUTHOR INDEX-SUBJECTINDEX

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,KENJI TAKAHASHI, AND TSUNEKOUCHIDA Chemical Effects of Ionizing Radiations on Nucleic Acids and Related Compounds

JOSEPHJ.WEISS The Regulation of RNA Synthesis in Bacteria

FREDERICKC. NEIDHARDT Actinomycin and Nucleic Acid Function

E. REICH AND I.H. GOLDBERG De Novo Protein Synthesis in Vitro

B. NISMANAND J. PELMONT

CONTENTS

O F PREVIOUS VOLUMES

Free Nucleotides i n Animal Tissues

P. MANDEL AUTHOR INDEX-SUBJECTINDEX

Volume 4 Fluorinated Pyrimidines

CHARLESHEIDELBERGER Genetic Recombination i n 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

SYOZO OSAWA 5-Hydroxymethylpyrimidines and Their Derivatives

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

H. G . ZACHAU

AND

H. FELDMANN

Uptake of DNA by living Cells

L. LEDOUX AUTHOR INDEX-SUBJECTINDEX Volume 5 Introduction to the Biochemistry of D-Arabinosyl Nucleosides

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

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

TATSUICHI IWAMURA Enzymatic Alteration of Macromolecular Structure

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

J. R. TATA

xix

xx

CONTENTS OF PREVIOUS VOLUMES

Nucleoside Antibiotics

JACK J. Fox, KYOICHIA. WATANABE, AND ALEXANDERBLOCH Recombination of DNA Molecules

CHARLESA. THOMAS, JR. Appendix 1. Recombination of a Pool of DNA Fragments with Complementary Single-Chain Ends

G. S. WATSON, W. K. SMITH,AND CHARLES A. THOMAS, JR. Appendix II. Proof That Sequences of A, C, GI and T Can Be Assembled to Produce Chains of Ultimate Length Avoiding Repetitions Everywhere

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

ROBERTWARNERCHAMBER^ The Biochemistry of Pseudouridine

EUGENE GOLDWASSER AND ROBERT L. HEINRIKSON AUTHOR I N D E X ~ U BINDEX JECT

Volume 6 Nucleic Acids and Mutability

STEPHENZAMENHOF Specificity in the Structure of Transfer RNA

KIN-ICHIRO MIIJRA Synthetic Polynucleotides

A. M. MICHELSON, J. MASSOULI~, AND W. GUSCHLBAUER The DNA of Chloroplasts, Mitochondria, and Centrioles

S. GRANICK AND AHARONGIBOR Behavior, Neural Function, and RNA

H. HYDI~N The Nucleolus and the Synthesis of Ribosomes

ROBERT P. PERRY The Nature and Biosynthesis of Nuclear Ribonucleic Acids

G. P. GEORGIEV Replication of Phage RNA

CHARLESWEISSMANN AND SEVERO OCHOA AUTHOR INDEX-SUBJECT INDEX

CONTENTS

OF PREVIOUS VOLUMES

xxi

Volume 7 Autoradiographic Studies on DNA Replication in Normal and leukemic Human Chromosomes

FELICEGAVOSTO Proteins o f the Cell Nucleus

LUBOMIRS. HNILICA The Present Status of the Genetic Code

CARLR. WOESE The Search for the Messenger RNA of Hemoglobin

H. CHANTRENNE,A. BURNY,AND G. MARBAIX Ribonucleic Acids and Information Transfer i n Animal Cells

A. A. HADJIOLOV Transfer of Genetic Information During Embryogenesis

MARTIN NEMER Enzymatic Reduction of Ribonucleotides

AGNE LARSSONAND PETER REICHARD The Mutagenic Action of Hydroxylamine

J. H. PHILLIPS AND D. M. BROWN Mammalian Nucleolytic Enzymes and Their localization

DAVIDSHUGAR AND HALINASIERAROWSKA AUTHOR INDEX-SUBJECTINDEX Volume 8 Nucleic Acids-The

First Hundred Years

J. N. DAMDSON Nucleic Acids and Protamine i n Salmon Testes

GORDON H. DIXONAND MICHAELSMITH Experimental Approaches to the Determination of the Nucleotide Sequences of Large Oligonucleotides and Small Nucleic Acids

ROBERTW. HOLLEY Alterations of DNA Base Composition in Bacteria

G. F. GAUSE Chemistry of Guanine and Its Biologically Significant Derivatives

ROBERTSHAPIRO Bacteriophage +Xl74 and Related Viruses

ROBERTL. SINSHEIMER

xxii

CONTENTS

OF PREVIOUS VOLUMES

The Preparation and Characterization of large Oligonucleotides

GEORGEW. RUSHIZKYAND HERBERTA. SOBER Purine N-Oxides and Cancer

GEORGE BOSWORTHBROWN The Photochemistry, Photobiology, and Repair of Polynucleotides

R. B. SETLOW What Really Is DNA? Remarks on the Changing Aspects of a Scientific Concept

ERWIN CHABGAFF Recent Nucleic Acid Research in China

TIEN-HSI CHENG

AND

ROY H. Do1

AUTHOR INDEX-SUBJECTINDEX

Some Articles Planned for Future Volumes Transcription and Translation i n Mitochondria

W. E. BARNETT The Ribosomal Cistrons of Higher Organisms

M. BIRNSTIEL N"-A’-lsopentenyladenosine: Biosynthesis, Metabolism, and ItsSignificance to theStructure of tRNA

R. H. HALL DNA Ligases

J. HURWITZ,M. GEFTER,AND A. BECKER X-Ray Diffraction Studies of Nucleic Acids

R. LANGRIDGEAND M. SUNDARALINGAM Induced Activation of Amino Acid-Activating Enzymes by Amino Acids and tRNA

A. H. MEHLER Natural and Artificial Regulation o f Purine Salvage

A. W. MURRAY,DAPHNEC. ELLIOTT, AND M. R. ATKINSON Synthetic Nucleotidopeptides-Models for Possible linkers in Nucleic Acids and Nucleotide Transferring Enzymes

Z. SHABAROVA Modifications of tRNA and Protein Synthesis

N. SUEOKA Species Specificity of Protein Synthesis

0. CIFERRIAND B. PARISI

xxiii

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The Roleof Conformation in Chemical Mutagenesis B. SINGER AND H. FRAENKEL-CONRAT Department of Molecular Biology and Virus Laboratory and Space Sciences Laboratory, University of California., Berkeley, California

I. Introduction . . . . . . . . . . . . 11. Chemistry of Action of Mutagens . . . . . . . . A. Nitrous Acid . . . . . . . . . . . . B. Hydroxylamine . . . . . . . . . . . C. Alkylating Agents . . . . . . . . . D. Nitrosoguanidine . . . . . . . . . . . E. Photochemistry . . . . . . . . . . . F. Brominating Agents . . . . . . . . . . G. Other Reagents . . . . . . . . . . . 111. The Reactivity and Mutability of Polynucleotides . . . . IV. The Reactivity and Mutability of Double-Stranded Polynucleotides . V. The Reactivity and Mutability of Protein-Encased Nucleic Acids . VI. Conclusions . . . . . . . . . . . . . Addendum . . . . . . . . . References . . . . . . . . . . . . .

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4 4 5 8 9 10 12

16 16 17 21 24 26 27

1. Introduction Recent reviews of the field of niutagenesis have confined themselves to selected aspects of this very extensive area of research (1-6). The limitations of the present article, derived from our chemical backgrounds, are to concern ourselves only with definitive in vitro chemical point modifications of nucleic acids, and the mutations detectable after such treatments. It then appears that the entire field, onIy 10 years old,' shrinks to a manageable volume of published data, particularly if one omits from consideration those papers in which the presumed reactions are derived from dogma, or from genetic evidence unsupported by chemical data. It is now becoming evident that reactions observed with one type of material (mononucleotide, singlestranded nucleic acid, etc.) do not necessarily occur, or represent the mutagenic event, in another type of material (double-stranded nucleic 1 We will not discuss the early isolated observations of mutants r e s d t h g from chemical treatments, such as the mustard gas studies of Auerbach etal., as reviewed recently by Auerbach [Science 106,243(1947); 168,1141(1967)l.

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B. SINGER AND H. FRAENKEL-CONRAT

acid, virus, etc.). Our experimental approach to the problem of chemical niutagenesis has impressed on us the importance of the conformation and milieu of a nucleic acid for two of its not necessarily related properties, namely its chemical reactivity and its mutability. It is these aspects of the field we particularly emphasize in this review. Before entering into a discussion of specific reagents, a few general comments on mutagenesis are in order. According to our present concept, all biologically active nucleic acids are templates, be it for replication or for transcription (DNA) or translation (RNA).Thus each change of a nucleotide in a part of the molecule carrying information represents a potentially mutagenic event. In such terms, mutagenesis can be studied without reference to biological materials. If poly C can be transformed chemically to poly U, and if in the process it loses its ability to bind poly G and instead binds poly A, then mutational events have been observed. Similarly, the ability of polynucleotides to act as templates for RNA polymerase has been used to detect chemical changes-the replacement of a C residue in poly C by a U (or T) leading to the incorporation of some A (from ATP), besides the predominant G (from GTP). Finally, the ability of polynucleotides to act as messengers and to direct the formation of polypeptides can be used, and changes in the nucleotide composition resulting from chemical mutagenesis can be deduced from the incorporation of a new amino acid. Since the base-pairing (coding) properties of each nucleotide are largely determined by the tautomeric state of the atoms bound to the 1 and 6 positions of the purines and the 3 and 4 positions of the pyrimidines2 (-N=C-XH t--) -NH-C=X), reactions that do not directly affect this

I

I

part of the molecule but that favor the unusual tautomeric form must also be regarded as mutagenic in the statistical sense, their effectiveness depending on the extent to which they favor the unusual tautomeric statea3 Other chemical modifications of the bases render them unable topair 2 Two different numbering systems have been used forthe pyrimidines, even by the 6ame authors and occasionally even in the same paper. We have translated all data into the official IUPAC-Chemical Abstracts-Ring Index system, even though this has the disadvantage, compared to the original Fischer system, that the corresponding positions on purines and pyrimidines carry different numbers. Thus the N-1 and C-6 positions of the purines correspond to the N-3and C-4 positions on the pyrimidines.

8

See also the article by Pullman and Pullman in this volume.

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3

with any other natural base, and these reactions can be defined as inactivating, broadening the meaning of this word from the customary biological to the chemical level, as we have already done with the term “mutagenic .” Finally there may exist harmless chemical changes in the bases, changes that neither interfere with base pairing nor alter its nature in either the absolute orthe statistical sense. The more classical manner of defining mutation is naturally derived from the action of mutagens on biologically active nucleic acids and the appearance of variants detected by biological or biochemical methods. The latter techniques have supplied by far the most data now at hand, but the use of simpler nucleic acids and of the isolated functions discussed above may become the more fruitful method for the study of mutagenesis. Whenever in the present discussion we use the term mutational or inactivating event we may be presumed to be thinking either in these molecular terms, or in those of the classical biologist. According to presentIy prevailing concepts, all the base triplets in information-carrying nucleic acids have phenotypic counterparts in amino acid sequences. Thus the ideal method for detecting all nucleotide replacements resulting in mutations, barring complete nucleotide sequence analysis, is to isolate all proteins coded by a nucleic acid, and determine their amino acid sequences. This has not yet been achieved for even the simpIest RNA-virus systems. However, the isolation of one gene product, namely the virus coat protein, is often easy, and the analysis for amino acid replacements in this protein has been a useful tool for the characterization of mutants in the TMV system. Unfortunately, considerable numbers of mutants must be isolated and studied before a sufficient number of amino acid exchanges is found to validate or establish the probable mutagenic event. This has been achieved only for nitrous acid deamination where by far the greatest number of amino acid exchanges support the established mutational mechanisms (C -+. U, A -+ hypoxanthine G G) (see Fig. I). The same types of exchanges have been detected after nitrosoguanidine treatment of TMV-RNA. Mutants produced by NHzOH and NH20CH3have, unaccountably, shown hardly any exchanges in the viral coat protein. The other reactions discussed in this essay present a random pattern of exchanges, suggesting the replacement of any base by any other. Since most of these reagents are rather ineffectual mutagens, the attribution of a given mutational event to the chemical employed is obviously quite dubious, and only when amino acid exchange data are available in statistically significant numbers for a given mutagen can one deduce the nature ofthe reaction from the analytical results. Such numbers of exchanges have notyet been obtained with any mutagen except nitrous acid. Such data as are available have recently been summarized (7).

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B. SINGER AND H. FRAENKEL-CONRAT

One aspect of mutagenesis that is generally not appreciated is the fact that not the absolute frequency of mutational events, but rather the ratio of mutational to inactivating events represents the true measure of mutagenesis. It must further be stressed that the observed chemical reactions need not necessarily be responsible for either the inactivating or the mutagenic events. Thus an average nucleic acid molecule may be modified in 100residues in a manner that has no biological consequence, and at the same time undergo one mutational and from 1 to 10 inactivating reactions. It would be then the level of the latter side reaction that would determine how powerful the reagent was as a mutagen. In discussing specific mutagens, we will point out examples in which such situations may exist. It should also be noted that mutagenic events are potentially coupled with lethality, since many mutations must be phenotypically lethal by producing inactive orincomplete enzymes. Thus all claims of having detected a noninactivating mutagen should be a priori discounted. Furthermore, the use of inactivation as a supposed measure of the mutagenicity of a reagent in a system where such mutagenicity has not been clearly demonstrated is obviously unjustified.

II. Chemistry of Action of Mutagens A. Nitrous Acid The dea.minating action of HNOt on adenine, guanine, and cytosine, under conditions where polynucleotides would not be degraded, was first studied by Schuster and Schramm (8). It appears that in TMV-RNA the three amino bases are deaminated at similar rates. Adenine and cytosine react normally, yielding only hypoxanthine and uracil, respectively. But guanine yields xanthine in poor yield (9), an additional unexpected minor product being 2-nitrohypoxanthine (10, ll), the others remaining unidentified. The reaction rate is a function of the concentration of undissociated HN02, and thus is greatly dependent on the pH of the solution. The discovery of the mutagenic action of HNOt by Gierer and Mundry (12) represents the beginning of our understanding of the nature of chemical mutagenesis. That deamination of cytosine t ouracil is a mutagenic event is selfevident (Fig. 1). The deamination of adenine to hypoxanthine is also mutag-enic, since hypoxanthine resembles guanine rather than adenine in its substituents at N-1 and C-6 and thus in its base-pairing properties (Fig. 1).The deamination of guanine does not affect these critical positions and thus would not be expected to be mutagenic. The data of Schuster and Vielmetter ( I S )suggest that the demination of guanine is actually inactivating. The findings that poly X is unable to hind polyC (I,$), that

ROLE

O F CONFORMATION

Inactivating events

Mutagenicevents

-

5

I N CIIEMICAI, MUTAGENESIS

HNO,

alsocrosslinking, depurination HNH

0

FIG.1. Reactions of nucleatides (anddeoxynucleotides) withnitrous acid.

xanthine (X) cannot replace guanine in reactions catalyzed by DNA polymerase (15), and that the deamination of guanine-containing polynucleotides renders them inactive as templates (16), all suggest trhat the presence of a carbonyl group in position 2 actually interferes with the binding of cytosine and therefore represents an inactivating event.

6. Hydroxylamine Hydroxylamine and its derivatives appear to react only with the pyrimidines. These reactions have been studied quite intensively. It appears very probable that only the reaction with cytosine is mutagenic. However, the nature of the chemical reaction that accounts for the mutagenicity of hydroxylamine is still somewhat in dispute. The reaction proceeds optimally and rapidly about pH 6.As shown by Fig. 2,NHzOH Probable mutagenic event

Probably inactivatine events

FIG.2.Reactions of nucleatides (anddeoxynucleotides) withhydroxylamine and methoxyamine.

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B. SINGER AND

H. FRAENKEL-CONRAT

tends to add t.0 the 5,6 double bond of cytosine2 (I), and to replace its 18).The first reaction path (reaction a and b) is amino group (11,111) (17, the faster, by a factor of about 4 (19),and reaction a had been thought until recently to be the obligatory primary reaction, as well as the mutagenic event on the basis of studies of the ability of hydroxylaminetreated poly C t o incorporate ATP along with the GTP into mixed polymers (20-2%)>. However, the recent finding of two laboratories that reaction c occurs directly and concurrently with reactions a and b (23, 24) has been confirmed by Brown and Hewlins, who determined the relative rates as about 1:4.These authors have also recently demonstrated that the preferred tautomeric form, by a factor of 10,of the hydroxyaniinocytosine is that corresponding formally to uracil oxime (111) (25). They also indicate that experimental evidence hasbeen obtained showing that this reaction product of NH20H with cytosine has the hydrogen bonding properties of uracil (25). In contrast, Janion and Shugar (26) report that neither polymers of N-4-hydroxyaminocytidine (111)nor of the dihydroxyamino compound (11) bind poly A, poly U, poly I, or poly C. However, they stress the importance of testing the activity of copolymers of cytidylic with a little N-4-hydroxyaminocytidylate as templates and, we might add, also as messengers. These authors also point out that the formation of I11 predvminates in acid solution. The conclusion, that reaction c represents the mutagenic event, is supported by the fact that in 5-substituted cytosines, in which reaction a does not occur under the usual reaction conditions (27), NHzOH reacts readily according to reaction c, and is an excellent mutagen in T-even phages (28). This reaction had been overlooked in earlier studies in which it was concluded that 5-methylcytosine does not react with NHzOH because no decrease in the typical absorbancy of the pyrimidine waa detected. While Phillips and Brown (6)are willing to accept reaction c as the mutagenic event for the T-even phages, they maintain their belief that, for nucleic acids containing unsubstituted cytosine, reaction a represents the mutagenic event. This preference is based in part on the results obtained in collaboration with Grossman (21, 29) on the effects of hydroxylamine and [W]NH20CH3on poly C. These studies showed a correlation between the extent of reaction a and the template-mutational data. However, the high and variable background of I4C-binding weakens the quantitative reliability of these data. Furthermore, since in poly C, as contrasted to monomeric cytosine compounds, reaction b isslow and product I, the “mutated” species, is believed to accumulate, one would expect this reaction to be reversible in both the chemical and biological sense, and yet

ROLE OF CONFORMATION IN CFIEMICAI> RIUTAGEKESIS

7

neither Phillips and co-workers (21) nor Wilson and Caicuts (22) were able to decrease the relative incorporation of ATP and GTP after acid treatment designed to reverse reaction a. Although Brown and Hewlins (19, ,L6)have obtained evidence that reduction of the 5,6double bond of cytosine greatly decreases the predominance of the amino form in the tautomeric equilibrium, there is no evidence nor reason to suppose that compound I would be mainly in the imino form, as would be required to account for the high mutagenic efficiency of NHzOH on the basis of reaction a. The availability of the recent data and the advantages of a unified hypothesis for all cases of hydroxylamine mutagenicity make us confident that reaction c leading to compound 111represents the mutational event. We have favored this hypothesis in discussing NHzOH mutagenesis at scientificmeetings since 1966, and we are happy to observe recent accumulation of experimental support coming from most laboratories engaged in the study of this r e a ~ t i o n We . ~ believe that our presentation of the mutagenic and inactivating events resulting from NHzOH treatment of RNA, as given in Fig. 2,has the support also of Kochetkov and his collaborators (24) (see page 403 of this volume). The action of NHzOH on uracil is most pronounced a t pH 9-10, although it occurs also at pH 6 (17,31). It results in an opening of the ring (Fig. 2) and must thus be presumed to be inactivating rather than mutagenic. There is no evidence that the reaction with uracil represents a mutagenic event. Poly U loses template activity for ADP upon NH20H Thus, the mutants treatment, but gains no new template specificities (21). found by us (32) when TMV-RN-4 was treated with NHzOH even at pH 9 are probably due to the cytosine reaction. Methoxyamine (HzNOCH3) reacts considerably more slowly with cytosine than does NHzOH (29), but it does not react with uracil (33). The mechanism of its action appears to be the same as that of NHZOH. N-Methylhydroxylamine (CH3NHOH) resembles NH20H in its reactivity (6) and its inactivating activity, but gave mutants only in one case and under exceptional conditions (see further discussion in Section IV) (34). Preliminary data indicate that this agent is not mutagenic on TMV-RNA (32). Hydroxylamine, as well as N-methylhydroxylamine, but to a much lesser extent methoxyamine, decomposes in aqueous solution and yields radicals that cause inactivation of biologically active nucleic acids (34,SS). These oxygen-dependent reactions become particularly significant a t low concentrations of NHZOH. They can be suppressed by addition of radical trappers (34, 36), but are best avoided by using fresh NHzOH solutions at

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B. SINGER AND H. FRAENKEL-CONRAT

high concentratioris and low temperature for niinirnal time periods. It appears that these side reactions cause mainly iiinctivating events (34).

C. Alkylating Agents Many reagents can introduce alkyl groups into nucleic acids in neutral aqueous solution. The extensive literature is reviewed comprehensively by Lawley in this series (4). The primary and often the only noticeable event upon treatment of nucleic acids near neutrality with dimethyl and diethyl sulfate, methyl and ethyl methane sulfonate, epoxides, mustard gas, etc., is the alkylation of guanine at N-7.Secondary sites of alkylation are adenine at N-1, N-3,and N-7 (the relative amounts of which differ for different types of nucleic acids and reagents, and are discussed in later Presumedmutagenic event

Probably inactivatlng events

J

no

R

Eventswithunknown effects: 3-alkyl-A ’I-alkyl-A Depurinatfon

FIG. 3. Reactions ofnucleotides (anddeoxynucleotides) withalkylrtting agents.

sections) and cytosine at N-3.Diazomethane a t the ether-water interphase inethylates preferentially the N-3of uracil and the N-1 of guanine, but in the squeous phase the saine products may result as with dimethyl $8).Whether diaaomethane or any of the allrylating agents sulfate (37, causes appreciable phosphate esterification remains uncertain, but doubtful. Alkylation might be expected to be, a t best, a low-level mutagen when causing substitution a t the 3 or 7 position of the purines, and to inactivate when involving their 1 position or the 3 position of the pyrimidines (Fig. 3). In studies concerning the effects of alkylation (with dimethyl sulfate, diethyl sulfate, and mustard gas) on the messenger activity of poly A

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9

(39-41) (see Addendum), the only effect noted was inactivation. The fact that poly A containing 14% of N-l-methyladenine can still bind one, but not two, strands of poly U (42)may supply a hint as to a, mechanism of inactivation of nucleic acids by alkylation. I n the case of TMV-RNA%,methylation was poorly mutagenic and all other substitutions tested (--CzF15,-CH2C€I,OH, -CH&OO-, -CH,CH,SCH&H?,CI), were nonmutagenic (43). Working with Newcastle disease virus, Thiry (&) reported dimethyl sulfate as well as ethyl ethane sulfonate to be mutagenic. When acting on T-even phsges, ethylating agents generally proved more mutagenic than methylating 4G), a fact further discussed in Section IV.With TMV and agents (45, TMV-RNA diethyl sulfate was found nonmutagenic but ethyl methane sulfonate was mutagenic (Table I), a finding that requires further study (32).

D.

Nitrosogomidine

The action of nitrosoguanidine on RNA has onIy recently been elucidated. Data primarily derived from the template activity of polynucleotides who suggested have been tentatively interpreted by Chandra et a2. (47), that alkylation occurs and that the order of susceptibility is G > A > C. Simultaneous and independent work in our laboratory has clearly demonstrated that 7-methylguanine is the main reaction product, and that much less of a methylated adenine \\;as also formed. The latter showed a white fluorescence as the nucleotide, but the site of the substitution has not yet been definitively identified ( 4 8 ,49)(see Addendum). Only under special conditions, discussed below, does nitrosoguanidine react with 48). 1-Methyladenine has since cytosine, methylating the 3 position (32, also been detected as B product of this reaction. The fact that alkylation of guanine by nitrosoguanidine is appreciable only in polynueleotides is a strong indication that its mechanism of action is not the same as that of typical alkylating agents, which readily alkylate mononucleotides. The relative levels of methylation of guanine and adenine also differed for these reagents (see Addendum) (48). The action of nitrosoguanidine on TMV-RNA caused a similarly low ratio of mutational to lethal events (@) as did dimethyl sulfate (32) (Table I), quite in contrast to the effectiveness of nitrosoguanidine when acting on metabolizing On the other hand, the finding that the latter was bacteria or cells (5U-52). a very good mutagen when acting on intact TMV rather than on its RNA represented a major stimulus to our interest in the topic of conformation dependence ofmutagenesis, and this aspect of the action of nitrosogunnidine is discussed in more detail in Sectioii V.

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H. FRAENKEL-CONRAT

TABLE I RELATIVEMUTAQENICITY OF MUTAGENS ACTINGON TMV-RNA IN VARIOUSCONFIQURATIONS (3.8) Mutagenicitya -~~~

Reagent Nitrous acid (pH 4.5) Hydroxylamine (pH 6) Methoxyamine (pH 6) N-Methyl hydroxylamine Dimethyl sulfate Diethyl suIfate Methyl methane sulfonate Ethyl methane sulfonate Ethyl ethane sulfonate Nitrosoguanidine Bromine Thiopyronin light Ultraviolet light

+

RNA in water

RNA in formamide

34 (8) 11 (8) 12 (8)

1 (2) 3.5 (7) 1 (5) 5 (6) 4 . 5(3) 2.1(5) 2 (6)

7 (3) 5 (3) 2 (16)

RNA in virus 110(5) 1 . 3(2) 1 . 6(1)

3 (4)

2 . 5(6) 1 (5)

4 (6) 7 (6) 9 (3)

8 (3) 3.6(3) 60 (15) 1 (2)

3 (2)

The following test was used: After reaction with the mutagen, at neutrality unless stated otherwise, the RNA was reisolated, or in the case of virus, separated from the protein, and tested for residual infectivity (with or without reconstitution with TMV protein) on the quantitative local lesion host, N . tJmcum var. Xanthi nc. Both the mutated sample and a mock-treated control were applied at equal concentration of infective particles (0.1-1 p g / d for fully active reconstituted TMV) to N . sybestris on which only certain TMV mutant strains produce local lesions (in numbers approximately proportional to concentration). The ratio of the local lesions found with the mutated RNA to the control (spontaneous or endogenous mutants) is termed “mutagenicity.” Such data have relative quantitative significance, but obviously do not measure all mutational events. The figures in parentheses are the number of samples, each ofwhich has been assayed 3 or more times on 9 leaves of N . sylvestris.

E.

Photochemistry3

Many dyes, particularly flavines, are effective mutagens for living bacteria or their phages, but these effects, largely deletions and insertions, are not within the scope of this review, since their mode of action is not clear, beyond the concept that many of the flavines may be readily intercalated between the bases of double-stranded DNA (53). Thiopyronin, methylene blue, and to a much lesser extent the flavines, serveas photocatalysts and sensitize nucleic acids toward visible light (54-68). These photoreactions are specific for guanine residues, causing the 68422).Illumination of various polynucleotides imidazole ring to open (65, leads to losses in amino acid incorporating activity only in the case of

ROLE OF CONFORMATION

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11

polymers containing guanine, but not specifically for amino acids requiring guanine in their codons (61, 62). However, these techniques would not necessarily reveal rare mutational changes in guanine residues. No degradation products are released from RNA even after extensive treatment (57). Since this reaction is somewhat mutagenic for TMV-RNA (57) (Table I) one may assume that the tautomeric state at N-1 and C-6 can be affected by this reaction, unless another, not yet detected, reaction actualIy represents the mutagenic event. It should be noted that the number of guanine residues lost exceeded the number of inactivating events in these experiments. The action of light in the presence of iron (Fe3+) also causes rapid inactivation and the appearance of some mutants (Sa, 63). I n chemical terms, no clear specificity of action has been detected, although the pyrimidines are more affected than the purines. Release of small-molecular material, partly in the form of bases, has been noted (63). It is possible that the mutants in this case should be attributed to base deletion although another possibility is considered below. The effects of Fe3f plus light could be greatly augmented by low levels of HzOz and are probably due to the formation of free radicals. These reactions do not seem to be of particular interest, because of their nonspecificity and the high ratio of lethal to mutagenic events. The action of ultraviolet light on nucleic acids, viruses, and microorganisms has been studied very intensively and has been reviewed in detail by J. K. Setlow (64). (See also the article by R. Setlow in Volume 8 of this series.) As far as its chemistry is concerned, the hydration of the 5,6double bond of the pyrimidines is one predominant reaction (6548), and the dimerization of neighboring pyrimidines another (69,70). Grossman (?I,72)studied the effect of UV irradiation on poly U and that on poly C with particular reference to the coding and Ono et al.(73) template activities of these polymers, respectively. As a first consequence of irradiation, the polymers lose much of their coding as we11 as their template activities, i.e. , they are inactivated. Surprisingly, irradiated poly U acquired a limited capability to stimulate the incorporation of serine, as that of phenylalanine declined. Conditions favoring the reversal of the reaction, dehydration of the 5,6 double bond (pH 8.3,85"C1 15 min) partly reversed both phenomena. No serine incorporation was observed in similar experiments by Logan and Whitmore [quoted by Johns et al.(SS)], and Setlow (64)suggests that the results might be due to the action of undesirable enzymes in the incorporation system. When the template activity of UV-irradiated poly C was tested, it appeared that the decrease in polymerization of GTP was followed, a t higher UV doses, by increased incorporation of ATP. This new capability was reported to be completely abolished by heating. The latter results

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B. SINGER AND H. FRAENKEL-CONRAT

would seem to givestrong support to the authors’ preferred interpretation of the data, namely that the hydration of the 5,6 double bond of cytosine represents the mutagenic event, possibly by causing the tautomeric shift at N-3,C-4. However, if hydration of the double bond caused the tautonieric shift of amino to imino in cytosine, one could hardly expect the same reaction to cause the opposite shift, keto to enol, in uracil to account for the serine coding activity detected by Grossman in irradiated poly U. Deamination has definitely been shown to occur upon UV irradiation of cytosine compounds and would be regarded as the most likely explanation for mutations resulting from such treatment (68, 74). On0 et al.(73)found no evidence for the formation of uracil under their conditions of inactivation of template activity, but it is not clear whether the formation of uracil had been eliminated a t the level of irradiation required to obtain “mutagenesis” (ATP incorporation). The deamination of cytosine hydrate or of cytosine dimers, which occurs very readily, was definitely not eliminated, and it would seem preferable to attribute the mutagenic events caused by UV to the known ready deamination of these photoproducts, were it not for the remarkable claim of Ono et a2. (73)that the mutation of poly C is completely reversed by dehydrating conditions. Since there is no evidence that the particular heated samples that had lost their ability to incorporate adenylate continued to act as template as far as guanylate is concerned, one wonders whether they could possibly have been overheated (70 , 15 min), and inactivated by extensive dimer formation, or degraded to a point where they lost all detectable activity. This question relates to that previously discussed, concerning the mutagenic event after NHzOH treatment. Since both UV and KHzOH readily came the saturation of the 5,6double bond, one would expect them to be equally good mutagens if this reaction were the mutagenic event. Since NHzOH is an excellent mutagen (IS, 28, ?5), whereas UV is a quite poor one in phage (64) and TMV-RNA (32) and has been found not to mutate DNA ( 76, (Tables 77) I and II), this consideration seems to supply additional support for the belief that reactions at (2-4 that occur readily with NHzOH (-NH? -+=NOH) and only as occasional side reactions after irradiation (-NH2 =O), represent the actual mutagenic events. An interesting combination of reactions was recently reported by Small and Gordon (SO), in that the UV product of cytidine, 5,6-dihydro-6-hydroxyoxycytidine1 reacts much more rapidly with hydroxylamine than cytidine, to yield the N-4-hydroxylamino derivative. KO tests of the mutagenesis of this reaction have as yet, been reported. --j

F. Brominating Agents Bromi~iation of TMV-RKA

was fourid to result in mutants (78). According to Rrammer ($9)the most clear-cut results were obtained with

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13

bromine, which a t pH 7 reacts r:q)idly md, in limitiiig amounts, almost exclusively with cytosine, while around p1-i 9, it affects mainly guanine. The first reaction is nil addition of BrOII to the 5,6double bond (Fig. 4). This product loses water at acid pH's, yielding, 5-bromocytosine which in turn can be further reacted to yield 5,.5-dibromo-6-hydroxy-hydrocytosine. In alkali, 5-bromo-0-hydroxy-hydrocytosine may lose HBr, forming 5-hydroxycytosine.

?

,

R

FIG.4. Reaction of cytosiiie with 1)romirie-water at pH 7.

A current comparative study of the mutagenicity of various agents on TMV-RNA indicates that broniinatioti is more mutagenic than all other reactions with the exception of HNOz and NHzOH (32)(see Table I).It remains to be established whetJherthis is due to tautomeric shift or ionization resulting from reaction a, b, or d, or t o other causes. Studies concerning the mutational specificity of these reactions are in progress. An extensive literature exists on the incorporation of Fi-deoxybromouridine into DNA and of 5-fluorouracil into RNA (1-,5). The substitution of thymidine by bromodeoxyuridine leads to many mutants while the sub-

TABLE 11

MUTAGENICITYAND Mutagen Nitrous acid

Hydroxylamine PH 6

ACTION

h f O D E OF CHEMICAL

TXV-RIiA

Denaturedb transforming DNA

Alkylation

TMV

Very strong (19) Deaminat ion G.A > C

Strong (88-91) Deamination G>>C > A cross-liking

Deamination C >A

Strong (76) C 4 substitution C-5.6addition

Very strong

Negative (SI)

Strong (52) C-4 substitution C-5,6addition Very weak (43,S.S) (metbylation) G-7 > A-1 > (2-3.A-7 > A-3

Verystrong (13)

3

,;:;~itu;m Methoxyamine

OF V A R t O U S

Strong (34, 96, 96) C-4substitution C 5 , 6addition

c

1

-

Negative (88)

MUTAGENS ACTINGON 813, +X174 Phage

VARIOUS NUCLFJC ACIDS IN VARIOUS STATESa

T2,T4 Phage

Positive (113,126) Strong (ffa,f23) [G,A > Cl Deamination G H M C >>A

>>

positive ($6, 116) C-4 substitution G5,6addition

c

1

Positive ($6) C-4 substitution C-5.6 addition

1

c

Very weak ($1) Weak (116) (methylation) (ethylation) G [G-7> A-1)

Vary strong (IS,Z8) C-4 substitution

Bacteria Positive (ISO)

(46, 46, 116)

(ethylation) G-7 > A-3

Denatured poly d(A-T) (template) positive (81), deamination. croas-linking Polynuoleotides (messenger) positive (16)

-

PoiyC (template) positive ($1, 32) C-3 substitution C-5,6addition

-

Poly C (template) positive (29) C-4substitution C-5,6addition

[C-4 substitutionl Strong (114) Strong

Polynucleotida

Positive (121)

Poly A (messenger) inactivated (39,41) (see Addendum) Poly U, G (messenger) inactivated (40) G-7 Poly C (template) positive (see Addendum)

Nitrosoguanidine Very weak (48,32) Positive (96) G-7 > A-1 > C-3, A-7 > A-3 Dyes and light

Weak ( 6 7 )

-

Strong (@.as) G-7,C-3 > A

-

Negative (32)

-

Negative (111)

Weak (218)

-

Very strong (60-68)

Polynucleotides (messenger) inactivated except poly A mutated (47) IG > A,Cl

Weak (116, 117)

Positive Polynucleotidea (messenger) (122-134) inactivated (67, 61,62,127)

Weak (64, 110)

Weak (64) Poly U (messenger) positive (71, 78), negative (68). hydration. dimeriaation Poly C (template) positive (78) hydration [deaminationl

G

G Ultraviolet

Very wcak (SI)

c.u

Negative (76. 77)

a The adjectives to denote mutagenic efficiency are based on our attempt to evaluate quite heterogeneous data [except for TMV-RNA4and TMV. where comparative data by one method are available (see Table I)]. The references refer only to the use of each reagent as a mutagen. The chemical reactions given are the major chemical events observed, a8 reported by various investigators, and not necessarily the mutational events. When no chcrnical data are available, we have used our judgment in giving the c1 probable reactions in brackets. 01 b The data quoted in this column refer to DNA that is more or less denatured by the use of low pH (4.2) or high temperature (60"-75OC). For reasons discussed in the text, no data on the modification of truly native DNA seem to exist in the literature, since the reaction conditions are generally not favorable for the maintenance of complete double-strandedness, except for the alkylation and UV irradiation of transforming DNA at neutrality and a t ambient temperature (91). Under these conditions, methylation is considerably more mutagenic than UV irradiation.

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stitution of much of the uracil of viral nucleic acids by fluorouracil produces very doubtful mutagenic results. Since these effects axe produced in vivo, with the analog usually being present during the intracellular replication of the DNA or RNA, these studies do not fall within the domain of in vitro mutagenesis to which we are confining this discussion. On the other hand, the study of the template activity of a poly d(A-BrU) cannot here be disregarded. Trautner etal.(80) found that DNA polymerase copying such a polymer caused the incorporation of a sniall but definite amount of deoxyguanosine. Howcver, nearest neighbor analysis did not show the expected scquences, with the deoxyguanosine being regularly adjacent to broinodeoxyuridine residues, since it was fourid next t o guanosine and adenosine in 42 arid 17y0of the cases. The interpretation of the nature of mutants resulting from DNL4 containing bromodeoxyuridine is similarly complex. It seems that a high coritent of the latter favors nonspecific errors in replication, and the same may be the case, with much lesser frequency, for fluorouracil in RNA. It appears reasonable that halogenation of cytosine gives it more of a uracil-like character, without the reverse (bromodeoxyuridine approximating cytjdine) being true or expected.

G. OtherReagents A considerable number of reagents used as mutagens have not yet been discussed. The chemical events for some of these have been elucidated. All of them seem to be less effective mutagens in vitro than those discussed in detail in the preceding sections. They all may be surmised to act mutagenically either by favoring the rare tautomeric form of the bases or by effectively deleting a base. Since none of these reagents have been found to show unexpected conformation dependencies, we list only some of the better known ones and their probable mode of action, without discussing them in detail. Most of these reactions have been discussed in other reviews (1-5). These agents are: peroxides and peracids (giving purine N-oxides) ;weak acid and high temperature (depurination, C -+ U) ; X-rays (pyrimidines); hydraeine arid other amines (cytosine); nitroso compounds (alkylation?); formaldehyde, manganese and other metal ions.

111. The Reactivity and Mutability of Polynucleotides The preceding discussion of potentially mutagenic reactions is in part based on model experiments performed with the four nucleotides, nucleosides or bases. Obviously the resct,ivities of tdie free bases, lnckjngthe pentose substitution at N-1 or N-9, respectively, can differ from those of the nucleosides and nucleotides, but these iieed riot coiiccrn us here.

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One clear exception to the usefulness of nucleosides and nucleotides as models for nucleic acids was our finding that nitrosoguanidine does not react to a detectable extent with guanosine, guanylic acid, GpU, or CpG, We will but does react readily with guanine in polymer linkage (48). therefore now consider the reactivity of the polynucleotides with various reagents. Polynucleotides are both larger and different in character from their monomers. Their long-chain nature with multiple diester bonds introduces chain breakage as a potential side reaction, which can complicate the interpretation of inactivation data. But more important is the fact, which until recently has been often overlooked, that the bases in all oligoand polynucleotides tend to stack themselves and interact with their neighbors by pi electron interaction.3 Thus all single-stranded homo- or heteropolynucleotides have more or less “structure.” The result is that most modification reactions are slower in the polymer than in the monomer. For instance, NHzOH causes no detectable loss in UV absorbance of poly C in several hours (20). Even though this was shown to be in part due to the fact that the initial loss of absorbance is compensated by the hyperchromic effect resulting from “denaturation” of the polymer by the reagent, the determination of the amount of product 111 (Fig. 2)after acid treatment confirmed the fact that the reaction rate is actually lower in the polymer. Other instances of this were reported by Kotaka and Baldwin (81) with HNOz and by Pochon and Michelson (8.2) for various alkylating agents. Quite in contrast to the generally slower response of polynucleotides, the action of nitrosoguanidine leading to methylation of guanosine was The fact that the initial observed, as stated, only in polynucleotides (48). rate of inactivation by nitrosoguanidine shows a maximum a t about 20°C and sharply declines at 37 ,as it does in dispersing solvents such as formamide and particularly dimethyl formamide, suggests that a binding of the reagent between the stacked bases is a prerequirement for the reaction (49). The question whether base-stacking favors or affects other reactions of nucleic acids is under continuing study. The observation that alkylation of adenosine and adenylic acid may yield products substituted at 1, 3,and 7,but that in poly A only 1-methyladenine has been reported to be formed in appreciable quantities, indicates that this reaction is also affected by base stacking (see Addendum).

IV. The Reactivity and Mutability of Double-Stranded Polynucleotides The most significant and singular structural feature in the field of nucleic acids is the double-strandedness resulting from the interaction of

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the complementary pairs of purine and pyrimidine bases (A with T or U, G with C or hydroxymethyl-C). The multiple bonds resulting from the interaction of two complementary strands, as it occurs in typical DNA and some viral RNA's, gives this structure considerable stability. The same type of interaction occurs in random fashion over short complementary segments of single-stranded RNA or DNA but it then has considerably less stability. As expected, the interaction of the substituents on the 1 and 6 positions of the purines with those on the 3 and 4 positions of the pyrimidines interferes markedly with all chemical reactions involving these atoms. This is most clearly demonstrated by a reaction we have not previously discussed because of its dubious mutagenic significance, i .e., that with formaldehyde. This reagent adds readily to all amino groups of RNA but fails to react with double-stranded DNA (83, 84) unless very high concentrations are employed (85).Thus reaction with formaldehyde has frequently been used as a means of ascertaining the extent of double-strandedness of polynucleotides, as well as for the purpose of preventing renaturation of complementary chains (86, 87). The most effective mutagen for RNA, HN02, has proved less useful for DNA because of the obvious fact that the amino groups of double-stranded molecules are not available to the reagent. However, HNOz is necessarily used in acid solution, i.e., under conditions where DNA begins to lose its structure. Thus the published results of nitrous acid treatment have been somewhat ambiguous. It seems that those who found HNOz not to cause many mutants in transforming DNA (88, 89) used a slightly higher initial pH and lower buffering capacity than customary, and noted that the reaction mixture became less acidic. On the other hand, HNOz was found to be highly mutagenic when a lower pH of 4.25 was maintained; these are conditions where the reaction may be demonstrated analytically (90, 91). I n relative reactivity, guanine is much more extensively deaminated than cytosine, and adenine is most resistant under conditions of incipient acid denaturation (90). Similar results were obtained by Schuster and Vielmetter (13) with calf thymus DNA. They are also in line with the finding of Kotaka and Baldwin (81) that poly d(A-T) could not be noticeably deaminated at pH 4.25 until the temperature was raised to over 60O"C.Free guanosine has recently been reported (11 )to react faster with HNO2 then does adenosine, while free cytidine reacts more slowly (A.M. Michelson, private communication). In contrast, the relative rates observed with transforming and phage DNA were G >> C >> A. This suggests that the third hydrogen bond of the G . C pair (Bainino .2-carbonyl) is much the most sensitive toward acid denaturation, followed by the typical G . C bonds, and that the A ' T linkage is definitely the most stable under these conditions.

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However, it should again be stressed that native DNA is quite resistant to nitrous acid and that side reactions, such as depurination (81), chain breakage, and cross-linking, as first pointed out by Geiduschek and coworkers (92,93), probably account for most of the biological effects resuking from nitrous acid treatment of DNA. This was also evident from the study of the nitrous acid reaction on poly d(A-T) (81). The frequently mentioned hydroxylaniine reaction is also greatly affected by double-strandedness. Thus, DNA reacts quite slowly (18). The single-stranded phage DNA 513 reacts much more rapidly than the double-stranded replicative form (9Sa). Hydroxylamine shows a very low level of mutagenesis on transforming DNA, unless conditions are chosen that tend t o eliminate base pairing in the DNA (94,91). Although this can be achieved by low ionic strength or dispersing solvents (ethylene glycol), heat is most frequently used t o render double-stranded DNA susceptible to hydroxylamine mutagenesis. This is an unfortunate choice, since obviously side reactions due to hydroxylamine decomposition, as well as deamination and depurination, may be greatly favored at 7O-8O0C,and may account not only for the observed inactivation but also for part or all of the mutagenesis observed under such conditions. This fact probably accounts for several surprising findings of Freese and Freese (34), which in our eyes have clouded the interpretation of the action of hydroxylamine. One of these findings, singular to heated DNA, is that NHzOH is most effective as a mutagen at pH 4.2(34). The other is that CHsNHOH is mutagenic, actually showing a greater frequency of mutation than does the generally accepted good mutagen, NHZOCHZ (methoxyamine), in these 95). It must be recalled that the latter reagent, lacking experiments (34, the free -OH group, is the only one of this group that does riot decompose to form radicals (35, 96). Since CH3NHOK has recently been found not to be mutagenic, while rapidly inactivating TMV-RNA (32)(Table I), it would appear advisable to omit its effects on DNA a t 70"from considerations of the mechanism of NHzOH mutagenesis. Another striking example for the protecting action of double-strandedness, in this case clearly affecting the "backside" of the pyrimidine molecule (the 5,6double bond) is the much slower rate of ultraviolet hydration of poIy C . poly I, or t,he double-stranded deoxy polymer, as compared with the corresponding single-stranded polymers (97, 98). As previously mentioned, transforming DNA is also not very susceptible to mutation by UV irradiation (76, ’27, 91). Another class of reactions showing a qualitative difference resulting from double-strandedness is alkylation. While the N-7 of guanine is always similarly highly reactive, the most reactive position on adenine in RNA and in poly A is N-1 and only that position is said to react with diniethyl sulfate (41) and methyl methane sulfonate (go), although the very similar

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chromatographic properties of 1-methyladenine and 7-methyladenine make it difficult to rule out the formation of the latter (4).When increasing amounts of poly U are added to poly A, complementary binding prevents the N-1 methylation of the adenine residues (99). In comparative experiments of the reactivity of different nucleic acids toward dimethyl sulfate by Lawley and Brookes ( I O O ) ,these authors found appreciable amounts of 3-methyladenine and only traces of 1-methyladenine in DNA, whereas in denatured DNA, like RNA, it is the N-1 of adenine and to a much lesser extent the N-3 of cytosine that is methylated. Why the 3 position of adenine is alkylated in DNA but not in poly A . poly U remains to be determined, but it must be noted that dimethyl sulfate was used for the DNA experiments, while methyl methane sulfonate was used with poly A . poly U (see Addendum). As previously mentioned, methylation of adenine at N-1 may be presumed to be an inactivating rather than a mutagenic event, since the part of the bases protected by double-strandedness is obviously also the template-active part during the replication of both single- and doublestranded nucleic acid. One is tempted t o use these facts to explain the observations that alkylation of RNA is weakly mutagenic compared to that of DNA. Thus one may assume that the alkylation of the N-7 of guanine is not inactivating but occasionally mutagenic, presumably by an increase in the unusual enol tautonier; however this mutagenic effect is depressed in RNA by the concurrent inactivatirig alkylations of the positions critical for replication In contrast, the latter on adenine and cytosine (N-1 and N-3,re~pectively).~ reactions do not occur in double-stranded DNA and thus the occasionally mutagenic effect of the guanine methylation becomes the predominant result of alkylation. On the basis of these considerations, one would predict alkylation to be the only effective mutagenic reaction for transforming DNA under conditions where it is completely double-stranded. This deduction has now been experimentally established by Bresler etal.(91), although it semis riot to be supported by the data of Wilhelm and Ludlum (40) , who found no significant incorporation of adenine-requiring amino acids by methylated poly U , G. It appears probable that an occasional miscoding, of the order of 1% of the 7-methylguanine, as postulated, would not be detectable by this method. There remains the question, why methylation was the most mutagenic of all alkylations in RNA (@), while the ethylating agents were generally found to be the most effective mutagens in double-stranded DNA. It has been suggested that the 7-ethyl group in deosygunnosine is more effective than the methyl group on the basis of greater stability, in that the deoxy‘See Addendum, p.26.

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riLosyl bond at N-9 is vcry lubile w l i ~ i :iL incthyl group quaternizes the N-7 positioti, but :tccording to ittost :iuthors it is less cffected by an ethyl 101). Thc f x t that l h s l e r el al.(91) found diniethyl substituent (100, sulfate to be a good mutagen for transforming DNA may be due to either the low pH used for reaction (pH S.2), or to more rapid testing of the alkylated samples. As a consequence of the protecting action of double-strandedness, side reactions play a much greater role in DNA and may account for a great part of the inactivating and mutagenic events obtained with reagents affecting the amino groups. One of these is the cross-linking caused by which niay lead to extensive depurination nitrous acid in DNA (92, 93), (81) and excisions of genetic material ( l o g )and , thus to mutations almost unrelated to the known chemistry of deamination. The cross-linking action of bifunctional alkylating agents (e.g., mustard gas) leads to similar consequences, unrelated to alkylation per se, which are favored in double stranded as compared to single-stranded DNA (4,103). Single-stranded DNA is more readily inactivated by bifunctional alkylating agents (104). In line with the above working hypothesis (Fig. 3),we suggest that alkylation of adenine may contribute inactivating events in addition to depurination (103). Nitrosoguanidine also seems to be a more effective mutagen on transforming DNA (96) than on single-stranded TMV-RNA suggests that native (48) although the use of elevated temperatures (60’) DNA is not mutated by this reagent. However, the difference in the reaction conditions employed with DNA and RNA do not justify analogies as regards the mechanism of mutagenesis, since nitrosoguanidine is quite unstable even at 25".

V. The Reactivity and Mutability of Protein-Encased Nucleic Acids We consider here the three classes of viruses that have been more or less intensively studied in terms of their reactivity and mutability, namely (a) TMV, (b) 'I'-even phages, and (c) small single-stranded DNA phages. Then we briefly discuss the mutagenic reactions to which resting bacteria have been subjected. We must realize, however, that it is always difficult to establish with certainty that the action of any chemical on a particle as complex as a bacteriophage, and particularly a microorganism, represents a true in vitro effect, as contrasted with conditions in which the reagent can be retained by, or bound in, the particle so as to produce its effects concurrently with subsequent metabolic activity. (a) The RNA encapsulated in the TMV particle is in general less reactive than free TMV-RNA, so that reactions usually must be performed under more rigorous conditions or for much longer time periods to obtain

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coinparable levels of inactivation. However, there is also at least one definite qualitative difference that has iiiteresting consequences, the nonreactivity of the guanine residues in the TMV particle toward HN02. Thus Schuster and Wilhelm (9) found the guanine content of TMV to remain unchanged at a time when about 20% of the adenine and over 30% of the cytosine had been deaminated (144 hours, pH 4.2), as contrasted to the deaminations in isolated RNA of 28,24, and 17% for G, A, and C, respectively, after 29.5 hours a t pH 4.2. Apart from the dramatic difference in the reactivity of guanine, i t is evident that the adenine is also less reactive, as compared to the cytosine in the virus than in the RNA. Since the action of HN02 on guanine is probably always inactivating, and the other two deaminations are mutagenic (Fig. l),one would expect the action of HNO2 on intact TMV to show a markedly higher ratio of mutagenic to inactivating events than results from the deamination of free RNA. This has been demonstrated by us in recent experiments, in which interference by the deaminated virus protein was ruled out by isolating the RNA from the treated virus and reconstituting it, prior to testing its residual infectivity and its level of mutagenesis, as compared to that of RNA directly (see Table I). deaminated arid reconstituted (3.2) The nature of the protecting action of the TMV structure on the guanine, and, to a lesser extent, on the adenine, residues is not known. Since only purine-containing polynucleotides seem to reconstitute readily one may surmise that these are the ones that with TMV protein (lob), interact with protein groups and become firmly bonded. It also appears logical that the protein groove holding the RNA should fit most tightly around the biggest nucleotide residues, and thus prevent access of reagents to it. This seeming nonreactivity of the guanine residues in the TMV particle may supply an explanation for the recent findings that nitrosoguanidine causes many mutations and little inactivation when acting on intact TMV, However, earlier studies quite in contrast to its action on TMV-RNA (48). (IOG’),recently confirmed by us, had indicated that the 7 position of guanine in the TMV particle, in contrast to its amino group, is available for alkylation. Since it was shown with TMV-RNA that this principal action of nitrosoguanidine, in contrast to typical alkylations, is dependent 49), it appears probable that the on the base-stacked conformation (48, alkylation of guanine in the TMV particle is inhibited by the absence of base stacking, rather than by the inaccessibility of its N-7 group to this particular reagent. As a consequence of this diminished reactivity of guanine, and possibly of adenine, to alkylation by nitrosoguanidine, it seems that a slower and highly mutagenic reaction is able to become predominant. The nature of this reaction has not yet been identified.

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Comparatively much 3-methyl cytidirie is formed in TMV, as it is formed if RNA (or poly C) is treated with nitrosoguanidine in dimethyl formamide, and the possibility that this is the mutagenic event has not yet been ruled out. On the other hand, the possibility that deamination of adenine is responsible for the mutagenicity of nitrosoguanidine is suggested by the results of Chandra etal.(47’). Hydroxylamine reacts so slowly with intact TMV, in contrast to that it is usually quoted as not inactivating several spherical viruses (lor), TMV at all. This reagent causes no mutants when acting on TMV (32). It is therefore suggested that the observed slow inactivation results from the decomposition products of NHzOH arising during several days at 37 C. We can offer no explanation for the failure of the cytosine residues in TMV to react with NHzOH (at pH 6), while being quite reactive to HNOs (at pH 4.5)and probably also to HC'HO (at neutrality). (b) In view of the strong protecting action of the TMV protein on its RNA, it is surprising that the T-even phages are quite susceptible to many mutagenic agents. This is also in distinct contrast to the unreactivity of double-stranded DNA or polynucleotides. Freese and Strack (94) have offered an interpretation for this phenomenon, namely, that the multiple folds necessary to accommodate the T-even DNA in a shell 1/600its length render small segments of the molecule effectively single-stranded. The guanine residues of T2 appear to react much more with HN02 than do the cytosine residues, and the adenine residues seem least reactive (IS). This is the same result obtained under similar conditions with transforming DNA (go), and thus indicates that the bulk of the DNA in T2 shows a pattern of variably low reactivities similar to that of double-stranded DNA. This is also indicated by the fact that the T-even phages are not noticeably affected by formaldehyde (83). Hydroxylamine is 1000 times inorc effective as a mutagen on T4 than on native B.subtilis transforming DNA (94) and even more effective than HN02. This can be explained on the basis that NHzOH acts atpH 6 only on the hydroxymethykytosine residues of the phage, and almost only according to reaction c (Fig. 2), and that therefore almost every chemical event is mutagenic, whereas with HNOz the concurrent deaminations of A, C, and G give comparable numbers of mutagenic and inactivating events. (c) The number of papers dealing experimentally with the chemical results of modification of double-stranded DNA's, including the T-even phages, is regrettably small, but this number is high compared to that dealing with the chemical modification of single-stranded DNA and/or the phages containing such DNA's. Mundry (108) arid Tessman (128) compared tjhe effect of H N 0 2 arid NI1,OH on intact $,X174and its DNA,

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and found no difference resulting from the presence of the protein coat. Otherwise we seem to have only deductions concerning the chemical events caused by agents used for reverse mutation, based on supposed primary mutagenic events. These deductions, often derived from in vivostudies, are frequently equated with chemical data, notwithstanding the fact that some of the conclusions based on genetic evidence are difficult to reconcile with known chemical facts. We do not review these papers, since we are not convinced ofthe validity of such deductions. We only advocate that more research effort be devoted to anaIytica1 studies of the results of mutagen treatment of single-stranded DNA, double-stranded DNA, and phages or viruses containing these two t-ypesof riucleic acid. We believe that the availability of such data would be of great benefit for our understanding of mutagenesis. (d) The invitro studies on mutagenesis in bacteria cannot readily be correlated with those on simple or more complex nucleic acids. Most typical mutagens have been successfully used with microorganisms with the seeming and surprising exception of NHzOH and its analogs (fO9). Although quantitative comparisons have not been made, it appears that the DNA in bacteria, like that in the phages, is more susceptible to mutagenic agents than isolated native transforming DNA (109). If this is true, it would suggest that DNA in its natural habitat is, in part, less “native” On the other hand, it is also possible that enough reagent than in vitro. remains in the cell to produce effects during the later growth and/or replicating phases, instead of acting strictly on noninetabolising bacteria. One clear example of the great difference in sensitivity of bacteria as compared to pure nucleic acids is nitrosoguanidine, which even at high concentrations is a poor mutagen for RNA (48). Under such conditions, it is apparently quite a good mutagen for transforming DNA (3), but is highly mutagenic for bacteria a t very much lower concentrations (51) (Table 11).

Vl. Conclusions I n Figs. 1-3 we have summarized the chemistry of the three most extensively studied types of mutagenic reactions to illustrate the working hypothesis proposed by Watson and Crick (11U) that mutagenesis requires a tautomeric shift in the N-1, C-6 region of the purines or the N-3,C-4 region of the pyrirnidine~.~ We have for the purposes of this argument listed our interpretation of the hydroxylamine and alkylation data, although we recognize that this is an arbitmry choice, and that future research may prove us wrong. We w i s h to stress ngairi in this context that the efficiency of a mutagen is dependrnt, not ordy on the extent of tautomeric shift it produces, but on the ratio of the frequencies of the mutagenic to the

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inactivating events occurring simultaneously, and such quantitative evaluations are not possible for most experiments. We have attempted to summarize in Tables I and I1 most of the mutagenesis, with particular reference to literature dealing with invitro nucleic acid conformation and milieu. In Table 11, we have attempted quantitative evaluations of the various mutagens acting on the various systems, but any such attempt leads to arbitrary choices, which should not be taken too seriously. We have also indicated the relative reactivity of the bases, without indicating the reaction mechanisms (which has been done in , when available, on the chemical data of the authors, and Figs. 1 4 ) relying, giving our judgment of the modified bases in parentheses when such data were not available. It must be stressed again that mutagenesis is usually observed at an early stage of the reaction, before the base changes reach the level of analytical detectability for most reactions. It will be noted that in Table I1 there is only one column for transforming DNA, and that this is headed “Denatured Transforming DNA.” The absence of a column for native transforming DNA is due to the lack of pertinent data. Double-stranded DNA cannot be treated with nitrous acid, which requires a pH below 5, without incipient denaturation, and its nonreactivity at pH 5 (IS)indicates that double-strandedness is critical in this respect. The same is true for NH20H, which has therefore generally been used under incipient denaturing conditions. Also nitrosoguanidine was used at elevated temperatures (96) presumably because it did not produce mutants at lower temperatures. The only reaction one would expect to be effectiveon genuinely double-stranded DNA, alkylation, has very recently been shown to be quite effective as a mutagen for transforming DNA (91). The last column contains results obtained with synthetic polynucleotides, acting as templates for RNA polymerase, or as messenger in the cell-free amino acid incorporating system. We have there used the terms “positive” and “inactivated” to denote when a polymer acquires a new function (a mutagenic event), and when it only loses some of its activity as template or messenger. From a survey of this table, it is evident that much more remains to be done. It is also evident that no extrapolation from one system to another (mononucleotides, polymers, or single-stranded nucleic acids, doublestranded polymers or nucleic acids, intraviral nucleic acid) is justified, since in many instances the differences in the behavior of nucleic acids in different milieus is not only quantitative but qualitative. We have seen some extraordinariIy sweeping and definitive statements about the mode of action of mutagens in our survey of the literature, many nil effort to avoid this attitude, and to of them erroneous. We havc nintle emphasize instead how iniich and what remains to be done, rather than what we, or others, believe to be known.

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We summarize the established facts pertaining to reactivity and mutability of various classes of nucleic acids as follows: Single-stranded polynucleotides or deoxypolynucleotides: generally less reactive than monomeric form, with the exception of methylation of guanine and possibly adenine by nitrosoguanidine, which requires basestacking; readily mutated by nitrous acid, hydroxylamine and methoxyamine, bromine, and to a lesser extent by many other agents. Double-stranded polynucleotides and deoxypolynucleolides: the hydrogenbonded regions (N-1, C-6 of purines; also C-2 of guanine; N-3,C-4 of pyrimidines) generally less reactive than in single-stranded polynucleotides; the N-7of guanine and the N-3of adenine equally and more reactive, respectively; the 5,6 double bond of pyrimidines less reactive than in single-stranded polymers; generally resistant to mutagenesis, except by alkylation under denaturing conditions. Protein-encapsulated single-stranded RNA ( T M V ):the amino groups of guanine and to a much lesser extent of adenine unreactive; the N-7 of guanine reactive to typical alkylating agents, but less to nitrosoguanidine; cytosine unreactive only toward hydroxylamine; readily mutated only by nitrous acid and nitrosoguanidine. Protein-encapsulated double-stranded D N A (Tb,T4):generally similarly resistant as free double-stranded DNA t o modification in the hydrogenbonded regions of the bases; reactive to typical alkylating agents; readily mutated by all highly mutagenic agents, including ethylation, probably because of the existence of single-stranded segments.

ADDENDUM Recent studies (32) of the alkylation of poly A using W-labeled dimethyl sulfate have shown that about 5 and 10% of the total alkylation is on the 3 and 7 positions, respectively. I n poly (A,U) the latter dkylations remain unchanged, only the 1position becoming quite unreactive. I n RNA the proportion of 3, and particularly, 7 substitution as compared to 1-methyl A, is much greater. Similar studies with [14C]nitrosoguanidinehave shown that this reagent acts similarly on poly A and the adenine residues of RNA, yielding allthree possible derivatives. A recent report on the template activity of poly C containing 6.3% of 3-methylcytosine showed incorporation of uridylic acid in the polyguanylic acid. The mechanism of this mutagenic event, which made cytosine code like adenine, is not understood [D. B. Ludlum and R. C. Wilhelm, J .B i d .Chem.243, 2750(196S)l.

ACKNOWLEDGMENTS We wish to thank Dr. L. Hirth of the University of Strasbourg, France, for his hospitality and interest during the gestation period of this review, during which we were partly supported by research grants GB 3107 and GB 6209X from the National Science Foundation and by National Aeronautics Space Adrninist,ration grant, NsG 479.

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We are also iiidebted to Doctors P. Brooltes, P. L). Lawley, T. H. Jukes, A. M. Michelson, and D. Shugar for reading the manuscript and making helpful suggestions.

REFERENCES 1. D. R. Krieg, This series 2, 125 (1963). 2.L. E. Orgel, Aduan.Enzymol. 27,289(1965). 3. E.Freese and E. B. Freese, Radiation Res.Suppl. 6, 97 (1966). 4.P. D. Lawley, This series 6,89 (1966). 6.U. Winkler, Fortschr. Botun. 28, 172(1966). 6. J. H. Phillips and D. M. Brown, This series 7,349 (1967). 7. H. Fraenkel-Conrat, in “The Molecular Basis of Virology” (H. Fraenkel-Conrat, ed.). Reinhold, New York, 1968. 8. H. Schuster and G. Schramm, Z. Nuturforsch. 13b, 697 (1958). 9.H. Schuster and R. C. Wilhelm, Biochim. Biophys. Acta68, 554(1963). 10. R. Shapiro, J. Am. Chem.SOC.86, 2948(1964). 11.R. Shapiro and S. H. Pohl, Biochemistry 6, 448 (1968). 12. A. Gierer and K. W. Mundry, Nature 182, 1457(1958). 13. H. Schuster and W. Vielmetter, J .Chim.Phys.68, 1005(1961). 14.A. M. Michelson and C. Monny, Biochim. Biophys. Actu129, 460(1906). f5. M. J. Bessman, I. R. Lehman, J. Adler, S. B. Zimmerman, E. S. Simms, and A. Kornberg, Proc. Natl. Acad.Sci. U.S. 44,633 (1958). 16.C. Basilio, A. J. Wahba, P. Lengyel, J. F. Speyer, and S. Ochoa, Proc. Natl. Acad. Sci.U. S. 48, 613(1962). 17. D. W. Verwoerd, H. Kohlhage, and W. Zillig, Nature192, 1038(1961). 18. D. M. Brown and P. Schell, J. Mol.Biol. 3,709 (1961). 19. D. M. Brown and M. J. E. Hewlins, J .Chem.SOC.( C )p. 1922(1968). 20.D. M. Brown and J. H. Phillips, J .Mol.Biol. 11, 663(1965). 21.J. H. Phillips, D. M. Brown, R. Adman, and L. Grossman, J. MoZ. Biol. 12, 816 (1965). 22.R. G. Wilson and M. J. Caicuts, J. Biol. Chem.241, 1725(1966). 23. P. D. Lawley, J. Mol.B i d .24, 75 (1967). 24.N. K. Kochetkov, E. 1. Budovsky, E. D. Sverdlow, R. P. Shibaeva, V. N. Shibaev, and G. S. Monastirskaya, Tetrahedron Letters p. 3253(1967). 26.D. M. Brown, M. J. E. Hewlins, and P. Shell, J .Chem.Soc.( C )p.1925(1968). 26. C. Janion and D. Shugar, ActaBiochim. Polon. 16,107 (1968). 27. C. Janion and D. Shugar, ActaBiochim. Polon.12, 337 (1965). 28. E. Freese, E. B.Freese, and E.Bautz, J. MoZ. Biol. 3,133(1961). 99. J. D. Phillips, D. M. Brown, and L. Grossman, J. MoZ. Biol. 21, 405 (1966). 30. G. D. Small and M. P. Gordon, d .Mol.Biol. 34, 281(1968). 31. H. Schuster, J .Mol.Biol. 3, 447 (1961). 32.B. Singer and H. Fraenkel-Conrat, In preparation. 33.N. K. Kochetkov, E. I. Budowsky, and R. P. Shibaeva, Biochim.Biophys. Acta68, 493(1963). 34. E. B. Freese and E. Freese, Proc. Natl. Acad.Sci.U.S.62, 1289(1964). 36. E. Freese, E. B. Freese and S. Graham, Biochim. Biophys. Acta123,17 (1966). 36. J.H. Van de Pol and G. A. Van Arkel, Mutation Res.2,466(1965). 37. A. K. Kriek and P. Emmelot, Biochim. Biophys. Acta91, 59 (1964). 58. A. K. Kriek and P.Emmelot, Biochemistry 2, 733(1963). 59. D.B. Ludlum, R. C. Warner, and A. J. Wahba, Science 146, 397 (1964).

28

B. SINGER AND

H. FRAENKEL-CONRAT

40. R. C. Wilhelm and D. B. Ludum, Science 163, 1403 (1966). 41. A. M. Michelson and M. Grunherg-Manago, Biochim. BiophTys. Acta91, 92 (1964). 42. J. H. Chen and F.F. Davis, Biochem. Biophys. Res.Commun. 20, 124 (1965).

43. H. Fraenkel-Conrat, Biochim. Biophys. Acta49, 169 (1961). 44. L. T. Thiry, ViroZugy 19, 225 (1963). 4.6. A. Loveless, Proc. Roy.SOC.B160, 497 (1959). 46.E. Bautz and E.Freese, Proc. Natl. Acad.Sci. U .S. 46, 1585 (1960).

47. P. Chandra, A. Wacker, R. Siissmuth, and F. Lingens, 2. Naturforsch. 22b, 512 (1967). 48.B. Singer and H. Fraenkel-Conrat, Proc. Natl. Acad.Sci.U.S. 68,234 (1967). 49. B. Singer, H. Fraenkel-Conrat, J. Greenberg, and A.M. Michelson, Science 160,1235 (1968). 50.J. B. Mandell and J. Greenberg, Biochem. Biophys. Res.Commun. 3, 575 (1960). 61. E. A. Adelberg, M. Mandel, and G. Chen, Biochem. Biophys. Res.Commun. 18,788 (1965). 62. F. Lingens and 0. Oltmanns, 2. Nuturforsch. 21b, 660 (1966). 65. L. S. Lerman, J .Mol.Biol. 3, 18 (1961). 54. D. Freifelder, P. F. Davidson, and E. P. Geiduschek, Biophys. J. 1, 389 (1961). 65. M. I. Simon and H. Van Vunakis, J. Mol.Biol. 4, 488 (1962). 66. A. Wacker, G. Tiirck, and A. Gerstenberger, Ndumissenschaften 60, 377 (1963). 67. B. Singer and H. Fraenkel-Conrat, Biochemistry 6,2446 (1965). 68. K. S. Shastry and M. P. Gordon, Biochim. Biophys. Acta129, 32 (1966). 69. K. S. Shastry and M. P.Gordon, Biochim. Biophys. Acta129, 42 (1966). 60.L. A. Waskell, K. S. Shastry, and M. P.Gordon, Biochim. Biophys. Acta129, 49 (1966). 61. M. I. Simon, L. Grossman, and H. Van Vunakis, J .Mol.Biol. 12, 50 (1965). 68.P. Chandra and A. Wacker, 2.Naturforsch. 2lb, 663 (1966). 65. B. Singer and H. Fraenkel-Conrat, Biochemistrp 4, 226 (1965). 04.J. K. Setlow, Current Topics Radiation Res.2,197 (1966). 65.R. L. Sinsheimer and R. Hastings, Science 110, 525 (1949). 66. K. L. Wierzchowski and D. Shugar, ActaBiochim. Polon. 8,219 (1961). 67.R. L.Sinsheimer, Radiation Res.6, 121 (1957). 68.H. E. Johns, J. C. LeBlanc, and K. B. Freeman, J .Mol.B i d .13,849 (1965). 69.H. E. Johns, S. A. Rapaport, and M. Delbruck, J .Mol.B i d .4,104 (1962). 70. K. B. Freeman, P.V. Hariharan, and H. E. Johns, J.Mol.B i d .13, 833 (1965). 71. L. Grossman, Proc. Natl. Acad.Sci.U .S. 48, 1609 (1962). 72. L. Grossman, Proc. Null. Acad.Sci.U.S. 60,657 (1963). 75. J. Ono, R. G. Wilson, and L. Grossman, J. Mol.Biol. 11, 600 (1965). 74.M. Daniels and A. Grimison, Biochem. Biophys. Res.Commun. 16, 428 (1964). 75. H. Schuster and H. G. Wittman, ViroZogy 19,421 (1963). 76.R. M. Litman and H. Ephrussi-Taylor, C m p t .Rend.Acad.Soi. 249, 838 (1959). 77. E. Cabrera-Juarez and R. M. Herriott, J .Bacterial. 86,671 (1963). 78. A. Tsugita and H. Fraenkel-Conrat, J .Mol.Riot. 4, 73 (1962). 79. K. W. Brammer, Biochim. Biophys. Acta72, 217 (1963). 80. T. A. Trautner, M. N. Swartz, and A. Kornberg, Proc. Natl. Acad.Sci. U.S.48,449 (1962). 81. R. Kotaka and R. L. Baldwin, J .Mol.Biol. 9, 323 (1964). 82. F. Pochon and A, M. Micheleon, Biochim. Biophys. Acta149, 99 (1967). 83. H. Fraenkel-Conrat, Biochim. Biophys. Acta16, 308 (1954). 84.M. Staehelin, Biochim. Biophys. Acta29, 410 (1958).

ROLE O F CONFORMATION I N CHEMICAL MUTAGENESIS

29

S. Zamenhof, H. E. Alexander, and G. Leidy, J .Exptl. Med.98, 373 (1953). R. Haselkorn and P. Doty, J .Bid.Ciient. 236, 2738 (1961). D. Freifelder and P. F. Davison, Biopfiys. J .2, 249 (1962). E. E. Horn and R. M. Herriott, Proc.Natl. Acad.Sci. L .S.48, 1409 (1962). S. H. Goodgal and E. H. Postel, Science 148, 1095 (1965). 90. R. M. Litman, J .Chim.Phys.68,997 (1961). 91. S. E.Bresler, V. L.Kalinin and D. A. Perumov, Mutation Res.6, 1 (1968). 92. E. P. Geiduschek, Proc. Xatl. Acad.Sci. U .S.47, 950 (1961). 93. E. F. Becker, Jr., B. K. Zimmerman, and E. P. Geiduschek, J .Mol.Bid. 8, 377

85. 86. 87. 88. 89.

(1964). 94. E. Freese and H. B.Strack, Proc.Null. h a d .Sci.U.S.48, 1796 (1962). 95. E. Bautz-Freese, J. Gerson, H. Taber, H.-J. Rhaese and E. Freese, Mutation Res.4,

517 (1967). 96.E. Freese and E. B. Freese, Biochemistry 4, 2419 (1965). 97.K.L. Wierzchowski and D. Shugar, Photochem. Photobiol. 1, 21 (1962). 98. R. B. Setlow, Proc. Xatl. h a d .Sci. U.S.63, 1111 (1965). 99. D. B. Ludlum, Biochim. Biophys. Acln96,674 (1965). 100. P. D. Lawley and P. Brookes, Biochem. J .89, 127 (1063). 101. P. Alexander, J. T. Lett, and G.Parkina, Biochim. Biophys. Acta48, 423 (1961). 102. I.Tessman, J.Mol.Biol. 6,442 (1962). 103. P. D. Lawley, J. H. Lethbridge, P. A. Edwards, and K. V. Shooter, Biochem. J. 106, 52P (1967). 104. N. Yamamoto, T. Saito and M. B. Shimkin, Can,cer Res.26, 2301 (1966). 105.H. Fraenkel-Conrat and B. Singer, Virology 23, 354 (1964). 106. P. Brookes and P. D. Lawley, Biochem. J .77, 478 (1060). 107. It. M. Franklin and E. Wecker, Nutzcre184, 343 (1959). 108.K. W. Mundry, impublished observations. 109.W. Hayes, “The Genetics of Bacteria and Their Viruses.” Wilcy, New York,1964. 110. J. D. Watson and F. H. C. Crick, Nature171, 964 (1953). 111. K. W. Mundry, 2. Vererbztngdehre 91,87 (1960). 112.W. Vielmetter and C. M. Wieder, 2. N n t i ~ ~ f o r14b, s c h312 . (1959). 113. I. Tessman, Virology 9, 375 (1959). 114. V. F. Chubukov and S. Tatarinova, Zliur. Mikrobiol. Epidemiol. Immunobiol. 42,80 (1965). 115. D. M. Green and D. R. Krieg, Proc.Null. Acad.Sci.U .S .47, 64 (1961). 116. D. A. Ritehie, Genetics Res.Cambridge 6, 168 (1964). 117.J. W. Drake and J. McGuire, .I.Virology 1, 260 (1967). 118. B. D. Howard and I. Tessmm, J. Mol.Hiol.9, 372 (1964). 119. J. W. Drake, J .Bacteriol. 92, 144 (1966). 120.F. Kaudewitz, Nature83,1829 (1959). 121.B. S. Strauss, J .Bacteriol. 83,241 (1962). 123.H. E. Kubitshek, Proc. Natl. Acad.Sci. Cr.S.62, 1374 (1964). 123.B. R. Webb and H. E.Kubitschek, Biochem. Biopfys. Ites. Commun.13, 90 (1963). 12.4. A. Zampieri and J.Greenberg, Mutation Res.2, 552 (1965). 125. I. Tessman, H. Ishiwa, and S. Kumar, Science 148, 507 (1065). 126.I. Tessman, R.. I<.Podder and 8. Kumar, J .Mol.Biol. 9, 352 (1964). 127.A. Wacker, H. Dellweg, 1,.Triiger, A. Ihrnhauser, E. Lodeman, G. Tiirck, R. Selzer, P. Chandra, and M. Ishimoto, Photochem. Photobiol. 3, 369 (1964). l d 8 .I. Tessman, Virology 36, 330 (1968).

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Pola rogra phic Techniques in Nucleic Acid Research E. PALEEEK Institute of Bwphusics, Czechoslovak Academyof Sciences, BTno,Czechoslovakia

I.Introduction . . . . . . . . . . . . 11. Principles of Polarography . . . . . . . . . A. Classical Polarography . . . . . . . . . B. Modern Polarographic Techniques . . . . . . . 111.The Behavior of Low-Molecular Weight Nucleic Acid Components . A. Adenine . . . . . . . . . . . . . B. Cytosine. . . . . . . . . . . . . C. Guanine . . . . . . . . . . . . . D. Other Purines and Pyrimidines . . . . . . . . E. Adsorption of Nucleic Acid Components on the Mercury Electrode F. Analytical Applications . . . . . . . . . IV. The Behavior of Deoxyribonucleic Acids . . . . . . A. Native and Denatured DNA’s . . . . . . . . B. DNA Containing Single-Strand Breaks . . . . . . C. Changes a t Premelting Temperatures . . . . . . D. Estimation of Denatured DNA in Native DNA Samples . . V. The Behavior of Polyribonucleotides . . . . . . . A. Synthetic Polyribonucleot.ides . . . . . . . . B. Ribonucleic Acids . . . . . . . . . . . VI. Concluding Remarks . . . . . . . . . . . Reference3 . . . . . . . . . . . . .

31 32

33 36 38 39 40 41 42 43 44 45 46

54 55 60 61 61 68 68 70

1. Introduction Most methods used for studying the physical properties of nucleic acids ( I ,2) provide information primarily on the regularities in their secondary structure. With the aid of these methods, a very important “static” picture of the structure of nucleic acids has been obtained. For a better understanding of the complicated function of the nucleic acids, it would be desirabIe gradually tody namize this picture. This process should include the elucidation of the states through which the double-helical structures pass in the course of temperature changes at premelting temperatures, during pH changes in the pH regions where denaturation does not take place. For this purpose it would be expedient to take advantage of methods “inverse” to some of those hitherto used, e.g., those ignoring the regularly arranged bases in the double-helical regions but registering the small number of bases not included within these regions or departing from them 31

32

E.

PALEFEK

under certain circumstances. Furthermore, it would be useful to be able to study nucleic acid structure not only in solution, but also under conditions approaching the environment existing in living cells. The recent paper by von Hippel and Printz on hydrogen exchange in DNA (3) and the study of Miller and Bach on the interaction of DNA with the electrically charged surfaces of synthetic polymers (4, 6) are among the first moves in this direction. Other methods will be needed, however, in order to obtain detailed information on the structure of nucleic acids with respect to its dynamics. Among the methods whose application might, in this respect, yield fruitful results are those of electrochemical analysis. These can provide information on the ease with which the substance in question yields or takes up electrons, on the accessibility of the potentially reducible or oxidizable groups for the electrode process, on the behavior and some properties of the substances adsorbed on electrically charged surfaces, etc. Such information may be of great importance as the interaction of nucleic acids with the surfaces of electrically charged structures, the presence of certain groups on the molecular surface, as well as electron transfer, come into play in the course of the functioning of nucleic acids invivo. Polarography, with its widely developed theory and instrumentation, represents a potentially very useful technique in this research area. However, until recently it has been used only in investigating substances of low molecular weight. An exception in this respect is BrdiEka's polarographic protein reaction (6), which is mainly used in clinical diagnostic work (7’). Owing to the high sensitivity of this reaction, conditioned by its catalytic character, the relatively high molecular weights and low diffusion coefficients of proteins present no drawback for direct current polarographic measurements. However, a reaction of similar qualities has not been uncovered for nucleic acids. I n recent years, some modern polarographic techniques have been found to be suitable for analyzing both natural and synthetic polynucleotides and to reflect sensitively changes in their secondary structure.

II. Principles of Polarography The polarographic method was elaborated by J. Heyrovsky (8) about 45years ago. For almost 20years from its invention, polarographic research was mainly directed to interpreting the current-voltage curves obtained in electrolysis by means of the dropping mercury electrode. Later other relationships were studied with this electrode, but the original method is still widely used, being referred t o as D.C. (direct current) or classical polarography.

33

POLAROGRAPHIC TECIJNIQUlGS

A. Classical Polarography The principle of D.C. polarography is illustrated by the simple diagram , shown in Fig. 1. The polarizable dropping mercury electrode ( P ) the surface of which is continually renewed, is immersed in the solution to be analyzed. A pool of mercury at the bottom of the vessel, or a separate electrode (e.g., a caIomel one) with a surface many times larger than that A

M LINEAR VOLTAGE SWEEP GENERATOR

FIG.1. Schematic diagram of the circuit formeasuring current-voltage curves with a dropping mercury electrode. R, reference nonpolarizable electrode; P, polarizable dropping mercury electrode; C, glass capillary; M , mercury reservoir; A , currenb measuring device.

of the mercury electrode, serves as the nonpolarizable reference electrode (R). A voltage that can gradually be increased (or decreased) is applied to the electrodes from an external source, and at the same time the current is measured (Fig. 2a). As long as the potential does not reach a value at which a transfer of electrons between the electrode and a substance in the solution begins, no substantial current is observed (the small current changes occurring under this condition are due to the so-called capacity or charging current, which is connected with tQc fact that earh mercury drop falling from the capillary niiist be charged tlo n potent]ial corresponding t o thc

34

E. ~

~

~~

Response obtained a

Time

b

C

Response obtoined

Signal applied

df ‘ Idf

n Time

tf

a02sec.

ifI

I

Rrne I

FIG. 2a,

b, and c. Forlegend see facing page.

PALEEEK

Signal applied and response obtained

Response obfained

d

Time 4 .

eE

G

E

Time

FIG.2.Summary of the signals applied and the responses obtained in different polarographic techniques. half-step potential; H , height of the step; (a) D.C.pohrography. Elpotential; El/* 8,supporting background electrolyte; B,background discharge. (b) A.C. polarography. Substances adsorbed on the electrode influence the capacity ofthe electrode double layer. The curve of the analyzed substance deviates from the in the potential region where the curve of the supporting background electrolyte (8) substance is adsorbed ( A )and the differential capacity decreased. From the difference between these two curves, the degree of coverage of the electrode surface can be calculated. The desorption of a substance may be manifested by a wave (T).If the substance is not adsorbed, but it is reduced electrolytically there appears a wave (F) not preceded by a deviation from the curve of the supporting electrolyte. The wave height (R)is characteristic for the concentration of the investigated substance and the wave potential (summit potential, E W ) ,for its nature. (c) Oscillographic polarography at controlled A.C.Signal applied: n, polarization with multiple current cycles; I ,polarization with a single current cycle per each drop of mercury; ZZ, polarization with two current cycles per each drop of mercury; j, time at which the drop falls. Response obtained: The nature of the substance under investigation is characterized by the potential of the indentation (EI) and its concentration by the depth (D)of the indentation; S, the curve of supporting background electrolyte; c, cathodic part of the oscillogram; A , anodic pnrt of the oscillogram; MP,left marginal point (at the potential of this point anodic dissolution of mercury takes place); G, right marginal point (at the potential of this point, background electrolyte discharge occurs). (d) Derivative pulse polarography. Signal applied: a,periodicity of the polarkation voltage; Response obtained: b,both capacity and faradaic current; c, capacity current only; d, faradaic current only; t, time at which current is measured; f, time at which the drop falls. The wave potential (EW)is characteristic forthe nature of the substance investigated, and the wave height ( H )for its concentration. S, supporting electrolyte. In the figtire conventionally used, times, potenlials, and current values are given.

36

E. P A L E ~ E K

applied voltage; the value of this cm.rent does iwt, :is u rule, exceed lo-’ A). As soon as the potential has rexchcd a value at whic.11 some suhst:mre in the solution is reduced or oxidized (such a substaiice is called iidepol:irizer, as it removes the polarization of the electrode), there ensues a sudden current increase; the increase continues until a potential is reached when all the depolarizer in the neighborhood of the electrode is exhausted and the current is limited by the transport rate of the depolarizer. A polarographic step’ is formed and its height ( H )depends on the concentration of the depolarizer, while the potential corresponding to the half-height of the step characterizes the quality of the investigated (half-step potential El,z) substance (Fig. 2a). As a rule, analysis is carried out within the concentration range of lW3 to l e 5 M ;several substances in the solution can be determined simultaneously provided their half-step potentials differ from one another sufficiently (as a rule, by more than 100 mV). It takes a few minutes to register a polarographic curve. More detailed information on the theory, instrumentation, and analytical application of polarography are 9-11). presented in various monographs (7,

B. Modern Polarographic Techniques The limits of sensitivity of D.C. polarographic analysis are given by the magnitude of the capacity current. If the faradaic current is of the same order of magnitude as the capacity current, or is less, then its exact determination is impossible. The application of modern amplifiers was therefore not able to increase the sensitivity of the method; it is the experiments aimed at modifying the method of the dropping mercury electrode polarization that have achieved success in this respect. The efforts made at raising the sensitivity, selectivity, and speed of the polarographic measurements have produced a number of techniques that differ slightly from classical polarography. It has been possible to suppress the capacity currents and to increase the sensitivity of the polarographic method by as many as three orders of magnitude. On the other hand, it has been found that a mere change in the capacity of the electrode double layer, conditioned by adsorption processes in which no transfer of electrons between the electrode and the analyzed substance takes place, can be utilized for analytical purposes. Moreover, the time required for analysis has been substantially shortened and the possibilities of analyzing substances with slightly differing half-step potentials (resolvability) have been improved. The same applies to the estimation of substances reduced at The term “step” refers to the curve obtained by means of D.C. polarography, and the A.C.polarographic and derivative pulse-polarographic curve is called a “wave.” The term“indentation” alludes to the effect observed on the oscillogram, dE/dtagainst E , which is obtained with the aid of oscillographic polarography at controlled A.C.

POLAROGRAPHIC TECHNIQUES

37

negative potentials in the presence of an excess of substances reducible at more positive potentials (separability). A review of modern polarographic methods has been given by Schmidt and von Stackelberg (12). Mentioned here briefly are only those methods that have been used in studying nucIeic acids. 1. ALTE~CNATING CURRENT I’OLAROGRAPFIY

The principles of alternating current (A.C.) polarography were proposed by several investigators, independently on one anot,her, in the years 1938-1944. It was Breyer who elaborated A.C. polarography for practical The method employs the superposition of sinusoidal, low application (13). frequency, small amplitude alternating voltage on the direct voltage applied to the cell and follows the dependence of the alternating current on the potential of the mercury electrode (Fig. 2b). The sensitivity of the method is approximately the same as that of D.C. polarography for reversible processes, and it is substantially lower for irreversible processes. Resolvability and separability are better than in the case of D.C. polarography. Contrary to the latter method, A.C.polarography is suitable for foliowing nonfaradaic adsorption/desorption phenomena. The waves due to these phenomena are called tensammetric. More information on A.C. polarography can be found in the literature (13-15). 2.OSCILLOGRAPHIC POLAROGRAPJ-IY AT CONTROLLED ALTERNATING CURRENT

Heyrovsk? proposed a method based on the polarization of the In 1941, eIectrode with A.C. and on the following of changes of its potential by means of an oscilloscope. The method was called oscillographic polarogra17). According to the terminology introduced phy at controlled A.C.(16, later, this method belongs to chronopotentiometry. A current of sinusoidal or rectangular shape is used for polarization (Fig. 2c). Commercially available apparatus operating at a frequency of 50 c/s displays the deon E (Fig. 2c), which is suitable for analytical purposes. pendence of dE/dt An important part of the method is the so-called “first curve technique” in which a single A.C. cycle is applied to each mercury drop at a definite time after the fail of the preceding drop. By means of this technique, primary processes can be differentiated from reactions conditioned by the product of electrolysis formed in the course of polarization by the first cycle. Moreover, the sensitivity of the oscillopolarographicmethod can be increased when the depolarizer accumulates on the electrode surface during the time preceding application of the current pulse (18-81). Compared with D.C. polarography, oscillopolarographic analysis is substantially

38

E. P A L E ~ E K

faster and approximately of the same sensitivity, but it is somewhat less accurate. Like A.C.polarography, it reflects changes in the capacity of the electrode double layer, due to adsorption and desorption of substances on the dropping mercury electrode. More details on oscillographic polarography at controlled A.C.are given in the literature (22-24).

3.SQUARE-WAVE AND PULSE POLAROGRAPHY Square-wave polarography was developed by Barker and Jenkins (25) over 15 years ago with the object of establishing a method that would possess the advantages of A.C.polarography but would be more sensitive for measuring faradaic currents. In principle, it works like A.C.polarography, but instead of the superposed alternating voltage of a sine wave form it employs a rectangular one with a frequency of 225 CIS. The current is measured after the surface of the mercury drop has almost ceased growing and toward the end of the polarization periods only, whereby the capacity current resulting from the growth of the drop and from the alternating polarization is eliminated. Pulse polarography is a modification of square-wave polarography, from which it diffem primarily by the fact that a single polarizing pulse of 40 msec duration is applied at a definite time after the fall of the preceding drop (26). The current is measured over the second half of the pulse duration, when the value of the capacity current decreasing with time is almost zero (Fig. 2d). Thanks to these modifications, the sensitivity of the method has been increased, especially for irreversible processes, the capacity currents have been suppressed, and measuring in a medium with smaller conductivity has been made possible. The method has two variants, the “normal” and the “derivative” (Fig. 2d). The latter is slightly more sensitive, allowing the determination of depolarizers even in concentrations of 10-8211 for reversible, and 5 X lW7M for irreversible, processes.

111. The Behavior of low-Molecular Weight Nucleic Acid Components An important condition for the study of the polarographic behavior of polynucleotides is suf€icient information on the processes to which their low-molecular components are subjected on the electrode. The following outline of the polarographic behavior of nucleic acid components has been written from this angle and does not therefore contain data primarily of interest in the polarography of organic substances. These aspects are dealt with in a review by Janfk and Elving (27)in which flavin and pyridine nucleotides are also considered.

39

TECHNIQUES

POLAROGRAPHlC

Of nucleic acid components, it is primarily bases, either free or bound in nucleosides or nucleotides, that are subject to electrolysis on the dropping mercury electrode. D-Ribose is polarographically reducible in its aldehydic form only. In neutral medium, its concentrated solutions yield a step of Ell2 = -1.81 V against a normal calomel electrode (28). Ribose and deoxyribose bound in nucleosides or nucleotides as well as the phosphate groups are polarographically not reducible (27). Sugars or sugar phosphates can, however, influence the electron density, diffusion coefficient, and adsorbability of the nucleoside or nucleotide, and consequently may somewhat change their polarographc behavior (e.g., the shift of Ellz).

A. Adenine The polarographic reducibility of adenine was described by Heath in 1946 (29)and confirmed by a number of authors (30-34). The protonked form2of adenine is reduced according to the following scheme (34):

R

R

H

R

Scheme I

The scheme does not include chemical reactions following the electroreduction (R = H, sugar or sugar phosphate). Adenine is reducible in an acidic medium only; at about pH 5 the step begins to diminish, and a t pH 6.5it disappears. E1,2becomes more negative At higher pH linearly with the increasing pH (EL12 = 0.975 - 0.084 P H ) . ~ values, a step of adenine is formed in the close vicinity of the background electrolyte decomposition. This is due to the lowering of the hydrogen The attachovervoltage caused by adenine and its reduction product (34). ment of the sugar or the sugar phosphate moiety to the adenine nucleus influences the polarographic behavior of the parent compound only slightly; the current-controlling factors and reduction mechanisms are common in principle for the entire series (35). The reducibility of adenine is not substantially influenced even when it forms part of a dinucleotide. At pH 3.7Ellzbecomes more negative in the series dAMP 5 adenine < 8 In the course of polarographic analysis, the substance that is reduced in the protonized form can yield a step even at a p H such that the nonprotonized form is predominantly present in the solution. The latter formis reducible owingto recombination with protons in the neighborhood of the electrode (see p. 360 in ref. 9). 8 All potentials were measured against a saturated calomel e1ect)rodeunless otherwise stated.

40

E.

PALEEEK

adenosine < deoxyadenosine < AMP < ATP, the difference in Ellzof adenine and AMP being about 25 mV. The A.C. polarographic behavior of these substances (35)and the results of measuring the differential capacity by the bridge method (36,SY) suggest a generally parallel order of increasing adsorbability, i.e., adenine < dAMP < deoxyadenosine < ATP < AMP. The oscillopolarographic and pulse-polarographic behavior of adenine and its nucleosides and nucleotides agrees in principle with the results obtained by the D.C. polarographic method. At higher pH, when the D.C. polarographic step is poorly developed, the oscillopolarographic indentation or pulse-polarographic wave can be measured much better; in a neutral medium, both adenine and the derived substances are inactive even when these methods are used (19).

6. Cytosine When the polarographic reducibility of adenine was established in 1946, it was claimed that the other bases and their nucleosides and nucleotides Later results obtained by were polarographically nonreducible (29). Cavalieri and Lowy (39, and Hamer el al.(30), as well as oscillopolaro39), indicated, however, that the question of graphic measurements (3.3, cytosine nonreducibility would have to be further investigated. Evidence for the polarographic reducibility of cytosine was presented in 1962 independently by PaleEek and Janik (40) and by Smith and Elving (41). Also reducible are 5-methylcytosine1 5-hydroxymethylcytosine, and the nucleosides and nucleotides derived from these bases and from cytosine (40, 42). All these substances yield a diffusion-controlled current. On the basis of polarographic measurements and the analysis of the product of macroscale electrolysis, a scheme of cytidine reduction on the dropping mercury electrode has been proposed (42, 43)(Scheme 11).

R

R

R

SchemeI1

The scheme does not include chemical reactions following the electroreduction (R = H, sugar or sugar phosphate). Cytosine and its derivatives yield a step in an acidic and a neutral medium; in an alkaline solution the step becomes lower, and at about pK 10 it disappears (40, 4% ). Below pH 3,the steps merge with the background discharge, whereas the oscillopolarographicindentations can be well

41

POLAROGRAPHIC TECHNIQUES

44). The half-step potential measured even in a medium of 2 M H2SOI (33, becomes more negative linearly with increasing p H (Ell2 cytosine = - 1.170 - 0.084 pH) (42). Polarographic (4.2) and oscillopolarographic show that the attachment of the sugar and the sugar measurements (3.9) phosphate moiety to the cytosine nucleus exerts a relatively greater influence on the ease of reduction of the parent compound than in the ,~ more negative in the order: cytidine adenine series. A t pH 6.5, E I becomes This fact has (-1.58 V) < dCMP < CMP < cytosine (-1.70V) (42). been explained by the electron-withdrawing effect of the pentose ring; in the adenine series, the distanc,e between the sugar ringand the reduction sites is too great t o withdraw electrons significantly from the reduction sites (2 7).

C. Guanine The polarographic nonreducibility of guanine has been established in several studies (27, 29,45). Very suitable for analyzing guanine and its nucleosides and nucleotides is oscillopolarography, which yields a sharp 39)around the anodic indentation on the curve of d E / d tagainst E (33: This indentation can be observed in a buffered potential -0.2 V (Fig. 3).

~

~~

~

~

~

E

-

FIG.3. Oscillogram of deoxyguariylic acid. , 2 X 10-4 M deoxyguanylic acid , background in 0 . 5 M ammonium formate with McIlvaine buffer (pH 5); electrolyte. The dropping niercury electrode was polarized with A.C. of - rectangular shape and frequency 50 c/s. The “first carye technique” was used. ME , right marginal point; a t the potential of this point the guanine moiety is reduced. ~

42

E. P A L E ~ E K

medium within the acid and the neutral regions; in the neighborhood of pH 7,the indentation begins to diminish and it disappears at about pH 9 (43). Nearly all measurements have so far been made in an ammonium formate medium, in which the anodic indentation is well developed. Investigations of the oscillopolarographic behavior of a number of purine derivatives have revealed that further substances having substituents in position 2 and 6, but with a free 7 and 8 position, such as 2,Gdioxypurine (xanthine), 2,6-diaminopurine, Zmethyl-6-hydroxypurine, and 2,6dioxy3-methylpurine (theophylline), yield similar anodic indentations. Substances that do not satisfy the above conditions do not yield indentations of this kind [e.g., 6-chloropurine, 6-oxypurine (hypoxanthine), 6-aminopurine (adenine), 2,G-diosy-3,7-dimethylpurine(caffeine), 3,S-dimethylxanthinel (43, 46). On the basis of oscillopolarographic studies and D.C. polarographic measurementswith a hanging drop, it has been presumed that a t a potential of the right marginal point (Fig. 3) a reduction product is formed that is adsorbed in the course of the polarization of the electrode to positive potentials and then oxidized while an anodic indentation is formed (46). The results of macroscale electrolysis and measurements by means of The products Kalousek’s switch have corroborated this presumption (47). arising by the reduction of guanine, guanosine, and guanylic acid are unstable, arid under certain conditions their oxidation regenerates the original substances. The site of reductiori has not yet been determined with certainty; the doubIe bond between N-7 arid C-8 of the imidazole ring is probably hydrogenated (48). The fact that guanine and its derivatives do not yield a D.C. polarographic step is evidently connected with the closeness of the potential of its reduction to the background electrolyte discharge. In agreement with this, there has been observed a shift of the background discharge to positive potentials by about 100 mV in the presence of these and a cathodic oscillopolarographic indentation in the close substances (4 7) vicinity of the right marginal point in a strongly acidic medium (46).

D. Other Purines and Pyrimidines Uracil and thymine are not reducible polarographically (4?9,41,45),but in an alkaline medium they yield an anodic step conditioned by the formation of a mercury conlpound (49). I n agreement with polarographic behavior in an alkaline medium, the two substances yield indentations in the cathodic and the anodic part of the oscillogram (dE/dt versus E ) close to the left marginal point (33, 39). Deoxyuridine, uridylic acid, thymidine and thymidylic acid do not yield such indentations (51). Hypoxanthine is polarographically reducible only in an acidic medium (30, 34); its step is found in the vicinity of the background electrolyte

POLAROGRAPHIC

43

TECHNIQUES

discharge and it is poorly developed (46). On the contrary, the sharp indentation yielded by hypoxanthine on the oscillopolarographic curve is The electrode process responsible for the step of well measurable (33). hypoxanthine proceeds according to Scheme I11 (34) :

I

H

I

H

Scheme III

The scheme does not include protonation and chemical reactions that follow the electroreduction. Xanthine is not reduced by the dropping mercury electrode within a normally available potential range (45), but on oscillopolarographiccurves, xanthine and xanthosine yield anodic indentations similar to that of guanine (33,44, 46).Compared with the indentation of guanine, the potential of that of xanthine is somewhat more positive. Derivatives of purine and pyrimidine that do not occur in nucleic acids naturally but that can be incorporated into them frequently contain W-Br, and polarographically reducible groups, such as -N=N-, others. However, the polarographic behavior of these substances has so far been little studied. 6-Azauracil and 6-azauridine give a reduction D.C. polarographic step; it is probabIy their protonized forni that is reduced by AzauraciI, azauridine, 9-asaguanine, the dropping mercury electrode (50). and 5-bromouridine yield characteristic indentations on oscillopolarographic curves (44,51).

E. Adsorption of Nucleic Acid Components on the Mercury Electrode

As a rule, molecules adsorbed 011 the surface of a dropping mercury electrode reduce the differential capacity of the electrode double layer. By measuring the differential capacity, it is possible to obtain information on surface activity, the orientation and association of molecules on the electrode surface depending on its potential (52). By means of the A.C. polarographic and the bridge methods, it has been established that all seventeen purine and pyrimidine derivatives under investigation are adsorbed on the dropping mercury electrode. Derivatives that are usual components of nucleic acids (36, 63)show the greatest tendency to intermolecular associations on the surface. Derivatives occurring only in sume nucleic acids (e.g., 5-hydroxy~nethylcytosine) or those that can be in-

44

E.

PALEEEK

corporated into nucleic acids (e.g., 8-azaguanine) show a much smaller tendency to intermolecular associations (64), while bases that are not incorporated into nucleic acids do not associate on the electrode surface (36). It has been found that the rate of association is much smaller than Nucleosides usually occurring in Iiucleic acids that of the adsorption (55). are also adsorbed. At a sufficiently high concentration, adenosine, deoxyadenosine, guanosine, deoxyadenosine, as well as deoxycytidine show a tendency to association on the surface of the dropping mercury electrode in the same way as the corresponding bases. On the contrary, no associations of this kind have been observed with uridine, thymidine, and cytidine (37). The adsorption of nucleotides has not yet heen &died systematically.

F. Analytical Applications The advantages of classical polarography cannot be fully utilized in the analysis of nucleic acid components. The steps of adenine, cytosine, and hypoxanthine arise in the vicinity of the background discharge so that it is rather difficult t o measure them, and consequently the accuracy of the method is decreased. Further disadvantages are due t o the D.C. polarographic inactivity of guanine and to the vicinity of Elpof the respective reducible substances, which renders their simultaneous determination impossible. Better result8 may be obtained by the method of oscillographic polarography at controlled A.C., which yields sharp indentations on curves of dE/dtagainst E and makes it, possible to determine two or more sybstarices simultaneously. For example, nlixtures of adenine and cytidine, of adenine and hypoxanthiiie, of cytosine and deoxycytidine, etc., can be analyzed. The presence of uracil and thymine does not interfere with the analysis. The anodic indentation yielded by guanine and its nucleosides and nucleotides ran be utilized in determining these polarographically nonreducible substances. Oxygen need not be removed from the solution prior to oscillographic measurement and the very act of measuring takes only a few seconds. If substances adsorbing on the electrode are present in the analyzed sample, they may influence the shape of the oscillopolarographic curve rather than that of the D.C. polarographic step. The pulse-polarographic technique is suitable for analyzing samples of this kind (if estimation of guanine is not involved). Although the application of polarographic techniques for analyzing nucleic acid components is in some cases more suitable than the more frequently employed optical density measurements in UV light (especially when analyzing mixtures of substances absorbing within the same region), these techniques have so far been relatively little utilized. Several studies have been devoted to de57), in different tissues (58, 59), and in termining adenine in the blood (56,

POLAROGRAPHIC

45

TECHNIQUES

RNA hydrolyzates (60) by means of D.C. polarography. Azauracil and its derivatives have been analyzed (51, 61, 62j, and the radiolytic splitting of has been followed (63) by the same method. The oscillocytosine in vitro polarographic method has been used for studying the enzymatic deaminatioii of adenosine, cytidine, and guanosine in bacterial cultures (64479, for analyzing solutions of purine and pyrimidine bases (52) and investigating changes induced in them by UV light (67-70), for determining adenine in , for identifying chromatographic spots (33) RWA hydrolyzates ( 7 1 )and and fractions (72).

IV. The Behavior of Deoxyribonucleie Acids The fact that the diffusion coefficients of nucleic acids of high molecular weight are very lowinduced some investigators to believe that the study of nucleic acids by methods of electrochemical analysis should be restricted 7.4). It has been proved merely to following their adsorption behavior (73, by means of various techniques, including classical polarography, that the above view was riot justified (76-79). If the electrode process ill classical polarography is controlled by diffusion, the height of the polarographic step can be calculated from the IlkoviE equation for the limiting diffusion current:

I d = 0.627x n x F

xcx

1Y2x m2/3 x t'/'j[A]

(1)

where n is the number of electrons taken up or delivered by a single depolarizer molecule during the electrode process, F the Faraday charge of 9.65x lo4coulomb, c the concentration of the depolarizer in mo1/cm3, D the diffusion coefficient of the depolarizer in cm2/sec, m the flow rate of mercury in gm/sec, and t the drop time in seconds. For native DNA with a sedimentation coefficient of 22S, a diffusion coefficient of 1.3X lo-* cm2/ see is reported, and that of denatured DNA is given as 7 X 10+ cm2/sec (80). Among nucleotides that are components of nuclleic acid, only dCMP is polarographically reducible at pH 7 (it is reduced by 4 electrons) (Section 111,B). Provided that DNA yields a diffusion-controlled current and that all residues reducible in a monomeric form are also reducible in DNA, then according to Eq. (1j bhe mean limiting diffusion current with 1 mM DNA-P would be approximately 0.15PA with native calf-thymus DNA and 0.36PA with denatured DNA (the following capillary constants were gm/sec, t = 2.8sec). considered: m = 3.76X Under favorable condit,ions, denatured DNA yields a D.C. polarographic step roughly corresponding to the calculated value, while native 7 7 )Because . the currents DNA is not reducible polarographically (’75, arising in the course of electrolysis of concentrated solutions of denatured

46

E.

PALEEEK

DNA are rather low, the D.C. polarographic technique does not appear to be suitable for practical analysis. Much better possibilities are offered in this respect by the pulse-polarographic technique and by oscillographic polarography a t controlled A.C. in conjunction wit,h the “first curve technique” (Section 11, B, 2).The latter method allows one t o follow, in addition to processes usually observed in D.C. polarography, the behavior of the guanine residues in the DNA molecule, which yield an indentation in the anodic part of the oscillogram.

A. Nativeand Denatured, DNA s The height of the D.C. polarographic step (around 0.6PA) yielded by denatured DNA in a concentration of 500 pg/ml in a medium of 0.5 M ammonium formate with 0.1 M sodium phosphate, pH 7.0(Fig. 4b) provides sufficient evidence for the electrolyt,ic nature of the process responsible for its formation (77). The behavior of denatured DNA haa ‘78) and oscillobeen studied also by means of pulse polarography (76, graphic polarography at controlled A.C., as well as by A.C. polarography (It?,? @. Allthese methods have produced further evidence for the fardaic character of the electrode process. The potentials at which DNA exhibits the respective effect (step, indentation, or wave) at neutral pH are roughly within the range of - 1.4to - 1.6V, depending on the method used as well as on the conditions of measuring. The magnitude of the current measured at the given DNA concentration depends on the quality and concentration of the salts in the background electrolyte. Most measurements have been carried out in a medium of 0.1-0.5 M ammonium formate with a phosphate buffer, which is suitable for all the techniques mentioned above. 1. D.C.

POLAROGK4PHY

At low salt concentrations and in the absence of ammonium formate 7.0) denatured DNA yields, even in a high concentration, only a very small D.C. polarographic step (ca. 0.1 PA), the presence of which may be conditioned either by a mere change in the capacity of the electrode double layer or by a very low faradaic current (74, 77). When the concentration of ammonium formate is increased, the step grows, Ellz shifts to more negative potentials, and the step Other salts, e.g., ammonium acetate, ammonium changes its shape (77 ). chloride, calcium chloride, act similarly to ammonium formate (19); however, the efficiency of divalent cations is much greater (e.g., 0.05 N CaClz produces roughly the same effect as 0 . 5 N HCOONH,). The dependence of the step height on the concentration of denatuied DNA is linear ( 77). Under the same conditions, concentrated native DNA solutions do not yield any reduction step (Fig. 4a). (e.g., in 0.1 M sodium phosphate, pH

POLAROGRAPHIC

47

TECHNIQUES

b

I

I

I

I

I

I

I

I

-0.8 -1.0

-1.2 -7.4 -1.6

-0.8 -7.0

I

I

-12 -1.4 -1.6

f ofenfial I VI

C

I

I

t

I

I

-0.8 -7.0 -1.2 -1.4 -1.6

I

-0.8

I

I

-1.0 -1.2

I

I

-1.4 -1.6

Potential I V I

FIQ.4. Polarograms of native and denatured DNA’s in 0.5M ammonium formate with 0.1 M sodium phosphate (pH 7).D.C. polarograms: (a)Native DNA at a concentration of 500 pg/ml. (b) Denatured DNA at a concentration of 500 pglml. A.C. , Native DNA at a concentration of 500 pg/ml; backpolarograms: (c) , Denatured DNA at a concentration of 500 pg/ml; ground electrolyte. (d) , background electrolyte. Measurements were performed on A.C. and D.C. polarograph GWP 564. The amplitude of A.C. voltage was 18 mV, frequency 78 c/s. Poteiitials were measured against the saturated calomel electrode.

-

-

~

48

E. P A L E ~ E K

D.C. polarography has been used also for studying DNA interaction with daunomycin (81), Cu2+, Cd2+ (80), and the anionic (Cu EDTA) complex (829, as well as for following ion transport through adsorbed monolayers (83). 2.A.C. POLAROGRAPHY AND THE MEASURING OF DIFFERENTIAL CAPACITY BY THE BRIDGEMETHOD

On A.C. polarogranis, both denatured and native DNA yields a wave in the neighborhood of -1.2V (wave 1) (Fig. 4c, d) for the formation of which neither the presence of ammonium formate nor a high salt concentra77,84,85). This wave is of a nonfaradaic character tion is necessary (73-75, corresponding to DNA desorption, as was observed earlier by Miller (86). Around - 1.4 V, denatured DNA yields another wave (wave 2) (74,84,85) that is, similarly to the D.C. polarographic step, better developed in the presence of ammonium formate (Fig. 4d) or at a higher salt concentration and is, partly at least, of a faradaic character (77). When the ammonium formate concentration is increased, wave 2 grows and shifts to more negative potentials, while wave 1 is shifted to more positive potentials, its height showing only a slight change. DNA adsorption on the dropping mercury electrode has also been studied by measuring the differential capacity of the electrode double layer by means of the bridge method (86). The measurements were carried out in a nonbuffered medium at pH 6.0. It has been found that DNA is adsorbed on the electrode and that it decreases the differential capacity of the electrode double layer within the region of potentials more positive than about - 1.2V. DNA is desorbed at - 1.2 V; at more negative potentials than about -1.3 V, DNA does not interact with the electrode surface and does not influence the capacity of the double layer. The areas per nucleotide were determined at full surface coverage: 93 A2 per nucleotide for denatured DNA irrespective of the polarization of the mercury surface, 35 per nucleotide for native DNA at a negatively per nucleotide at a positively charged charged mercury surface, and 86 iz surface. These results indicate that native DNA preserves its doublehelical structure when adsorbed on a negatively charged surface, whereas unfolding occurs on a positively charged mercury surface.

A2

3.OSCILLOPOLAROGRAPHY AND PULSE POLAROGRAPHY The behavior and character of the indentation CI-1 yielded by both native and denatured DNA and of the indentation CI-2 yielded only by denatured DNA on oscillopolarographic curves (Fig. 5) roughly corresponds The so-called to the A.C.polarographic wave 1 or wave 2 respectively (75). “first curve technique” (Section 11,B, 2)can be profitably used for studying

b

a

E

-

FIG.5.Oscillograms of native and denatured calf-thymus DNA’s. , (a) Native D N A a t a concentration of 100pg/ml in 0.5M ammonium formate with 0.1M sodium phosphate (pH 6.8). (b) Denatured D N A a t a concentrationof 100p g / m l in 0.5M ammonium formate with 0.1M sodium phosphate (pH 6 . 9 , Measurements were carried out on the universal polarograph (75). The dropping mercury electrode was polarized with A.C.of rectangular shape and frequency 50 c/s. The “first curve technique” was used. CZ, cathodic indentation; AZ, anodic indentation; AZ,, anodic indentation produced by native D N A not being - present on the “first curve’’ but appearing due to repeated polarization by A.C.MP, right marginal point; if the electrode is not polarized to the potential of this point, the indentation AZ or A I ,does not appear. dE dt

1

a

b

E FIG.6. Right section of the cathodic part of the oscillogram (dE/dtversus E ) . (a) Deoxycytidylic acid a t a concentration of 160p g / d in 1 M ammonium formate with 0.05M sodium phosphate (pH 7); (b) Denatured calf-thymus D N A at a concentration of 200 pg/rnlin 1.0M ammonium formate with 0.05 M sodium phosphate (PH 7). , First curve; . . . . . . ., second curve; ___, background electrolyte only (first curve). The dropping mercury electrode was polarized with A.C.of rectangular sha,pe and frequency 50 c/s. After PaleEek (76).

-

50

E. PALEEEK

the indentation CI-2as its depth on the first curve is substantially larger than on the second and successive curves (Fig. 6 )This . technique facilitates measuring at lower DNA concentrations than those required for D.C. and A.C. polarographic measurements (18. 76). So far, pulse polarography (26)(Section 11,B, 3)has proved to be the most sensitive and accurate technique for following faradaic currents produced by denatured DNA. The lowest concentration ofdenatured DNA still yielding a measurable wave is around 1 rJg/ml (76). IfDNA is polarographed at a low sensitivity ofthe pulse polarograph and a high DNA concentration, the obtained results are analogous to the D.C. polarographic measurements. Under these conditions, denatured DNA yields only one wave (wave 111)and native DNA appears to be inactive. A t high sensitivity, additional waves can be observed. Concentrated solutions of native DNA produce a wave (wave 11)that is slightly more positive than wave 111produced by denatured DNA (Fig. 13), and another wave around - 1.2V (wave I). Denatured DNA produces, in addition to wave 111,also wave I.The nature ofwave I has not yet been studied in detail; the behavior of this wave roughly corresponds to that of nonfaradaic A.C. polarographic wave 1. Wave 111,which is specific for denatured DNA, can be compared with the D.C. polarographic step. The height ofwave 111depends on the concentration of ammonium formate as does the height ofthis step. A comparison between the pulse-polarographic waves and the effects obtained by means of other polarographic techniques is given in Table I.This table shows that while wave I and wave I11can be compared with effects yielded by other techniques, no such anaIogy is found for wave 11;more data concerning this wave are given in Sections IV,B and IV,C, 2. TABLE I COMPARISONBETWEEN THE PULSE-POLAROORAPEIIC WAVES OF NATIVEAND DENATUREDDNA AND THE EFFECTBOBTAINED BY MEANS OF OTHER POLAROGR4PHIC TECHNIQUES Technique

NativeDNA

Pulsepolarography D.C.polarography

Wave I -

Wave I1

A.C.polarography

Wave 1 (c)

-

Oscillopolarography at controlled A.C.

Identation GI-1(c)

-

-

Denatured DNA Wave I -

Wave I11 Reduction step(f)a Wave 1 (r) Wave 2 (f) or (fc) Indentation Indentation GI-1(c) CI-2(f)

Abbreviations in parentheses: (f)electrode process of faradaic character; (c)electrodeprocess of nonfaradaic (capacity) character; (fc)combmed faradaic and nonfaradaic process. Q

POLAROGRAPHIC

51

TECHNIQUES

Native DNA does not yield a wave I11at pH 7, but a wave4appears on the polarogram a t about pH 6 (Fig. 7);the potential of this wave corresponds approximately to the potential of denatured DNA in the same medium (19). The wave of native DNA grows slightly up to about pH 4 and then increases sharply with decreasing pH. On the contrary, the height of the wave of denatured DNA shows a relatively small change

3

I

I

I

4

5

6

7

P* FIG. 7.Dependence ofheight ofpulse-polarographic wave on pH.Calf-thymus DNA, 40 pg per ml in 0.2M ammonium formate with Britton-Robinson buffer. 0 0, Native DNA; @0, denatured DNA. The sensitivity ofSouthern-Harwell Pulse Polarograph was 1/40 or higher and the number of divisions was calculated for t’he sensitivity 1/40.

under the influence of pH within the acid region (Fig. 7). Similar results have been obtained by the oscillopolarographic “first curve technique” when both the cathodic (19) and the anodic indentation (Section IV,A, 5) was measured. 4 The term “wave 111 refers o n l y to the wave yielded by denatured DNA at pH 7. When the pH is changed, the wave potential is shifted and the role of the groups participating in the electrode process may also change.

52

E.

4.THE NATURE OF

PALEEEK

THE REDUCTION CURRENT PRODUCEDBY DENATUREDDNA The faradaic character of the process to which denatured DNA is subject under certain conditions on the dropping mercury electrode hm been demonstrated. The mechanism of the process is, however, not known in detail. The behavior of apurinic acid indicates that cytosine residues ’77). The dependence of the depth of indentaparticipate in the process (7’5, tion CI-2 on the G C content in denatured DNA samples analyzed at high temperature support this idea. The fact that no dependence of this kind however, may be explained on the has been found at room temperature (7’5)) one hand by the masking of reducible cytosine residues due to aggregation of denatured DNA and by the participation of other residues (e.g., adenine) in the electrode process on the other hand. The possibility of adenine residues also participating in the process cannot be excluded, in spite of the fact that monomeric adenine is reducible in an acid medium only (Section 111,A). The polarographic behavior of synthetic polyribonucleotides appears tto suggest the participation of cytosine and adenine residues (Section V, A, 1). The unambiguous solution of this problem by macroscale electrolysis of denatured DNA a t a controlled potential and the analysis of the reduction product is complicated due to adsorption of the reduction product on the electrode (19). The polarographic reducibility of denatured DNA is conditioned by the accessibility of the reducible groups for the electrode process. The nonreducibility of native DNA has been explained by the inaccessibility of these groups in the regular arrangement of bases in the Watson-Crick double helix (75). The role of background electrolyte ions consists perhaps in their influence on the adsorbability of denatured DNA, which in turn affects its reducibility. With rising concentration of ammonium formate (or NH4C1, MgC12, CaC12, etc.) the difference between the potentials of A.C.polarographic wave 1 and wave 2 is becoming greater linearly with the increase of the D.C. polarographic step (19). This fact might be explained by the presumption that only unadsorbed polynucleotide is reduced. The definitive classification of the process to which denatured DNA is subject, on the dropping mercury electrode, as a diffusion, adsorption, kinetic, or catalytic process is complicated, as the criteria used in classifying electrode processes in which low-molecular substances take part are in most cases not suited for macromolecular denatured DNA. The results so far obtained suggest that the process to which denatured DNA is subject on the electrode is in principle diffusion controlled, but it. is complicated by 19). the adsorption of the reduction product (77,

+

POLAROGRAPHIC

53

TECHNIQUES

5. ANODICINDENTATION DUE T O

GUANINE

RESIDUES

I n multiple polarization of the dropping mercury electrode with alternating current, native DNA in a niedium of ammonium formate yields an anodic indentation AI, on the curve of d E / d tversus E around -0.2V (Fig. 5 ) . The indentation becomes deeper after DNA denaturation (87,88). For the indentation to appear, the electrode - niust be first polarized to the potential of the right marginal point ( M P )corresponding to the hackground eIectroIyt,e discharge (Fig. 5 ) , as in the case of the anodic indentation of guanine and its derivatives (Section 111, C). Of the DNA degradation products, indentaticm A I is yielded by apyrimidinic acid only, while apurinic acid and other products not containing guanine do not yield this indentation (89).Consequently, guanine residues in DNA are probably responsible for the indentatinn AI. This conclusion has been confirmed by the oscillopolarographic behavior of synthetic polyribonucleotides (19). If the dropping mercury electrode is polarized by a single cycle applied to each drop (“first curve technique”) (Section 11, B, 2) instead of by multiple A.C.cycles, the indentation A1 appears only on the oscillogram of denatured DNA, while it does not appear on that of native DNA (90). The indentation of native DNA appears after more polarization cycles are applied to one drop and it grows unt,il it reaches a limiting value, which is, however, lower than the depth of indentation of denatured DNA. The depth of the indentation of denstmed DNA does not become greater with an increase in the number of polarization cycles. The behavior of native DNA indicates that on the application of the first polarization cycle to the dropping mercury electrode, the guanine residues contained in its molecule are not a,ccessible for the electrode process or the number of accessible groups is too low. A certain amount of these residues is released for the electrode process under the influence of additional polarization cycles. This release might take place either within the region of positive potentiah in agreement with Miller’s data (Section IV, A, 2),or a t very negative potentials near the right marginal point, where the guanine residues are reduced. Studies of the oscillopolarographic behavior of DNA at different pH values show that native DNA does not yield an indentation A1 on the first curve in the neutral and alkaline region. In an acidic medium, an anodic indentation can be observed, which points to the accessibility of the guanine residues for the electrode process in the course of the first polarization cycle (44). I tis interesting that in native DNA both groups responsible for the reduction process (Section IV, A ,3)and guanine residues become accessible

54

E.

PALEEEX

at acid pH values far from the pH at which DNA denaturation takes place. This phenomenon might be explained by DNA conformation changes due to the protonation of some bases. The character of these changes could be analogous to the changes found at premelting temperatures at neutral pH (Section IV, C, 2). It is only in the case of denatured DNA that the depth of indentation AI, at pH 7.0 depends on the G C content of the DNA. On the other hand, no such dependence is found in the case of native DNA samples. C contents Investigations of native DNA with approximately equal G obtained from various bacteria of the genus Bacillus show a correlation between the genetic relatedness of the bacteria and the depth of the indentations (90). By a more detailed analysis of DNA’s isolated from Bacillus subtilis and B .brevis, greatly differing from each other in oscillopolarographic behavior (Section IV, C ,l), it has been demonstrated that a difference in the depth of their indentations AI, is not due to the presence of impurities such as proteins, polysaccharides, RNA, or denatured DNA (91) or to a difference in molecular weights or the presence of single-strand breaks in the DNA double helix (90, 9%).On the electrode, the guanine residues are probably more easily released from the weak parts of the molecule, which might be represented by regions where G C pairs are surrounded by a great number of A . T pairs (90, 44).Consequently, the DNA and B. brevis different oscillopolarographic behavior of B. subtilis DNA could be explained by great differences in their nucleotide sequences

+

+

(911-

B. DNA Containing Single-Strand Breaks Concentrated solutions (0.5-1 mg/ml) of native DNA isolated from various bacteria and mammalian organs have been studied by the pulsepolarographic technique. Although some samples did not yield a wave I11 (whereby the presence of more than about 0.5% of denatured DNA was excluded), all the analyzed samples did yield a higher or a lower wave I1 (Section IV, A, 3). The latter wave was not removed either by ft thorough purification of the DNA samples or by changes in DNA isolation techniques. On the other hand, the height of wave I1 increased markedly under the influence of agents such as y-rays and DNase I, which introduce singleThe increase in the height of strand breaks into the DNA molecule (79). this wave permitted the detection of changes due to the y irradiation of DNA in a concentration of 500pg/ml with a dose of several hundred rads (93).After treatment with low concentrations of DNase I, wave TI increased in its dependence on time to a limiting value (79). The negative wave I11 appeared approximately at the same time, indicating a more

POLAROGRAPHIC

TECHNIQUES

55

complete DNA degradation. A relatively small number of single-strand breaks introduced into the DNA molecule by mild DNase I treatment (roughly several hundredths of a percent of all phosphodiesteric bonds broken), which did not result in a drop in the viscosity of the DNA solution, produced an increase in wave I1as well. However, such small changes in t.he DNA mvlecule could not he detected by the oscillopolarographic technique. Aftter a more extensive degrdation of DNA by DNase I, there appeared on the oscillopolarographic "first curve" an indentation a.pproximately corresponding by its potential t80the indent&on CX-2. The sufficient height of wave I1at elevated te~nperat~ures suggests that this wave is probatbIy of a fwadair character. The groups responsible for its format'ion could be bases in t,he virini t.y of t8hebroken phosphodiest'er bond t.hat become accessible for reduc*t,itrna t the dropping mercury electrode. This accessibility could he due to local changes in the regular arrangenient of the bases in the DNA double-helix. Such changes in the neighborhood of single-strand breaks niight either ttakc place in the bulkof the solution . these results it does not follow or seconda,rily un t.he electrode ( 7 9 )From tha.t t.he single-strand breaks arethe only source of changes in the DNA structme responsible for wave I!.Shearing nf the DXA solution in the capjlla,ry did not affect, the height o f wave 11.However, the possibility canriot be excluded that., with greakr changes in molecular weight, when the number of t,he double-&and breaks reaches a, value coniparable to the polarographicttlly detectable number of single-strand breaks, the influence of the free ends of t,he riiolecules might come into play. Moreover, the height of wave I1might be influenced by other factors locally labilizing the DNA double-helix. The introduction of a larger number of single-strand breaks into the DNA molecule also results in the deepening of oscillopolarographic indentation AI, (92). This phenomenon has been explained by an easier release of guanine in the neighborhood of the interrupted phosphodiester bond owing to niultiple polarization of the mercury electrode with A.C. (Section IV, 4, 5). The sensitivity of the oscillopolarographic detect.ion of singlestrand breaks is substantially lower than t,hat of t'he pulse-polarographic t,echnigue.

C. Changesat Premelting Temperatures 1. THE COIJESEOF DNA DENATURATION FOLLOWED BY

POLAROGRAPHIC TECHNIQUES

When following the course of thermal denaturation of DNA by measurements at room temperature after quick cooling of the sample, the de-

56

E.

PALECEK

naturation curve obtained by polarographic methods is very similar to that 90,941. obtained by observing optical density a t 260mp (Fig. 8)(18, 75,78] If, however, the measurements are performed at elevated temperatures, changes can be observed at temperatures below the melting temperature (Fig. 9). These changes have been detected in all the DNA samples isolated

c

.o

u

0

u

E

U

40 Temperature

(TI

60

80

Temperature ( ‘C

100

1

FIG.8 (Left). Thermal transition of calf-thymus D N A followed by pulse-polarographic technique atroom temperature. DNA samples were heated for 10 minutes a t thc temperature given in the graph and quickly cooled. Pulse-polarographic measurements were carriedout in the concentration of 32 &nl in 0.3 M ammonium formate with 0.1 M sodium phosphate (pH 7)at room temperature. The height of wave 111was followed at the sensitivity 1/5 or lowerand the number of divisions was calculated for thesensitivity 1/5. FIG.9 (Right). Thermal transition of DNA followed by oscillopolarographic and spectrophotometric methods a t elevated temperatures, Calf-thymus D N A a t a concentration of 95p g / d in 0.1 M ammonium formate with 0.02 M sodium phosphate (pH 7). XX, Spectrophotometry; 0 0, oscillopolarography; 00 , oscillopolarography (cooling of the sample). In oscillopolarographic measurements, the dropping mercury electrode was polarized with A.C.of rectangular shape and frequency 50 c/s. The “first curve technique’’ was used. After PaleEek (75).

from various bacteriophages, bacteria, and mammalian organs that have been investigated by means of uscillographic polarography after multiple polarization of the dropping mercury electrode with A.C.(94) and 95). Measurements were carried out in by the “first curve technique” (75, a medium of ammonium formate with sodium phosphate at pH 7.0. At temperatures 20°C or more below the temperature at which the

57

POLAROGRAPI-IIC TECHNIQUES

optical density at 260 mp begins to rise, there appears the CI-2 indentation, and its depth increases with temperature (Fig. 9). The depth of the CI-2 indentation at premelting temperatures is not influenced by DNA degradation by shearing in a capillary and is not proportional to the DNA G C content. For example the indentation of B. brevis DNA appears at substantially lower temperatures and attains greater depths than the DNA, even though the difference in the G C indentation of B.subtilis contents of these DNA s is onIy slight. Consequently, in this case there exists a certain analogy with the behavior of the anodic indentation AI,of these DNA sat room temperature (Section IV, A, 5),but contrary to this the formation of the indentation CI-2 at premelting temperatures is not, conditioned by multiple polarization with A.C. Changes in the polarographic behavior of DNA at premelting temperatures can be also observed A.C.(77,85), and pulse polarography (78). From oscilloby D.C.(77), polarographic, D.C.and pulse-polarographic measurements, it is evident that the effect under investigation is of a faradaic character, like the process to which denatured DNA is subject on the dropping mercury electrode (Section IV, A).However, the phenomena produced on polarographic curves by native and denatured DNA at premelting temperatures differ to a certain degree from one another. For example, the pulse-polarographic wave of native DNA appears at more positive potentials than the wave of denatured DNA, so that both the substances can be followed simultaneously, provided their waves are approximately of the same height (78). It has been concluded that the wave produced by native DNA at elevated temperatures is identical with the wave I1 (Sections IV, A, 3, and IV, B).

+

+

2. THE NATURE OF PREMELTING CHANGES The changes in the polarographic behavior of DNA at premelting temperatures can be explained by the release of reducible groups that, at lower temperatures, are not accessible for the electrode process (75). Conformation changes conditioning this release could occur in the bulk of the solution or later in the course of interaction of the DNA molecule with the electrode. The fact that changes in DNA properties have been detected recently by means of other rionelectrochemical techniques [such as viscircular dichroism (99), small-angle X-ray scattering coinetry (96-98), (IOO), hydrogen exchange analysis (3) , and hydrolysis by micrococcal nuclease (IOl)]suggests that contact between the DNA molecules and the electrode is not a necessary precondition for changes in the behavior of DNA at premelting temperatures. The results of polarographic mensiirements support this idea. If the structure of native DNA should reinsin intact in the bulkof the solution a t premelting temperatures and the DNA should

58

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PALEEEK

unfold only in the course of polarization with forced current, then it could be expected (provided that the unfolding is sufficiently slow) that the formation of the indentation would depend on the time for which the electrode is polarized by the forced current. A shortening of this time to

lo2

10

I 40

60 Temperature

80

100

tcl

FIG. 10. Thermal transition of irradiated calf-thymus DNA followed by pukepolarographic method. DNA was irradiated by ?-rays a t a concentration of 500 p g / d in 0.015 M NaCl with 1.5 mM sodium citrate (pH 7)with a dose of2000 fads. Irradiated DNA: Xx; (logarithmic scale). Control: 00; 00 (logarithmic scale). Pulse-polarographic measurements were carried out at a DNA concentration of 4%pg/ml in 0.2 M ammonium formate with 0.05 M sodium phosphate (pH 7).The sensitivity of the Southern-Hamell Pulse Polarograph was 1/5 or lower, and the number of divisions was calculated for the sensitivity 1/5. After PaIeEek (78).

+-

+

less than sec resulted neither in the disappearance of indentation CI-2 on the “first curve” nor in its deepening on the second and third curve (10%’).A t prernelting temperatures, changes have been registered whether a potential of about - 1.5V was applied to the electrode throughout the lifetime of the drop (pulse polarography) (78)or whether the current

POLAlWGRAPHIC

59

TECHNIQUES

circuit was opened prior to application of the current pulse (7 6,95) (oscillopolarographic “first curve technique”). DNA is not adsorbed on the electrode at the potential of about - 1.5V, and there is little probability that its structure is disturbed prior to the reduction process. At more positive potentials, arising on the dropping mercury eIectrode prior t o application of the current pulse in oscillopolarographic analysis, DNA is adsorbed on the electrode and its structure might be disturbed due to interaction of DNA with the electrode surface. Nevertheless, the results obtained by the pulse-polarographic and by the oscillopolarographic methods at premelting temperatures are analogous. The conformation changes responsible for the changes in the polarographic behavior of DNA at preineltirig temperatures probably occur in the parts of the niolecule niost Iabile thermally, i.e., where the bases loop out of the double-helical structure at broken phosphodiester bonds, in A T-

+

i i Siqle -strand breoknq

Risein tempemfure

agent

0

b

C

FIG.11. Schematic presentation of thepossible DNA conformation changes in the region of a single-strand breakatpremelting tempewtures. AfterPaleEek(78).

rich regions, etc. In agreement with this assumption, the polarographic behavior of DNA at premelting temperatures has been influenced by the introduction of a small number of single-strand breaks into the DNA molecule, by means of DKase I or by 7-radiation (Fig. 10). Spectrophotometric measurements at 2GO m p have, under similar conditions, not revealed any difference in the optical density of the DNA samples into which single-strand breaks had been introduced and the control samples (78). If changes in DNA structure occur at premelting temperatures in the bulk of the solution, then the fact that these changes cannot be detected spectrophotonietricaily could be explained by the idea that the vertical stacking of the bases is not substantially influenced. The local untwisting of the doubie helix could include the interruption of hydrogen bonds, changes in angles and distances between adjacent bases, as well as changes

GO

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PALEEEK

in hydration; the bases remain vertically stacked as schematically illustrated in Fig. 11, without losing their stacking energy. In the region of singlestrand breaks there might occur even at room temperature certain conformation changes of influence only in the close vicinity ofthe interrupted phosphodiester bond (Fig. l l a , b) (Section IV, B). A rise in temperature in the premelting zone could then result in the extending of these changes to greater distances from the break (Fig. llc). As is evident from the preliminary results, premelting changes can be influenced also by other attacks on the DNA molecule. For example, with cross-linked DNA the extent of these changes was found to be smaller than with the control sample (19).

D. Estimation of Denatured DNA in Native DNA Samples The fact that denatured DNA is polarographically reducible, while native DNA is, under the same conditions, inactive has been exploited for elaborating methods for the estimation of denatured DNA in the presence of native DNA. Oscillopolarographic estimation of denatured DNA is based on measuring the depth of indentation CI-2on the “first curve” at room temperature ( 18). The lowest concentration of denatured DNA that can be oscillopolarographically estimated is about 10 pg/ml; measurements can be made in a volume of 0.2 ml, or even in a smaller volume. The amount of proteins and RNA usually occurring in DNA samples, or even a greater amount (approximately up to 200/,), does not interfere with this estimation. If the content of denatured DNA in a native DNA sample is small (around lo%), the accuracy of the estimation is reduced, because the small indentation of denatured DNA may be slightly deformed in the presence of native DNA. The presence of denatured DNA in the native sample is not indicated specifically by the indentation CI-2; a similar indentation is also produced by DNA partially digested by DKase I (79). Higher accuracy, sensitivity, and selectivity can be achieved by the pulsepolarographic technique, which allows the estimation of several tenths of a percent of denatured DNA in a native DNA sample (76). By means of this technique it is possible to differentiate between denatured DNA in the native DNA sample and single-strand breaks in the DNA double helix (79). The accuracy of the estimation of denatured DNA is only slightly influenced by the presence of native DNA (Fig. 12). The time required for pulse-polarographic measurement is several minutes, whereas oscillopoIarographic analysis requires only a few seconds. The presence of proteins and RNA in amounts usual in purified DNA samples does not interfere with estimation (76). The oscillopolarographic technique can also be used for estimating the DNA G C content. Basically it involves the determination of the

+

POLAROGRAPHIC

61

TECHNIQUES

e

E 0

f

4

DNA cuncentrotion f p g / m l ) FIG.12. The dependence of the height of pulse-pobrographic wave I11 on the 0 , Denatured DNA only; 00, concentration of denatured DNA. 01 m g of native DNA/ml plus denatured DNA in the concentration given in the graph. Measurements were carried out in 0.3M ammonium formate with 0.05M sodium phosphate (pH 7). The sensitivity of the Southern-Harwell Pulse Polarograph was 1/5 or lower, and the number of divisions was calculated forthe sensitivity 1/5. After Pale6ek and Frary(76).

temperature at which the indentation CI-1 disappears. This temperature lies close to the spectrophotometrically established T,. This technique is faster than the spectrophotometric method, but it is less accurate and requires well-purified DNA samples (105).

V. The Behavior of Polyribonucleotides The polarographic behavior of DNA suggests that polarography could be also useful in RNA research, where many problems connected with secondary and tertiary st,ructure of RNA’s have riot yet been solved. The polarographic study of synthetic polyribonucleotides can provide information that may be of importance to this investigation and might also he used for a better interpretation of the polarographic behavior of DNA.

A. Synthetic Polyribonucleotides Synthetic polyribonucleotides containing bases that are polarographically reducible in a monomeric form (poly C, poly A, and poly I) behave like denatured DNA; i.e., they produce effects on polarographic curves due toelectrode reduction processes. Polynucleotides not containing polarographically reducible bases [poly IT,poly rT, poly G, and poly (U,G)] do not produce such effects. In agreement, with oscillopolarographicbehavior

62

PALEEEK

E.

80

80

70

70

60

60

50

50

40

3c

- 1.6 - 1.4

-1.4

-1.6

I

-1.2

&

80

80

70

60

-1.2

60

50 40

I

20

-1.4

-1.2

I

70

1 1 1 1 .

-1.4

90

70

1

-1.6

-1.6

I

90 90

m

I

I

I

-1.2

- 1.6

~

30

1

- 1.4

-1.6

-1.2

-1.4

-1.2

Potential (V)

FIG.13.Pulsepolarograms ofdouble-stranded and singlestranded polynucleotides. The upperrow;double-stranded polynucleotides: (a)Nativecalf-thymus DNA a t a concentration of470p g / d in0.3M ammonium formate with0.1 M sodiumphosphate (pH 7);sensitivity 1/20(wave11). (b)1 X 10-4M poly(I) poly(C)In0.3M ammonium formatewith0.1M sodiumphosphate (pH 7);sensitivity 1/20. (c)1 X lo-' M poly(A) poly(U)in0.3M ammonium formate with0.1M sodiumphosphate (pH 7); sensitivity 1/40. The lowerrow:single-stranded polynucleotides: (d)Denatured calf-

-

POLAROGRAPHIC TECHNIQUES

63

of guanine, polynucleotides contailling guaniiie residues produce a characteristic anodic indentation on the curves of dE/dlagainst E. 1. POLYNUCLEOTIDES CONTAINING REDUCIBLEBASES a. PolyC. At neutral pH, poly C yields a D.C. polarographic step at about -1.4V (19). Like the step of denatured DNA (Section IV, A, l), this step depends on the presence of ammonium formate and attains heights of the same order of magnitude. Owing to the inadequate sensitivity of the D.C. polarographic method, the behavior of poly C has been studied primarily by pulse-polarographic and oscillopolarographic methods allowing the analysis of solutions containing a few micrograms of polynucleotide per milliliter. M ammonium formate with 0.1 M sodium phosIn a medium of 0.3 phate (pH 7),poly C yields, in addition to a pulse-polarographic wave a t about -1.5V, another more negative wave around -1.7 V against a The fact that no wave corresponding to mercury pool electrode (Fig. 13). wave I of native and denatured DNA was found was attributed tentatively to a too low concentration of poly C used in the pulse-polarographic measurements (the same is true of poly A and poly I). A t higher concentrations of poly C, there appears a distinct A.C.polarographic desorption wave at about -0.9 V and another more negative wave corresponding in its potential t o the D.C. polarographic step of poly C. Similar results have been obtained by means of the oscillopolarographic method in connection with the “first curve technique.” At neutral pH, poly C is a random coil with continuously fluctuating regions of helicity, whereas in acidic solutions it forms a double-helical structure with a system of hydrogen bonds involving a shared proton between each pair of cytosine bases (104). When the pH is decreased, the more negative wave disappears and the height of the single pulse-polarographic wave and/or the oscillopolarographic indentation decreases (19) approximately in agreement with the transition of poly C to the doubleThe shielding of the potentialIy reducible double helical form (Fig. 14). bond of cytosine in the double helix of the acid form of poly C evidently prevents its reduction on the dropping niercury electrode, as in the case of thymus DNA at a concentration of 50 pg/ml in 0.3M ammonium formate with 0.1 &I sodium phosphate (pH 7); sensitivity 1/40 (wave 111). (e) 5 X M poly C in 0.3M ammonium formate with 0.1 M sodium phosphate (pH 7 ) ;sensitivity 1/80. (f)5 X lO-‘M poly A in 0.3M ammonium formate with 0.1 kf sodium phosphate (pH 7); sensitivity 1/40. Molar concentrations of polynucleotides are based on the phosphorus content. Measurements were performed on a Southern-HarweI1 Pulse Polamgraph, Mark XI. The potentials were measured against the mercurypool at the bottom ofthe polarographic vessel.

64

E.

PALEEEK

e

P z .a E

P)

s

0’ 2

3

4

5

6

PH

FIG.14.Dependenceof height of pulse-polarographic wave of polyC and cytidine on pH. 00, 3 X lo-' M polyC in0.2M ammonium formate withtheBrittonRobinson buffer sensitivity 1/40. 00, 3 X 10- M cytidine in 0.2M ammonium 1/80.Southern-Harwell Pulse formatewiththeBrittori-Robinson buffer; sensitivity Polarograph, Mark 11.

poly (I) poly (C) (see below). The oscillopolarographic melting curve shows a sharp increase in the depth of the indentation within the narrow temperature range at pH 5.0 (Fig. 15) in contrast to the curve obtained at pH 7.0(Fig. 16).These results agree with the presumed structure of poly C at pH 5 and pH 7 (104, 106). b. PolyA . I n a medium of 0.03 M NaCl with 0.18M phosphate at pH 7, poly A yields only a very small D.C. polarographic step, which induced Berg et al.to conclude that poly A was not reducible polaroIn the presence of ammonium formate, the polarographic graphically (74). At pH 7.0, poly A yields behavior of poly A resembles that of poly C (19). a D.C. polarographic step with around - 1.4V, thus differing from its low-molecular coinponents (adenine, adenosine, adenylic acids, and adenosine dinucleotide), which are reducible only in an acidic medium (Section 111, A).The shift in reducibility of poly A to the neutral pH may be explained by the higher pK value than that found for adenosine (104). In agreement with D.C. poIarographic behavior, poly A yields a pulsepolaragraphic wave (Fig. 13)and a cathodic indentation on the oscdlogram (19). The faradaic currents produced by poly A in a neutral medium are not due to contamination of the sample by another reducible substance, because they are produced by poly A samples of different origin and do not disappear after purification. Alkaline degradation of poly A resulted in a decrease in its pulse-polarographic wave and finally in its disappearance; +

20

‘

20

I

40

60

Temperature I‘C)

VIG 15.Thermal trarisitiori of poly C a t pH 5 followed oscillopolurograyliically and spectrophotometrically. Poly C in 0.2 M ammonium formate with the McIlvaine buffer p H 5. In oscillopolarographic measurements the dropping mercury electrode was polarized with A.C. of rectangular shape and frequency 50 c/s. The “first curve technique” was used. 00 , Oscillopolarography ; 0 0, spectrophotometry.

0.55 30

F

0.50

-F c .p

-c

I

=k

E

20

U

+ w (0

B

0.45

.-c < a

-

8 c U

5,

a

5

2

10

w

3

0.40

Q

0

2

0.35 40

a0

60 Temperature I‘CI

-

FIG.16. Thermal transition of poly C and poly (I) poly (C) nt pH 7 followed oscillopolarographically and spectrophotometrically. Xx, 5 x 10-6 M poly c in 0.1 M ammonium formate wit,h 0.1 M sodium phosphate (pH 7); oscillopolarography; 1X M poly (I). poly (C) in 0.1 M ammonium formate with 0.1 M sodium phosphate (pK 7). 00, Oscillopolarography; 00,spectrophotometry. In the oscillopolarographic measurements, the dropping mercury electrode was polarized with A.C. of rectangular shape and frequency 50 c/s. The “first curve technique” wa8 used.

66

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PALEEEK

no further wave was formed in the course of degradation. On the other hand, the product of the alkaline or enzymatic degradation of poly C yielded one wave (at approximately the same potential as the wave of CMP), which was higher than the waves of undegraded poly C. The transition from the single-stranded form of poly A to the doublestranded “acid” form was followed by means of polarographic techniques. The suppression of the polarographic activity in the “acid” form of poly A was, however, less marked than in the case of poly C. The measurements could be performed at relatively low ionic strength, because in acidic media (from about pH 6) the pulse-polarographic wave or the D.C. polarographic step appeared even in the absence of ammonium formate. The A.C. polarographic behavior of poly C and poly A suggest that besides bases (86) other groups might take part in the adsorption of a polynucleotide on the mercury electrode (19). c. PolyI. In an acidic medium, poly I yields two D.C. polarographic around - 1.0 and - 1.2 V (19). In weakly acidic (above ca. steps with El,t pH 5) and neutral media, poly I appears to be not reducible polarographically. On oscillopolarographic curves, there may be observed some indication of an indentation even at neutral pH at very negative potentials; however, this indentation is not suitable for analytical purposes owing to the close vicinity of the right marginal point. The inflexion arising in the region of the background discharge on the pulse-polarogram is a little more suitable for these purposes. I n a neutral medium, hypoxanthine is D.C. polarographically inactive ($3, 46) and does not yield an inflexion or a wave on pulse-polarograms (19). 2.POLYNUCLEOTIDES CONTAININGNONREDUCIBLE BASES

Poly U and poly (U,G)do not yield a reduction D.C.polarographic step in 0.5 M ammonium formate with 0.1 M sodium phosphate pH 7, even at about 1 mM concentration (19). Poly U, poly rT, and poly (U,G) produce pulse-polarographic waves only at a high sensitivity of the instrument. These waves are substantially smaller than the waves of equally concentrated poly C or poly A and could be of capacity character. From A.C.polarographic measurements, it follows that poly U and poly (U,G) are adsorbed and desorbed within a potential region similar to that of DNA (Section IV, A, 2), and give rise to tensammetric (nonfaradaic) waves. The oscillopolarographic cathodic indentations of poly U and poly (U,G) arise within the same potential region where the tensammetric waves appear. At neutral pH, poly (U,G)and poly G yield an anodic indentation of a character similar to that of denatured DNA.

3. POLYNUCLEOTIDECOMPLEXES At neutral pH, the polarographic behaviors of the double-helical complexes of polvnucleotides differ considerably from the behaviors of

POLAROGRAPHIC

67

TECHNIQUES

-

constituent single-stranded polynucleotides. In the complexes poly (A) poly (U), poly (A). poly (rl'), and poly (I) . poly (C), the reducibilities of poly A and poly C is suppressed and their adsorption behavior is greatly changed (19). The polarographic mixing curves can have the form of the letter V, as is the case for the spectrophotonietric curves, only if both the polynucleotides are polarographically reducible. Poly (I) . poly (C) in a sufficiently high concentration yields a small oscillopolarographic indentation or a pulse-polarogaphic wave that does not disappear even in the presence of excess of poly I (Fig. 17)and whose

Mole % of polyI

-

FIG.17.Formation of the 1: 1 complex of poly (C) poly (I) followed by pulsepolarographic method. Homopolymers were mixed in 0.1 M NaCl with 0.01 M sodium phosphate (pH 7).After 2 hours of incubation at room temperature the supporting background electrolyte was added. The pulse-polarographic measurements were carried out in 0.3M ammonium formate with 0.1 M sodium phosphate (pH 6.9). The sensitivity of the Southern-Harwell Pulse Polarograph was 1/80. Concentration of poly C (4 X 10-6M)was held constant in all samples while the amount of poly I varied as indicated in the figure. The height of the more positive pulse-polarographic wave of poly C was measured.

potential is somewhat more positive than the potential of the poly C wave (Fig. 13).The wave of poly (A) . poly (U) differs in potential from the wave of free poly A to such extent that both the substances can be determined simultaneously provided that their waves are approximately of the same height. Addition of polyU in a concentration equivalent to that of free poly A leads to the disappearance of the poly A wave. I n agreement with the results obtained with native DNA (Section IV, C, I), changes are exhibited in t.he polarographic behavior of poly (I) . poly (C) at temperatures below the temperature of melting (Fig. 16). The height of the mnve of poly (I) . poly (C) atid poly (A) poly (U) increases aftcr irradiation

68

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PALEEEK

with low doses of y-rays, as is the case with wave I1 of native DNA (Section IV, B).

B. Ribonucleic Acids Information on the polarographic behavior of RKA is still rather scanty. The references in the literature merely supplement the studies with DNA. The RNA preparations employed have not been well-defined in most cases so that the results obtained do not allow us to draw definitive conclusions. I n the presence of ammonium formate, RNA produces a small, poorly developed D.C. polarographic step within the potential region where the step of denatured DNA is formed (19). A better developed step observed by Luthy et al.in 0.2M KCl and 0.1M HClOd (SO) was perhaps caused by RNA degradation products. On A.C. polarograms of RNA there appears a wave (’74) resembling to some extent wave 1 of DNA. In the presence of ammonium formate, no other wave is formed (19). RNA yields an anodic indentation on the oscillopolarographic first curve (90). tRNA produces a pulse-polarographic wave at a potential similar to that of denatured DNA; the wave of tRNA is, however, considerably smaller than that of equally concentrated denatured DNA (76). As expected, a difference in the polarographic behavior of tRNA and rRNA has been found, but the results are only of a preliminary character (19). Substantially more information is available on the influence of RNA on the polarographic behavior of proteins. These problems, however, exceed the scope of the present article and are therefore mentioned here only briefly. Although the presence of nucleic acids influences the shape and height of the polarographic steps of proteins in NH4CI-NH,0H buffer in thgpresence of cobalt salts (7), polarography can be usedfor determining traces of proteins in RNA (106) and DNA samples (87, 88). If the protein is active enough polarographically (activity being connected with the presence of -SH and 44- groups in the molecule), as little as 0.3% of protein can be determined in the RNA sample (106). The polarographic behavior of tobacco mosaic virus has also been subjected to investigation on the basis of the polarographic activity of its protein (107-111). Owing to the fact that the protein that is a constituent of the virus differs in its behavior from the other proteins, the polarographic technique could be used for determining very small amounts of proteins (around 0.2%) contaminating the purified virus preparation (107-109).

VI. Concluding Remarks The progress achieved in recent years in the use of polarography for studying the properties and t,he structure of nucleic acids has been pri-

POLAROGRAPHIC

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marily due to two factors: (a) the finding of a suitable medium in which polynucleotides containing polarographically reducible bases yield faradaic currents; and (b) the application of modern polarographic techniques allowing accurate measurement of these very low faradaic currents. The medium for analyzing nucleic acids has been found empirically. The results hitherto obtained suggest that the main role of the salts present in the analyzed solution consists in their influence on polynucleotide adsorption. The pulse-polarographic technique, which has been found best suited for studying nucleic acids, has been designed with the aim of enhancing the sensitivity of the polarographic analysis of low-molecular substances. It appears, however, that it will find application especially in the study of macromolecular substances. The use of classical polarography for these purposes cannot be successful, as the concentrations in which the macromolecular substances under investigation could yield a sufficiently high D.C. polarographic step cannot be usually reached either because of the limited solubility of these substances or because, at higher concentrations, the aggregation of molecules may exert an unfavorable influence on measurement. The results of polarographic analysis of nucleic acids, interesting from the point of the study of their secondary structure, may roughly be summarized as follows : Random-coiled polynucleotides containing polarographically reducible bases yield faradaic currents, whereas in polynucleotides having a double-helical structure in which the reduction sites are hidden, these currents are suppressed. However, a small amount of the bases contained in the double-helix remains accessible for the electrode process and can yield a pulse-polarographic wave differing in its potential from the wave of the parent random-coiled polynucleotide. These bases are probably located in the labile regions of the double helix, e.g., at broken phosphodiester bonds, a t loops out of the double-helical structure, at free ends of the molecules, etc. The length of the labilized region depends on the conditions in which the molecule finds itself. An increase in temper+ ture, or more generally perhaps the approaching of conditions at which helix-coil transition takes place, probably leads to the extension of these regions. I n order to obtain more accurate ideas about the changes taking place in the labilized regions of the double helix, we should know more about the details of the mechanism of the electrode processes to which the bases contained in the polynucleotide are subject on the electrode. For the time being, it seems probable that the bases in the polynucleotide are reduced on the electrode just aa are the bases in monomeric forms, the process, however, being complicated by adsorption. The elucidation of all the processes to which the polynucleotide is subject in the neighborhood of the electrode requires a great deal of further investigation.

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The polarographic reducibility of the disulfide groups in proteins, recently discovered by Cecil and Weitzman (1l S )represents , an interesting analogy with the reduction of bases in the polynucleotide. It has been found that the ease of reduction of the -S4- bonds depends on their position in the molecule so that intrachain and interchain -S-Sbonds can be differentiated. The results were obtained by means of macroscale electroreduction and D.C.polarographic measurements; however, the polarographic currents were very low. By applying pulse polarography or other modern polarographic methods, the polarographic determination of -S4groups could become a very important technique in protein research. From the point of view of the chemical analysis of nucleic acid the polarographic techniques can be especially useful for the determination of nucleic acid components in mixtures and for the estimation of denatured DNA in the presence of an excess of native DNA. The possibilities offered by methods of electrochemical analysis for investigating nucleic acids and for molecular biological research in general have not hitherto been fully exploited. The complicated nature of the processes to which the substance under investigation is subject on the electrode may at first appear as a drawback, but later it can be made use of for obtaining information that other, basically more simple, methods cannot furnish. A deeper understanding of the processes to which polynucleotides are subject on the dropping mercury electrode and the application of the techniques of electrochemical analysis for studying more Complicated systems, such as pure nucleoproteins, ribosomes, viruses, etc., may yield highly interesting results in the future.

ACKNOWLEDGMENTS I should liketo thankDr.B. Janik, Dr.V. Vetterl, and otherworkersfrom my laboratory who allowed me to quoteexperimental results inadvance ofpublication. 1 am Dr.R. Kalvoda, Dr. Z. Pechan, and my colleaugues also grateful toDr. M. Heyrovskf, attheInstitute ofBiophysics bothfornumerousdiscussions and forcritical reading of themanuscript.

REFERENCES I. J.Josseand J. Eigner, Ann.Rev.Biochem. 36,789 (1966). 2.G. Felsenfeld and H. T.Miles, A m . Rev.Biochem. 36,407 (1967). 3.P.H. von Hippel and M. P. Printa, Federation Proc.24,1458(1965). 4. I.R. Miller, Biochim. Biophys. Acta103,219 (1965). 6. D. Bachand I.R. Miller, Biochim. Biophys. Acta114,311 (1966). 6.R. Brdibka, Collection Czech. C k m . Commun. 6,112 (1933). 7.M. Bfezina and P.Zuman, Polarography in Medicine, Biochemistry and Pharmacy, p.585.Interscience, New York,1958.

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8. J. Heyrovskf, Chem.Listy 16, 256 (1922). 9. J. Heyrovskf and J. Kuta, “Principles of Polarography.” Publ. House Czechoslovak Acad. Sci., Prague, 1965. 10. L. Meites, “Polarographic Techniques.” Interscience, New York, 1965. 11. P. Zuman, “Organic Polarographic Analysis.” Pergamon Press, Oxford, 1964. 1%. H. Schmidt

and M. von Stackelberg, “Modern Polarographic Methods.” Academic Press, New York, 1963. IS.B. Breyer, i n“Progress in Polarography” (P. Zuman and I. M. Kolthof, eds.), Vol. 11,p.487.Interscience, New York, 1962. 1 4 .B. Breyer and H. Bailer, “Alternating Current Polarography and Tensammetry.” Interscience, New York, 1963. 15. D. E. Smith, i n “Electroanalytical Chemistry” (A. J. Bard, ed.), Vol. 1,p. 1. Dekker, New York, 1966. 16.J. Heyrovskf., Chem.Listy 36, 155(1941). 17.J. Heyrovskf. and J. Forejt, Z.Physik. Chem.(Leipzig) 193, 77 (1943). 18. E. PaleEek, Biochzm. Biophys. Acta94,293 (1965). 19. E. PaleEek, unpublished results. 20. R. Kalvoda and G. Budnikov, Collrction Czech.Chem.Commun. 28, 838 (1963). 21.R. Kalvoda, Collectzon Czech. Chem.Commun.in press. 2% .J. Heyrovskf and R. Kalvoda, “Oseillographische Polarographic mit Wechselstrom.” Akademie Verlag, Berlin, 1960. 23. R. Kalvoda, “Techniques of Oscillographic Polaropraphy.” Elsevier, Amsterdam, 1965. 24.M. Heyrovskf. and K. Micka, in“Electroanalytical Chemistry” (A. J. Bard, ed.), Vol. 2,p. 193.Dekker, New York, 1967. 25. C. C. Barker and I. L. Jenkins, Analyst 77,685 (1952). 26.G. C. Barker and A. W. Gardner, Z.Anal.Chem.173, 79 (1960). 27.B. Janik and P. J. Elving, Chem.Rev.68,295 (1968). 28. S. M. Cantor and Q. P.Peniston, J .Am. C hem.SOC. 62 2113(1940). 29.J. C. Heath, Nature168, 23 (1946). SO. D. Hamer, D. M. Waldron, and D. L. Woodhouse, Arch.Biochem. Biophys. 47,272 (1953). 31.N. G. Luthy and B. Lanit), J .Phnrrn. Pharmacol. 8,410 (1956). SZ. F. A. McGinn and G. B. Brown, J .Am Chem.Soc.82,3193(1960). 3s. E. PaleEek, Collection Czech. Chem.Comniun. 26, 2283(1960). S4. D. L. Smith and P. J. Elving, J .Am. Chem.SOC.84,1412(1962). 56. B. Janik and P. J. Elving, in preparation. S6.V. Vetted, Collection Czech. Chem.Commun.31, 2105(1966). S7.V. Vetted, J .Electroanal. Chem. 19, 169 (1968). 58. L. F. Cavalieri and B. A. Lowy, Arch.Bzochem.Biophys. 36, 83 (1952). 39. E. PaleEek, Natumuissenschaften 46,186(1958). 40.E. PaleEek and B. Janik, Arch.Biochem. Biophys. 98,527 (1962). 41.D. L. Smith and P. J. Elving, J .A m . Chem.SOC. 84,2741(1962). 42. B. Janilc and E.Palerek, Arch. Bzochem. Biophys. 106, 225 (1964). 4.5’. B. Janik and E. PaleEek, Abhandl. Deut.A kad.Wiss. Berlin, KZ.Med.p. 513(1966). 44.E. PaleEek, Abhandl. Deut.Akad.I.c”ss. Berlin, Kl.Med.p. 501 (1966). 45.D. L. Smith and P. J. Elving, Anal.Chem.34, 930 (1962). 46.E. PaleEek and B. Janik, Chem.Zwsti16, 406 (1962). (In Russisn.) 47.B. Janik and E.PaleEek, Z.Natztrforsch. 2lb, 1117(1966). 48. B.Janik, in preparstion.

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0.Manouiek and P. Zuman, Chem.Listy 49,669(1955). 60.J.KrupiEka and J. Gut, CoUectwnCzech. Chem.Commun.26, 592 (1960). 61.A. I f d o v h , Collection Czech. Chem.Commun.29, 182(1964). 66. P. Delahay, ‘‘Double Layer and Electrode Kinetics,” p. 81.Interscience, New York, 1965. 63. V. Vetted, Experientia 21, 9 (1965). 64. V. Vetterl, Abhandl. Deut.Akad.Wiss. Berlin, K1.Med.p .493 (1966). 66.V. Vetterl, Collection Czech. Chem.Commun.,in press. 66.W. Leyko and H. Panusz, Bull. SOC.Sci. Lettres Lodz,Cl.IZI 6, 1 (1954). 67.S. Eguchi, Nichidai ZgakuZdgshi20, 2368(1961). Chem.Abstr. 61, 3496a(1964). 68. B. Filipowicz and W. Leyko, Bull. SOC. Sci. Lettres Lodz,Cl.I I I4,1 (1953). 69. S. Fiala and H. E. Kasinski, J .Natl. CancerInst. 26,1059(1961). 60.8. Matsushita, F. Ibuki, and A. Aoki,J .Agr.Chem.SOC. Japan57, 67 (1963). 61. V. Bulant, M. Urks, and H. Pafizkovfi, Antibiotiki 9, 545 (1964). 62. F.$om, J. Berfinek, J. Smrt, J. KrupiEka, and J. skoda, CoUection Czech. Chem. Commun.27, 575 (1965). 63. R. Pleticha and J. J. Weiss, Anal. Biochem. 16, 510 (1966). 64. D. Kalhb,Chem.Zvesti 14,823 (1960). 65. D. KalBb, Experientia 17,275 (1961). 66. D. KalBb, Abhandl. Deut.Akad.Wiss. Berlin, Kl.Med.p. 333 (1964). 67.D. Kalhb, Experientia 19, 392 (1963). 68. D. KalBb, Chem.Zvesli 18, 435 (1964). 69.D. Kalhb, Abhandl. Deut.Akad.Wiss.Berlin, Kl.Med.p. 519(1966). 70. D. KalSb, Experientiu 22, 23 (1966). 71. J.BohfiEek, Ezpe-rientia 19, 435 (1963k 7.2. V. Habermann, E. MaidlovS, and R. Cernf, Collection Czech. Chem.C o m m m . 31, 139(1966). 73.H. Berg and F. A. Gollrnick, ‘‘Vortrkge des I. Seminars uber Molekularl>iologit., Reinhardsbrunn,” p. 74.Inst. Mikrobiol. Exptl. Therap., Jena, 3965. 74.H. Berg, H. Bar, and F. A. Gollmick, Biopolymers 6,61 (1967). 76.E.PaleEek, J.Mol.B i d .20, 263(1966). 76.E. PaleEek and B. D. Frary, Arch.Biochem. Biophys. 116, 431 (1966). 77.E. PaleEek and V. Vetterl, Biopolymers 6,917 (1968). 78.E. PaleEek, Arch.Biochem. Biophys. 126, 142(1968). 79. E. PaleEek, Biochim. Biophys. Acta146, 410(1967). 80.D. Bach and I. R. Miller, Biopolymers 6, 161(1967). 82. E. Calendi, A. Di Marco, M. Reggiani, B. Scarpinab, and L. Valentini, Biochim. Biophys. Acla103, 25 (1965). 86. I. R. Miller and D. Bach, Biopolymers 4,705(1966). 83.I. R. Miller and D. Bach, Proc.4thInter. Congr.Polarography, Prague, 2966p.64. 84.H . Berg and F. A. Gollrnick, Abhandl. Deut.Akad. Wiss.Berlin, Kl.Med. p . 533 (1966). 86. H. Berg, Monatsber. Deut.Akad.Wiss.,Berlin 7,210(1965). 86. I.R.Miller, J .Mot.Biol. 3, 229 (1961). 87. E. PaleEek, Nature188, 4751(1960). 88. E. PaleEek, Biokhimiya 26, 803 (1960). (In Russian.) 89. E. PaleEek, Biochim. Biophgs. Acta61,1 (1961). 90.E. PaleEek, Collectiwn Czech. Chem.Commun.31, 2360(1966). 91. E. PaleEek and E. Lukfikvfi, Biophysik 3, 272 (1966). 92. E. Lukfi~ov&Biophysik 6, 183 (1968).

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POLAROGRAPHIC

TECHNIQUES

73

5 3. 8. KuEan, unpublished results. 9.4. E. PaleEek, 2. Chem. 2, 260a (1962).E. PaleEek, Abhandl.Deut.Akad.Wiss. Berlin, Kl.Med.p.270 (1964). 96. E. PaleEek, J. Mol.Bioi. 11, 839 (1965). 96. A. M.Freund and G. Bernardi, Nature200, 1318 (1963). 97.T. I. Tikhonenko, G. A. Perevertajlo, and E. N. Dobrov,Biochim. Biophys. Acta

49, 160 (1961). 98. M. Boublik, L. Pivec, J. fiponnr, and Z. &ormovS, Collection Czech. Chem.Commun.

30, 2645 (1965). 89. J. Brahms and W. H. F. M. Mommaerts, J .Mol.Biol. 10, 73 (1964). 100. V. Luzzati, A. Mathis, F. Masson, and J. Witz, J. Mol.Biol. 10, 28 (1964). 101. P. H.von Hippel and G. Felsenfeld, Biochemistry 3, 27 (1964). 10%.E. PaleEek, Intern. Symp.Macrornol. Chem.Prague,1965,Preprints Sci.Comm. p. 26. 103. J.Boh4Eek and E. PaleEek, Collection Czmh.Chem.Commun. 30,3456 (1965). 104. A. M.Michelson, J. MassouliB, and W. Guschlbnuer, This series, 6, 83 (1967). 206. W.Guschlbauer, Proc.NaU. Acad.Sci.U .S.6, 1441 (1967). 106. V. Blazsek and L.Bukaresti, Biochim. Biophys. Acta61, 970 (1962). 107.G.Ruttkay-Nedeckfr, Biochim. Biophys. Acta26,455 (1957). 108. G. Ruttkay-Nedeck4, Collection Czech. Chem.Commun. 26,3363 (1960). 109. G.Ruttkzy-Xedeckp, Collection Czech. Chem.Commun. 28,585 (1963). 110.G. Ruttkay-Nedeckp, Collection Czech. Chem.Commun. 29, 1809 (1964). 111. G. Ruttkay-Nedeckfr and A. Anderlov4, Nature213, 564 (1967). 11%. R. Cecil and P. D. J. Weitzman, Biochem. J .93, 1 (1964).

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RNA Polymerase and the Control of RNA Synthesis JOHN P. RICHARDSON Institiit de Biologie Mollculaire, Universitt! de Genkve, Geneva,Switzerland

I. Introduction . . . . . . . . . . . . 11. General Concepts of the Regulation of RNA Synthesis . . . 111. Purity and Physical Properties of RNA Polymerase Preparations . A. Purity . . . . , . . . . . . . . B. Physical Properties . . . , . . . . . . IV. The Transcription Mechanism . . . . . . . . A. The Role of DNA , , . . . . . . . . B. De Noao Synthesis of RNA Chains and the Direction of Chain Growth . . . . . . . . . . . C. Release ofRNA Molecules . . . . . . . . D. Growth Rates of RNA Chains . . . . . . . V. Selective Transcription . . . . . . . . . . A. Asymmetric Transcription . . . . . . . . B. Further Restrictions on Transcription . . . . . . C. Preferential Chain Initiation with Purine Nucleotides . . VI. The Mechanism ofRNA Chain Initiation . . . . . . A. The Binding ofRNA Polymerase to DNA . . . . . B. The Nature of the RNA Polymerase Attachment Site . . C. The Initiation Reaction . . . . . . . . . D. Initiation Sites . . . . . . . . . . . VII. Termination Signals . . . . . . . . . . VIII. Inhibitors . . . . . . . . . . . . . A. Inhibitors of Polymerization . . . . . . . . B. Inhibitors of Initiation . . . . . . . . . IX. Conclusions . . . . . . . . . . . . References . . , . . . . . . . . . Note Added in Proof . . . . . . . . .

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1. Introduction RNA polymerase plays a key role in the functioning of a cell. It is the enzyme that is directly responsible for the synthesis of all types of cellular RNA. Under the direction of DNA, it catalyzes the sequential assembly of four ribonucleoside triphosphates into RNA molecules. In this way, the genetic information stored. in the nucleotide sequences in the DNA is transferred into a form that can be used to direct the synthesis of specific 75

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proteins. It is also in this way that the other RNA components of the cell, the RNA of the ribosomes and the transfer RNA’s, are produced. The existence of RNA polymerase [systematic name: Nucleosidetriphosphate :RNA nucleotidyltransferase (DNA dependent) ; EC 2.7.7.61 has been recognized for more than ten years (I). It has been found in a wide variety of sources and surely exists in all cells in which RNA synthesis occurs ( 2 )However, . the purest preparations have been isolated from bacteria. Even though advances have been made in the purification of RNA polymerase from animal tissues (3-5), the best preparations from these sources have specific activities that are even less than that of a bacterial crude extract (4). Eventually, total purification from higher organisms will be achieved, and it is most likely that these polymerases will have properties very similar to those of the enzymes isolated from bacteria. This review is restricted almost entirely to a discussion of the enzyme from Escherichia colibecause it is the most highly studied and because, eonsequently, the most is known about it. Many of the properties of the reaction catalyzed by E. coliRNA polymerase have already been given in the review by Hurwitz and August in the first volume of this series (6). Since that review, much progress has been made in understanding the properties of the enzyme itself as well as its mechanism of action and its specificity. It is these subjects that are given the main consideration in this review and, in particular, emphasis is placed on the mechanisms by which RNA synthesis ‘may be regulated.

II. General Concepts of the Regulation of RNA Synthesis The amount of any given species of RNA present in a cell seems carefully regulated t o meet the requirements of the cell’s metabolism (7,8). Since many RNA’s are metabolically unstable, the regulation of their levels may be achieved by control of their rates ofdegradation as well as their rates of synthesis. There is good evidence that most RNA species are made directly by transcription from DNA templates (at least in bacteria) (6, 8-12); thus whatever mechanism exists to control the synthesis of a particular RNA must also be a mechanism that in some way controls the action of RNA polymerase. The amount of a particular RNA species found in a cell at any given moment is not directly related to the relative amount of genetic information in its DNA. For example, ribosomal RNA’s comprise as much as 80% of the RNA in rapidly growing E. coli( l S )yet , these RNA’s are complementary to less than 0.4% of the E . coligenetic material ( I d )Further. more, the complete genome contains considerable information that is not needed by the bacteria for growth under particular conditions. There are

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genes that code for enzymes including those needed to produce amino acids that may already be present in the medium or for enzymes needed to metabolize certain carbon sources. Thus, it seems logical that the cell has some means of allowing some genes to be transcribed independently of others and also of determining whether a certain gene or set of genes may be transcribed at all. In terms of mechanism, these propositions imply that there are specific sites on the DNA where RNA polymerase can initiate and terminate the synthesis of RNA molecules. This would make it possible for certain transcriptive units to be very active while others are inactive. The particular activity of each transcriptive unit could then be regulated by elements specific to that unit. Both positive and negative control elements have been postulated to explain the regulation of various types of genetic units. Considerable evidence has been presented by Jacob and Monod in support of their analysis of the factors that control the synthesis of enzymes for lactose metabolism in E .coli(15). The lactose regulatory system contains a gene tthat makes a product called a repressor, which in its active form blocks the synthesis of all the enzymes for lactose metabolism. The repressor is considered a negative control element because in its absence the enzymes whose synthesis it normally coiitrols are produced at maximum rate. In molecular terms, Jacob and Monod postulated that the repressor could act by binding to a site on the DNA and thus prevent RNA polymerase from transcribing the adjacent structural genes for the enzymes. TWOobservations are in good agreement with this model: first, in the cell the levels of RNA from the lactose genes directly reflect the activity of these genes as expressed in terms of rates of enzyme synthesis (16); second, the lactose and has been shown to bind specifically to repressor has been isolated (17) the DNA region it is hypothesized to affect (28).It thus seems quite likely that an important general mechanism of control is one in which the transcription of a particular section of a DNA molecule can be specifically blocked by a regulator product. An example of a genetic unit regulated by a positive control element is the unit coding for the synthesis of the enzymes for arabinose metabolism in E.coZi(19). The regulator gene in this case makes a product necessary for the synthesis of these enzymes. One possibIe function of the product of the regulator gene might be to activate the site where transcription of the arabinose genetic unit is initiated. Thus, this regulator gene might directly affect RNA polymerase molecules or sume of the sites on the DNA where the enzyme acts. However, no strong evidence has been presented to support this particular model, and it is possible that the control occurs at the level of protein synthesis. Mechanisms of positive control are probably very important in the

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regulation of the activity of the genes of many viruses. In these cases, there is good evidence that transcription is regulated directly. Many viruses possess whole blocks of genes that do not start to function until after the For the phages T4 and A, at least, viral DNA has begun to replicate (20). the product of a particular “early” gene is required for the expression of In both cases, it has also been shown that the the “late” genes (21-23). levels of RNA for such genes also reflect the gene activity; RNA for these genes is not detectable until after viral DNA synthesis has started, and it becomes detectable only if the regulator genes have made their products (25-28). It has been postulated, therefore, that the regulator genes make products that in some way allow RNA polymerase to initiate transcription of the late genes. In addition to systems that control the activity of particular genes, it has also been suggested that there may be mechanisms that control the 30). The possibility that the cell has activity of all genes coordinately (29, such systems arises from considerations of the rate of total RNA synthesis as a function of changes in cellular environment. For example, when a cell is transferred from a very rich growth medium to one that is poorer, net RNA synthesis (measured by the incorporation of precursors into total RNA) stops very quickly (8). However, because of the fact that messenger RNA is unstable, measurements of the net incorporation of precursors indicate only the effect of the changes on the synthesis of the stable species of RNA (ribosomal and transfer RNA’s). Hence, what is often considered coordinate control may only be the control of these stable species of RNA. A model for RNA control that would have a general application is one 32). He has suggested that most if not all RNA proposed by Stent (31, synthesis may be controlled by a type of feedback in which the rate of synthesis of an RNA species would depend upon its rate of utilization. The principal hypothesis of this model is that something is necessary to pull the RNA product away from the complex of RNA polymerase and DNA in order for the RNA to be synthesized efficiently. Under this system, the synthesis of messenger RNA’s would depend upon the ability of ribosomes to use the messenger for protein synthesis; the synthesis of ribosomal RNA’s might depend on the attachment of the RNA to a pool of free ribosomal proteins; and the synthesis of transfer RNA’s on the attachment of each RNA to its activating enzyme. One of the special features of this model is that it implies a close coupling of the rate of RXA production to the rate of protein synthesis, and therefore could explain the results of all experiments in which the level of RNA from a particular gene reflects the rate of synthesis of the protein encoded by that gene. Support has been given to this model by the finding that there is a very

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close physical coupling between nascent RNA and ribosomes (33-35). A given messenger RNA appears to be used in the synthesis of a protein even The finding th a t the direcbefore its own synthesis is completed (36,37). tion of transrription of the messenger (58, 39) and the direction of its trailslation are the same indicates the feasibility of such a mechanism (4G4.2). Further, the finding that the mRNA’s for some genes with strong polar amber mutations are shorter than the RNA’s from the wild-type genes (45-45) also is compatible with the Stent model because amber mutants are known to give rise to chain termination in protein synthesis (46); but per se they have no direct effect on the transcription step (47). Thus the synthesis of the RNA beyond the amber mutation night be affected because the ribosome cannot proceed past the point on the RNA containing the defect. Although these facts, as well as others having to do with RNA synthesis are all after ribosome depletion during starvation for magnesium ions (48), easily interpretable in terms of the Stent model, there are others that do not seem to fit so well. One problem occurs in considering what happens to RNA synthesis in various cells starved for an amino acid required for growth. I n most cases, as measured by incorporation of precursors, net RNA synthesis stops. If this measure of RNA synthesis really represents a coordinate inhibition of the synthesis of all types of RNA synthesis, this observation would be quite compatible with the Stent hypothesis. However, the synthesis of messenger RNA continues unabated under these conditions, even after the synthesis of total RNA seems to have stopped (49). Thus, the very species of RNA whose synthesis one would expect to be blocked first, by its inability to be used, is the one species that is still synthesized a t a near normal rate. Although the rate of synthesis of the stable species of RNA, rRNA and tRNA’s, still may be responding to changes in the rates with which these RNA’s are pulled away from the polymerase, these stable species also could be responding to some other mechanism t ha t acts by blocking initiation. The question then arises whether there really is any coordinated regulation of RNA synthesis. There may be none a t all except in the trivial case of starvation for an energy source, which leads to a depletion of the nucleoside triphosphate substrates. Nevertheless, there may be some important regulatory mechanisms that are coordinate. Thus it is of interest to study all factors that affect the total rate of RNA synthesis by RNA polymerase. Further, the Stent hypothesis may still be true in some special cases, and thus it is of interest t o investigate those factors that control the release of RNA molecules as well as those that might lead to a n increase in synthesis rates by coupling with protein synthesis.

111. Purity and Physical Properties of RNA Polymerase Preparations A. Purity Before discussing the mechanism and specificity of RNA polymerase action, it is important to consider the purity of the enzyme preparations used for these studies. There exist several methods for isolation and purification of RNA polymerase from E. coli(60-59), most yielding very purepreparations of enzyme. However, this variety of procedures and the difficulties inherent in repeating any one of them inevitably lead to variabilities between the preparations used. The simplest criterion of purity of any enzyme is its specific activity. For RNA polymerase, unfortunately, this criterion is not very reliable, especially when comparisons are made between preparations from different laboratories, because there is no generally accepted standard assay. Since the quality of the DNA that is used in the assay is of particular importance, it is necessary tochoose one DNA as a standard. Because of its availability and the ease with which it can be prepared, T4 DNA is recommended as the standard (60). Under the direction of this DNA and with the assay conditions described by So etal.(61), a very active preparation of RNA polymerase catalyzes the synthesis of as much as 9.1 pmoles of RNA (expressed as nucleotides incorporated) per mg of protein1 in 10minutes at 37 C (62). Physical purity of a protein is indicated by its sedimentation and electrophoretic homogeneity. Like many other proteins, RNA polymerase has a tendency to aggregate. However, by raising the salt concentration to 0.5 M with KCI (or NaCl or NH4Cl), most of this aggregation can be avoided (53,64, 63) ; thus, under the proper conditions, a very pure preparation is at least 95% homogeneous on sedimentation analysis. Electrophoretic purity is harder to achieve. Preparations that show only one peak in the ultracentrifuge often include several minor components with different electrophoretic mobilities. It is possibIe to reniove most of these impurities selectively by using zone electrophoresis as a step in purification (53). Similar purification has also been achieved by chromatography on phosphocellulose columns (59). As a result of electrophoretic analysis, it is lriiown that RNA polymerase is an acidic protein; at pH 8, the pH optimal for enzymatic activity, the enzyme migrates to the anode (54). 1 The method used to measure the protein concentration is also of importance. In this w e the amount of protein used was based on the absorbance of the enzyme based = 6.5(64). on an extinction coefficient of E g , , ,

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The ratio of absorbance at 280 m p to that at 260 mp of a purified preparation of RNA polymerase is 1.65, which suggests a nucleic acid 58). content of less than 0.1% (54, For studies of the physical properties of the enzyme, the above criteria may be sufficient. However, for certain enzymatic studies, there are other tests that are even more important. The presence of as little as 0.001% of a nuclease in a preparation is not detectable by physical means, but such a contamination could greatly affect the activity of the enzyme. However, it is possible to rule out the presence of extremely small amounts of such interfering activities by appropriate assays. Applying very sensitive tests, it has been possible to exclude the presence of DNases or RNases in purified preparations to the extent that 100 pg of RNA polymerase breaks less than one out of 10,OOO nucleotide linkages in 10 minutes at 37 C.Most of the better enzyme preparations used to study the properties of RNA polymerase have less nuclease activity than can be measured in the assays used. Maitra and Hunvitz also tested their preparations for the presence of several other enzyme activities involved in polynucleotide and mononucleotide metabolism, and were able to detect only one significant conAlthough this enzyme may have taminant: polyphosphate kinase (58). been present to the extent of O . l ~ o ,its activity could be completely inhibited mM) to the reaction mixture. by adding ADP (0.02 Therefore, it can be concluded that it is possible to make preparations of RNA polymerase that are physically homogeneous and that are virtually free of the most important contaminating activities. However, it cannot be said that all preparations used are of the same quality. Even in the same laboratory, there are large variations from one preparation to the next. Ideally, each preparation should be checked by every possible criterion, or a t least by activity, electrophoretic purity, and nuclease contamination. Because the enzyme is very stable when stored under the proper conditions, it is naturally advisable to prepare the enzyme in large batches that can be analyzed properly and then used for many experiments. Even with the use of very pure enzyme preparations, it is always possible that a given property may be the result of a contaminant not detectable by the assays used. It may occasionaly be necessary to invoke such considerations to explain apparent differences in results obtained with “pure” RNA polymerase prepared by different procedures. In addition, it will be of interest eventually to know whether the enzyme is composed of units that are necessary for different steps in RNA synthesis. For example, the component that catalyzes the poIynierization step might be separable from one that determines the specificity of initiation. In such a case, the purity of the enzyme would be dependent upon what is defined to be the major catalytic property of RNA polymerase.

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B. Physical Properties The fact that R N A polymerase has such a strong tendency to aggregate has made it difficult to measure its molecular weight. Only in buffers of sufficientlyhigh ionic strength (e.g., 0.5 M KCI) is the enzyme homogeneous enough to attempt accurate measurements. The sedimentation coefficient of the unaggregated form, extrapolated to zero concentration, is 12.8S (64); for convenience, i t is referred to as the 13S form. An earlier measurement of the molecular weight of this form, based on the method of sedimentation equilibrium in a very short column, yielded a value of 4.4 f 0.8X lo5(54). More recently, Priess and Zillig combined measurements of sedimentation and diffusion to calculate a molecular weight of 3.6X lo5 (64). This value is in very good agreement with the sedimentation equilibrium result of 3.7x lo5obtained by Maitra and Hurwitz (58). The earlier estimate is higher because the enzyme may still have some tendency to aggregate in solutions of high ionic strength and especially at the high protein concentrations used in sedimentation equilibrium measurements. Several values for the sedimentation coefficient of the larger forms of RNA polymerase have been reported. However, a systematic analysis of the sedimentation coefficient as a function of the ionic strength of the buffer used shows that the value can vary continuously between 13 and 26 S (68, 63,66). Molecular weight estimates for some of the larger forms indicate that they are true aggregates, not just different conformational Because so many forms exist, it is extremely difficult to decide forms (54). what the true functional unit of RNA polymerase is. It certainly should not be inferred that all the various forms are active as such. Under conditions of the assay, all forms are probably in equilibrium with a form that can successfully initiate R N A synthesis on a D N A template. The continuous variation observed in the sedimentation coefficient as a function of ionic strength is most likely a manifestation of such a rapid equilibrium between several forms of the enzyme. Deciding which is the functional unit is not simplified by the fact that the enzyme is active in assay conditions that give rise to all forms from 13 to 26 S (62,G6). Nevertheless, the following observations do tend to support the likelihood that the 13 S form is the functional unit. (a) Some preparations show little or no tendency to aggregate (58). (b) Dilute solutions favor dissociation, and the enzyme usually is assayed in dilute solutions. (c) The binding of polynucleotides to the enzyme also favors its dissociation (67, 68) ; under conditions in which free enzyme sediments at 21 S, complexes between R N A polymerase and small polynucleotides sediment at 13S in buffers without Mg2+ions and at 15-16S in buffers with Mgz+ions (62). The results of these experiments are particularly relevant because some of the poIynucleotides used, for example, poly dT and poly dC, are

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fairly good templates for RNA polyiiierase. However, the biologically important template is double-helical DNA, and it is still not known what effect it has on enzyme structure. estimated the molecular weight of the smallest of the Stevens etal.(63) poly U dissociated forms, using the amounts of enzyme and poly U in the complex and the number average molecular weight of the poly U used. The This result suggests that the 13 S form can value estimated was 390,000. bind by itself to a polynucleotide. It is unlikely that the active enzyme is a structure smaller than the 13S forni. Although it is possible to dissociate the enzyme further with protein denaturants such as urea, detergents, or alkali, none of these smaller units is active (59). It has also not been possible yet to renature the further dissociated enzyme. The finding that different preparations of enzyme have different properties of aggregation has led to the suggestion that RNA polymerase However, because certain undefined factors may be heterogeneous (67). such as storage and aging as well as defined factors such as the presence of oligonucleotides have profound effects on the sedimentation properties of the enzyme, it is easy to explain many of the differences in preparations without resorting to the suggestion of heterogeneity. There also may well be several different types of RNA polymerase in a cell, but at present there is no strong biochemical evidence to support this suggestion. However, there is some evidence that the RNA polymerases isolated from different strains of E.coliare not identical. This is indicated from the differences in the amino acid compositions that have been published for RNA polymerase 58,64). from E.coliK12, E.coliW, and E.coli B (56, Since RNA polymerase is quite large, it seemed very likely that something could be learned about its structure from electron microscope studies. So far, however, these studies have been disappointing. Objects with a very regular shape are observed in some preparations (69, 7 0 )but , with further 7 2 )Negatively . stained purification these objects become much rarer (71, objects seen in the purest preparations have no obvious structural features (7.2, 7 3 )Since . it is very likely that RNA polymerase is denatured by the staining procedures used, it would be worthwhile trying some other procedures to visualize the enzyme. An even more interesting aspect of an electron microscope study would be to determine the size, shape, and fine structure of RNA polymerase bound to DNA. Enzyme bound to DNA has already been visualized, but only by staining techniques that either denature the enzyme or distort its . what is needed is a technique that will dimensions (69, 73,7 4 ) Again, permit visualization of both DNA and the fine structure of the enzyme without distorting either. Although little has been learned about the shape and fine structure of

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RNA polymerase, good progress has been made recently in studying the chemical subunits of RNA polymerase. Enzyme denatured either with 8 M urea or 0.1%sodium dodecyl sulfate exhibits two major protein bands upon acrylamide gel electrophoresis and in the ultracentrifuge (69). Burgess has been able to isolate these two components and determine their molecular weights. One component has a molecular weight of 38,000 and the other 160,000 (66). These two components are separate polypeptide chains that are quite dissimilar both in amino acid composition and on analysis of the peptide fragments produced by cyanogen bromide cleavage. From the molecular weights of the two components and the finding that they exist in equimolar quantities, Burgess has suggested that the 13 S form of RNA polymerase may be composed of two small subunits and two large subunits. This would suggest a molecular weight of about 400,000 for the 13S form. More recently he has resolved the larger of the two subunits into two components that differ very slightly in molecular weight (66). It remains to be seen whether they are two completely different proteins or whether they differ only in the addition of a small number of amino acids to one chain. It will also be of interest to determine whether it is possible to associate one of the subunits with a particular activity of the enzyme.

IV. The Transcription Mechanism A. The Roleof DNA The importance of DNA in the RNA polymerase reaction became fully evident as soon as enzyme preparations were pure enough to show an absolute requirement for DNA. In the earliest studies, the nucleotide composition ofthe RNA product appeared to be the same as that of the DNA used to direct the reaction (6). However, a much more sensitive test, was used by Geiduschek el al.(76) to prove the template function of DNA. They showed that the RNA product from a T2 DNA-directed RNA polymerase reaction could form RNA-DNA hybrid helices very efficiently with heat-denatured T2 DNA, but could not form such hybrids with other DNA’s, including a DNA with a base composition similar to that of T2 DNA. This test shows that the RNA product is complementary to the DNA used to direct its synthesis. Another example of this test is given in Table I. The nucleotide compositions of T7 DNA and h DNA are nearly the same (76); nevertheless the RNA synthesized in a reaction mixture containing h DNA hybridizes less than 1% as well with T7 DNA as with X DNA; likewise, the RNA synthesized in a reaction mixture containing T? DNA will not hybridize with X DNA but will with T7 DNA. The fact that only 50% of the total input RNA is able to hybridize with the DNA

ItNA POLYMERASE

AND THE CONTIWL

OF RNA

TABLE I HYBRIDIZATION OF RNA POLYMERASE PRODUCT RNA TEMPLATE A N D NONTEMPLATE DNA5 DNA template for RNA polymerase reaction

T7

x

85

SYNTHESIS

WITII

Percent of RNA hybridized with

T7 DNA 50 0.3

A

DNA 0.2 49

Blank filter 0.3 0.4 ~

RNA was synthesized invitro from the two DNA templates using a modification of the T7 DNA assay mixture described in reference 54.[3HHJUTP (100 pCi/pmole) was used to label the RNA. Synthesis was allowed to proceed for 30minutes at 37"C,and the RNA was isolated by treatment of the reacbion mixture with sodium dodecyl sulfate (0.4%) and hot phenol (172). Unincorporated nucleotides were removed by dialysis for16hours against 0.3 M NaCl containing 0.03 M Nar citrate. Hybridization of RNA to DNA was performed as described by Gilleapie and Spiegelman (f73).Filters with 2 r g of the indicated DNA (or without DNA) were incubated for 15 hours a t 66 Cin 1 ml of 0.3M NaCI, 0.03 M NaDcitrate buffer containing 0.12 pg (-5000cpm) of RNA synthesized from the indicated DNA template.

used as template reflects the efficiency of the method used to measure hybridization. One of the first questions asked about the mechanism of RNA polymerase action was: How can the enzyme use a double-helical DNA as a template? During RNA synthesis, the double-helical structure of the template DNA is conserved (76,77,78). In this respect, the RNA polymerase reaction differs from the DNA polymerase reaction. Two very plausible models have been suggested to explain how RNA polymerase can use the information in the sequences of nucleotides in a double-helical DNA to direct the ordered synthesis of RNA molecules without permanently destroying the DNA structure. I n the first model (78, 79), it is postulated that the DNA strands separate over a short region to allow RNA polymerase to use one of the strands as a template in the Watson-Crick sense. As transcription proceeds, the finished end of the RNA chain is displaced from a temporary hybrid helix with the template strand by the re-formation of the DNA helix. In the second model (8&82), the DNA strands do not unwind. Instead, genetic information is transferred by a new set of hydrogen bonding rules, during which both strands take part in directing the synthesis of the RNA product strand. A clue to understanding which of these models might be correct is provided by the fact that RNA polymerase is also able to use singlestranded DNA’s as template. The product of the reaction in which the single-stranded DNA from 4x174viris2 is used as template was shown by

* See article by R. L.Sinsheimer in Volume 8 ofthis series.

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Chamberlin and Berg to be a RNA-DNA helix (78). The base composition of the RNA product is coniplementary to that of the template DNA strand. More recently, fibers of the RNA-DNA hybrid made by RNA polymerase with f 1 DNA (another naturally occurring single-stranded DNA) were analyzed by X-ray crystallography (8s). This analysis showed that the RNA-DNA hybrid is indeed a double helix in which one strand is antiparallel to the other. This evidence strongly indicates that RNA polymerase uses a single-stranded DNA as a template for synthesis of an RNA strand in the Watson-Crick sense. It was also shown by Chamberlin and Berg (78) and by Sinsheimer and that this RNA-DNA hybrid can be used as a template for Lawrence (84) RNA synthesis. When further synthesis is allowed to occur, it is found that the RNA strand in the hybrid helix can be displaced by the newly synthesized RNA strand, but that the displacement is not obligatory. Using [ 3H]RNA in the hybrid and 14C-labeledsubstrates, Sinsheimerand Lawrence found that the ratio of the two labels in the free RNA after 1.2-fold further synthesis is consistent with a probability of displacement of the preexisting RKA strand by the newly synthesized strand of 0.5. Thus, unlike the transcription of the DNA double-helix, the transcription of the RYADNA helix can be either conservative or semiconservsJt’we. Since the transcription of single-stranded DNA and RNA-DNA hybrid involves the formation of Watson-Crick base-pairs between the nascent RNA and the DNA, Chamberlin and Berg concluded that it is very likely that such base pairing is also involved in the transcription of doublehelical DNA (78). Thus, it seems likely that the mechanism of transcription of double-stranded DNA resembles that predicted in the first model. If the second model were correct, it would mean that RXA polymerase would use one mechanism to copy single-stranded DNA and a very different mechanism to transcribe double-helical DNA. However, a problem still remains. If the same nieehanisni is used for transcription of double-helical DNA and of RNA-DNA hybrid, why is the transcription of one template completely conservative and of the other partially semiconservative? One possible explanation was proposed by Chamberlin and Berg (78), based on the fact that the DNA-RNA helix is less stable than thc equivalent DNA double helix. They postulated that when double-helical DXA is transcribed, template DNA strands would always rewind aridthus displace RNA being synthesized because the DNA helix is more stable than the alternative RNA-DNA helix. I n contrast, when the RNA-DNA hybrid is template, the growing RNA strand and the preexisting RNA strand would have equal probabilities of remaining in the helix because there would be no difference in the stabilities of the respective helices.

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Recently, Chamberlin presented evidence that supports the model in which the stability of the two possible helical products determines the mode of replication (85).A model DNA template consisting of poly dI as one strand of the double helix and poly dC as the other [(dI), . (dC),,] was used as a template. This copolymer can direct the separate incorporation of either CTP or ITP with RNA polymerase. Denaturation temperatures of the three possible helical products showed that (rI), . (dC), is more . (dC),, and that (dI), . (rC), is less stable. stable than the template (dI)?& When CTP was the subst,rate, free poly C was formed and the template (dI), . (dC), was left intact, but when ITP was used, poly dI was displaced and the double-stranded product was the hybrid (rI), (dC),. These results show that the mode of transcription of the same template can be either conservative or semiconservative, and that the choice of the mode seems to depend upon the relative stability of the possible helical products. Thus, experiments with transcription of single-stranded DNA and with model DNA polymers are consistent with the model in which the DNA helix unwinds to expose a single strand for transcription. Another line of evidence that supports this model is the finding that it is possible to isolate a very stable complex between the DNA template and the growing RNA chain. In studies with native T4 DNA, Bremer and Konrad found that nascent RNA and the DNA template exist in a complex stable enough to be isolated by zone centrifugation (86). However, this stability is strongly dependent upon the integrity of the enzyme; treatment of the complex with a detergent that denatures the enzyme but has little or no effect on polynucleotide structure completely dissociates nascent RNA from its complex with T4 DNA. Similar results have been found in studies of the complexes between nascent RNA and other linear DNA templates such as T7 DNA and X DNA (87,62). By themselves, the results of Brenier and Konrad (86) are compatible with both of the proposed models of transcription. However, the results of similar experiments by Hayashi with the replicative form of 4x174DNA2 (a naturally occurring double-helical form of 4x174 DNA found in host cells after infection with the virus) do support the hypothesis that the DNA strands unwind to expose one strand as a template and that the nascent When this small [M.W. = RXA forms a temporary hybrid helix with it (88). 3.4x lo6 (89)] “circular” (cyclic) DNA is used as template, most (although not all) of the RNA remains athched to the DNA even after treatment with detergents and phenol. Further, trhe complex is stable even during banding in a CsCl gradient. Hayashi also showed that part of the nascent RNA is resistant to ribonuclease, but that, once labeled, this resistant portion could be “chased” by the addition of excess unlabeled substrate. This result indicates that the ribonuclease-resistant portion of the RNA is

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near the growing point of the RNA molecule. Thus, it appears that, with the continuous, double-helical form of 4x174DNA, a significant portion of the RNA is very stably attached to the DNA template, a result that is in strong contrast with the findings with linear double-helical templates. It is likely then that the difference in stability of the nascent RNA may be related to the important difference in structure between the acyclic and cyclic forms of double-helical DNA s.The 4x174RF-DNA preparation used by Hayashi consisted primarily of what are known as twisted cyclic forms. Tertiary twisting of many if not all such DNA s (including 4x174 RF-DNA) is believed to result from a deficiency of turns in the basic double helix (90-92)). I n other words, there are constraints on the configuration of a cyclic DNA that do not exist with a linear DNA. Thus, it is possible that re-formation of an unwound portion of a DNA helix may be less favorable for a cyclic DNA than for an acyclic DNA. This in turn might mean that the region of attachment between a nascent RNA chain and its template is longer and more stable with the former DNA than with the latter. It would be of interest to see how results with 4x174RF-DNA compare with those with other twisted cyclic DNA s,and to see whether the t,emplate properties of a DNA are changed when the DNA is converted from this form to a simpler cyclic form by introduction of a single-strand break.

8. De Novo Synthesis of RNA Chains and the Direction of Chain Growth

A characteristic of RNA moleculcs synthesized by purified RNA polymerase is that there are three phosphates attached to the 5‘ hydroxyl groups. Thus, alkaline hydroIysis of the product RNA releases the 5 terminal nucleotide as a nucleoside tetraphosphate (pppNp), the 3’ terminal nucleotide as a nucleoside (N)and the internal nucleotides as 2 and 3’-monophosphates (Np) (93, 38, 39). Since these three different products can be easily separated electrophoretically, it has been possible to show that both ends of enzymatically synthesized RNA molecules receive Iabel from substrates. This indicates that RNA polymerase does not require RNA primers in order to initiate RNA chain synthesis. Bremer et al.also were able to use this method of identifying end If a labeled groups t o determine the direction of RNA chain growth (38). substrate is added under conditions in which there is rapid growth but very little initiation, it will be incorporated preferentially into the growing end. Bremer etal.found conditions t o label RNA molecules in that way and, from their analysis of the products of alkaline hydrolysis of the labeled RNA molecules, concluded that, chain growth occurs by addition of new nucleotides to the 3’-hydroxyl end of nascent RNA chains. I n other words,

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the direction of chain growth is froni the 5'-hydroxyl end to the 3'-hydroxyl end. Maitra and Hurwitz used another technique to arrive at the same 32Plabel in the third (y) phosphate of an incorporated conclusion (39). nucleoside triphosphate can only be incorporated at the 5’end of an RNA chain. Once such a substrate is incorporated into a growing RNA chain it cannot be removed by the addition of excess unlabeled substrate. Thus, end of the newly synthesized chain addition must occur at the opposite (3’) RNA molecule. A third fact that confirms this conclusion is that 3'-deoxyadenosine 5'-triphosphate is incorporated into RNA chains in an RNA polymeraseThis analog of ATP has no hydroxyl group at its catalyzed reaction (94). 3’position; therefore it cannot be added to the 5' end of a chain. There are at least two important consequences of the fact that RNA chains are synthesized in the 5' to 3’ direction. The first is that, since it is possible messenger RNA is translated in the same direction (40-429, that a ribosome can attach to a nascent RNA chain and start to use the RNA as a messenger for protein synthesis even before the chain is completed. The second is that since the known RNA exonucleases in E.coli that might be responsible for RNA degradation (in the cell) are specific for the 3’terminus of RNA chains, nascent RNA may be immune to degrada96). tion until it is completed and released (95,

C. Release of RNA Molecules In Section IV, A, evidence was presented to show that during its synthesis the growing RNA chain is coinplexed in some way to the DNA template. It is of interest to ask whether this RNA chain is ever released and whether one enzyme molecule can initiate the synthesis of more than one RNA chain in vitro. Under the conditions used by Bremer and Konrad, newly synthesized RNA molecules are not released during synthesis on a T4 DNA template (86). Even after synthesis in a reaction mixture ceases, most of the RNA can be isolated as part of a complex with enzyme and DNA. This result suggests that each enzyme can catalyze the synthesis of only one RNA chain. Bremer and Konrad presented other evidence to support such a mode of action. After the first few minutes, which is the time needed for complete initiation, the number of nascent RNA molecules is constant throughout the reaction (86). However, recent, experiments indicate that the conclusions of Bremer and Konrad may not hold for all conditions of synthesis. I n reaction mixtures with salt concentrations as high as 0.2 M , each enzyme can catalyze

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the synthesis of several RNA molecules. The effect of salt is apparent in the kinetics of RNA synthesis. In reaction mixtures that contain little or no added salt (other than the divalent cation and the buffer), synthesis stops completely after only 1 hour a t 37 C (86, 97).In reaction mixtures with a much higher salt concentration, however, such as those described by So etal.(61) and by Fuchs etal.(97), synthesis continues for several hours with only a slight decrease in rate. From this change in kinetics alone it seems quite likely that either each enzyme makes more than one RNA molecule or that each RNA molecule is able to become much larger. The following experiments favor the first of these suggestions. Complexes between RNA polymerase and DNA are retained by membrane filters (98), whereas free RNA is not (99). If proper conditions for synthesis are chosen, the amount of RNA that remains in a complex with enzyme and DNA can be determined directly from the amount of RNA that is retained by a membrane filter. Figure l a demonstrates the use of this technique to confirm the findings of Bremer and Konrad under the conditions of low salt (0.05 M KC1). Up to 60 minutes, at least 80% of all the RNA made, which in this case is essentially all the RNA that will be made, is in the enzyme-DNA complex (curve B ) .Control experiments (not shown) demonstrate that no RNA is retained by the filter if the reaction mixture is treated with detergent; and free RNA added exogeneously after initiation of synthesis is not retained. The latter control indicates that the compIex detected by this technique is not formed spontaneously. However, the experiment shown in Fig. l a does indicate that RNA molecules are gradually released: by 180 minutes only 50YG of the RNA is still retained by the filters. This decrease is accompanied by a similar decrease in the amount of DNA retained, which indicates that the complex does eventually dissociate even in low7 salt reaction mixtures. Figure l b shows the results of a similar experiment in which 0.2M KCI is present in the reaction mixture. Initially all the RNA is retained by the filter, but after 10 minutes of incubation over half of the RNA is no longer attached to the complex, and by 45 minutes there is no further increase in the amount of nonfilterable RNA even though synthesis continues for several hours. I n contrast, all the DNA is still retained by the filter, a t least during the first 90 minutes, which indicates that most of the enzyme molecules must still be involved in the complex with DNA. This interpretation is consistent also with the observation that the maximum amount of nonfilterable RNA in Fig. l b is nearly the same as the total amount of RNA that can be made in the low salt conditions (Fig. la). It is quite possible that the release of RNA from the complex that occurs in the presence of high salt is just the result of the action of a

(a) 0.05 M KCI; 5 rnM MgCI, A, t o t a l RNA

( b ) 0.20 M K C I ; 12 rnM MgCI,

B, nonfilterable RNA

d

I

1

1

20

40

60

I

80

100

120

140

I

160

I

IN

Time ( m i d

FIG.1. Effect of salt concentration on RNA synthesis and of fraction of RNA in mplex with enzyme and DNA. Each reaction mixture contained 0.0484Tris-HC1 1.6 m M each of ATP, GTP, CTP and [aHjUTP (6pCi/pmole), 4.2 mM of ffer, pH 7.9, nercaptoethanol, 50 p M EDTA, 6 pg of a*P-labeled T4 DNA (tL500 cpmlpg), and pg of hydroxylapatite-pursed RNA polymerase (64)in a volume of 0.5 ml. I n dition, one reaction mixture (a) contained 0.05 M KC1 and 5 mM MgClz and the other 1 0.2M KC1 and 12 mM MgC12. Incubation at 37 C was commenced after adding syme to the mixture. At the indicated times, two 25-pl portions were removed from :h reaction mixture. One portion was assayed for acid-precipitable material using !mbrane filters to collect the precipitates. The other was diluted into 2 ml of cold 11 M Tris-HC1 buffer pH 7.9 containing 0.05 11.1NaC1, filtered through a membrane .er (Sartorius MF 50, Gottingen), and the filter was washed with 20 ml of the same ution. Filters were assayed for radioactivity by scintillation counting as described :viously (1%). Parts (a) and (b) of the figure give the results for reaction mixtures a and b, respecely. The 32P-labelin the T4 DNA was used asil control to determine the amount of VA bound to filters. In both experiments, retention of DN.4 by the filters was quanative up to 90 minutes of synthesis. After 90 minutes, the amount of nonfilterable DNA 0, :lined linearly to values of 80% in (a) itlid 65% in (b) by 180 minutes. 0a1 acid-insoluble material; A--A, radioactivity remaining on f i h rafter filtran in Tris-NaC1buffer.

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JOHN P. RICHARDSON

nuclease. If this were the only reason for release, there should be no increase in the number of RNA chains initiated de novoafter completion of the first round of initiation. RNA chain initiation can be measured quite simply by the incorporation of nucleoside [-y-”P] triphosphate into RNA (59). Thus, it is possible to determine whether high salt allows continued initiation of RNA molecules from the course of incorporation of [-y-3zP] ATP. The results of such an experiment indicate that high salt conditions allow T4 RNA chains to be initiated throughout the incubation period and therefore for long periods after initiation would have stopped in a low salt reaction mixture (100, 62). Thus, it seems that high salt conditions allow RNA polymerase to release RNA chains in a way that enables the enzyme to initiate the synthesis of new RNA molecules de novo.

D.

Growth Rates of RNA Chains

Increasing the ionic strength of the RNA polymerase reaction mixture has another important effect: it increases the rate at which RNA chains are polymerized. Growth rates of RNA chains can be estimated from the sedimentation distributions of RNA isolated from reaction mixtures at various times. Using this method, Bremer and Konrad found that the growth rate of T4 RNA chains is 2.5nucleotides per second under their conditions of synthesis (86). At a saturating concentration of nucleoside triphosphates, this value extrapolates to 4 nucleotides per second (101). However, in the presence of 0.2M KC1, the conditions of So etal.(el), a T4 chain growth rate as high as 16 nucleotides per second has been measured (62). Because of the rapidity of growth in 0.2M KC1, and the likelihood that RNA chain initiation is not synchronous, the sedimentation profiles actually give a slight underestimate of the growth rates. Thus, it is likely that in 0.2M KCl, RNA molecules grow at a rate of more than 20 per second. This invitro growth rate is almost as high as the value of 28 that Bremer has measured for the growth rate of T4 RNA invivo(102). These results bring into question the relevance of Stent’s hypothesis (31,32) that RNA synthesis is controlled directly by a feedback mechanism in which RNA synthesis is dependent upon the attachment of the nascent RNA chain to other factors in the cell. The findings of Bremer and Konrad that the growth rate of RNA molecules is slow and that nascent RNA molecules are not released invitroquite naturally suggest that other factors are needed to increase the growth rate and to release newly synthesized RNA chains. In fact, the addition of ribosomes and the factors needed to bind inRNA to ribosomes does lead to some stiniulation of the growth rate and to some release of RNA molecules under conditions in which the ionic strength of the incubation mixture was 0.1 or less (103-106).

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However, the results obtained with 0.2A2 KC’1 show that it is possible to achieve both of these ends merely by inrrcasing the ionic strength of the reaction mixture. These two results lead to a dilemma. Which is more important-the added factors? or the higher ionic strength? It is quite possible that the high ionic strength used in the reaction mixtures that give RNA release and high growth rates may be too high for other cellular functions including protein synthesis. However, measurements of intracellular ionic conditions indicate that the KC1 concentration may be as high as 0.2M in E.coli(106). There isgood evidence to indicate that RNA polymerase ceases synthesis SO quickly in low salt conditions because the enzyme is inhibited in some way by its product. This is best demonstrated by the experiment of Krakow in which the rate of RNA polymerase activity in the presence of ribonuclease is measured by the release of pyrophosphate (107). Krakow found that the enzyme remains active for several hours in the presence of ribonuclease. This is a direct indication that the nascent RNA chain is in some way inhibitory to RNA polymerase. On first analysis, this product inhibition seems to be just a manifestation of the well-known fact that many types of RNA are quite potent However, for inhibition of RNA inhibitors of RNA polymerase (108, 109). polynierase by an exogenous RNA, the RNA must be added before synthesis begins (108). RNA added while the reaction is in progress has little or no effect on the reaction rate. Thus, it is concluded that there is some special feature of the product RNA that allows it to inhibit the activity of the enzyme that made it. Further, the inactivation of RNA poIymerase by its nascent RNA is not a simple function of the size of the RNA molecule. Reduction of the growth rate of RNA chains by lowering the concentration of nucleoside triphosphates does not change the rate at which the enzyme is inactivated (97, 101). This result means that the rate of RNA growth is not inversely proportional to the size of the newly synthesized RNA molecules. Instead, it seems likely that inactivation is n random process in which the RNA molecule is in some way involved. One action of higher salt Concentrations is to prevent this inactivation from occurring. It is possible that in the presence of the salt, an RNA molecule is aIways released before it can inactivate the enzyme molecule that catalyzed its synthesis. However, the inactivation that occurs in a low-salt reaction mixture is not irreversible; RNA polymerase activity is recovered merely by raising the ionic strength (97). Thus, it is more likely that high salt concentrations makes RNA polymerase immune to endproduct inhibition. In fact, So etal.have pointed out that in 0.2 M KCl RNA polymerase is much less sensitive to inhibition by exogeneously

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added RNA (6 1). This could be related to the loss of product inhibition. The effect noticed by So etal.at least explains why RNA polymerase is able to make large quantities of RNA i n the presence of 0.2M KC1 and to reinitiate synthesis on DNA molecules without becoming inhibited by the RNA made and released during the reaction.

V. Selective Transcription A. Asymmetric Transcription

The transcription of many native, double-helical DNA’s by RNA polymerase is asymmetric; in any particular region of the DNA, only one of the two complementary DNA strands is transcribed (1iU,111). However, the conditions in which this important property of RNA polymerase can be observed are very stringent. Denatured DNA’s are transcribed symmetrically, as are DNA’s that have been degraded to such an extent that many single-stranded regions are exposed. Further, the exact ionic conditions of the reaction mixture may be important with some if not all enzyme preparations; the conditions required for proper strand selection with M . lysodeikticus RNA polymerase are very different from those found to be appropriate for the E.colienzyme (118). The clearest way to show that transcription is asymmetric is to demonstrate that the product RNA is complementary to only one of the two strands of the DNA used as template. This can be shown particularly well for transcription of 4x174 RF-DNA. The DNA found in the mature 4x174 virus is single-stranded, but upon infection of a host cell this single-stranded form is converted by partial replication to a cyclic doublestranded form known as (9x174 RF-DNA? As mentioned in a previous section, 4x174 RF-DNA can be isolated intact. Hayashi et al. used (9x174 RF-DNA as a template for E. coliRNA polymerase and found that the product RNA had a base composition identical to that of the DNA strand found in the mature virus (110). Furthermore, this RNA could not form an RNA-DNA hybrid with the viral DNA strand but could with denatured replicative form. Thus, RNA polymerase must have transcribed only one strand of $X174 RF-DNA. Comparison of these results with the hybridization properties of 4x174 RNA synthesized in vivoindicated that the same DNA strand is transcribed in vivo as in vitro. Previously, other workers had used another form of double-helical (9x174 DNA as template and had found the opposite result: transcription of that DNA was symmetric (50, 113). The double-helical DNA used in those studies had been prepared in vitro by extensive enzymatic replication of the single-stranded form of 4x174 DNA. It is known now that the

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product of extensive replication by DNA polymerase in vitro has several structural peculiarities (114). It is possible that these structural peculiarities could perturb the selection process in RNA polymerase transcription, and could therefore be the reason that enzymatically replicated DNA, unlike 4x174RF-DNA, is transcribed symmetrically. Another clear case of asymmetric transcription involves the Bacillus megaterium phage a (115). The two strands of the linear phage DNA molecule can be separated on the basis of the difference in their densities; therefore, a-specific RNA can be analyzed by determination of how well it hybridizes with the two different DNA strands. Such an analysis shows transcription of native a DXA that a RNA made in viuoor during in vitro is coinwith a crude RNA polymerase preparation from B. megaterium plementary to only one and the same DNA strand. The results with a DNA thus show that acyclic as well as cyclic DNA can bc transcribed asymmetrically. The use of 4x174 RF-DNA and a DNA as templates was fortunate because in both cases the same DNA strand is transcribed in all regions of the DNA that are open to transcription. However, with other DNA’s, such as h DNA, one strand is transcribed in some regions and the other in Thus, for these DNA’s not all the RNA formed is other regions (116,117). complementary to the same strand. Nevertheless, it is possible to demonstrate that such DNA’s are transcribed asymmetrically. One way is to show that the RNA product is not self-complementary.Since complementary RNA molecdes are able to form stable RNA-RNA helices when it is possible to show lack of annealed under the proper conditions (118), self-complementarity by a simple test based on the fact that RNA in a double-helical form is more resistant to ribonuclease than single stranded However, this test has two limitations: One is that the RNA RNA (111). must be free of fragments of single-stranded DNA that could also form helices with the RNA; the other is that t,he formation of the RNA-RNA double-helix by annealing is a concentration-dependent reaction and it is difficult technically to use a high enough concentration to achieve saturation in the formation of the helices. Another test based on the same principle can be used to determine the that is complementary to RKR transcribed fraction of RNA made in vitro from the same DNA in vivo. After incubating a small fixed amount of labeled RNA synthesized in vitro with increasing amounts of RNA synthe labeled RNA that is complementary to the cellular thesized i n vivo, RNA becomes resistant to ribonuclease. This test thus gives some indication of the fidelity with which RNA polymerase has copied the same DNA strand in uitro that is copied in viuo.Thc following are two csaniples of results obtained from this test: less than 2(xl of RNA made 11-y trnrtscription

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of T4 DNA invzfro becomes riboiiuclease resistant when annealed with a large excess of RNA extracted from cells harvested 20 minutes after infecin contrast, as much as 38% of the RNA made tion with T4 phage (62); from transcription of denatured T4 DNA becomes ribonuclease resistant by the same test (119).

B. Further Restrictions on Transcription Transcription with RNA poIymerase invstrois restricted to specific regions of the DNA template. In the cases of phages T2, T4, SPOl, and X, little or no RNA is made from DXA regions containing the “late” genes of the viruses (119-124). In most cases this conclusion has been reached from the results of hybridization studies. For instance, labeled RNA made by RNA polymerase from T4 DNA is completely diluted from hybrid formation with T4 DNA by a large excess of “early” (5minutes after infection) invivo T4 RNA. If the T4 RNA synthesized invitro contained a significant fraction of “late” RNA, the latter RNA would still appear in the hybrid with DNA. Further, even when RNA synthesis from T4 DNA is allowed no significant amount of late to proceed for as long as 90 minutes invitro, RNA is made (61). Regional selection in h DNA transcription has also been shown by another technique. With X DNA it is possible to separate and isolate the two halves of the DNA and thus determine what fraction of a X RNA preparation is complementary to each of the halves. Approximately 7580% of the RNA made from transcription of whole native X DNA is complementary to the half that contains most if not all of the genes for early 124). function (the right hand half of the genetic map) (123, Results of experiments with SPOl DNA indicate that selective transcription is not an exclusive property of homologous RNA polymerase-DNA bacteriophage, yet E.coliRNA polycombinations. SPOl is a B. subtdis merase is able to transcribe only early SPOl RNA from SPOl DNA (119). Changes in the structure of DNA that allow a template to be transcribed symmetrically also lead to a loss of regional selectivity. RNA synthesized from denatured T4 DNA contains T4 RNA species made This suggests that both early and late and their complenients (119). whatever mechanism exists to restrict transcription to certain regions of the DNA is probably the same as that restricting transcription to one of the two strands within those regions. RNA polymerase does not transcribe late RNA The fact that E.coli from the DNA of the viruses of T2, T4,SPO1, and X suggest that other factors are needed if these regions are to be transcribed. Since the purified DNA used in the invitro transcription espcrimerits contains less than 0.4%

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contamination with protein, it is unlikely that each of the transcription sites for the late genes is covered by some repressor molecule attached to the DNA of mature phage. It is more reasonable to postulate that RNA polymerase isolated from the host bact.eria is unable to initiate synthesis from the DNA regions of the late genes. Thus, it may be that, for late RNA to be synthesized, the product of one or more of the early phage genes is necessary. This product would be a positive control element: it might be a new RNA polymerase, a factor that alters the specificity of the host RNA polymerase, or a n enzyme that makes new regions of the DNA recognizable to the host RNA polymerase. Recently, Snyder and Geiduschek (125) have been able to denionstrate of one of the putative positive control elements of T4, the action in vitro the product of gene 55.They have devised crude enzyme systems in which I n one of these late T4 RNA can be synthesized asymmetrically invitro. systems, the synthesis of late T4 RNA depends upon the addition of components that must include the product of gene 55.

C. Preferential Chain Initiation with Purine Nucleotides Further evidence that transcription by RNA polymerase is selective comes from measurements of nucleotide sequences a t the 5‘-phosphate end of RNA chains. The distribution of nucleotides appearing a t this end of RNA chains is most easily determined by an indirect method, in which the relative incorporation of each of the four nucleoside triphosphates into the 5’ terminal position is measured. One procedure is to make four reaction mixtures that are identical except for the label in the nucleotide. In each mixture, one of the four nucleoside triphosphates is labeled with a2Pin the y (and/or p ) phosphate. Label in these positions of the triphosphate is incorporated only into the 5’ terminus of RNA chains. Table I1 presents data obtained by Maitra and Hurwitz, who used this method to examine The most the product of transcription of several DNA templates (39). striking result of this set of experiments is that there is a strong preference for purine nucleotides for chain initiation. In most cases, less than 15% of the chains were initiated with a pyrimidine nucleotide. This preference is also evident with the denatured DNA’s. However, denaturation of a DNA does change some of its template properties. After denaturation, the number of RNA chains initiated from a DNA in a 30-minute incubation period is increased even though total RNA synthesis is decreased, and the ratio of chains initiated with GTP to those initiated with ATP is increased. I n general, there is a preference for initiation with GTP, and the higher the guanine plus cytosine content of the DNA, the greater is this preference. Only in some of the cases in which the adenine plus thymine

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TABLE I1 INCORPORATION OF r-31P-NUCLEOSIDETRIPHOSPHATES WITH DIFFERENTDNA PREPARATIONS

DNA primer

T2

RNA r-3zP-Nucleotide incorporated (pmoles) synthesis (pmoles) ATP GTP UTP CTP

4800 1000 T5 4000 SP3 5480 Clostridiurn perfringens 2800 Escherichia coli 2000 Escherichia coli heat-denatured 1300 Micrococcus lysodeikticus 2560 Calfthymus 5700 Calfthymusheat-denatured 2000

T2 heat-denatured

Q

2.3 3.6 1.80 1.25 1.60 0.6 2.4 0.36 1.0 3.8

1.2 5.1 1.4 1.0 2.1 1.5 8.1 2.5 1.8 10.1

0.13 0.88 0.41 0.39 0.28 0.13 0.56 0.10 0.33 0.82

0.10 0.40 0.23 0.12 0.25 0.10 0.44 0.12 0.20 0.66

DatafromMaitraand Hunvitz(39).

content of the DNA is greater than the guanine plus cytosine content are more chains initiated with ATP (for example with DNA from T2, T5, and SP3,but not with calf thymus DNA). Bremer et al.(38)used a different procedure to measure the relative incorporation of nucleotides into the 5 end of RNA chains and found that native T4 DNA also directs the synthesis of primarily ATP-initiated chains. Their results showed that 70% of the chains were initiated with ATP, 17% with GTP, and the rest with UTP or CTP. Travers (1.26)’ in determining the sequences at the 5‘ end of RNA molecules synthesized invitro with T4 DNA as templates, observed that all of the chains initiated with either ATP- or CTP have a uridine in the second position and either a uridine or a cytidine in the third position. Since the codons specifying initiation of protein synthesis in E. eoli are Travers’ results indicate that translation may not AUG and GUG (127), commence with the first nucleotides at the 5’ end of T4 RNA chains synthesized in vitro. The results presented in this section indicate that RNA polymerase can be very selective in its transcription of native, double-helical DNA’s. After denaturation, a DNA is transcribed much less selectively. Nevertheless, initiation from denatured DNA’s is not completely random: most RNA molecules are still initiated with B purine nucleotide, preferentially a guanosine nucleotide. Thus, purine initiation probably reflects only one level of restriction in transcription. The other level of restriction is reflected in strand selection, regional selection, and by the uniqueness of the sequence pppR-U-- at the 5’ termini of T4 RNA synthesized in vitro.

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VI. The Mechanism of RNA Chain Initiation The important findings concerning the selective nature of RNA trandescribed in the previous section indicate that the initiascription invitro tion of RNA synthesis by purified ltNApolymerase must be a very specific process. Thus, it is of great interest to understand the mechanism of RNA chain initiation and particularly to understand how RNA polymerase recognizes those sites on the DNA at which transcription is to begin.

A. The Bind,ing of RNA Polymerase to DNA A necessary first step in the initiation process is the interaction between the enzyme and the DNA. It can be shown by several methods that RNA polymerase can bind to DNA even in the absence of the four nucleoside 98, 128-132). Under certain conditions, the association triphosphates (74, is sufliciently strong enough to allow separation of bound enzyme from free enzyme by slow procedures such as zone sedimentation or zone electrophoresis (128, 129). With the use of these methods as well as faster ones, the following observations concerning the binding of RNA polymerase to DNA have been made. 1. The reaction is fast. A complex between native T4 DNA and RNA polymerase is detectable less than 15 seconds after the addition of enzyme to the DNA solution (62). This has been shown by the membrane filter technique (98). 2.The binding is reversible. Once bound to one DNA molecule an Thus, addition of enzyme can become free and bind to another (131). unlabeled enzyme will dilute labeled enzyme already complexed to DNA (132). However, there may be some sites on the DNA to which the enzyme especially when the ionic strength is less than binds irreversibly (1S3, 134), 0.1. 3.The association is very sensitive to the ionic strength and to the pH of the buffer used (131). The higher the ionic strength, the greater the dissociation constant, and a t concentrations of KaCl or KCl that aregreater than 0.35M it is no longer possible to detect complex formation between This result corresponds well with the fact enzyme and T7 DNA (98, 130). that the enzyme also is not able to initiate synthesis from T7 DNA in buffers with such high molarities (62). There is also some indicatioii t h t at very low ionic strength (r/2 < 0.05) the binding reaction is very slow (U2). This could be a reflection of the fact that both the polymerase and the DNA have net negative charges at pH 8 and thus, without shielding by enough counterions, the two molecules may have trouble interacting.

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The effect of pH is less studied, but higher pH does lead to lower Most . binding experiments are done at the pH binding affinities ( i 3 1 ) optimum for the emyme, which is pH 7.9 to 8 (measured at 20°C) in Tris-HCI. 4. Magnesium ions are not required for binding (129, 131). 5.The binding reaction is by itself specific under appropriate conditions (74, 131,132,98,66). This is indicated by the finding that the amount of enzyme that can bind to a particular DNA species is much less than that expected if the binding reaction were completely nonspecific and were limited only by the number of enzyme molecules that could be closely packed on the DNA. However, under some conditions, the binding does not seem to be limited by the specificity of attachment. The enzyme does seem to have a small affinity for all regions on the DNA and as the ionic strength of the binding mixture is lowered, this weaker attachment is also detected by the techniques used to measure binding (66,131). It is likely, therefore, that some of the initial measurements of binding saturation yielded values too high for the number of specific sites present because the conditions in those experiments were chosen to favor the binding reaction; thus they could easily have included the measurenient of the weaker binding. The values from these earlier experiments were in the range of 2-3pg of polymerase bound per microgram of DNA, for several different species of DNA (T7, polyoma, papilloma, A). Recently, Pettijohn and Kaiiiiya have found that most, if not all, of the weaker, nonspecific attachment of RNA polymerase to polyoma DNA is eliminated if the ionic strength of the buffer is as high as 0.18 (66). I n a buffer having this ionic strength, using the sucrose gradient technique, they measured a saturation value of 0.64.9 pg of IWA polymerase bound per microgram of polyoma DNA. Since the molecular weight of the active unit of RNA polymerase appears to be between 300,000 and 370,000, this saturation value suggests that there is an average of one binding site per 330,000 to 600,000 molecular weight units of DNA, or between five and nine sites per polyonia DNA molecule. Such a value is consistent with an estimate of one binding site per cistron. 6. Binding sites appear to be situated throughout the DNA. Electron polyoma DNA, micrographs of RNA polymerase bound to T7 DNA (73), and papilloma DNA (74) show that the binding sites are discrete. However, it has not been possible to show whether the distribution is random or not. The experiments with cyclic polyoma and papilloma DNA’s do show, in addition, that RNA polymerase can attach to a DNA with no free ends. 7. RNA polymerase also binds t30single-stranded DNA. In fact, it can be shown by direct binding studies that there are more sites per unit weight on a single-stranded DNA than on a double-helical DNA (131). As

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much as 12 pg of RNA polymerase can bind to 1 pg of the single-stranded DNA of the phage f l under conditions in which 1 pg of T7 DNA is saturated with 2 pg of enzyme. Since niost single-stranded DNA’s studied (including denatured DNA’s) are siniilar in enzyme binding properties and it is likely that there are as inany binding template properties (113, 135), sites per unit weight in the other single-stranded DNA’s. In certain other respects the binding of enzyme to single-stranded DNA and to double-helical DNA is similar. The attachment of RNA polymerase to single-stranded DNA is reversible, is sensitive to the salt concentration, and does not require Mg2+ ions (131, 62). However, competition studies between f lDNA and T7 DNA suggest that on the average the affinity of RNA polymerase persitemay be as much as three times greater for the sites on f l DNA. Further, comparison of the shape of the binding saturation curves for these two DNA’s suggests that there is a greater variation in the affinity of the enzyme for different sites on f l DNA than for those on T7 DNA. It is possible, therefore, that transcription by RNA polymerase of denatured DNA is less specific than that of double-helical DNA because RNA polymerase is able to bind to many more sites on a denatured DNA and can use many, if not all, of the attachment sites as points of RNA chain initiation.

B. The Nature of the RNA Polymerase Attachment Site Since RNA polymerase appears to be able to recognize specific signals that determine the points of attachment of the enzyme to DNA, it is of interest to determine what these signals are. One possibility is that RNA polymerase can recognize particular nucleotide sequences. However, since the backbone of a double-helical DNA appears t o be very regular, it is not completely obvious how an enzyme could recognize sequences of bases that are hidden in the interior of the double-helix. Since the structure of DNA is dynamic (136), there might be sufficient unwinding and rewinding of the helix at any given locus to permit the enzyme to recognize a sequence while the DNA is temporarily unwound. Not enough is known yet about the structure of DNA to be certain whether this could happen. However., evidently are able to other very specific enzymes, such as methylases (137), recognize particular nucleotide sequences, so there must be a way for this to be done. The fact that RNA polymerase binds t o single-stranded DNA, but with much less specificity than to double-stranded DNA, prompted Chamberlin, 138)to suggest that all binding sites may be in Wood, and Berg (135, single-stranded regions. The smaller number of sites on normally doublehelical DNA’s would be those rare places of single-strandedness, such as at

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the ends of the molecule, at regions of single-strand breaks, and at points where there is a collapse of the normally helical structure. Of these three possibilities, only the last is likely to be of importance in determining natural binding sites because the enzyme binds perfectly well to closed cyclic forms of DNA, forms that have neither free ends nor single-strand Breaks and ends, however, could well be sources of artificial breaks (74). binding sites. If so, this would explain why much of the selectivity of transcription is lost when a DNA is badly damaged or degraded. If an unwound region is the signal for an RNA polymerase attachment site, some other special feature is needed for the enzyme to recognize which of the two strands are to be copied. It is still quite likely that this might be done by a special nucleotide sequence or, as the following experiments suggest, by certain types of sequences. Studies of RNA synthesis with certain synthetically prepared DNA homo and copolymer pairs such as (dG,. dC,), (dA, . dT,), and d(A-G), . d(T-C), have indicated that RNA polymerase has a strong preference for theuse of the pyrimidine strand 139, 140). This preference has also been of these pairs as the template (50, reflected in binding studies with single-strand components of the homopolymer pairs; RNA polymerase is able to bind more strongly to poly dT than to poly dA (98). Since these are homopolymers, this difference in binding probably reflects a difference in affinity rather than a difference in number of sites. Thus, it is possible that the polymerase could make its choice by binding to the strand with the greater pyrimidine content in the unwound region. Since it is likely that RNA polymerase actually binds to both strands of double-helical DNA, this must mean that the pyrimidinerich strand would be bound by the site on the enzyme specific for the template strand. Various studies have shown that stretches of pyrimidine nucleotides and purine nucleotides on opposite strands of double-helical DNA do exist (141-143). RNA homopolymers are able to form stable complexes with single-strands of DNA (1.64)and many DNA species show a difference in the capacity of the two different strands t o bind these RNA homopolymers (l&-147). The ability of a DNA strand to form such complexes is thought to result from the esistence of regions in the DNA where there may be as many as ten or more identical nucleotides in a row. Thus, a DNA strand that forms complexes with poly G is considered to have at least one if not several regions containing stretches or clusters of deoxycytidine residues. This property has made it possible to fractionate the two strands of various DNA’s on the basis of complex formation. The separated strands then can be compared for their ability to form RNA DNA hybrids with natural RNA’s. With T7 DNA, this has led to a striking result: only one of the two strands forms complexes with poly G and it is this strand that is comple1

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nientary to all the natural T7-specific RNA (148). With X DNA, both strands form complexes with poly G, but again the strand that binds the greater amount of this homopolymer is also the strand that is complemenThus, in tary to the greater number of different A-specific mRNA’s (116). these two cases, it appears that the DNA strand with the greater number ofdeoxycytidine clusters is the same strand that has more regions complementary to RNA from that DNA. This is consistent with the interpretation that these pyrimidine-rich clusters, which are presumed to be responsible for the complex formation with poly G, are those that orient the direction of RNA polytnerase transcription. Since other DNA’s appear to have similar clusters of thymidine, it is possible that this enzyme orientation could bc a general feature of pyriniidine clusters (145-147). Another attractive feature of these putative homopolymer runs in natural DNA’s is that such regions in a double-helical DNA could have a n important difference in structure that might make them recognition points for RNA polymerase attachment. X-Ray crystallographic studies of DNA homopolymer pairs [e.g., (dG), . (dC), and (dA), . (dT),] show that these particular types of DNA do have structures that are different Thus, a region in a double-helical from natural double-helical DNA’s (149). DNA containing a run of cytidine (or thymidine) nucleotides on one strand and guanosine (or adenosine) nucleotides on the other could have a structure different from the rest of the DNA, made of mixtures of nucleotides. It would be of considerable interest to isolate regions of DNA that bind RNA polymerase and determine whether there is something special about the nucleotide composition and sequences in such regions. One method that has been tried is to digest away the unbound portions of DNA with DNA nucleases (for example, a mixture of pancreatic deoxyribonuclease and venom phosphodiesterase), and then to isolate those regions rendered Unfortunately, the resistant by the presence of the polymerase (62, 150). difficulty associated with this approach is that the binding of the RNA polymerase is reversible; thus, during the digestion, an enzyme molecule may leave a “natural” binding site and go to any of the single-stranded regions created by the nuclease action. What is needed is some method to attach the polymerase more firmly to the DNA-irreversibly during the course of the digestion-but only at a specific binding site. The addition of nucleoside triphosphates, to start thc reaction, irreversibly binds the polymerase to the DNA; but atI the same time the enzyme molecules move away from starting sites as synthesis begins. Moreover, it has not yet been possible t o make the reaction start with complete synchrony. However, it does seem likely that some method will be found to attach the polymerase firmly to its specific binding sites. Then it should be possible to do this importmt experiment.

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C. The Initiation Reaction The kinetics of chain initiation can be measured by the rate of appearance of newly synthesized RNA chains. Under conditions in which each enzyme makes only one RNA molecule from a double-helical DNA template (e.g., low salt concentrations), it has been found that it takes nearly 2 minutes a t 37 C for half the chains that are eventually to be made to be 39, 151). However, the binding reaction is complete in less initiated (38, than a few seconds, and once initiated, RNA chains grow at, a rate of a t least 3 nucleotides per second. Thus, there seem to be one or more ratelimiting steps between biridiiig of RNA polymerase to DNA and the commencement of normal polymerization. Walter et al.(152) have suggested that i t might be neccssary, after enzyme attachment, for RNA polymerase to unwind the DNA in ordcr to expose a strand for use as a template and that this might be the ratelimiting step of initiation. This suggestion, of course, is based on the presupposition that the enzyme does not normally bind to an already unwound region. The basis of this hypothesis was an analysis of factors that influence initiation. Normally, at 37"C,there is no obvious lag in the rate of RNA synthesis, but at lower temperatures a lag is evident and the duration of the lag increases with decreasing temperature. Since the binding reaction itself is not greatly inhibited by lower temperatures, Walter et al.suggested that the strong temperature dependence of the unwinding of the DNA helix might be responsible for the apparent temperature sensitivity of initiation. The fact that a similar lag is not observed when denatured DNA is used as template is consistent with the suggestion that DNA unwinding is the rate-limiting step for initiation of transcription from double-helical DNA templates. To prove this point, it would be helpful to have accurate physical measurements showing that RNA polymerase can destabilize a region of DNA merely by binding to such a region. Some attempts t o make such measurements have not been successful, most probably because the effects, if there are any, are too small to be detect,ed by the methods employed (131,62). Also, the presence of the enzyme on the DNA may produce other physical changes that could easily mask a change resulting from local denaturation. An example of this has come from attempts to measure hypochromicity of the DNA produced, it is assumed, by the presence of the I n this case, there appeared to be other spectral changes that enzyme (62). were more important; these changes probably resulted from changes in light-scattering following the interaction of enzyme and DNA to form a larger, complex particle. Initint,ion of RNA cahains commences with the cnndensation of the

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second nucleoside triphosphate to the first one. This can occur only after both nucleotides have been bound in their proper sites in the enzymeDNA complex. It is quite likely that the binding of the very first nucleotide is different from the binding of the rest because there is no 3’hydroxyl group to which the first nucleotide can be added. It has been suggested by Anthony etal.that the rate-limiting step in initiation may be the binding of this first nucleotide (153). They have found that, when certain nucleoside triphosphates are present in low concentrations, there is a noticeable lag in initiation. This Iag is more evident when the purine nucIeoside triphosphates are limiting. Since RNA chains are known to be initiated preferentially with a purine nucleotide, this observation would be consistent with the proposal that initiation is more sensitive to the substrate concentration than subsequent steps iu synthcsis. As Anthony et al.have pointed out, to a much higher I<,for however, the rate limitation could be due either the binding of the first nuclevtide, or to the possibility that the initiation step is a biinolecular reaction for the first two nucleotides. An important change occurs in the stability of attachment of RNA polymerase to DNA sometime during or after the initiation of polymerization. In the absence of RNA synthesis, the binding of enzyme to DNA is reversible; when an RNA chain is being synthesized, binding is irreversible a t least until the RNA molecules have reached a minimum length (36, 130, 131,37,93,154). There is a good reason for the stability of the enzyme attachment during the polymerization: it prevents the release of unfinished chains. What is not known yet is at what poirit after initiation the attachment becomes stabilized. It could be that the binding of the first nucleotide triphosphate is sufficient for stabilization of the attachment of the polymerase to the DNA. Anthony et al.have presented some evidence that incubation of the polymerase with two nucleoside triphosphates leads t o a However, significant stabilization of the enzyme to the complex (153). since it is difficult to account for all polymerase molecules in their experiments, it is not certain whether this stabilizabion represents merely the stability of the very few enzyme molecules that might have started synthesizing a small chain. In contrast, the complexes made bctwecn RNA polymerase and singlestranded DNA templates never become as stable as those made with double-helical templates. This is evideiit from the fact that the RNA molecules made from single-stranded DNA templates are much smaIIer (133, 151,39) and that, unlike the process with double-stranded DNA as template, new molecules continue to be initiated throughout the reaction, These observations suggest that either even in low salt buffers (39, 155). the nonteniplat,e strand in doutde-helical DNA or the RNA strand being

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displaced is very important for the stability of the complex of RNA polyniera.se, nascent RNA4and double-helical DNA.

D. Initiation Sites It has been noted, in several investigations of the binding of RNA polymerase to DNA, that the sites on many DNA species are not homogeUnder some circumstances, neous with respect to binding affinity (l.%t?-IS4). the enzyme can be bound irreversibly to sites having unusually high affinity for the enzyme. Stead and Jones estimated that as many as 8 of the 50 sites on T7 DNA and about 5 sites on X DNA are vf this sort (194). It is possible that these sites for which RNA polymerase has such a high Because of this possibility, affinity are the only true initiation sites (256). it is of interest to firid methods for determining the number of true initiation sites on a DNA molecule. Brerner etal.estimated the number of initiation sites on T4 DNA by a method that is based on the fact that some RNA polymerase molecules in These enzyme molecules, known czs their preparations are defective (lS3). “early quitters,” are able to initiate the synthesis of a RNA chain hut cease to function within a minute or two after initiation (38).It is believed that after an ‘(early quitter” has stopped functioning it effectively prevents that site to which it is attached from being used for transcription of large (M.W. > lo5) RNA molecules. In fact, Bremer etal. found that even with excess RNA polymerase, no more than 180large RNA molecules could be Using this figure as well as an made for each T4 DNA molecule (133). estimate of the fraction of enzyme molecules that were “early quitters” in their preparations, these authors calculated that there are about 180 initiation sites on T4 DNA. It is very likely that there are as many as 180 cistrons 011 the T4 genome. However, more than half of the total number of cistrons probably specify late functions (21, 157). Since RNA polymerase transcribes only “early” RNA from native T4 DNA, one would expect to find at most RNA polymerase on T4 DNA; between 80 and 90initiation sites for E.coli thus, the value obtained by Bremer and his colleagues seem rather high. Nevertheless, considering the assumptions involved both in their calculation and in the “genetic” estimate of the iiwnber of initiation sites, the agreement between the two values is not bad. Certainly, no conclusions about the mechanism of enzyme action can be derived on the basis of such a difference. used a second method to measure the number of initiation Sentenac etal. They devised a technique to measure the number sites on T7 DNA (258). of GTP-initiated RNA chains synthesized in the presence of 3’-deoxy-

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adenosine 5’-triphosphatc, an ATP analog, which, as is mentioiled in Section IV, B of this review, is incorporated into RNA chains but prevents further elongation. From the data of Sentenac etal.it is estimated that a maximum of 50 GTP-initiated chains can be synthesized per T7 DNA molecule in the presence of this inhibitor. Assuming that each site could be used only once in this situation, this value indicates the number of GTP sites on T7 DNA. Under normal conditions of RNA synthesis (without inhibitor and with DNA in excess), two-thirds of the T7 RNA chains commence with GTP and one-third with ATP. If this ratio represents the relative frequency of sites for the two kindsof chains, then there should be R total of 75 initiation sites per 1’7 DNA molecule.

VII. Termination Signals The fact that the RNA found in the cell is the product of transcription of distinct genetic units on the DNA suggests that there must be specific mechanisms for terminating the transcription of RNA molecules as well as for initiating them. Thus, it has been postulated that there may be sites on the DNA that are recognized by the RNA polymerase as points where RNA polymerization is to be terminated. Presumably such a site would also cause the newly synthesized RNA and the polymerase to be released from the DNA at that point. It has not been possible to obtain convincing evidence that such termination signals exist. I n fact, the little evidence that does pertain to this subject supports the opposite conclusion, that RNA polymerase is not able to recognize specific “stop” signals. It appears attain sizes that are much larger that the RNA molecules made invitro than those normally found for RNA molecules made on the same kind of DNA inside the cell. The bulk of T4 RNA synthesized invitro isolated after 20 minutes of reaction sediments faster than 25 S (86) whereas the peak in the sedimentation profile of T4 RNA synthesized invivois only at 14 S (159). These differences could result merely from degradation of the invivoRNA during isolation or from a prefereiitial synthesis invitro of the longer transcriptive units. It is also possible that the units of transcription are large for T4 DNA and that tjhereis a special mechanism for cutting the T4 messengers into smaller pieces after synthesis. Not enough is known yet about the existence of operons in T4 to rule out such a hypothesis. For this reason, differences in the size of the RNA’s made invitro and invivo is not sufficient to prove that RNA polymerase does not recognize termination signals invitro. In their analysis of T4 transcription in vitro, Bremer and Konrad found that the RNA molecules are not released from their complex with the enzyme and DNA during the course of the reaction (86). This could

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also be construed as an inability of the polymerase to recognize termination signals. However, in reaction mixtures with higher salt concentrations the RNA molecules are released during the reaction. This may mean that termination signals are recognized only under some conditions, but it is also possible that release of the enzyme and the RNA stimulated by high salt is a random process; once nascent RNA molecules reach a certain size, their attachment may become very unstable. If termination signals exist on DNA, what might they be? One possibility is a rare nucleotide that is recognized in DNA synthesis but is not recognized by one of the ribonucleoside triphosphates during RNA synthesis. Another, more likely, possibility is that there is a certain sequence of nucleotides in the DNA that, when it enters the enzyme, greatly lowers the stability of enzyme attachment. As was pointed out earlier, the nontemplate strand may be very important in stabilizing the enzyme-DNA complex; thus, some sequence on that strand may determine release. Although it is easy to propose models for termination, the problem still remains to demonstrate that the enzyme can even recognize a signal, Unfortunately, proof of the existence of such signals may have to rest on negative results; it is necessary to show that an RNA molecule that is in part homologous to one genetic region is not also homologous toan adjacent, genetically distinct region. Such a negative result has already been found in the case of T4 transcription. Geiduschek et al.have shown that nearly all, if not all, tlhe species of “early” T4 RNA are represented in the invitroproduct, whereas levels of “late” T4 RNA are very low (122). Although the early and late genes on the T4 genome are clustered in large groups, a few (perhaps 4 or 6)early genes are adjacent to late genes; thus something may prevent the passage of RNA polymerase from these early gene regions to adjacent late gene regions. However, the initiation sites for the early genes may all be a long way from the start of late genes, and thus there may always be a good probability that the enzyme cannot transcribe everything in between without being detached randomly at some stage. It cannot be ruled out, from the data of Geiduschek et al., that RNA from the early genes just adjacent to the late genes on the genetic map are also made in very low quantities. If termination signals do not exist, it is necessary to postulate other factors to separate RNA molecules of one operon from those of another. Jones etal.(106) presented evidence that the ribosomes can catalyze what appears to be a specific release of RNA molecules from the enzymeDNA complex. This could be the manifestation of a specific RNA detaching mechanism. However, there remains the problem of whether RNA polymerase can proceed to synthesize RNA from adjacent operons under the control mechanism of the first operon. Ifoperons are oriented in the same

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direction, a repressor on the second could prevent the polymerase from proceeding far beyond the end of the first. If operons are oriented in opposite directions then the RNA made from the second operon would be ‘Lanti-message”and therefore probably would be nonsense. Although this anti-messenger” RNA might be degraded rapidly, such a mechanism seems wasteful. It therefore remains likely that termination signals do exist and are read by R.NA poIymerase. It is only necessary to find the appropriate system to demonstrate their existence. ((

VIII.Inhibitors A. lnhibitors of Polymerization One of the best-known inhibitors of RNA polymerase is actinomycin (160), which acts by blocking polymerization of RNA chains. At a concentration of 1 p M , actinomycin has little effect either on the binding of

RNA polymerase to DNA (131) or on the initiation of RNA chains (155); however, the RNA chains made in its presence are much shorter than usual (87, 155). Since this antibiotic is known to bind very strongly to DNA (161), it could block polymerization by preventing RNA polymerase from moving through a region of the DNA to which it is bound. Even though actinomycin binds very specifically t o G . C pairs in double-helical DNA (f60),its low level of inhibition of initiation does not necessarily imply that there are no G . C pairs in the initiatioii site. At much higher coneentraHowever, at the tions (10 p M ) ,actinomycin also inhibits initiation (162). low concentration that blocks polymerization selectively, the probability that actinomycin binds to an initiation site might be too low to be noticeable. It is also possible that a difference in structure at the RNA polymerase attachment site on DNA may make it less probable that an actinomycin molecuIe can bind there. Another inhibitor of polyinerization is 3‘deoxyadenosine 5’-triphosphate. Because this analog of ATP can be incorporated into RNA chains in pIace of ATP, it inhibits by preventing further chain elongation (94).

B. Inhibitors of Initiation Of the known inhibitors of irrititttion, most appettr to act by preventing the attachment of RNA polymerase to DNA. For example, proflavine inhibits attachment of RNA polynierase to DNA (131) and reduces the number of RNA molecules that, :ireinitiated in a given time (155), but it has very little effect on the rate with which those chains that we initiated Proffavine is known to bind very efficiently to DNA are polymerized (87). (163) and in so doing to alter the secondary structure of the DKA (164). It

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is likely that it is this change in DNA structure that allows proflavine to prevent RNA polymerase binding. However, it cannot be excluded that proflavine also binds to RNA polymerase and alters its ability to attach to DNA. In contrast to proflavine, other inhibitors are known to prevent initiation by directly altering RNA polymerase. In one class of such inhibitors are those that change the affinity of the enzyme for DNA. For example, in a reaction mixture with a salt concentration of 0.5 M , RNA polymerase is unable to initiate transcription (153, 154, 131, 97). However, if synthesis has already started at a lower ionic strength and the salt concentration is then raised to 0.5M , there is very litt81einhibition of further synthesis. Since it is not possible to detect the binding of RNA polymerase to DNA (131, 132,98)or to other polynucleotides (62)(e.g., denatured DNA, tRNA, poly U) at such an ionic strength, it appears that the effect of salt is to lower the affinity of the enzyme for DNA in the initiaI attachment step. The second class of inhibitors that block initiation by altering the enzyme include the many compounds that, by binding to enzyme, prevent it from binding to DNA. This class includes many polynucleotides (108, 109,151, 139, 98, 67)and possibly several other polyanions, including heparin (152) and polyethylene sulfonate (165). The best-studied example of this type of inhlbitor is tRNA. The binding of tRNA and DNA to RNA polymerase is mutually exclusive (131); thus it is likely that tRNA binds either to the same site in the enzyme to which DNA binds or to an overlapping site. At one time it was suggested that tRNA inhibition might be very important as a mea~isof regulation (29, 30). This suggestion was supported by the finding that there is some difference in the capacity of aminoacyl166). However, the tRNA and free tRNA to inhibit RNA polymerase (108, differences in inhibition are not very great, and both forms of tRNA have 6’7). Further, the about the same ability to bind to RNA polymerase (131, inhibitory properties of tRNA are greatly reduced in reaction mixtures Thus, it now seems more likely that inhibition containing 0.2A4 KC1 (61). by tRNA does not play an important role in the regulation of R.NA synthesis. Very recent work has led to the description of two inhibitors that may affect initiation, but not binding. One is the antibiotic known as rifamycin (167). I n potency and specificity it is more effective than actinomycin in inhibiting RNA polymeraae (let?), but it differs from actinomycin i n that it does not bind to DNA (169). This is best indicated by tlie fact that mutants of bacteria have been found that are resistant to this antibiotic, and the RNA polymerase isolated from such a mutant is likewise insensitive to it (169). Thus, the action of rifainyciri seems to be specific far RNA polymerase itself.

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Preliminary evidence indicates that rifainycin affects neither the binding nor the chain polymerization properties of sensitive enzyme (169, 270).Therefore this antibiotic is believed to block some step in initiation after binding. A similar result has been reported for the antibiotic streptovaricin R44P (171).

IX. Conclusions The purpose of this review is to consider the mechanism of RNA polymerase action in terms of the regulation of RNA synthesis. The evidence presented indicates that accurate and selective transcription of DNA can be achieved with very pure RNA polymerase. It also can be concluded that the step that is most important in determining the selectivity of transcription is initiation. However, it is not known whether some initiation sites are preferred over others, or whether transcription rates vary from region to region. Both factors could affect the degree of selectivity and therefore the relative abundance of RNA from different transcriptive units. Nevertheless, the most important aspect of the regulation of gene activity is the absolute control of transcription. It is certain that regions under positive control, such as the Iate genes of T4 and A, depend on the presence of some additional protein factor to be transcribed at all. It also is likely that regions that are subject to negative control, like the lactose are restricted in the initiation of mRNA synthesis. The operon in E.coli, lactose repressor is known to bind t oa region of DNA just adjacent to the region where transcription of the lactose operon is presumably initiated ( I S ,1 74). It appears, therefore, that the action of this repressor is to prevent initiation of transcription a t the site to which it binds. However, this conclusion has not been confirnied by experiment, and it is possible that the repressor could block transcription by RNA polymerase at some step immediately following initiation. I n spite of the advances that have been made in our understanding of RNA polymerase and the reaction it catalyzes, there exist several very fundamental properties that still are not understood. We do not know yet whether RNA polymerase can recognize signals for the termination of RNA synthesis. Further, although much is already known about the mechanism of initiation and transcription, the models that have resulted from this knowledge are vague and imprecise. Finally, we are not even certain of the molecular weight of the functional unit of the enzyme. ACKNOWLEDGMENTS I want to thank Dr. Gary G w i n very much for his help and advice in preparing this review, Professor Alfred TissiBres for the hospitality of his laboratory, and Dr. Edward Brody forhis helpful discussions. The author is a postdoctoral fellow of the American Cancer Society.

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REFERENCES 1. S. B. Weiss and L. Gladstone, J .Am. Chem.Soc.81,4118(1959). 2. D. Elson, Ann.Rev.B w c h .34, 449(1965). 3. D. Ballard and H. G. Williams-Ashman, Nature203, 150(1964). 4.J. J.Furth and P. Ho, J .Bid.Chem.240,2602(1965). 5.M. Ramus, J.Doly, P. Mandel, and P.Chambon, Biochem. Riophys. Rex Commun. 19, 114(1965). 6. J. Hurwits and J. T. August, This series 1, 59 (1963). 7. F. C.Neidhardt, This series 3, 145(1964). 8. 0. Maalne and N. 0. Kjeldgaard, ‘‘Control of Macromolecular Synthesis.” Benjamin, New York, 1966. 9. S. A. Yanofsky and S. Spiegelman, Proc. Natl. Acad.Sci.U .S.48,1069(1962). 10. H. M. Goodman and A. Rich, Proc.Nail. Acad.Sci. U .S.48, 2101(1962). 11.F. Gros, W. Gilbert, H. Hiatt, P.F.Spahr and J. D. Watson, ColdSpringHarbor Symp.Quant. Biol. 26, 111(1961). 12. J. Humits, J.J.Furth, M. Malamy, and M. Alexander, Proc. Natl. Acad.Sci. U.S. 48, 1222(1962). 13.R. B. Roberts (ed.), “Studies of Macromolecular Biosynthesis,” Section VI, Carnegie Inst. Washington, Publ. 624,Washington D. C., 1964. 14. G. Attardi, P. C. Huang, and S. Kabat, Proc. N d l .Acad.Sci. U .S.63,1490(1965). 16. F. Jacob and J. Monod, J .MoZ. Biol. 3,318 (1961). 16. G. Attardi, S. Naono, J. Rouvibre, F.Jacob, and F. Gros, ColdSpringHarbor Symp.Quunt.Biol. 28, 363 (1963). 17. W. Gilbert and B. Muller-HU, Proc. JVutZ. Acad.Sci.U .S.66, 1891(1966). 18. W. Gilbert and B. Muller-Hill, Proc. Natl. Acad.Sci.U.S. 68,2415(1967). 19.D. Sheppard and E. Englesberg, ColdSpringHarborSymp.Quant.Riol. 31, 345 (1966). 20.S. P.Champe, Ann.Rev.MicrobioZ. 17,87 (1963). 21. R. H. Epstein, A. Bolle, C. M. Steinberg, E. Kellenberger, E. Boy de la Tour, R. Chevalley, R. S. Edgar, M. Susman, G. H. Denhardt, and A. Lielamis, Cold SpringHarbm Symp.Quant. Biol. 28, 375 (1963). 22. J.S. Wiberg, M. Dirksen, R. H. Epstein, S.E. Lurk,nnd J. M. Buchanan, Proc. Nail. Acad.Sci.U .S.48,293 (1962). 23. W. F. Dove, J .MoZ. Biol. 19, 187(1966). 24. A. Bolle, R. H. Epstein, W. Salser, and E. P.Geiduschek, J .MoZ. Biol. 31, 325 (1968). 26.A. Joyner, L. N. Isaacs, H. Echols, and W. S. Sly, J .MoZ. BioZ.19, 174 (1966). 26. A. Skalka, Proc. Natl. Acad.Sci. U .S.66, 1190(1966). 27.S. Naono and F. Gros, J .MoZ. Biol. 26, 517 (1967). 28. L. P.Gage and E. P. Geiduschek, J .Mol.Biol. SO, 435 (1967). 29.G. S. Stent and S. Brenner, Proc. Natl. Acad.Sci. U.S. 47, 2005(1961). 30. C. G. Kurland and 0. Maal~e,J .MoZ. Biol. 4,193(1962). 31.G. S. Stent, Science 144, 816 (1964). 32. G. S. Stent, Proc. Roy.SOC. B164, 181(1966). 33.M. Schaechter and K. McQuillen, J .Mol.Biol. 22,223(1966). 34. H. K. Das, A. Goldstein, and L. I.hwney, J .MoZ. Biol. 24,231(1967). 35.G. Mangiorotti and D. Schlessinger, J .MoZ. Biol. 29, 395 (1967). 56. D.H. Alpers and G. M. Tomkins, J .BioE.Chem.241, 4434(1966). S7. L.h i v e and V. Kollin, J .Mol.Biol. 24, 247(1967).

RNA

POLYMERASE

AND

T H E CONTROL

O F RNA

SYNTHESIS

113

38.H. Bremer, M. W. Konrad, I<. Gaines, and G. S. Stent, J .Mot.B i d . 13,540(1965). 39.U. Maitra and J. Hurwitz, Proc. Natl. Acad.Sci. U .S.64,815 (1965). 40. G. Streisinger, Y. Okada, J. Emrich, J. Newton, A. Tsugita, E. Terzaghi, and M. Inouye, ColdSpringHarborSymp.Quant. Bid.31,77 (1966). 41. R. Thach, T. A.Sudararajan, K. F. Dewey, J. C. Brown, and P. Doty, ColdSpring HarborSymp.Quant. BioE.31,85 (1966). 42.A. J.Wahba, M. Salas, and W. M. Stanley, Jr., ColdSpringHarborSymp.Quant. Biol. 31,103(1966). 4s. G. Contesse, S. Naono, and F. Gros, Compt.Rend.Acad.Sci.263,1007(1966). 44.F. Imamoto and C. Yanofsky, J .MoZ. Biol. 28,1 (1967). 46.F. Imamoto and C. Yanofsky, J. MoE. Bid. 28,25 (1967). 46.A. S.Sarabhai, A. 0. W. Stretton, S. Brenner, and A. Bolle, Nature201,13 (1964). 47.S. Brenner and A. 0. W. Stretton, 6.Mol.Biol. 13,944 (1965). 4.9.S. Naono, J. Rouv2re and F. Gros, Biochim. Biophys. Acta129,271 (1966). 49.D. W. Morris and N. 0. Kjeldgaard, J .M o l .Biol. 31,145(196s). 50.M. Chamberlin and P. Berg, Proc. Natl. dcud.Sci.U .S. 48,81 (1962). 61.J. J. Furth, J.Hurwitz, and M. Anders, J .Biol. Chem.237,2611(1962). 52.A. Stevens and J. Henry, J. Biol. Chem.239,196(1964). 53.W. Zillig, E. Fuchs, and R. Millette, i n “Procedures in Nucleic Acid Research” (G. L.Cantoni and D. R. Davies, eds.), p. 323.Harper and Row, New York, 1966. 54.J. P. Richardson, Proc.Natt. Acad.Sci. U.S.66,1616(1966). 65.C. Babinet, Biochem. Biophys. Res.Commun. 26,639 (1967). 56.R. L. Ratliff, D. A. Smith, and T. T. Trujillo, Los Alamos Scientific Laboratory, University of California, Los Alamos, New Mexico, Report LA-3638,1967. 67.A. Ishihama and T. Kameyama, Biochim. Biophys. Acta138,480 (1967). 58.U. Maitra and J. Hurwitz, J .Biol. Chem.242,4897(1967). 69.R. Burgess, Federation Proc.27,295 (1968). 60.C. A. Thomas, Jr. and J . Abelson, in “Procedures in Nucleic Acid Research” (G. L.Cantoni and D. R. Davies, eds.), p. 553.Harper and Row, New York, 1966. 61. A. G. SO,E. W. Davie, R. Epstein, and A. Tissikres, Proc. N d l .Acad.Sci. U .S.68, 1739(1967). 62.

J. P. Richardson, unpublished observation.

63.A. Stevens, A. J.Emery, Jr., and N. Sternberger, Biochem. Biophys. Res.Commun. 24,929(1966). 64.H. Pdess and W. Zillig, Biochim. Biophys. Acta140,540(1967). 66. R. Burgess, personal communication. 66. D. Pettijohn and T. Kamiya, J .Mol.Biol. 29,275 (1967). 67.H. Bremer and G. Stent, ActaBiochim. Polon. 13,367 (1967). 68. D. A. Smith, A. M. Martinez, R. L. Ratliff, D. L. Williams, and F. N. Hayes, Biochemistry 6 ,3057(1967). 69. E. Fuchs, W. Zillig, P. H. Hofschneider, and A. Preuss, J .Mol.Biol. 10,546 (1964). 70.A. J. E. Colvill, E. F. J. van Bruggen, and H. Fernhndez-Morhn, J .Mol.B i d .17, 302 (1966). 71.E. M. Crawford and L.V. Crawford, personal communication. 72.M. Lubin, personal communication. 73.H. S. Slayter and C. E. Hall, J .Mol.Biol. 21,113(1966). 74.L.V. Crawford, E. M. Crawford, J. P. Richardson, and H. S. Slayter, J .Mol.Biol. 14,593(1965). 76.E. P.Geiduschek, T. Nakamoto, and S. B .Weiss, Proc.Natl. Acad.Sci. U .8.47, 1405(1961).

114

JOHN

P. HICHARDSON

Y6. R. L.Sinsheimer, in“The Nucleic Acids” (E. Chargaff and J. N. Davidson, eds.), Vol. 3,p.187.Academic Press, New York, 1960. 77.M. Chamberlin, R. L. Baldwin, and P. Berg, J. Mol.Biol. 7 ,334 (1963). 78.M. Chamberlin and P. Berg, J .Mol.Biol. 8,297 (1964). Y9. K. Psigen, 6.Theoret. Biol. 3,268 (1962). 80. G. Stent, Advan.VirusRes.6, 95 (1958). 81. A. Rich, Proc. Natl. Acad.Sci.U .S.46,1044(1960). 8.9. G. Zubay, Proc.Natl. Acad.Sci.U.S.48,456 (1962). 83. G. Milman, R. Langridge, and M. J. Chamberlin, Proc.Natl. Acad.Sci. U .S .67, 1804(1967). 84. R. L. Sinsheimer and M. Lawrence, J. Mol.Biol. 8,289(1964). 85. M. J. Chamberlii, Federation Proc.24,1446(1965). 86.H. Bremer and M. W. Konrad, Proc. Natl. Acad.Sci.U .S.61,801 (1964). 87. J. P. Richardson, J. Mol.Biol. 21,115(1966). 88. M. Hayashi, Proc.Natl. Acad.Sci. U.8. 54, 1736(1965). 89. A. K. Kleinschmidt, A. Burton, and R. L. Sinsheimer, Science 142,961(1963). 90. J. Vinograd, J. Lebowitz, R. Radloff, R. Watson, and R. Laipis, Proc.Natl. Acad. Sci. U.S.63,1104(1965). 91. L. V. Crawford and M. J.Waring, J. Mol.Biol. 86, 23 (1967). 92.P. H. Pouwels, C. M. Knijnenburg, J. van Rotterdam, and J. A. Cohen, J .MoZ. Biol. $2, 169 (1968). 93. U. Maitra, A.Novogrodsky, D.Baltimore, and J. Hurwitz, Biochem. Biophys. Res. Cmmun. 18,801(1965). 94.J. T. Shigeura and G. E. Boxer, Biochem. Biophys. Res.Commun. 17,758 (1964). 96. M. Grunberg-Manago, This series 1,93 (1963). 96. P. F.Spahr, J .Biol. Chem.239, 3716(1964). 97. E. Fuchs, R. L. Millette, W. Zillig, and G. Walter, EuropeanJ. Biochem. 3, 183 (1967). 98. 0. W. Jones and P. Berg, J. Mol.Biol. 22,199(1966). 99.A. P. Nygaard and B. D. Hall, Biochem. Biophys. Res.C m m u n . 12,98 (1963). 100. R. L. Mdette, Referred to in Fuchsetal. (97). 99, 362 (1967). 101. H. Bremer, Mol.Gen.Genet. 102.H. Bremer and D. Yuan, J. Mo2. Biol. 34,527(1968). 10.9. D. H. Shin and I(.Moldave, J. Mol.Biol. 21,231(1966). 104. M. Revel and F. Gros, Biochem. Biophys. Res.Commun. 27,12 (1967). 106. 0. W. Jones, M. Dieckmann, and P. Berg, J .Mol.Biol. 31,177 (1968). 106. M. Lubin and H. L. Ennis, Bwchim.Biophys. Acta80,614 (1964). 107. J.S. Krakow, J. Biol. Chem.241,1830(1966). 108. A. Tissihres, S. Bourgeois, and F.Gros, J .Mol.Biol. 7, 100(1963). 109. C. F.Fox and S. B. Weiss, J .Biol. Chem.239, 175(1964). 110.M . Hayashi, M. N. Hayashi, and S. Spiegelman, Proc. Natl. Acad.Sci. U .8.61, 351 (1964). 11i. A. J. E.Colvill, I,. C. Kanner, G .P.Tocchini-Valentini, M. T. Sarnat, and E. P. Geiduschek, Proc.Natl. Acad.Sci.U .S.63,1140(1965). 118. E. Brody, personal communication. 119. J. Hurwitz, J. J. Furth, M. Anders, and A.Evans, J. Biol. Chem.237,3752(1962). 114. C.L.Sehildkraut, C. C. Richardson, and A. Kornberg, J. Mol.Biol. 9, 24 (1964). 116. E. P. Geiduschek, G. P. Tocchini-Valentini, and M. T. Sarnat, Proc.Natl. Acad. Sci.U .S .62,486 (1964).

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116.K. Taylor, Z. Hradecna, and W. Szybalski, Proc. .Vnll. .lend. Sci.U.S. 67,1618 (1967). 117.S. N. Cohen and J. Hunvitz, Proc. Natl. h a d .Sci.U . S.67, 1759(1967). 218.E. P.Geiduschek, J. Moohr, and S. R. Weiss, Proc.Nail. h a d .Sci. U.S .48, 1078 (1962). 219.E. P. Geiduschek, E. N. Brody, and D. L. Wilson, in"Molecular Associations in Biology" (B. Pullman, ed.), p.163.Academic Press, New York, 1968. 120.R. B. Khesin, M. F. Shemyakin, Zh. M. Gorlenko, S. L. Bogdanova, and T. P. Afanasevs, Biokhimiya %7, 1092(1962). 121.E. P. Geiduschek, Bull. SOC.Chin.Bid. 47,1571(1965). 19%.E. P. Geiduschek, L.Snyder, A. J. E. Colvill, and M. Sarnat, J .Mol.Biol. 19, 541 (1966). 123.S. Naono and F.Gros, ColdSpringHarborSymp.Quant.Biol. 31, 363 (1966). 12.4. S. N. Cohen, U. Maitra, and J. Hurwitz, J. Mol.Biol. 26, 19 (1967). 126.L. Snyder and E.P.Geiduschek, Proc. Natl. Acad.Sci. U. S .69, 459 (1968). 126.A. Travers, personal communication. 187.K. A. Marcker, B. F. C. Clark, and J. S. Anderson, ColdSpringHarborSymp. Quant.Biol. 31, 279 (1966). 128.M. Kodoya, H. Mitsui, Y. Takagi, E. Otaka, H. Suzuki, and S. Osawa, Biochim. Biophys. Actu91, 36 (1964). 129.C. F. Fox, R. I. Gumport, and 5. B. W e k , J .Biol. Chem.240, 2101(1965). 150.P. Berg, R. D. Kornberg, H. Fancher, and M. Dieckmann, Biochem. Biophys. Res. Commun. 18, 932 (1965). 131.J. P. Richardson, J. Mol.Biol. 21, 83 (1966). 152.N. Sternberger and A. Stevens, Biochem. Biophgs. Res.Commun. 24, 937 (1966). 133.H. Bremer, M. W. Konrad, and R. Bruner, J .Mol.Biol. 16, 104(1966). 134.N. W. Stead and 0. W. Jones, J .Mol.B i d .26, 131(1967). 136.W. B.Wood and P. Berg, J. Mol.Biol. 9, 452 (1964). 136.P.H. von Hippel and M. P. Printz, Federation Proc. 24,1458(1965). 197. M. Gold and J. Hurwitz, J. Biol. Chem.239, 3866(1964). 158.M. J.Chamberlin and P. Berg, J. Mol.Biol. 8,708(1964). 159. M. J.Chamberlin, personal communication. 140.S. Nishimura, D. S. Jones, and H. G. Khorana, 3.Mol.Bzol. 13, 302 (1965). 1.41. J. H. Spencer and E.Chargaff, Biochim. Biophys. Acta68, 18 (1963). 142.K. Burton, M. R. Lunt, G. B. Petersen, and J. C. Siebke, ColdSprinQHarborSymp. Quant. Biol. 28, 27 (1963). 143.J. B.Hall and R. L.Smsheimer, J. Mol.Biol. 6,115(1963). 144.Z. Opara-Kubinska, H. Kubinski, and W. SzybaIski, Proc. Natl. Acad.Sci. U .S. 62, 923 (1964). 145.H. Kubinski, Z. Opara-Kubinska, arid W.Szybalski, J. Mol.Biol. 20,313 (1966). 146.W. Szybalski, H. Kubinski, and P. Sheldrick, ColdSpring HarborSymp.Quant. Biol. 31, 123(1966). 147.P. Sheldrick and W. Szybalski, J. MoZ. Biol. 29, 217 (1967). 148.W. C. Summers and W. Szybalski, Virology 34,9 (1968). 149.R. Langridge, 7thIntern. Congr. Bwchem.,Tokyo,1967. Abstr. 1, 57 (1967). 160.F. R. Blattner and C. A. Thomas, Jr.,7thIntern. Cmgr.Bwchem.,Tokyo, 1987. Abstr. 4, 656 (1967). 101, 6 (1968). 151.H. Bremer and R. Bruner, Mol.Gen.Genet. 16%.G. Walter, W. Zillig, P. Palm, and E. Fuchs, EuropeanJ. Biochem. 3,194(1967).

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165. D. D. Anthony, E. Zeszotek, and D. A.Goldthwait, Proc. Null. Acud.Sci. I J .S. 66, 1026(1966). 164. P.Chambon, M. Ramuz, and J.Doly, Biochem. Biophys. Res.Commun. 21, 156 (1965). 156.U. Maitra. Y. Nakata, and J. Hurwitz, J. Biol. Chem.242, 4908(1967). 166.E. J. Freeman and 0. W. Jones, Biochem. Biophys. Res.Commun. 29, 45 (1967). 167.R. H. Epstein, personal communication. 168.A. Sentenac, A. Ruet, and P. Fromageot, EuropeanJ .Biochem. 6, 385 (1968). 159.K. Asano, J .Mol.Biol. 14, 71 (1965). 160.E. Reich and I. H. Goldberg, This series 8, 183(1964). 161.J. M. Kirk,Biochim. Biophys. Acta42, 167(1960). 162.A. Sentenac and P.Fromageot, in preparation. 165.R. I.De Mars,S. E. Luria, H. Fisher and C. Levinthal, Ann.Inst. Pasteur 84,113 (1953). 164. L. S. Lerman, J .Mol.BioZ.3,18 (1961). 166.P. Chambon, M. Ramur, P. Mandel, and J. Doly, Biochim. Biophys. Acta149, 584 (1967). 166.H. Bremer, C .Yegian, and M. Iionrad, J .Mol.B i d .16, 94 (1966). 167.G. Hartmann, K. 0. Honikel, F. Kncisel, and J. Niiesch, Biochim. Biophys. Acta 146, 843(1967). 168.W. Wehrli, J.Nuesch, F. Knusel, and M. Staehelin, Biochim. Biophys. Acta167,215 (1968). 169.J. Nuesch, personal communication. 170.A. Sippel and G. Hartmann, Biochim. Biophys. Acta167, 218 (1968). 171.S. Mizuno, H. Yamazaki, K. Nitta, and H. Umezawa, Biochem. Biophys. Res. Commun.30,379(1968). 179.K. Scherrer and J .Darnell, Biochem. Biophys. Res.Commun. 7 ,456 (1962). 173.D. Gillespie and S. Spiegelman, J .Mol.Biol. 12, 829 (1965). 174.J. R. Beckwith, Science 166, 597(1967).

NOTE ADDEDI N PROOF Recent experiments in several laboratories show that an important protein factor can be separated from RNA polymerase by chromatography on phosphocellulose ( N lN,2 ) . The resulting enzyme, which contains the two subunits previously described by Burgess (69),is apparently a core that is able to transcribe calf thymus DNA but can only transcribe T4 and T5 DNA well when the factor is added back. Since only the “core” retains any RNA polymerase activity, it is likely that the factor separated by phosphocellulose is a subunit required forinitiation of transcription from certain DNA sites. N1. R. R. Burgess, A. A.Travers, J. J. Dunn, and E. K. F. Baiitz, Nature 221,43 (1969). N2. G. Hager and B. D. Hall, personal communication.

Radiation-Induced Alterations in the Structure of Deoxyribonucleic Acid and Their Biological Consequences D. T. KANAZ~R

‘

Faculty of Sciences, University of Belgrade Boris KidrichInstitule jor Nuclear Sciences-Vintcha, Belgrade, Yugoslavia

I. Introduction . . . . . . . . . . . . 11. Evidence Favoring the Idea That DNA Is an Important Macromolecular Target for Lethal Radiation Effects in Living Systems . . . . . . . . . . . . . A. Lethal Changes in Bacteriophage DNA Produced by Radiation B. Radiation-Induced Damage to DNA and Inactivation of Microorganisms . . . . . . . . . . . C. Relationship between Mammalian Cell Death and Damage toDNA . . . . . . . . . . . . 111. A General Survey of the Physical and Chemical Nature of Radiation-Induced Damage to DNA . . . . . . . . . A. The Nature of Physical and Chemical Damage t.o D N A B. Radiation-Induced DNA Breakdown . . . . . . IV. Radiat’ionAction on DNA Replication A. Brief Survey ofthe Biochemistry ofDNA Replication . , B. Radiation Action on DNA Template Function in Vi itro. . . . . C . Radiation Effects on DNA Synthesis in Vivo V. Radiation Effects on DNA Transcription and on the Biosynthesis

.

. . . . . . . .

117

120 121 123 127

132 132 138 142 143 143 149

. . . . . . . . . . . . . 166 A. Radiation Effects on Priming Activity of DNA in the RNA Polymerase System . . . . . . . . . . 167 B. Radiation Effects on Genet.icTranscription i nVivo . . . 171 VI. Biological Consequences Resulting from Radiation-Induced Damage to DNA . . . . . . . . . . . 204 VII. A Working Hypothesis . . . . . . . . . . 207 References . . . . . . . . . . . . . 214 ofRNA

1. Introduction From the beginning of this century, the effects of ionizing radiation on Iiving systems have been studied a t different levels of organization: subcellular structures, whole cells, tissues and organs, and whole organisms, unicellular as well as multicellular. (For a more comprehensive coverage of 117

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these topics see refs. 1-4.) Much information has accumulated, but fundamental problems of radiation biology have not yet been solved. The basic problems, such as the physicochemical nature of the mechanisms(s) leading to radiation-induced death, the site of primary radiation action and the nature of the primary lesion produced by radiation, and the problem why different strains of the same species and different species of organisms respond to radiation in different ways, are still not well understood and Difficult theohave been the subject of considerable speculation (5-10). retical and experimental problems, such as the penetration and random release of energy within the cells, intramolecular and intermolecular transfer of absorbed energy, distribution of energy absorption events, and the effects of oxygen radicals induced by radiation, have been encountered and have been of continuing interest to radiation biologists in the last 50 years. These early studies led to simplified concepts of the existence of , although recent critical and noncritical cellular target elements (5-7) information on the radiation effects and survival of irradiated microorganisms is in clear conflict with the target theory (10-18). Nevertheless, from these early radiation biology results, collected over the years on a wide variety of living systems, there has emerged an important observation on the quantitative relationship between DNA content and radiation sensitivity of viruses, bacteria, mammalian and plant cells. This finding has been of great importance for later progress in the field of radiation biology since it stimulated the study ofradiation action upon the macromolecular constituents of living systems. The results of these studies showed clearly that radiation may produce, in the DNA of living systems, chemical lesions (8-26) that may block cell division (mitosis) and cause the death of living systems without affecting significantly their metabolism, in the sense that ionizing radiation does not disturb markedly the synthesis of other cell constituents in cells that have lost their capacity to proliferate. Energy production and synthesis of the bulk protein are almost unaffected in these cells (86).The most pronounced lesion is inhibition or delay of DNA Consequently, in recent research, attention has synthesis (8-18,15-56,86). been focused on the action of radiation on the structure and metabolism of DNA (10-18, 54-122). Thus, radiation biology, influenced by the new concepts of molecular biology, is rapidly becoming molecular radiobiology, with the prospect of furnishing a better understanding of the fundamental mechanisms of radiation action upon the cellular structure and function. Current experimental evidence suggests strongly that radiation produces, directly and/or indirectly, changes in the primary structure of DNA leading to a loss of secondary and tertiary configuration, i.e. , depolymerization, which may prevent both DNA replication (84-88) and transcription (84,85,88-90). The biological consequence of such damage to DNA may result in mutagenesis and/or death.

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Thus, DNA seems to be an important site of radiation action in all living systems. This appears to be a phenomenon of universal importance and of considerable interest to our understanding of radiation action upon living systems. Recent results, however, show also that DNA carries genes determining the capacity of living systems to repair radiation-induced damage and to repair structural defects occurring within its own structure (20-18,91 ,92). Consequently the molecular and biological consequences of radiation-induced lesions in the structure of DNA depend to a large extent on the genetic constitution of the DNA itself and that of the living system as such. Recent investigations show that in a number of living systems there is a multienzyme repair mechanism by which a variety of molecular and structural alterations induced in DN4,not only by ionizing radiation but also by UV irradiation, chemical mutagens, and carcinogens, are recognized and corrected in such a way that the genome is completely reconstructed. The final outcome of radiation action, therefore, depends greatly on the amount and nature of the DNA damage, on its locationi.e., the probability that the damage will be produced in functional segments of DNA (functional genes)-and on the capacity and effectiveness of the repair of the radiation-induced damage to the DNA. It is becoming clear that the responses of living systems to radiation depend not only on the type of radiation and the number of direct andindirect physicochemical events producing disorganization of the cell and structural and informational defects in the DNA of living systems, but also on the efficiency of the repair of these DNA defects. I t is, thus, very likely that the principal lethal events are caused only by the presence of unrepaired defects in DNA. These may result in (a) an irreversible blockage of normal DNA replication and/or (b) alterations in transcription of the genetic code giving rise to failure of translation, or to faulty synthesis of specific proteins, or to the synthesis of incomplete, nonfunctional protein chains. Accordingly, it appears that radiation damage to DNA, unless repaired, is a basic event that, if it results in inhibition of DNA replication and transcription, may be lethal. The objective of this review is (a) to summarize present knowledge on some aspects of the chemical nature of the main lesions induced by irradiation in DNA and on their interference with DNA replication and transcripand in uieo, (b) to assess, if possible, the bioIogics1consequences tion in vitro of this impairment of the DNA template functions, and (c) to propose a simplified overall picture of radiation action in the frame of a unified mechanism based on the integrated DNA-RNA-protein functions, which seem to be intrinsically entwined in such a way as to influence the genetic makeup of the cell, cell division, and the expression of the cell genome. It should, finally, be emphasized that many contributions to this field unfortunately contain results obtained under such a variety of experimental

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conditions that integration into an overall picture is exceedingly difficult. There is no satisfactory theory of radiation action, and the ideas proposed are numerous and very often conflicting. Nevertheless, it is commonly believed that the main radiation effects are at some stage mediated through DNA. This also may explain why this review pays particular attention to the impairment of DNA template functions and its biological consequences. Since this review is limited only to some aspects of radiation molecular biology, topics such as radiation mutagenesis, radiation chemistry, and repair mechanisms are beyond its scope. These topics have been dealt with extensively in recent reviews by Freese and Freese (94), Witkin (95), and Dubinin (93);by Weiss (W), Wacker (22),and Setlow (96);and by HowardFlanders and Boyce (97), respectively.’

II. EvidenceFavoringthe IdeaThat DNA Isa n Important

Macromolecular Targetforlethal Radiation Effects in living Systems When a living system is exposed to ionizing radiation, the absorbed energy may cause a complex interplay of biochemical processes that stem from one primary lesion, occurring probably in the cell genome, and that, ultimately, may cause its death. Although the precise nature and location of a primary lesion are not yet well established, at present it is commonly believed that the DNA of living systems is the principal target for radiation action. There are several lines of evidence that DNA is an important, and perhaps the principal, target for inactivation ofmost living systems by UV and ionizing radiation, but there is no indication that DNA is the only target. However, there is good evidence that radiation-induced damage to 48DNA may be lethal to most unicellular living systems (10-18, 25-33, 62,54-74, 86) and to mammalian cells in vitro(‘74-81, 105, 1.21, 122). In addition, the experimental evidence leaves little room for doubt that, even in higher organisms, the impairment of DNA structure and function may be responsible for death after whole-body exposure to ionizing radiation (S4-47, 81-8s). However, it should be emphasized that the metabolic stability of living cells or higher organisms after irradiation depends upon a balance between biochemical damage and repair processes. Furthermore, at many levels of organization, “homeostatic” mechanisms produce responses to environmental stresses that tend to preserve the integrity of the living systems. Although the primary mechanism responsible for the radiation-induced death of multicellular organisms remains unknown, it still may be assumed with fair certainty that direct or indirect radiation 1

See also article by Setlow in Vol.8 ofthis series.

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effects on DNA structure and metabolism represent the primary lesion in a series of molecular events leading to the death of higher organisms. The starting point of this discussion is to enumerate the l i e s of evidence indicating that damage induced in the DNA structure by radiation, unless repaired, may lead to cell death.

A. Lethal Changes in Bacteriophage DNA Produced by Radiation The bacteriophages seem appropriate for the study of the physicochemical nature ofprimary DNA structural changes produced by ionizing radiation and their relationship to phage inactivation. Highly purified phage samples may be irradiated and phage survival-i.e., the capacity of a bacteriophage to reproduce itself in a sensitive host bacterium-can be followed. In addition, phages have the advantages that their DNA is easily extracted and that radiation damage is repaired only slightly, if at all (51). This allows the detection of primary chemical lesions induced by irradiation and the assessment of their relative biological significance. One possible complication in any study with bacteriophage infection is that inactivation after irradiation may come about by loss ofthe ability ofthe phage to adsorb to the appropriate host cell (bacterium). However, evidence from phage adsorption after X-irradiation indicates that this is >. not the case (51 Viral radiobiology indicates strongly that (a) phages suspended in buffer are inactivated by X-irradiation owing to the production ofdoublestrand breaks in the phage DNA; (b) single-strand breaks are not lethal; (c) strand breakage cannot explain all the lethal effects ofionizing radiation and phages are probably inactivated by both double-strand breakage and by DNA base damage; and (d) the most important base damage may be to thymine. In addition, the substitution of thymine in phage DNA by 5-bromouracil renders it more susceptible to X-irradiation and enhances phage inactivation (48-52, 98-100, 10.2). Since this substitution does not affect the rate of X-ray-induced single-strand breakage, the enhanced phage inactivation is probably due toDNA base damage. This means that sensitization of phages by bromouracil is probably the result of an increase in the rate of DNA base disintegration. These results strongly suggest that the loss of the ability ofa bacteriophage to reproduce itself in a sensitive host cell can be directly related to the structural alterations (defects) produced in the DNA by ionizing radiation. Since phage is probably the most unique system from which DNA undamaged by the isolation procedure can be obtained, and since it lacks a repair mechanism, the results mentioned above support directly the view that the DNA is the principal target of lethal radiation effects in viruses (Fig. 1, A, B, and C).

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Time. minutes

FIG.1. (A) Percent survival of plaque-forming ability of phages T4,T5,T7 and asfunction oftime of X-irradiation. The phages were suspended in buffered histidine and irradiated. Survival was determined by plating on Difco nutrient agar (T5and T7), X agar (1% tryptophan, 0.5% NaCl,1.5% agar) forphage A, or T4 agar [M.Chase and A. H. Doermann, Genetics, 43,332 (1958)]. P l a h were scored after 16 hours a t 37 C except for T7 (room temperature). Figure 1shows that the sensitivities ofT7,A and T5 increase roughly with the molecular weights of their DNA, except for phage T4 (see legend forFig. 1 B).X-ray resistance ofphage T4 is probably due to some base damage, which is in some way repaired ID. Freifelder, personal communication (1968)l.

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0

2

4

8 10 Time, minutes

6

12

14

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FIG.1. (B)Percent survival of intact native DNA molecules isolated from phages T4, T5,T7,and X irradiated with X-rays. Phages were X-irradiated for the indicated times, followed by neutralization. The DNA was released by incubating the phage at pH 13.3 amount of unbroken DNA was determined by analytical ultracentrifugation. The sensitivity of each DNA is proporlional to the DNA molecular weight (T4DNA mol. wt. = 120million, T5 DNA mol. wt. = 76million; X DNA = 33million; T7 DNA = 25 million) [D. Freifelder, personal communication (1968)].

123

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Dose

-

.8MR

FIG.1. (C)Surviving fraction of infectious DNA isolated from phage 4x174. Phage DNA was prepared and assayed in an E. coliK12 spheroplast system according to Cuthrie and Sinsheimer, Biochim. Biophys. A d a 72,290 (1963). BrU-+X174phages were purified by CsCl density-centrifugation. The irradiations, with 6OCobalt gammr-source, were done in air on DNA suspended in 4% Difco-nutrient broth. The survival of normal (O), and bromouracil-substituted (A), DNA in the presence ( O ) ,and absence (A)of M/10 cystearnine hydrochloride atpH 7.5is shown. From Hot2 (102a).

B. Radiation-Induced, Damage to DNA and Inactivation of Microorganisms

I n recent years, considerable effort in radiation biology has centered on the lesions induced in DNA and their relation to cell death and mutation. These results suggest strongly that reproductive death of irradiated bacteria, that is, inability of the cell to divide and to produce an unlimited number of progeny cells, seems to result from radiation-induced structural defects in DNA that, unless repaired, inhibit cell multiplication by blocking the normal template activity of the DNA. It is likely that the principal lethal event caused by the presence of unrepaired damage in DNA is an irreversible blockage of normal DNA replication and its transcription. Thus, it appears that the radiation-induced damage may act as either a permanent or a, temporary block to normal DNA replication, depending on the amount and extent of structural lesions in the DNA and on the functional capacity and efficiency of the repair mechanism(s). Consequently, the completion of repair and resumption of normal DNA synthesis lead to the formation of a visible colony or cell clone. There is evidence for the existence of repair mechanisms in bacteria other than Escherichiu coli (11-13, 16-17,91, 92)and Bacillus subtitis (103). Micrococcus radiodurans exhibits extreme resistance to ionizing radiation because of a highly efficient

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DNA repair mechanism (14,18, lo,&106). A repair mechanism was recently found also in Salmonella typhimurium (106)and in the unicellular green alga Chlamydomonas reinhardi (207). In support of the view that DNA is an important site for the inactivation of microorganisms, the following evidence may be cited: 1.The best evidence for the biological consequences of damage induced in DNA structure by radiation comes from studies on the sensitizing effect of incorporated halogenated DNA base analogs into bacteria (10, 108-1l a ) . There is now abundant evidence that the halogenated pyrimidines or their deoxyribonucleosides(5-bromocleoxyuridine, 5-iododeoxyuridine) can under appropriate conditions become extensively incorporated into DNA in place of thymine with a concomitant increase in the sensitivity of bacterial 66a,119). cells (Fig. 2) to both ionizing (110, 111) and UV radiation (10,

Dose in Krads

Dose in Krods

FIG.2.(Left) Dose-survival curves forcontrol and bromodeoxyuridine-sensitized E. coli K 12, substrain JE-850. (Right) A semilog plot ofrelative sedimentation distance of DNA from control ( 0 ) and sensitized ( 0 )cells in pH 7 sucrose gradients (indicating double-strand scission of DNA) us. X-ray dose. A threefold increase in slope for DNA from the sensitized cells us. that fromcontrol is observed. From Kaplan (17).

That this radiosensitivity is specifically attributable to the incorporated analogs into the DNA is supported by the facts that (a) the increase of radiosensitivity is proportional t o the extent of analog incorporation, and it is competitively inhibited by thymine or thymidine; and (b) there is a quantitative resemblance in the degree ofanalog radiosensitization between cells and transforming DNA extracted from the same intact B. subtilis culture (10, 108). Recent results indicate that radiosensitization by DNA-incorporated halogenated pyrimidine analogs is based on two properties of these compounds: they are somewhat more sensitive to radiochemical events induced by radiation than are natural pyrimidines, and the incorporated analogs

RADIATION-INDUCED ALTERATION I N DNA

125

make DNA more susceptible to radiation-induced cross-linkage (10) arid to X-ray-induced double-strand breaks (17). This incorporation gives also rise to radiochemical products that affect certain steps of the repair processes (118, 113). In addition to pyrimidine analogs, the purine analog 2-amino-6mercaptopurine is also one of the most effective sensitizing agents (210). Several purine analogs have now been shown to sensitize one or more purine-requiring auxotrophs to lethal effects of ionizing radiation (110, 111). The sensitized state is completely reversible by purine supplementation (110, I l l ) . Furthermore, according to Kaplan (Fig. 3), a striking degree of I^^

FIG.3. Percentage of survival versus X-ray dose for E . coli JE-850 cultures subjected to thymidine starvet.ion under various conditions preventing thymineless death. From Kaplan (118). Control, prior to starvation 0 Thymidine starved plus asauridine, 2 hours.

a Thymidine refed, minus uridine,

14 hours.

(> Thymidine refed, plus uridine, 3 hours.

0 Thymidine refed, minus uridine, 3 hours.

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radiosensitizationcan be induced in thymine auxotroph strains of E.coli by thymine starvation (ria). The radiosensitized state is reversible by thymidine supplementation. The results presented on the development of increased radiosensitivity during thymine starvation and its selective reversal by thymidine suggest strongly that the sensitivity to ionizing radiation is dependent on the macromolecular structure ofDNA. Support for this view is provided by the altered physical properties of the DNA from thymine-starved bacteria (114). Such structural damage to DNA would presumably result in an increase of the relative yield of nonrepairable X-ray-induced defects in the DNA, which can even be mutagenic (115). 2.Bacterial sensitivity to UV and ionizing radiation and to nitrogen mustards can be correlated with the DNA base composition. Thus, Kaplan and Zavarine (63) have reported that sensitivity of eight different species of bacteria to X-rays increases linearly with the guanine plus cytosine (G C) content of their DNA. 3.Information concerning the action of radiation upon the transforming ability of DNA suggests that the loss of biological activity of transpneumoniae and B. subtilis forming DNA isolated from Diplococcus depends on the size of the genetic marker and the base sequences and composition of the marker loci, and upon the molecular weight of the DNA being irradiated (110, 117). Kaplan et al.(110) have reported, however, that the radiosensitivity of different markers in transforming DNA from B.subtilis may be correlated with differences in their average G C content (Fig. 4). 4. It is suggested with reasonable certainty that there is probably no real distinction between “lethal” and “mutagenic” DNA defects produced in the DNA structure by UV radiation. According to Witkm (95, 118), a bacterium will survive irradiation if (a) the residual DNA damage (after repair) does not exceed a certain strain-specific limit still compatible with DNA replication, and (b) at least one DNA strand is free of induced lethal mutations, which probably originate fromDNA replication errors due to unrepaired UV damage present in the replicating DNA, although errors in repair cannot be completely excluded. The excision of these DNA defects induces the decline in mutation frequency (218). From his studies on mutation induction by ionizing radiation in Paramecium, Kimball has observed that the closer t o the time of DNA synthesis X-rays are applied, the more mutations are produced (119). This supports again the view that errors in replication of DNA rather than errors in repair are involved in radiation-induced mutagenesis. However, Eckstein etal.(86) have demonstrated that X-irradiation (50krad) can prevent the proliferation and destroy the colony-forming ability of yeast cells. The energy producing processes as well as the syn-

+

+

RADIATION-INDUCED

127

IN DNA

ALTERATION

thesis of the bulk protein in these cells were practically unafYected, whereas DNA replication was inhibited. These results suggest very strongly that DNA may be considered as a primary macromolecular target for radia tion-induced lethal and mutagenic damage in microorganisms. It appears that the structural integrity of DNA is essential for the survival of microorganisms after irradiation.

1.0

I 0

25

80 100 '120 X-RAY DOSE (krods)

50

150

FIG.4. Relative X-ray sensitivities of three different markers in B. subtilis-trans; formingDNA, assayed on mutant strains auxotrophicfor histidine (SB-400), 0-0 tyrosine (SB-65), AA ;and indole (SB-l68), 00. D-10 values: histidine, 105,000 rads; tyrosine, 143,000rads; and indole, 195,000 rads. From Kaplan etd.(110).

C. Relationship between Mammalian CellDeath a n d Damage to DNA Many studies have sought to determine the effects of ionizing radiation and in vieto and to on the DNA metabolism of mammalian celIs in vitro show whether radiation-induced lesions in DNA structure and metabolism can somehow be correlated with cell death, i.e., with the loss of capacity to form clones of progeny cells. Recent investigations have indeed shown a close relationship between damage to cell DNA and cell death. Thus, DjordjeviE and Sxybalski (7'4) demonstrated that mammalian cells are sensitized to the lethal action of X-rays if grown in media containing 5-bromodeoxyiiridine under condit,ionsin which R part of DNA thymine is

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D. T. KANAZIR

replaced. This fact is frequently invoked as an argument in favor of the hypothesis that the cell DNA is a primary target for biological action of ionizing radiation. It has also been established that mammalian cells are more sensitive to radiation during the DNA replication period, i.e., the S period of mitosis ( 7 5 4 0 122). , If synchronous cell populations

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.-t

Hours

FIG.5. (A)Idealized curves representing: I. Color (acridine orange) intensity of the cytoplasm in L-strain at different developmental phases The intensity of staining was measured with microdensimetry. 11. The rate of RNA synthesis in L-strain cells during different developmental phases followed by autoradiography. [*HI Adenosine and [aH]cytidine were used as precursors. The developmental phases are designated by symbols describing approximately the morphological appearance of the cells: M , cells in the phase of mitosis; N , new cells originating from M cells, these are transformed into symmetric . are chronologically cells (8)which then develop into asymmetric cells ( A s )These followed by La,wide cells (which are divided into La,and Lazcells by their differences in staining and in rates of RNA and DNA synthesis). In the last step of development, ovoid cells (0)become transformed into M cells. The morphology and synthetic capacity of the offsprkg of a given cell was followed by photography, staining, and autoradiography. From Neskovid. (207). (B)The rate of incorporation of tritiated thymidine into synchronous populations of HeLa cells at various times after mitosis. In all cases, the cells were irradiated with control; 300 rads; - A , various doses immediately after mitosis. -O-J 600rads; -O-, 900rads. From Brent el al.(79). (C) The rate ofincorporation of tritiated thymidine in synchronous populations of HeLa cells at various times after mitosis. -o-] control experiment; -0-, culture irradiated with 1000 rads 11hours after mitosis. From Brent et al.(79). (D)The rate of incorporation of thymidine in synchronous populations a t various times after mitosis. Irradiations with 1000 rads were carried out at the following times (arrows); -A-, mitosis; -0-, 7 hours after mitosis; 11 hours after mitosis. Control culture, -0-. I h m Brent, etnl.(70).

-n-,

Time after mitosis (hr) C

Time after mitosis (hr)

30

a 520

.-c x

:10 0

Time after miitosis (hrl

Fro.5 B, C, and D.

130

D. T. KANAEIR

(Fig. 5 C, D) are irradiated in the S phase, DNA replication is largely inhibited. This prevents cell division and ultimately leads to cell death. If the cells are at mitosis, in the hT stage or in the GI period, when irradiated, DNA replication in the same cell cycle apparently proceeds normally, but cell division is delayed and depressed and only a limited amount of From this it DNA synthesis occurs in the second cycle (Fig. 5D) (79). has been concluded that the cell is more sensitive to radiation action during the replication stage of DNA than during other mitotic stages (75-80, 121). “Suicide” experiments with mammalian cells show a more direct relationship between radiation-induced damage to DNA structure and cell death. The experiments were performed with the use of a variety of tritium-labeled precursors including thymidine or uridine, and histidine or lysine. The results showed that, of all the tritium-produced disintegrations, the most effective, i.e., the most lethal, were those that occurred in the DNA, whereas with tritiated amino acids the killing efficiency was very low (122). It is also well established that radiation is mutagenic and capable of There is a good deal of producing chromosomal aberrations (1.23-125). evidence supporting the view that chromosomes represent the principal target for the radiation-induced loss of reproductive capacity of mammalian cells. It appears also, at least in plant cells, that the variation in radiosensitivity may be correlated with the chromosome number and nuclear characteristics (nuclear size and structure, chromosome complement and DNA content). Although the results appearing in the literature for mammalian cells are conflicting, some reports indicate an increase in the radioresistance of cells with increased ploidy (125). Unfortunately, we still do not know the precise molecular nature of the damage produced by ionizing radiation to chromosomes of the mammalian cell. The reasons for this are twofold: first, ionizing radiation is not selectively absorbed; and, second, the molecular structure and organization of the chromosome is still poorly understood. Nevertheless, it would seem reasonable to accept that, in mammalian cells, the primary damage producing chromosome aberrations may be single scissions in one of the two DNA strands. Mortality in higher organisms presumably results from cell death and, more specifically, from cell death in the intestine and in the hematopoietic system. It appears that the sensitivity of higher organisms to radiation, as estimated by survival after singIe or multiple radiation exposure, is not determined by variations in the extent of the damage from a standard dose of radiation, but by the ability to survive the damage once it is produced. This ability is determined by the genotype. It is therefore clear that the cell genome is the likeliest site of lethal injury, and it has been proposed

RADIATION-INDUCED

ALTERATION

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131

that cell killing results principally from damage to the chromosomal DNA. Much information is consistent with this concept, although there are some difficultiesin applying it generally. Furthermore, the possibility exists that the net chromosomal injury in somatic cells may vary between strains because of differences in the efficiency or effectiveness of repair processes. This is consistent with the fact that inbred mouse strains differ widely in Although the mechanism of the cell death of radiation sensitivity (125). higher organisms is not yet well established, the mechanism proposed above remains, for the present, an attractive working hypothesis. Although no other macromoleculax target has so far been explicitly identified, some of the lethal action of radiation may be attributed to other, %on-DNA” targets. They probably play an important role in the so-called late effects of radiation. It has become increasingly obvious that the structural integrity of the cell is essential for normal integration of its metabolic activity. I t has been suggested ( l ) however, , that cell damage resulting from exposure to radiation may be due to a disruption of cellular organization. This may give rise to physiologically and biochemically indirect effects that can obscure the primary effects of radiation. Thus, some evidence for the formation of peroxides in the tissues of irradiated The precise sites of peroxide formaanimals has been obtained (120-128). tion in irradiated cells are not known. The study of peroxide formation after X-radiation (10 krads) in the subcellular components of liver shows clearly that little peroxide formation occurs in any fraction immediately after irradiation and that microsomes and lysosomes form peroxides readily, whereas nuclei and mitochondria form relatively small amounts of peroxides. Furthermore, it is difficult to assess the importance of peroxide formation as a consequence of irradiation in vivo because relatively little is known about lipid peroxide formation in normal tissues. But possibly extensive spatial disorganization of membranes and of bound enzymes within cells, or extensive leakage of enzymes and other important cell constituents (precursors) may occur. Radiation damage to membranes is strictly time dependent, and it appears that radiation causes in the membrane some change that subsequently leads to its rupture. Since the late effects of radiation are beyond our scope, such damage is not discussed here in detail. In summaryl the information concerning the radiation-induced death of cells of higher organisms suggests that death in higher organisms, as well as in bacteria, may be determined by two distinct types of primary radiochemical lesions, namely, damage to some critical structure, possibly DNA or DNA-protein, :~nd interference with some of the metabolic systems responsible for the repair of that critical structure.

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D. T. KANAZIR

111. A GeneralSurveyof thePhysical and ChemicalNature of Radiation-Induced Damage toDNA

A. T h e N a t u r e of Physical a n d Chemical D a m a g e to DNA Although radiochemical lesions in DNA appear to be responsible for the loss of viability of irradiated cells, the precise chemical nature of such lesions has not yet been established. We do not discuss here the chain of intermediary events leading to primary and final chemical damage to DNA. The purpose of this section is only to summarize the major lesions produced by radiation in the DNA molecule in order to assess their possible biological importance. Almost all studies in radiation biology have been carried out either with particulate radiation or with electromagnetic radiations of wavelengths less than those of the visible spectrum. These can be divided into two classes, ionizing and nonionizing radiations, according to how they release most of their energy. X- and y-rays exert their effects by producing ionizations within the cells, whereas UV-radiation produces mainly molecular excitations that absorb the radiation energy. Highly penetrating ioniziig radiac tions lose their energy along the tracks of individual ionizing particles as they slow down within cytoplasm. UV radiation, however, can release energy only where it is absorbed, and since various chemical compounds absorb UV at various wavelengths differentially, this type of radiation promises to be far more selective than ionizing radiation for gaining an understanding of the molecular events that result in molecular damage. Since ionizing radiation is nonselectively absorbed, we still do not have a clear picture of the chemical basis of its effects because they are relatively manifold and unspecific. Although both UV- and ionizing radiation act in fundamentally different ways, the point of attack in the cell is the same, i.e., nucleic acids. At present, the overall chemical changes induced by ionizing radiation in the DNA and in its secondary and tertiary structure are not known in exact detail. Another obvious and serious problem is that chemical damage to DNA may be repaired in living systems. This could obscure any relationship between induced primary chemical lesions and the structural status ofthe DNA a t the time after irradiation when DNA must carry out a critical function. The experimental difficulties experienced in attempting to detect biologically relevant damage are also not insignificant because of the problem of measuring the small yields of radioproducts. Therefore, in order to raise the concentration of damaged DNA constituents to measurable levels, it has been common to resort to very high doses, i.e., well above the

RADIATION-INDUCED

ALTERATION

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“biological” doses. With these difficulties in milid, I wish now to discuss the main physical and chemical defects induced in DNA by radiation. 1. RADIATION-INDUCED PHYSICAL CHANGES IN DNA

One approach to the problem is to detect chemical changes in the DNA by their physical consequences. Since DNA is a long polymer, any chain break could be detected as a large change in molecular weight and in any supramolecular organization of DNA that may be followed by physical methods. However, problems arise when one attempts to isolate DNA from irradiated cells as the isolation of DNA from most living systems is usually accompanied by substantial degradation by hydrodynamic shear forces. The resulting DNA sample often contains many more breaks than may have been produced by the radiation. In order to overcome these expenBut mental difficulties, DNA has been isolated and then irradiated invitro. here again the heterogeneity in molecular weight resulting from the action of shear forces so much reduces the sensitivity of the usual physical methods (viscosimetry, ultracentrifugation) used to determine molecular weight changes that very high doses, “nonbiological doses,” are again required. No solution to these difficulties is apparent, at least for more complex living systems, although they might be avoided in bacteriophage systems as reported by Freifelder (129).I n any case, the available data on the physical properties of DNA can be summarized as follows. (a) The results of invitro experiments show a significant decrease of viscosity of DNA after ionizing A similar radiation due to a decrease in its molecular weight (130-136). depolymerization is observed when DNA is irradiated in the dry state (137). A decrease of DNA viscosity is also observed in invivosystems, such as bacteria (32,138) and cells of different tissues of higher organisms (139-144). (b) By ultracentrifugal analysis of irradiated DNA samples, it was demonstrated that ionizing radiation produces single- and doublestrand breaks in viral DNA (51, 52, 129,145)as well as in bacterial DNA (17, 146). Radiation also causes both partial and complete denaturation of the DNA macromolecule (130-136, 147-1511. These changes suggest strongly that DNA is depolymerized by irradiation both invitroand invivo.This conclusion was confirmed by light149), which, in addition to demonstrating a scattering experiments (147, decrease in molecular weight, indicate that DNA probably accumulates single-strand breaks before double-strand breakage occurs. However, the interpretation of the data obtained by viscosimetry, ultracentrifugation, and light scattering may be complicated for several reasons. A decrease in viscosity could result from depolymerization caused by partial denaturation or single-strand breakage. Ultracentrifugal

134

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analysis may have, in principle, a capacity to detect degradation of a small fraction of a homogeneous material, but since DNA as usually prepared is always heterogeneous with respect to molecular weight, such analysis yields onIy average changes in molecular weight. I n order to avoid these complications, a new approach was made using sucrose density gradient centrifugation a t neutral and alkaline pH. By this technique, it is possible to detect primary single- and double-strand breaks in DNA (16, 1'7). It appears that double-strand breakage of DNA is a major structural change produced by ionizing radiation and so may contribute to lethality (16, 17,1.29).

2.CHEMICALCHANGES INDUCEDIN THE STRUCTUREOF DNA BY RADIATION Radiation can destroy DNA directly and indirectly (Fig. 6).Direct damage is the splitting of the bonds between the bases, and the sugar and X

- rays

DNA

H

FIG. 6. Diagrammatic representat,ion of the three types of X-ray damage to DNA molecules suspended in an aqueous environment. (1)Direct damage, probably a relatively rare event, which results in double-strand or single-strand breaks. (2)Indirect damage mediated by RI radicals, whichcause double-strand or single-strand breaks and which are quenched by radical scavengers-for example, histidine orSH-containingcompounds. (3)Indirect damage mediated by another class of RZradicals, the formation or action of which depends on the presence of oxygen and leads to chemical damage to the pyrimidine bases. This damage is depressed by the absence of oxygen, or by the addition of SHcontaining compounds, but not by radical scavengers of the histidine class. (From Szybalski, see Ref.129.)

RADIATION-INDUCED

ALTEltATION IN DNA

135

phosphate, by high-energy radiativii. The indirect one occui-s by the attack of peroxides or radicals formed from water or organic compounds by ionizing radiation, which may act on nucleic acids and their precursors within the cell. The overall chemical changes induced by irradiation in DNA bases and in DNA secondary and tertiary structure are not known in detail a t present. The chemical nature of the products formed by radiation have been investigated in three ways: (a) irradiation of free bases, nucleosides, and nucleotides in water, (b) irradiation of DNA in water, and (c) irradiation of DNA in dilute salt solutions (see refs. 3,21,23). From such studies, much information has been accumulated, but it is difficult to evaluate its biological significance. In any case, three main types of changes can be distinguished in irradiated DNA, namely, base destruction, sugar-phosphate bond cleavage, and chain breakage, all leading to the changes in secondary structure (radiation-induced hyperchromicity) . The precise effects these radiation products may have on the template function of DNA, i.e., on base pairing, is still unknown, so it is difficult to evaluate their biological significance. It is clear, therefore, that none of the results from in vitro systems can readily be applied to the situation prevailing in more complex living systems. For that reason, in some experiments DNA was irradiated in vivoand then extracted for chemical analysis. Since thymine destruction waa to be measured, high doses were used. Such experiments with bacteria or mammalian cells are complicated because of the difficulty encountered in extracting DNA from irradiated cells. This problem may be solved by using bacteriophage systems from which, by avoiding hydrodynamic shear, native DNA molecules may be obtained.

3.RADIATION-INDUCED DAMAGE TO PHAGE DNA Since bacteriophage seems to be the most appropriate system for the study of primary physicochemical damage to DNA induced by radiation, and since the doses used are in the biological range, these experimental results assume an important biological significance, in that the damage to DNA can be directly correlated with phage inactivation except for phage T4 (Fig. 1, A, B, and C). The X-ray resistance of phage T4 probably involves base damage, which is repaired in some way although, if so, the repair system is not the one that repairs UV damage (D.Freifelder, personal communication). When bacteriophages are X-irradiated, plaque-forming ability (viability) is lost (48-52, 129, 145). The specific chemical or physical changes responsible for this inactivation havc recently been positively identified. The number of X-ray-induced single- and double-strand breaks per

136

D. T. KANAZIR

coliphage has been measured at high (20-100%) survival leveIs by ultracentrifugal analysis of the DNA (Fig. 7,A and B). After irradiation in buffer, at least 90% of the nonviable phage particles were found to have been inactivated by double-strand DNA breaks. The induction ofsingle52,129). The principal base damage may strand breaks is not lethal (51, have been to thymine (129). DNA breaks were also observed when coliphage T7 was irradiated with argon nuclei (151). On the basis of these

67

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FIG.7. (A)Optical trace of the sedimentation velocity boundary for T7 DNA extracted from phage irradiated to various survival levels, as indicated. The arrow refers to the direction of sedimentation. M is the meniscus. D N A concentration, 20pg/ml, in 1 M NaCI. Speed, 33,450 rpm. Time of sedimentation, 30 minutes. The verlical bomdary (A) is that ofunbroken molecwles. The 1railing portion (B) represents trhebroke11 molecules. From Freifelder (289). (B)Percentage of unbroken DNA molecules (phage B3) following X-irradiation of DNA in lop3 hl IAiididine InifTered at pH 7.8with lOP A'l Na phosph:tte. Vrom Freifelder (189).

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FIG.8 Sedimentation patterns of D N A from control and irradiated bacteria ( ~ ~ c ~ c& e r KU, ~ c substrain ~ ~ ~ a JE-850) in sucrose gradients (pH 13 and p H 7,indicating single- and double-strand DNA scissions, respectively). Sedimentation distance, D, from meniscus of gradients (fraction 40)is expressed as a percentage of tho value of the DNA of unirradiated cells. From Ilaplan ( 1 7 ) . = 21.4 mm

1715 A

rn

40 36 32 28 24 20 16 12

FIG. 9. Hadioactivity of titrated material in E. coliB/r, lysed and centrifuged on alkaline sucrose gradients, as a function of distance sedimented in 90 min: (A) unirradiated; (B)20 kR,no incubation; (C)20 kR,20 min incubation; (D) 20 kR, 40 min incubation. Approximately 5,000 cpm were placed on each gradient. Average distance from the meniscus average sedimentation constant and average molecular weight are given for each plot. Each curve was fitted by eye to the combined data from three experiments (each indicated I)y 0, 17,A).Double arrows indicate that portion of the curve used to determine s 2 0 , ~and Froin McGrath and Williams (16).

(a). (m)

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KANAZIR

data, Freifelder (129, 1 2 9 ~ correlated ) the inactivation of X-irradiated phages with the yield of double-strand scissions and base damage in DNA (Fig. 1, A, B, and C). The same kind of damage seems also to be lethal in bacteria. Kaplan (17) reported that irradiation of E.coli induces a decrease in the sedimentation rate of alkali-denatured native DNA attributable to single- and double-strand scission (Fig. 8).To rule out the interference of preparative analyzed procedures known to break DNA, McGrath and Williams (18) DNA in alkaline sucrose gradients and obtained pieces of single-stranded DNA by direct hydrolysis of cells placed on the top of the gradients. From their results, single-strand scissions induced by X-rays are reparable in radiation-resistant strains of E.coli(Fig. I)) but not in the sensitive BS-1 drain, whereas neither double-strimd scissions nor viability exhibit repair during reincubation. Bromodeoxyuridine increased the yield of doublestrand scissions per unit dose to an extent proportional to its effect on radiation lethality (Fig. 10, A and B). These correlations suggest that even in X-irradiated, radioresistent bacteria, double-strand scissions are the major radiochemical lesions leading to loss of viability. However, calculations indicate that single-strand scissions could account quantitatively for lethality in the highly radiosensitive strain BS-1 although double-strand scissions, produced in lesser yield, would also be expected to contribute to some extent. Therefore, single-strand scissions are probably important in radiosensitive mutants lacking a repair system, and perhaps also in cells grown under conditions in which repair is inhibited.

B. Radiation-Induced DNA Breakdown One of the first and probably most important indications that ionizing radiation drastically affects the structural integrity of the DNA molecule in bacteria is the degradation of DNA following radiation. Stuy (57,58) observed vigorous degradation of DNA in Hemophilusinjluenzae, and Miletii: etal.( 5 9 4 1demonstrated ) radiation-induced DNA degradation in a number of strains of E . coli. Similar degradation of DNA in E . coli 15T-L- was induced by fast proton bombardment (152). It has also been observed by Kuzin in mammalian cells (153). Furthermore, irradiated bacteria incubated under starvation conditions or in the presence of chloramphenicol (59-61) exhibit an increase in both the rate and extent of DNA loss. Degradation is enzymatically mediated after an initial primary effect. Although the question as to the nature of the enzyme is still open (156), it may have the characteristics of an exonuclease (70). TrgovEeviE and KuEan (70) showed that y-irradiated DNA (E.coliB) was broken down a t a higher rate than nonirradiated DNA by unidentified enzyme(s) present in crude extracts of E.coli.

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Furthermore, Pollard and Achcy (154), studying the kinetics of DNA degradation in y-irradiated E.coli15 T-L-, demonstrated that the amount of degradation increases Iinearly with the dose. They observed tl potentiating effect of oxygen and an increased degradation in cells grown on 5bromouracil. While DNA degradation is proceeding, DNA synthesis continues if the degradation does not exceed approximately 40%, but it is slower. This suggests that incomplete DNA components are unable to serve as “templates” for continued DNA synthesis (154). In a comparative strains, BS-1 and B/r, McGrath etal.(59) observed study of two E.coli that the initial rate of degradative loss of DNA from X-irradiated cells is the same for the two strains, but the BS-1 strain degrades its DNA much more extensively than the resistant B/r. Chapman and Pollard (165) showed that DNA throughout the degradative process is double-stranded DNA, similar to the unirradiated control DNA with respect tomelting temperature, annealing, and buoyant density as measured by CsCl densitygradient centrifugation. The results of Grady and Pollard (156) also showed a considerable is irradiated in the increase in the amount of DNA degraded if B.subtilis presenceof actinomycin D (Fig. 11,Aand B). No degradation was observed in the unirradiated controls, regardless of whether actinomycin D was present or absent. The concentration of actinomycin D used was in the range where RNA synthesis is drasticaIIy reduced while DNA synthesis is not greatly affected. The effect of actinomycin D on DNA degradation suggests that some relationship between it and the synthesis of RNA should be sought. The effect of actinornycin D might be due to inhibition either of existing repair systems or of the synthesis of any repair enzymes that would otherwise be induced by radiation action. Whatever the explanation of these results may be,they suggest that the action of actinomycin D on radiation effects in mammalian cells might well be in the potentiation of radiation-induced DNA degradation. I n connection with this degradation of DNA, several questions may be raised: Does the total degradation of DNA observed after irradiation involve those cells destined t o die or the large populations of still-viable cells undergoing repair? IS degradation the actual process that alters the vital functions of DNA-that it, does i t act to kill the cells? What are the properties of the DNA newly synthesized after the degradation? The first question may be answered by observations suggesting that the observed DNA degradation after low doses of radiation is a population effect in which only the DNA of those cells suffering a radiation lesion is degraded. This was also confirmed by the autoradiographic studies of Shaffer and McGrath on irradiated E. coli(15 ). On the question as to whether DNA degradation is a cytopathological

5 Fraction number

10

20 25 30 35 40 Fraction number

15

FIG. 10.(A)Sedimentation patterns of DNA from control and irradiated bacteria (Eschem’chia coli K12,JE-850) in sucrose gradients pH 7. There is a progressive decrease of sedimentation rate of undenatured DNA from irradiated cells with increasing X-ray dose. The sedimentative distance, D, from meniscus of gradients (from fraction 40) is expressed as a percentage of the value of DNA from unirradiated cells. From Kaplan (17’). (B) Sedimentation patterns (sucrose gradients, pH 7) of DNA from unirradiated and irradiated bacteria grown on bromodeoxyuridine (sensitized cells). There is again a progressive decrease of sedimentation rate with increasing dose, but of greater magnitude for any given dose compared with sedimentation rate of DNA from irradiated nomemitized cells (see Fig. 10A).From Kaplan (17).

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FIG. effect of actinomycin D. (A) Degradation of DNA a t two different doses (25and 65 krads). To cultures of Bacillus subtilis 168T- ind-, actinomycin D was added to a final concentration of 1.0rg/ml. After 10 minutes at 37 C with aeration, the samples were oxygenated for 1.0 minute and exposed to the desired dose in a 6oCosource. One-milliliter samples were removed and added to 1.0 ml of cold 10% trichloroacetic acid at intervals up to 120minutes. After 1 hour in a n ice bath, samples were filtered through Schleicher and Schuell Type B-6 filters, rinsed with 10 ml of cold 5% acid, glued t o planchets, dried, and counted. The loss of radioactivity from the acid-precipitable fraction a s a function of time after irradiation was taken as a measure of DNA degradation. Curves have been normalized by setting the acid-precipitable radioactivity in the sample removed before irradiation equal to 100%. (I3) Dose-response of DNA degradation in 8, subtilis 186T- ind-. From Grady and Pollard (166).

142

D. T. KANAZIR

event leading to lethality, one could give a negative answer since most irradiated bacteria survive this degradation, especially if their DNA repair mechanisms are sufficient. Therefore, one can conclude that the most dramatic consequence of Xor y-irradiation of bacteria is a degradation of DNA initiated by some primary damage to the structure of DNA. The differences in the level of degradation among related bacterial strains suggests the existence of genemediated controls. It may be of interest to mention here that DNA degradation is not observed in Salmonella typhimurium even after very high doses (20krad) of radiation (158). This again supports the view that the degradation is probably gene-controlled. The mechanisms responsible for these DNA degradation processes remain to be determined. It should be noted also, as the answer to the third question raised above, that postirradiation DNA synthesis gives rise to DNA molecules that are more difficult to degrade by radiation than is normal DNA (154). This finding suggests very strongly that the structure and integrity of DNA molecules synthesized de novo in irradiated bacteria are altered. The observations of Billen et al.(159, 160) indicate that the new DNA synthesis takes place in a manner different from the normal sequential process, resulting in a DNA molecule that in itself is different. This may, however, be due to damage to the template properties of inZrivo irradiated DNA. Hudnik-Plevnik and Stocken (161) also observed some differences in the structure of DNA synthesized de novo in UV-irradiated bacteria. All these observations suggest clearly that ionizing radiation may produce, both in vivo and in vitro, different types of changes in DNA structure: base destruction, sugapphosphat,e bond cleavage, chain breakage, cross-linking of DNA strands, and DNA degradation. These changes may impair DNA template functions in replication and transcription.

IV. Radiation Action on DNA Replication There are two major functions of DNA in cellular metabolism: selfreplication in the DNA polymerase system and the synthesis of RNA by the DNAdependent RNA polymerase system. Any alteration in the DNA structure induced by ionizing radiation may interfere with either of these functions. Recent work shows that ionizing radiation does indeed reduce the synthesis of DNA and of RNA. The factors involved in this radiation action on the DNA are not yet wholly understood, but they are of great importance in understanding the nature of radiation action on living systems. I n any case, according to the dogma of contemporary molecular biology, one should expect that the st,ructural integrity of DNA, which

HADIATION-INDUCED

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acts as the primary template containing the genetic code from which all cellular properties and attributes stem, must be held inviolate if correct genetic information is to be passed into the ceIl and its progeny.

A. BriefSurvey ofthe Biochemistry of DNA Replication The available evidence on the replication of DNA in living systems suggests that one strand of the parent DNA is passed to each of the daughter molecules, whereas the complementary strand in each of these molecules is synthesized from nucleotide units (162,163).From genetic and autoradiographic experiments, it appears that synthesis of the two new strands of DNA invivoproceeds unidirectjonally and simultaneouJy from a single starting point in the original molecule (16.4). The replication of DNA takes place within a fraction of the generation time. The enzymes involved in DNA replication, the DNA polymerases or DNA nucleotidyltransferases, have been isolated from microbial cells (165) and animal tissues (166-168). They catalyze the formation of polydeoxyribonucleotides from the 5'-triphosphates of deoxynncleosides in the presence of MgZ+ ions and DNA (template) (169-173). Samples of DNA from widely different sources are equally effective primers of DNA polymerase, and small oligonucleotides also behave as primers (174). However, the end products formed by highly purified enzyme preparations differ from native DNA in certain physical and biological properties in spite of being presumably exact copies of the template with molecular weights similar to that of the primer (1 ’73,175). These nnd other results lead to the conclusion that DNA polymerase in viva either exerts the function of a repair enayme, as already demonstrated invitro(1761, or represents the "replicase," an enzyme specific for the synthesis of only one strand of DNA in the direction of the 3'-hydroxyl end. Since Cairns (164) has shown that both strands of DNA in the bacterial chromosome grow simultaneously in one direction, another enzyme corresponding to DNA polymerase is postulated. Recently, two DNA polymerases of different template specificities have indeed been isoIated (176). I n spite of the vast literature on structure and synthesis of DNA, little is known about the molecular properties of the replicating machinery itself of which DNA polymerase is assumed to be an essential constituent. This is again a serious difficulty encountered in the attempts t o study the action ofradiation on DNA replication.

B. Radiation Actionon DNA TemplateFunction in Vifro Using UV-irradiated DNA as a primer (template) in the calf thymus polymerase system, Bollum and Setlow (177)found that UV-induced

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dimers ofthymine between the adjacent residues in polynucleotide chains are the major, but not the only, photoproducts responsible for the loss in priming ability of irradiated DNA. The loss in priming activity at high doses is somewhat correlated with the number of thymine sequences in the DNA. Because the lesions induced in DNA structure by ionizing radiation are, as mentioned earlier, very heterogeneous, and nonselective as compared with those produced by UV, no specific lesions of DNA can be clearly identified as the cause of the similar loss of DNA template function after ionizing radiation. However, the experiments of Harrington (84,85), Zimmermann et al.(88), and Hagen et al.(88a) suggest that the priming activity of DNA for DNA- and RNA-polymerases is severely depressed even by low doses of radiation. Harrington demonstrated that irradiation is effective whether delivered t o the whole cells or to dilute solutions of DNA. 1000, and 10,000 R) of DNA in dilute solution deIrradiation (500, presses markedly its priming ability for both DNA- and RNA-polymerases (Fig. 12,A-C). Irradiation with 500R reduced the priming ability of DNA in a DNA-polymerase reaction to about 50%,whereas loo0 R reduced it to 26-27% of the control (84, 85). The priming activity of irradiated DNA appeared to decrease more when DNA-polymerase preparations ofhigher purity were used in the reaction mixture pig. 12A). The DNA irradiated in vivo (extracted from irradiated cells) also showed reduced priming activity in a purified DNA-polymerase system. The results suggest very strongly that X-irradiation in the dose range of 500-1000R inhibits priming activity of DNA, i.e., DNA replication in vitro, when the DNA (dilute solutions). primer (template) is irradiated either in v w o or in vitro The dose effect curve concerning the DNA priming activity, when DNA is presented in Fig. 12 D. It should be noted that was irradiated in vitro, the amount of irradiation necessary to inhibit cell division ranges between 50 and 500 R. Recent work on the physicochemical properties of DNA shows that these doses may produce structural changes that in turn may suppress DNA-template functions. In addition, Zimmermann el al.(88) showed that the priming ability of calf thymus DNA for RNA synthesis is much more radiosensitive than the priming ability for DNA polymerization (Fig. 13). They also found indications of an alteration in DNA structure after low doses of X-rays. An inhibitory action of X-rays on primer activity of DNA is also suggested by the work of Stacey (178), who used a partially purified calf thymus DNA polymerase. It should be emphasized here that the results mentioned above are in 180), some of them contrast to those obtained by earlier investigators (179, having shown that DNA primer activity is unaffected by X-radiation

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(173, 180). This discrepancy may be due either to the properties of primer (native or denatured DNA) or to the source and purity of polymerase preparations use. The recent results (84, 85) concerning the reduced priming ability of DNA in poIymerase systems in which the polymerases are free ofnueleases strongly suggest that the reduced priming activity of DNA probably arises from some alterations produced by radiation in the structure of the DNA. Although, there is no evidence of extensive degradation of DNA under the conditions in which its priming ability is reduced, the reduction in viscosity, the lowering of the thermal transition temperatures (TnJand sedimentation rates, and the increasc in sensitivity to hydrolytic enzymes of D N A irradiated in viva and in vitro indicate rupture of both single and These changes double strands with small regions of denaturation (85, 88). may be accompanied by destruction or alteration of DNA bases causing branching and/or cross-linking of DNA. The breakage of DNA strands could result either from a direct action of radiation on DNA or indirectly as a consequence of oxidation of the sugar moiety. Free radicals may alter the structure of DNA bases, but these altered bases may still remain attached to the DNA chain. Thus, for example, hydroxyperoxides of thymidine may be produced by X-irradiation. It is supposed that dihydroxythymine might be lethal to DNA, because it has been established that saturation of the 5,6double bond of uracil impairs the pairing (182) and replication (182) properties of polyuridylic acid serving as a template in an in vitro system. It may, therefore, be concluded that the loss of the priming ability of irradiated DNA in a polymerase system seems to be caused by chain breaks and alterations in the base structure of the DNA primer, which block the assembly of nucleotides along the template. The idea of the existence of base defects in DNA niolecules is supported by the finding that the ability of irradiated DNA to form specific hybrids with isologous mRNA decreases suddenly after irradiation once a critical point of chemical “defects accumulation” is reached (185). At present it is very difficult to evaluate the biological significance of experiments. First of all, the invitropolymerization of these in vifro labeled DNA precursors does not necessarily mean orderly sequential synthesis of DNA or RNA as observed in the living cells. It is still not possible to determine whether DNA formed in an invilrosystems is a faithful copy of the original DNA, or whether changes in base sequences have taken place. The problem is still more complicated because the enzyme preparations used in these systems are not sufficiently defined to exclude the possibility that the observed polymerization in vitro may be ascribed to the action of repair enzymes known to exist in many living systems and types of cells. However, what, still may be an important conclusion from

50-100 250

500 irradiation dose, r

1000

6 700-

500-

Control DNA 300

Minutesof incubailon

FIG.12.(A) Effect of irradiation on the priming activity of DNA for crude and purified DNA-polymerase. The incubation mixtures contained: dATPJ4C (4.3gM; 7200cpm/mpmole), dGTP, dCTP, dTTP 8.6r M each), MgCL (2.4 mM), Tris buffer pH 8.4 (30.8mM), 8-mercaptoethanol (1.2 d), DNA (50gg unheated), PO4 b d e r pH 7 (5.7 a), enzyme: HC DNA polymerase (0.025 ml, 12.9fig protein) in a final volume of 0.875ml. I Crude enzyme (HC DNA polymerase, 0.02 ml, 10.3pg protein, 50 pg of DNA/tube); - - -, purified enzyme (AK VII, 0.02 ml, 5.6pgprotein), DNA = 25 gg/tube. From Harrington (84). (B) Time courseof clATP-"C incorporatioil using control arid irradiated DNA as primer. Incubation was carried out under the condition described underFig. 12 A. Enzyme = AT< VII, 0.02 ml,5.6gig prot,ein. DNA = 50px/(.ul)e. Fn~mHsrrington (84). 140

-

500 500r

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Dose

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FIG. 12. (C) Effect of enzyme concentration on the incorporation of dATP-14C using unirradiatedandirradiated DNA’sasprimers. Theincullation conditions as in Fig. 12 A. From Harrington (84). (D) Dose effect curve of relative DNA priming activity. For determination of priming activity, samples of 0.25 ml contained, in pmoles: Tris buffer (pH 7.9), 10; MnC12, 1.0; MgCI,, 5.0;[S-14C]ATP(120-150 counts/min per mpmole), 0.10; nonlabeled CTP, GTP, U T P each, 0.10; 0-mercaptoethanol, 0.2; protein, 0.1mg; DNA, 0.01 mg. A M P incorporation is related to the AMP incorporation on unirradiated DNA. The symbols indicate various independent experiments. From U. Hagen, &’I. Ullrich, 13. Kroger and E. Petersen, manuscript in preparo.tion (1968). 147

3.01

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loot a z

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.-E B

oc--O

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52 nmoles D N A - P

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0 1 I

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FIG.13 (A). Priming ability of irradiated DNA in the RNA polymerase system. Samples of0.25ml contained in pmoles: Tris buffer MgCle,1.0; [& C]ATP (1.56 x 106counts/min per pmole), 0.10; nonlabeled CTP, GTP, UTP, each 0.10; (pH 7.91, 10;MnC12, 0.25; 8-mercaptoethanol, 2.0; 0.02 mg protein and DNA as indicated. From Zimmermann etal.(88). (B).Effect of irradiated DNA on nucleotide incorporation from nucleoside triphosphates into RNA. Labeled nucleoside triphosphates : p X, [&14C]ATP (1.56 lo6counts/min per pmole); A, [8-14C]GTP (1.83 X lo6counts/min per pmole); 0, [8-14C]UTP(2.02 X lo6 5 counts/& per pmole). Incubation mixtures as in Fig. 13 A with addition of 0.1 pmole of the labeled nucleotide; 0.1pmole of each nonmg protein in the case of[W*C]ATPor 0.04mg in the other cases. From Zimmermann etal. labeled nucleotide; 52mpmoles DNA-P;0.02 Z (88).

x

R Y

a

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these in vitro studies is that no effect can be ascribed to the total inability of irradiated DNA to act as a primer for D N A synthesis in vivo. It may rather be supposed that through the persistence of synthesis with damaged DNA primers, a population of DNA molecules unable to support the survival of the irradiated organism may be produced. This damage, if not repaired-as seems to be the case with radiation-sensitive strains-may prevent cell multiplication and cause cell death.

C. Radiation Effects on DNA Synthesis in Vivo AIany physical and chemical agents may produce inactivating DNA alterations, which, in contrast to mutagenic alterations, block DNA replication and thereby prevent cell multiplication and cause ceIl death. The study of DNA template function in vivois complicated by the fact that radiation-induced structural defects in DNA, which otherwise inhibit normal DNA replication (synthesis), may be repaired. In almost all living systems, complex enzymatic mechanisms exist that serve in a general way to protect the informational character and template structure of DNA against change. It is, therefore, very likely that the principal lethal events in almost all living systems are caused by the presence of unrepaired defects in DNA structure which may, irreversible, block normal DNA replication (i.e., DNA synthesis) and its transcription (i.e., synthesis of

RNA) .

1.RADIATION ACTIONON DNA REPLICATIONOF VIRUSES

Viruses seem to be appropriate systems for attempts to relate DNA damage to a particular effect, such as a loss of the capacity of a virus to reproduce itself in a sensitive host. Since the reproduction of DNA viruses depends upon DNA replication, it seems possible, by following viral replication, to follow the replication of its DNA. Thus, it has been demonstrated that pha.ge 4x174,containing single-stranded DNA, is more ~ ) to ionizing radiation (1Ct"a) after replacing sensitive to UV-light ( 1 0 1and thymine by 5-bromouracil in the phage DNA. However, this correlation between radiation-induced damage to DNA and phage replication became even more evident when experiments were performed with infectious DNA isolated from phage BU+X 174and irradiated as free molecules in solution (10.2~). As shown in Fig. 1 C, the surviving fraction of infectious DNA (giving rise to phage replication) is significantly decreased after gamma irradiation when phage DNA thymine is substituted by 5-bromouracil. Cysteamine gives a protective effect in normal infectious DNA and in BU-DNA (10%~). Direct irradiation of frozen or dried single-stranded DNA viruses

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(4174and 513)yields exponential or one-hit survival curves. This may indicate that one random hit anywhere in single-stranded molecule of DNA or RNA can inactivate the ability of a virus to reproduce itself. The protein coat, which should absorb about the same amount of energy per unit of mass as the nucleic acid, appears radiation-insensitive at least for the function being measured, the ability of the phage to infect the cell (61). The results also appear to indicate that there is little or no redundancy in genetic material for coding information in the single-stranded phage DNA (184). The effect of ionizing radiation on the replication of single-stranded DNA is in agreement with 32P“suicide” resdts, which show that the alteration of one bond in the single-stranded DNA can lead to destruction or to mutation (186-187). I n double-stranded DNA Viruses, only simultaneous double breaks in 56,129). the DNA, and presumably base alterations, seem to be lethal (61, Single-strand breaks seem not to be lethal, and were assumed to be of little or no consequence to the survival of these viruses ( l a g )Sublethal . damage may be manifested as a dosedependent decrease in the ability of a phage to reproduce the normal number of progeny (burst size) in its first life cycle (188).Hence sublethal damage to phage DNA decreases the rate of phage DNA replication. This view is supported by the finding that radiation of complexes of T2 and E.coliB acts on phage DNA but not on the mechanisms utilized for viral DNA replication (189). It suggests also that the bacterial enzymes involved in the synthesis of the precursors needed for viral DNA replication, as we11 as the enzymes involved in the last step of DNA polymerization, and those involved in phosphorylative oxidation in bacteria are not damaged even a t very high dose levels. Therefore dose-dependent lethal events, disturbances in burst size, and the increase in the latent period of the survivors may be directly related to definite structural changes in the DNA produced by ionizing radiation. From the foregoing, it has become clear that viral radiobiology may be directly related to the damage induced by ionizing radiation in the structure of viral DNA molecules. Damage to DNA that prevents DNA replication is lethal for viruses (Figs. 1 and 7). 2.RADIATION EFFECTS ON DNA REPLICATION IN BACTERIA In proceeding from viruses to bacteria, the working hypothesis that radiation-induced structural defects in DNA, unless repaired, are the major cause of lethal events must be considered more carefully since the bacterial cell is a more complex system. I n addition to molecular damage, the “cytoskeletal” structure, so esseiitial in preserving theinternal organization of the cell, has to be considered in all the events that may lead to

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death after irradiation. For exainple damage to the cell membrane causing a release of enzymes and/or precursors of macromolecules may play an important part in the lethal events that follow ionizing irradiation. The attempt to correlate cell death with radiation damage to the two template functions of DNA is complicated by the extensive DNA degradation and repair. On this basis, survival of bacteria after ionizing radiation should be determined jointly by the probability of formation of DNA structural defects and the subsequent repair or restoration of the capacity for replication and transcription (Fig. 9). Hence the principal lethal events ofunrepaired defects in DNA structure are the cessation of normal DNA replication and/or inhibited or abnormal transcription. It should be 1?7). emphasized that this seems to be the case after UV irradiation (161, Thymine dimers and other photoproducts within DNA strands appear to cause both in vivoinhibition of DNA synthesis and in vitro inhibition of DNA replication (177, 190). DNA replicated on the damaged template seems to have an altered structure (177). Moreover, Setlow and Setlow ( 1 9 0 ~discovered ) that thymine d5mers in transforming DNA can block its biological functions.’ Recent work,as already mentioned, shows that ionizing radiations produce drastic effects on the DNA of bacterial cells, causing reduced synthesis, reduction of genetic transcription, and DNA degradation (10-19, 54-74). These lesions in DNA appear to be responsible for the loss of viability in X-irradiated cells. The physicochemical and radiochemical nature of such lesion? in DNA has not been clearly established. Alldata available up to date suggest that ionizing radiation reduces the DNA synthesis (10-19, 54-74). This effect is dose dependent; higher doses in some strains may completely prevent DNA synthesis. The reduction of DNA synthesis is apparent both in cultures grown in oxygen and in nitrogen atmospheres. (Fig. 14,A and B) (264). This reduction in the rate of DNA synthesis seems not to be due to the lack of precursors (32, 79,86)or to a decreased activity of the enzymes but rather to damage to DNA template involved in DNA synthesis (86), functions. McGrath and Williams (16)and Kaplan (17)observed, in a decreased alkaline sucrose gradients of DNA from X-irradiated E .coli, sedimentation rate of denatured and native DNA attributed to single- and double-strand scission (Figs. 9 and lo). Irradiation-induced DNA damage may involve also alterations of DNA-basestructures. Such chemical alterations may result in errors in pairing and a consequent, slightly altered, replication may then give rise to new abnormal DNA molecules with altered sequences contributing to the production of mutations and reproductive cell death (Fig. 27).Furthermore, cross-linking between two complementary bases or between a base

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7000

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aooo

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FIG.14. Effects of irradiation on DNA synthesis in E . coli15 T- L-.(A)The cultures were oxygenated and then irradiated with various doses (5500 R/min) The postirradiation DNA synthesis was followed by [l*CJthymineincorporation into DNA. For lowdoses or radiation, DNA synthesis is slower than in controls, whereas a t doses of 20 kR, DNA synthesis isstopped. From Pollard and Ashey (154). (B) The synthesis ofDNA in bacteria irradiated under conditionsas in Fig. 14A,but with cultures nitrogenated priorto irradiation. The rate of DNA synthesis under these conditions is relatively less reduced than ifthe cells are oxygenated prior to radiation. From Pollard and Achey (154).

and protein moiety, orbetween two bases on the same strand could obstruct or block DNA replication and thus hinder cell multiplication. It is reasonable to assume that the probability ofcross-linking increases with increasing reactivity of the radicals induced by radiation. The electron spin resonance

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data of Phil and Sanner (191) indicate that the radicals formed in guanine and cytosine are more reactive and more easily formed than those induced in adenine and thymine base pairs, suggesting that these bases may be particularly involved in cross-linking reactions. It is evident that no given lesion induced by ionizing radiation in the structure of DNA can be clearly identified as the cause of loss of DNA template functions. The observations on reduced DNA synthesis after ionizing irradiation do not provide information of a qualitative nature as regards alterations in the pattern of DNA synthetic processes. One can suppose that some more subtle changes in DNA replication may occur even after doses of radiation that appear to have little effect on net DNA synthesis. Thus, it has been suggested by Billen et al.(?2,169, 160) that after doses of X-rays or UV light that kill most of the exposed cells, new DNA synthesis is carried out in a manner different from the normal, producing a DNA that seems different from the normal. Billen elul. note that the effect of X-rays on the DNA replication system is such M to alter the normal course of the chromosome replication sequence. Radiation appears to bring about an eventual initiation of additional replication sites. It could be speculated that some DNA fragments undergo limited replication. Since the rate of DNA synthesis in E. coliis decreased after X-irradiation, replication at the original growth points may be excluded and synthesis at new replication sites apparently does not take place simultaneously. If it were not so, one would expect increased rates of DNA synthesis after irradiation. The results could also suggest that certain portions of the chromosome are replicated twice and other portions not at all. Thus the DNA may be only partially replicated. If this is the case, one would expect a heterogeneous DNA profile in CsCl density gradients. This seems indeed to be the case since the DNA synthesized by X-irradiated cells exhibits a more complex and heterogeneous banding pattern than does DNA from unirradiated cells (72, 169, 160). This density heterogeneity reveals that after exposure to higher doses, all the DNA synthesized after irradiation shows a broad band with a mean buoyant density close to that expected for single-strand fragments made up of light DNA (72,169,160). Relatively few studies have been directed at the molecular mechanism of this aberrant DNA replication and the physical nature of the DNA synthesized subsequent to irradiation. It should be noted that Pettijohn and HanawaIt (IZ), studying DNA replication in bacteria after UV irradiation, also observed in irradiated cells newly synthesized DNA molecules banding in CsCl at a position intermediate between the heavy and the light DNA. I n conclusion, there are at least several immediate effects of ionizing radiation on bacterial cells: degradation of DNA, delay of damaged DNA replication resulting in the synthesis of modified D 3 A molecules, and

154

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T. KANAZIR

cessation of transcription. But in addition to these changes in the structure and function of DNA, the cells incubat,ed after irradiation become “leaky.” This effect is time dependent and suggests an effect of radiation on the structure of the cell membrane. The cessation of genetic transcription may cause a failure of the cell to form the enzymes necessary for the growth of a11 the components of the cell wall and membrane; the resulting alteration may cause a steadily increasing permeability in growing cells. Excess radiation can completely stop transcription and therefore produce permanent damage to the cell wall which leads to cell death. Therefore, one can also suppose that the initial lesion of ionizing radiation may be a rupture of the attachment of the DNA growing point to the cell membrane and/or the loss or inactivation of the replicating enzymes bound to the cell membrane. This may be followed by degradation of DNA and cessation of transcription at least in that part of the bacterial chromosome near the rupture. Restoration of the cell wall might facilitate the attachment of DNA growing points to the cell membrane and thereby prevent degradation of DNA and allow DNA replication and transcription. In any case, in the light of our present knowledge it is commonly believed that replication death in bacteria treated with ionizing radiation arises primarily through the formation of structural defects in DNA that block normal DNA replication and transcription. The survival after irradiation should therefore be dependent on the repair of these DNA defects, Although it has long been known that X-ray damage to bacteria can be reversed, evidence for “cut-and-patch” repair mechanisms similar t othat observed following W exposure is not conclusive. McGrath and Williams (16) have demonstrated an enzyme repair process that joins together most of the broken pieces of DNA (Fig. 9). Death could result, therefore, from a failure of the cell to repair all the breaks. The outcome of incomplete repair would be reproductive death. However, Dean etal.(18) have shown in M . radiodurans a DNA degradation followed by a resynthesis of DNA very similar to the pattern of excision, leakage, and resynthesis described by Setlow and Carrier (12) and by Boyce and HowardThe repair enzymes are sensitive to iodoacetate (192). Flanders (13). Therefore the resistance to radiation shown by different bacterial strains reflects repair capacity. Billen etat.(72) found no evidence for a repair synthesis of the type seen in W-irradiated cells. Since repair of DNA breaks appears to occur (16, 7 2) after X-irradiation, the following possible explanations are apparent: (a) repair involves a simple rejoining of breaks in the DNA backbone, which may or may not involve phosphate exchange; (b) repair synthesis is less extensive than that following UV exposures; or (c) relatively long fragments of DNA are excised as compared to UV. These results suggest that the dose required to kill resistant cells is

RADIATION-INDUCED

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155

IN DNA

determined not only by the intrinsic sensitivity of the DNA, but also by the repair capability. The chemistry of these repair mechanism(s) may differ depending on the nature of the damage to DNA.

3.RADIATION EFFECTON DNA REPLICATION IN MAMMALIAN CELLS The available data suggest that the synthesis of precursors and their phosphorylation immediately after irradiation (1-6 hours) are not altered (40, 79,193). At the time when the synthesis of DNA is depressed or blocked, irradiation does not interfere with the biosynthesis of DNA precursors or with the activity of the kinases phosphorylating thymidine or thymidylic acid (42, 76,193,194). Some workers consider inhibition of nuclear (195) and/or mitochondria1 oxidative phosphorylation (196-200) as the principal disturbance responsible for cell death and inhibition of w

LL 3

0

?

z W LL

a W

m

w 100 200 300 TIME (rnin)

t w

loo

200 NO

TIME (min)

IRRADIATED

100 200 TIME ( m i d

FIG.15.Effects of irradiation on DNA polymerase activity, DNA synthesis, and protein in synchronized yeastcells. The cells were irradiated with50 kR. The soluble dataarefrom 200ml aliquots of theculture containing 10s cells attimezero. For details concerning theDNA polymerase activity assays, seeref. (86).From Eckstein et al. (86).

DNA synthesis, but from the more recent observations of Betel (2Ol), it may be concluded that the activitjy of the enzyme system involved in oxidative phosphorylation of lymphatic cell nuclei is not inhibited after lethal doses of whole-body radiation. It seems also unlikely, as shown in Fig. 15, that irradiation interferes directly with the activities of the polymerase enzymes. A11 recent results suggest that radiation acting on the DNA itself impairs its template (primer) function. The exact nature of the primary effect of ionizing radiation on DNA in mammalian cells remains still unknown. Unlike bacteria, in which continual DNA synthesis takes place in the logarithmic phase, the replicative DNA process in the cells of higher living

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organisms is confined to only a part of the division cycle (74,80,202-207). In mammalian cells, the period of DNA synthesis (S period) is preceded by a GI stage, i.e., by the time of preparation for DNA synthesis (74, 80, 202207,910, Z l l ) ,and followed by a premitotic (G2) stage, i.e., the time of preparation for the impending division. Although little is known about the seem processes essential for mitosis, the observations of NeSkovi6 (207) evidence that synthesis of some mRNA and specific proteins takes place in the G) stage of mammalian cells (Fig. 5A). Mammalian cell cultures offer many advantages for studies related to growth multiplication and survival. The advantages stem from the fact that mammalian cell cultures are comparable to cultures of microorganisms in which each cell has an independent life cycle, and genetic and environmental control are readily attained. Many investigators have recognized these advantages and have used mammalian cell cultures in radiobiological research. The inhibitory effect of ionizing radiation on cell division (74-80, 202-207), protein and nucleic acid synthesis (74-80,193,194) and population growth as well as its promotion of chromosomal and cytological aberration (76,208,209) have been studied, but it is hazardous to integrate the cytological studies with physiological and biochemical ones carried out in different laboratories and with different objectives. In attempting to interpret the results obtained from many laboratories, the following variables must be taken into account: sensitivity of different cell lines, dose rate, radiation source, temperature at the time of irradiation, growth media, manipulation of cells prior to, during, and after irradiation. Unfortunately, these parameters, which may affect the interpretation of the results obtained, vary from experiment to experiment. Hence the aim of this section is not to discuss the numerous observations of cellular radiobiology but to attempt to point out the evidence that may help to create an integrated picture of the mechanisms of radiation affecting DNA replication in mammalian cells. It is well known that after moderate doses of X-radiation in asynchronous populations, the rate of DNA synthesis is at least temporarily depressed. The cells remain longer in the synthetic period while the bulk of RNA and protein synthesis remains substantially unaffected (74-80, 204207,193,294). The nonirradiated cultures multiply exponentially, whereas cell culture irradiated in the logarithmic phase do not mdtiply a t all. As the rate of thymidine incorporation falls immediately after irradiation, it follows that the rate of DNA synthesis in the cells, which are in the S phase, must be affected immediately by the radiation (Fig. 5,C, D).The data of Mak and Till (205) suggest that nearly all the cells irradiated in the GI phase can initiate DNA synthesis, but the onset of DNA synthesis is delayed. The rate of DNA synthesis of cells irradiated in the S phase was

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depressed very shortly aftcr irratliut ion (ill the close range 580-10,000 rack). Using synchronously dividing cells (HeLa S3), Terasima and Tolmach (2%’)demonstrated that the capacity of these cells to synthesize DNA is more affected by radiation when the cells are irradiated in the S phase than when irradiated in another phase of the cell cycle. Dewey and Humphrey (209) showed that L cells irradiated in the S phase display a higher frequency of chromosomal aberrations than cells irradiated in the GI or G:! stages. The lesion(s) in the molecules responsible for these various radiation responses remain to be identified. One could imagine that the irradiation damage to DNA prevents DNA replication and/or transcription, i.e., the synthesis of RNA necessary for the formation of a protein that is in turn required for the repair of the damaged DNA and/or its replication. This view is supported by the fact that in logarithmically growing cultures the rate of DNA synthesis decreases essentially to zero in the 9- to 10-hour period following inhibition of protein synthesis with puromycin or of that 2l%%’l4). of RNA synthesis with actinomycin (207, That the inhibition of DNA synthesis results from a direct effect on the progress of synthesis itself, and not from an artifact, produced by an alteration in the composition of the pool of DNA-precursors, was confirmed by the examination of the acid-soluble constituents of the cells. The nature and size of this pool were found to be unaffected by irradiation of the mammalian cells (76, 79, 195, 194). This finding was also confirmed by irradiating a synchronous population of cells in suspension during the period of DNA synthesis and measuring the rate of thymidine uptake at various times during the cycle. An apparently immediate major effect on the rate of DNA synthesis was observed. Cell division and the terminal stages of the same cell cycle were markedly delayed and depressed in a dose-dependent fashion. In experiments in which two complete cell cycles were studied, the irradiation (dose range 500-1000 rads, dose rate 100 rads/ min) was applied during the GI and S periods of the first cycle (Fig. 5, C, D). Only when the cells wcre irradiated during the S period was a marked effect seen on DNA synthesis during the first cell cycle (Fig. 5 , C, D). In some experiments, a small recovery in the rate of DNA synthesis toward the end of the S period was noted. In the case of cells irradiated during the S period of the first cycle, autoradiographic studies showed that for the remainder of the first cycle the percentage of labeled cells was the same as in the control culture (90% at the peak of S), but the irradiated thymidine incorporation into the DNA was reduced. During the second cycle, both the percentage of the labeled cells and the amount of labeling were reduced as compared with the control cultures (79). These results clearly indicate two important and separable effects of radiation: (a) if the cell is irradiated in the S phase, DNA replication is

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largely inhibited; this prevents cell division and ultimately leads to cell death; (b) if the cell is at mitosis or in GI when irradiated, DNA replication in the same cycle apparently proceeds normally, but cell division is nevertheless delayed and depressed and only a limited amount of DNA synthesis occurs in the second cycle (Fig. 5 D). It follows that the DNA replication mechanism is more radiosensitive during the S period than during other cycle periods. A possible explanation for this is that any damage within a replicating stretch of DNA twin helix will rapidly block the replicating process. One could assume that synthesis cannot proceed beyond a damaged region. The lower radiosensitivity of the replication system when exposed to radiation prior or after the S period may be due either to greater compactness of the DNA at that time, or/and to the existence of repair mechanism(s), such asthose observed in bacteria. The latter seems to be the case found “non S-phase” incorporation since -Painter and Rasmussen (77,125) of thymidine in the GI and Gz periods. If the phenomenon of repair is to be accepted as a fact, it would mean that the repair mechanisms have to operate rapidly enough to eliminate the damage caused by irradiation in the it would stages preceding DNA synthesis. According to Brent etal.(79), appear that damage produced 1-2 hours before the synthetic period is reached can be repaired. Furthermore, Elkind and Sutton (616-218) demonstrated that mammalian cells are able to repair sublethal radiation There have been several reports describing similar results damage in vitro. in a variety of mammalian cell systems both in vitro and in vivo(223-224). The results presented show that X-irradiation of mammalian cells causes an inhibition of uptake of precursors into the DNA molecule. No similar results occur with bulk RNA or proteins. Since the pool of phosphorylated DNA precursors remain unaffected, the observed effect seems to be specific for DNA and therefore does not appear to involve initial phosphorylation of the labeled precursors (nucleotides). The question that may be raised concerns the exact site of the radiation effect. The polymerases, already present in the cell, probably are not the radiosensitive site because, in general, fully formed enzymes require relatively high Thus, only the DNA remains as doses fortheir inactivation (86, 193, 226). a candidate for the primary site of the radiosensitive reaction. This view is strongly supported by the findings of DjordjeviE and Szybalski (74), who found that mammalian cells, if grown in media containing 5-bromodeoxyuridine under conditions in which part of the thymidine in DNA is replaced by the analog, become more radiosensitive to ionizing radiation. Szybalski (146) claims that radiation-induced scission of the main chain of DNA may be facilitated by the introduction of one bromine atom into the DNA molecule. He showed also that bromouracil-labeled DNA is more

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susceptible to X-ray induced double-strand breaks. Lett etal.(80) suggest alternatively that bromouracil incorporation renders radiochemical damage more difficult to repair by subsequent me ta b o h processes. However, the repair system may itself be damaged. From these observations, it follows that radiation-induced death of mammalian cells is probably determined by two distinct mechanisms, damage to DNA or DNA-protein and interference with some metabolic processes responsible for repairing the DNA damage, which, unless repaired, are transcribed and translated. Damage to structure of DNA and/or repair mechanism(s) may have as biological consequences : teniporary delay in entering mitosis; death of cells before entering mitosis (interphase death); death of cells after a few mitoses (reproductive death).

4.RADIATIONEFFECTON DNA SYNTHESIS IN HIGHER ORGANISMS Acute mortality of animals results presumably from cell killing and more specifically from cell killing in the intestinal tract and hematopoietic system. Because of the close correspondence between the cell survival curves 2n vitro and inv i m ,it seems reasonable to suppose that the mechanisni of cell death is similar in both cases. Despite the extensive st8udywith mamthe mechanism of cellular death invivois not yet well malian cells inuitro, understood. I n higher organisms, this problem seems to be more complex because of the multiplicity of biochemical changes that have been described in animals following lethal or sublethal exposure to X-irradiation. The response to radiation that may be detected in an intact animal is complicated by the presence of dead cells and by adjustments occurring within the tissues following X-irradiation. These adjustments may include migration of cells, proliferation of surviving cells, and repair of radiation damage. Although it is unlikely that whole body irradiation of an animal injures only one type of molecule, it is not unreasonable to assume that the alterations of only a few types of macromolecules by even relatively small doses may be critical and that these may cause death. One could also assume that the damage caused by X-radiation results from alterations to one or a few types of macromolecules regardless of the complexity of the living system. For the moment, however, an attractive working hypothesis is that among the many effects produced by ionizing radiations in cells of different tissues and organs, the most characteristic is a temporary or permanent inhibition of DNA synthesis. More than 30 years ago Hevesy (226) reported that X-irradiation inhibits the incorporation of radioactive phosphate into the DNA of Jensen sarcoma, a fiiidiiig interpreted as an inhibition of DNA synthesis. It was established subsequently that inhibition of DNA synthesis represents a universal effect of ionizing radiation on living systems.

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However, it should be emphasized that the sequence of niacromolecular alterations following ionizing radiation and leading to a block in DNA synthesis arid to cell death inmammalian tissues is far from beingelucidated. Unfortunately, the many contributions t o this field contain data obtained under such a variety of experimental conditions that its integration into a coherent picture of the mechanisms of radiation action on higher organisms is exceedingly difficult. Moreover, one should take into account not only intracellular regulatory mechanisms, but also the functional states of many interrelated systems of tht. whole organism, in particular the hypot halainus-pituitary-adrenocorticalsystems, since it is knuivnthat radiation, as n stress factor, may act on the functioning of these systems. Radiatioii might, cause thc release of hormones interfering with the rate of genetic transcription and translation. Therefore, this section is concerned mainly with the immediate radiation effect on DNA synthesis, prior to cell death and necrosis. Among the observations, some of them controversial, made by the early investigators who studied the effect of total-body irradiation on DNA synthesis, there is considerable agreement that there is a reduction in the rate of DNA synthesis, as first demonstrated by Hevesy etal.(226) and later confirmed by many others (34-38, 81-83). Much of this work was concerned with the DNA content of various tissues and with the rate of incorporation of various labeled precursors into the DNA. More recently, labeled thymidine became the precursor of the choice for studying transitory changes in the synthesis of DNA after irradiation. Thus, in a series of experiments with rats irradiated with 50, 100,200,400, and 800R, using thymidine-2-14C as the precursor 60 minutes after irradiation, Nygaard and Potter (81) showed a rapid and extensive inhibition of the incorporation into DNA of thymus, spleen, and small intestines (Fig. 16). With lower doses this depression was less evident, as expected. These observations are in agreement with the results of previous studies on the effects of irradiation of incorporation of various precursors into DNA of the thymus (37, QO), spleen (39, 41),and intestine (227). The common feature of all available evidence is that a decrease in the rate of incorporation is observed soon after the irradiation. It should be noted that the tissues studied have high rates of cell division and hence high rates of DNA synthesis. Furthermore, the inhibition of incorporation by X-radiation has been observed with all precursors of DNA so far tested (37, 58, 81,228). The interpretation oflabeling data is complicated by the fact that few tissues consist of a single population of cells so that one obtains the summaafter irradiation tion value for the radiation effect on various cell types. Also, wc often deal with a changing of ~ c lpopulation l owing to the lossrof the niow sensitive cells, affcci ing tlw relative aliuntlnnccs of thc various typcs

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I N DNA

SPLEEN THYMUS

7

6-

50

5-

400 r

8

4

12

16

20

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HOURS AFTER IRRADIATION

FIG.16.Effect of X-irradiation on DNA synthesis in various tissues of the rat. The rats were irradiated with various doses of X-rays and examined at 15,30,60, 120,and 240 minutes after irradiation. Synthesis of DNA in different tissues was followedby determining the DNA specific activity. [14C]Thymidine was used as DNA precursor. Variations of specific activities that reflect the changes in the rate of synthesis ofDNA in thymus, spleen, and small intestine after different doses of total body X-irradiation are presented in (A),(B), and (C), respectively. From Nygaard and Potter (81).

1

I

1

I

8 12 16 20 HOURS AFTER IRRADIATION

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and numbers of cells in the tissue. Different approaches to the problem have studied been made to overcome these difficulties. Myers and Skov (193) the effects of X radiation on DNA synthesis in rat thymocytes in vitro. These cells are readily isolated from the thymus, irradiated and kept alive in suspension for at least one day. A dose of 1000 rads of X- or y-radiation causes an inhibition of DNA labeling. The irradiated cells incorporated thymidine at almost the control rate during the first 15 minutes of incubation at 37”C, but the rate of DNA synthesis was slower. Nygaard and 230)gave rats whole-body irradiation with 6oCoy-radiation Potter (229, for1minute, the doses ranging from 125to 4600reps, using the incorporation of thymidine-14Cinto DNA in 8 minutes as a measure of the rate of DNA synthesis. These short “pulses” were selected to permit detection of relatively rapid changes in the rate of synthesis and to reduce the likelihood of changes in the cell populations during the observations. The results showed an immediate, dose-dependent effect of radiation on the incorporation of thymidine into the DNA of the spleen and small intestine. The determination of free thymidine triphosphate ruled out changes in the studied, DNA synthesis pool of precursors as the cause. Fausto etal.(83) in nuclei isolated from spleen after irradiation (800 R). The results obtained in vivoand in vitro showed that X-irradiation inhibits the incorporation of thymidine-’4C into DNA of the spleen and into DNA of the nuclei of the spleen homogenates in the early time intervals after irradiation, during which there is no release of nucleases. Thymidine incorporation into DNA was inhibited about 50% at 1, 3,and 6 hours after irradiation. In of thymidine and TMP into TTP addition, the phosphorylation in vitro by the spleen supernatants obtained from irradiated mice was not inhibited 1 hour after the administration of 800 R. All these results suggest that, irradiation interferes directly with DNA replication. This view is supported 232), who observed an immediate inhibition of also by Van Lancker (231, th~midine-~H incorporation into the DNA of homogenates of regenerating liver. Furthermore, in order to assess the effects of irradiation on DNA synthesis in hepatocytes of regenerating liver, parallel quantitative autographic and biochemical studies of the same samples of liver have been The animals were locally irradiated 19 hours after partial made (233). 6000, and 12,000R, at hepatectomy and doses of 188,375, 750, 1500, 3000, a rate of 300 R/min, were applied to the liver immediately before the injection of thy~nidine-~H and the animals were sacrified 1 hour later. DNA synthesis was depressed to 35% of the control value with no decrease in the percentage of labeling of hepatocytes and no change in acid-soluble precursors or in DNA content. This strongly suggests that the inhibition of the incorporation ofthymidine into the DNA of hepatocytes is due to the radiation effect on the final steps of DNA replication, i.e., on polymerization.

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These results strongly suggest that irradiation inhibits the synthesis of DNA in the cells of various tissues. This seems to be a universal effect of ionizing radiation on living cells regardless of the complexity of organization of the living system as a whole. The finding raises several questions: What is the cause of inhibition of DNA synthesis? What is the nature of DNA synthesized de novoafter irradiation? Is the radiation damage in higher organisms reparable or not? Since the inhibition of DNA synthesis was observed within a few hours after irradiation, it cannot be ascribed to the increased activity of nucleases (83), to cell necrosis, or to changes in cell populations in the tissue of irradiated animals. Other factors must be involved. The block in DNA synthesis may be caused (a) by a change in the amount of DNA precursors or change in the activity of enzymes synthesizing them; (b) by a decreased rate of oxidative phosphorylation, i.e., ATP resynthesis; (c) by radiation damage to the enzymes essential for DNA replication (polymerases) ; and/or (d) by damage to the template DNA itself so that it cannot be replicated. a.Radiation Effects on thePoolof D N A Precursors. Bishop and Davidson (234) found a detectable increase of acid-soluble DNA precursors in rabbit appendix and thymus 1 hour after total-body X-irradiation (1000 R) and a marked increase after 4 hours. Ord and Stocken (40) observed, 1hour after total-body exposure t o lo00 R, an accumulation of both mono- and triphosphates of deoxyribonucleosides and some ribonucleoside triphosphates in rat thymus. These observations support the view of early inhibition of DNA synthesis not for lack of DNA precursors, but rather from failure of the polymerization processes or breakdown in the essential energy-generating mechanisms in the nucleus. b. Radiation Eflect o n theActivity of EnzymesInvolved intheSynthesis of DNA Precursors and on theResynthesis of ATP.The preformed kinases, polymerases, and glycolytic enzymes seem unaffected by X-radiation The observations that up to 1hour between 250and 10,000rads (179,180). of after irradiation there is no interference with phosphorylation invitro th~rnidine-~H ort h ~ m i d y l a t e - ~ suggest ~ c that irradiation does not interfere with the activity of thymidine or thymidylate kinases (83).These findings systems are in good agreement with observations concerning from invitro the activity of these enzymes in the spleens of irradiated mice (230). The fact that all the phosphorylating enzymes and polymerase activity can be recovered, after irradiation, in the cytoplasmic fraction of liver homogenates (231, 236)indicates that the “biological” doses of radiation do not inactivate the catalytic sites of the existing enzymes. However, the formation of enzymes seems to be a process sensitive to radiation. In mammals, the iqatbesis and activity of several enzymes appear to be closely correlated

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with the cell proliferative rate. Among these enzymes are deoxycytidylate deaminase, thymidine and thymidylate kinases, thymidylate synthetase, and possibly others. These enzymes may be identified by the fact that they usually exhibit a high activity in rapidly proliferating tissues, such as many tumors, regenerating liver, and embryonic tissues, and little or no activity in nongrowing tissues and organs, such as adult liver (210, 236239,240). The available information indicates that the synthesis of dCMP deaminase is sensitive to y-and X-irradiation in the thymus (lo00 R), in 5-day-old chick embryo (510 R), and in regenerating liver (1500R). Furthermore, the synthesis of thymidine kinase in regenerating liver is also very sensitive to radiation. A similar decrease in the activities of these enzymes was observed after actinoniycin D treatment (239). These results seem to be consistent with the view that a decrease in enzyme formation arises from an alteration in the DNA molecule by radiation as well as by actinomycin D treatment, leading to a cessation of genetic transcription of DNA and an inhibition of the production of the mRNA’s needed for enzyme synthesis (238, 239,240). Since ATP, as the energy source, may be involved in the synthesis of DNA precursors, it may be of interest to summarize briefly our information on the action of radiation on oxidative phosphorylation. Whole-body irradiation (700R) uncouples oxidation and phosphorylation in the mitochondria of spleen and thymus (241). However, on the basis of experiments with liver homogenates, prepared from rats exposed to 650 R 3 hours earlier (242), it was concluded that resynthesis of ATP was unaffected. These early results on ATP resynthesis in mitochondria are 241). However, a decreased resynthesis of ATP controversial (196-201, in nuclei from thymus cells and other lymphoid tissues was observed shortly after exposure to low doses of X-rays (195), and inhibition of nuclear oxidative phosphory lation was considered to be the principal lesion associated with interphase death (195). This hypothesis was further extended by Whitfield and Youdale (243, 244), who suggested that inhibition of nuclear oxidative phosphorylation gives rise t o accumulation of inorganic phosphate in nuclei. This should in turn induce dissociation of the nucleoprotein complex leading to the disintegration of the nucleus. However, recent experiments of Betel (201) show that neither the overall rate of ATP synthesis nor the rate of ATP utilization is affected in rats given whole body irradiation with 875 rads. It can therefore be concluded that nuclear oxidative phosphorylation cannot be one of the main factors limiting the rate of DNA replication, i.e., the rate of DNA synthesis, in the cells of higher organisms. c.The Eflect of Radation o n D N A Polymerase. Extracts from molds and from yeast cells exposed during interphase to lo5 R of X-radiation (Fig. 15)

RADIATIOK-INDUCED

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IN DNA

lG5

shows no decrease in DNA po1yiner:tse activity (86).The enzyme seems insensitive to ionizing radiation. Therefore, lack of polymerase activity cannot be the cause of the inhibition of DNA synthesis. This finding may have biological significance, since polymerases may play a role in the repair process. d. T h eEfects of Radiation on DNA Template Function inVivo. Although the exact site of the primary effect of X-irradiation on the DNA biosynthetic pathway in cells of higher organisms remains unknown, it seems quite probable that radiation-induced damage to the DNA of such cells may decrease its priming capacity. The findings of Meyers and Skov (193) indicate that the synthesis of DNA in thymocytes may result from the disruption of the deoxyribonucleoprotein of the cell. This immediate effect was observed at a dose level at which all other cell functions remain unaffected. Loss of deoxyribonucleoproteinintegrity, therefore, may be considered as a possible cause of inhibition of DNA synthesis. Voiculetz etal. (245) found that the chromatographic profiles of DNA synthesized de novo in bone marrow cells, intestines, and testes of mice irradiated with 1350R differ from the profiles of the corresponding DNA’s from nonirradiated animals. Kritskii etal.(43, 139) have shown that X-irradiation (2000 R) applied locally to the legs of rabbits depresses the synthesis of DNA, and decreases the viscosity of bone marrow DNA. Furthermore, immediately after irradiation a rise in proportion of readily precipitable DNA (fraction I) was observed, probably indicating a change in the population of DNA molecules. The collapse of the DNA-protein (246) or the loss of DNA 247a)induced by radiation in mammalian cells may be due integrity (247, to the destruction of the -SH groups of nuclear histones or to partial denaturation of DNA, which could reduce the template capacity of DNA. These changes in the integrity of DNA protein may also alter the DNA priming capacity for RNA polymerase. These results are consistent with the idea that irradiation produces functional damage to the DNA template. If so, one should then expect that some changes in the composition of newly synthesized DNA would occur. observed tbat irradiation causes a marked Berenbom and Peters (248) change in splenic DNA composition, resulting in relative increases in guanine and adenine and relative decreases in cytosine and thymine. However, this coufd be attributed to changes of cell population. Observations made in our laboratory also support the idea that the composition of the DNA synthesized after irradiation may be altered. Bebarevii: etal.(249) showed that, on the fourth day after irradiation, the specific radioactivities of intestinal DNA purines and thymine were significantly higher than the specific radioactivities of purines and thymine of intestinal DNA of nonirradiated animals. This change may indicate either a faster synthesis or a more rapid turnover of intestinal DNA in the irradiated animal, although

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the change in the intestinal cell population induced by irradiation may also be one of the causes for the changes in specific radioact>ivities.The same change was evident even on the ninth day after irradiation, i.e., the specific radioactivities of DNA guanine, adenine, and thymine from irradiated animals were considerably higher than those of nonirradiated controls. Although the biochemical meaning and nature of these changes remain unknown, they could be due to damage to the DNA template, resulting in synthesis of a faulty DNA, the properties and composition of which seem to differ significantly from the normal. Similar changes were observed in intestinal RNA purines (249). Tokarskaya (250) demonstrated that the point of radiation injury on the DNA of dry seeds is the DNA thymine. This change in DNA seems to be potential, finding its expression in the course of development. Using 14C-labeling,she showed that the changes in the DNA template capacity resulted in the synthesis of faulty DNA. Thus, DNA synthesized after irradiation has a qualitatively different composition. Although more studies are needed in order to explain the effects of ionizing radiation on DNA replication in intact animals, it seems vepy probable that ionization of DNA molecules in the cells of different tissues produces alterations to electron configuration along the DNA molecule that result in the prevention of DNA replication and/or transcription of the genetic code. If this proves to be the case, then we are dealing with a universal phenomenon, one that is valid for all cells regardless of the structure of tissues. It s e e m unnecessary to invoke the presence of different mechanism of action to explain the effect of radiation on DNA synthesis in different tissues and organs.

V. Radiation Meets on DNA Transcription and on the Biosynthesis of RNA Finally, in attempts to assess the immediate biochemical damage in animal cells and the response of tissues in intact animals to radiation exposure, cell population kinetics and the characteristic pattern of metabolism of different types of cells at different stages in their life cycles must be taken into consideration. In addition to these factors, the fact that in animal cell nuclei large amounts of protein are associated with DNA should be also taken in account, since the presence of thiol proteins (histones and other proteins) may be especially useful in protecting DNA against ). radiation-induced damage by low doses of radiation ( 2 4 7 ~However, damage to these proteins, such as oxidation of thiol groups on the histone F3-1fraction or the inhibition of phosphorylation of the lysine-rich histone (Fl fraction), may result in such conformational changes which may

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prevent chromosomal DNA from carrying out its normal functions. (See article by Hnilica in Volume 7 of this series.) As already mentioned, it is widely accepted that DNA is the main target of radiation action. Since DNA is endowed with two basically vital functions, replication and transcription, it seems logical t o consider here the problem of the effects of radiation on the transcription of genetic information. Radiation effects on transcription were earlier studied by following the induced synthesis of enzymes. In the earlier work with UV and ionizing radiations (651-256), it was shown that radiation action depresses the rate of induced enzyme synthesis, leading to the conclusion that this arises from a decrease in the rate of transcription. This must be closely, if not directly, related to radiation damage to DNA and/or degradation of the DNA template. Today radiation effects on genetic transcription may be studied namely, in a DNA-dependent RNA polymerase system, and/or (a) in vitro, (b) within the living cell. This section is concerned with radiation effects on transcription in both systems.

A. Radiation Effects on Priming Activity of DNA in the RNA Polymerase System2 If the synthesis of all classes of RNA is DNA-dependent, any structural alteration produced by radiation in the DNA cistrons coding for RNA should be transcribed and expressed in the structure of newly synthesized RNA. The priming activity of DNA in RKA synthesis is reduced after irradiation of DNA in dilute solution (84, 85, 88, 88a). This can be seen even after exposure to 1000 rad and it is more pronounced after higher doses. Harrington (84, 85) showed that the priming ability of DNA for RNA polymerase is significantly depressed regardless of whether the irradiation was delivered to the living cells or t odilute solutions of DNA. From these results, it follows that 50-1000 R of X-irradiation inhibits the priming activity of DNA in the RNA polymerase system. I n the cells of higher organisms, the genome is riot composed onIy of free DNA. It is an intricate nucleoprotein complex containing several classes of proteins, such as histones and residual acidic proteins (256). Therefore one could expect that radiation damage may vary according to whether deoxyribonucleoprotein (DNP) or DNA is irradiated, as the DNA may be protected by the protein in t,he complex. In the case of irradiation of calf-thymus DNP, the major part of the radiation damage orciirs i n the Seearticle by Hichardson in this volume

1ti8

D. 1. KANAZIR

protein moiety of the DNP (266). Hence it is obviously important to examine the effectsof radiation on the template activity of DNA and DNP. For that purpose, DNA and DNP prepared from calf thymus were irradiated in aqueous solution and tested for their priming capacity in systems with RNA poly’merase prepared from Micrococcus lysodeikticus and [3H]CTP as labeled precursor (256). A dose of 2000 rads reduced the priming capacity of free DNA by 5oy0 whereas the priming activity of DNP was only about 25% of that of an equivalent amount of “free” DNA. Irradiation of DNP with a dose of 12,000 rads caused only a 30% reduction in RNA synthesis, whereas the same dose lowered the template activity of free DNA to 2% of the original DNA activity. With higher rads its activity doses, the priming activity of DNP rose until at 72,000 was approximately 40% above that of the unirradiated DNP. These results can be explained by assuming that in the normal state the protein of DNP is not complexed with DNA throughout its length and that only noncomplexed DNA regions serve as primer sites for RNA synthesis as suggested by Huang and Bonner (257) and Georgiev ( 2 5 7 ~The ) . radiation may, therefore, cause two different affects. First, it may attack free, noncomplexed regions of DNA causing an initial drop in the rate of RNA synthesis. Secondly, further increase of the radiation would cause, t o a lesser or greater extent, the release of DNP proteins and uncover regions of free DNA that may then serve as new sites for DNA transcription, increasing the rate ofRNA synthesis. Furthermore, Ord and Stocken (247a) have demonstrated that exposure of rat thymus nuclei in vivoor in vitroto 1000 rads may cause oxidation of the thiol groups of the histone F3-1fraction. This oxidation is likely to contribute to a reduction of DNA transcription, since the oxidized F3 histone fraction is more effective in depressing DNAdependent RNA synthesis than its reduced form ( 2 4 7 ~ ) . These results suggest that radiation causes a decrease in priming capacity of DNA for RNA synthesis and that the rate of transcription decreases. But from these results it is not possible to decide whether the RNA formed on an irradiated DNA template is merely reduced in quantity, although still a faithful copy of the original DNA, or whether alterations in its base composition have taken place. I n order to answer these questions, Weiss and Wheeler (258) subjected the RNA newly synthesized on irradiated DNA templates to a detailed analysis of base composition. Their results showed again that the rate of synthesis of RNA had been significantly depressed and that the ratios of cytosine to adenine and of guanine to uracil together with the nearest neighbors to adenine in the RNA ivvcre :iltercd. This Ruggests that if transcription is taking place on irradinted DNA, some errors i n the base composition of the newly synthesized RNA may occur.

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IN DNA

These results may, however, serve as a liiie of evidence that radiation to DNA may change (a) the rate of DNA-dependent RNA synthesis in zritro and (b) the base composition and probably the nucleotide sequences in RNA thus synthesized. It is difficult to explain the results observed in the various i n vitrosystems as the overall chemical changes induced by ionizing radiation in DNA are not known in detail (see Section 1x1). In addition to the destruction of DNA bases, breaks in single and double strands, as well as cross-linking between the DNA molecules and splitting of hydrogen bonds are observed (84, 85, 88,88a) after irradiation, and the melting temperature for DNA &rands separation is lowered from 75 C

t

a W c

Q (L

g

8z a

f 8

25

id. I

0

P'

-G-C)(

Mn

a/

-

FIG.17.Priming activity and molecular weight of degraded DNA. For determination of priming activity, samples of 0.25 ml contained, in pmoles: Tris buffer (pH 7.9), 10; MnC12, 1.0; MgC12, 5.0; [8-14C]ATP(12&150counts/min per mpmole), 0.10; nonlabeled CTP, GTP, UTP, each, 0.10; 8-Mercaptoethanol, 0.2; Protein, 0.1 mg;DNA, 0.01 mg. AMP incorporation related to the AMP incorporation on untreated DNA. From Hegen et al.(88~). 0, +-radiation; A, W light; 0 , deoxyribonuclease; 0 , ultrasound.

to 67 Cowing to the breaks and partial denaturation of DNA. It should be stressed that alterations such as breaks and/or base destruction could be the direct or indirect cause of the reduced priming activity of irradiated DNA in RNA synthesis (Fig. 13). The evidence suggests that the reduced priming activity of DNA after irradiation is not caused by an extensive degradation of DNA by nucleases contained in most polymerase preparations (84). It may, therefore, be assumed that the loss of the priming activity of DNA in the RNA polymerase systems is directly impaired by alterations produced in DNA structure by radiation. Hagen etal.(88a) made an extensive study of the priming activity of DNA samples degraded and denaturated by various agents (Fig. 17). After

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gamma irradiation (with 6oCo), the molecular weight of DNA and its priming activity decreased simultaneously, while the single breaks produced by enzymatic degradation affected the transcription process only very slightly, if a t all. Hence a loss of priming activity after gamma irradiation may depend on some damage other than breaks. The importance of base alteration is obvious in the case of UV irradiation where loss of priming activity is observed even before any degradation of DNA molecule is measured, probably because of alterations in DNA structure. Hence one could assume that the loss of priming activity after gamma irradiation is also attributable mainly to alterations in the base structure and only to a smaller extent to the breaks in DNA strands. Bollum and Setlow (27Y) showed that formation of pyrimidine dimers by UV irradiation may decrease the rate of RNA synthesis. The model they suggested assumes a bridging of the gaps formed by the excision of dimers by “end addition” of noncomplementary bases, after which normal transcription may resume. The rate of RNA synthesis seems to be resumed, but the structure of this RNA may be altered. Alternatively, the breaks caused by radiation may produce additional sites to which the RNA polymerase can be bound. This may explain why denatured Tz DNA (259)and heat-denaturated calf thymus DNA (660) are significantly more effective in binding RNA polymerase than is native DNA. Furthermore, UV irradiation of native E .eoli DNA has no effect on the rate of formation of the DNA-RNA polymerase complex, (261) while the priming capacity of DNA is significantly reduced ( I W ) .At the same time, it seems that RNA-polymerase becomes much more strongly bound to irradiated DNA (258). Thus, one could envisage that irradiation produces new sites capable of binding RNA polymerases at a normal rate, but not capable of supporting the synthesis of RNA. These sites could be associated with regions of chain breakage. We cannot presently differentiate between the two possible mechanisms involved in the radiation-induced inactivation of DNA priming activity. The first mechanism assumes a decrease in the rate of incorporation of labeled precursors into RKA due to a cessation of movment of RNA polymerase along the DNA strand caused by an alteration in the base structure. The second mechanism assumes that radiation-induced DNA strand breaks produce new extra sites to which RNA polymerase is bound, but that these sites are not capable of supporting RNA synthesis. The nature of the new sites remains unknown, and it cannot be excluded that base alterations may also be involved in their formation. As mentioned above, base composition and nucleotide sequence seem to be impaired in the RNA newly synthesized on DNA templates damaged by radiation. However, the experiments of Weiss and Wheeler (258) indicate that the only changes common to all RNA polymerase systems in which the

RADIATION-INDUCED

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IN D N n

17i

priming ability of irradiated IlNA s was studied are a decrease in the rate of cytosine incorporation and an increase in incorporation of one or both of the purine bases. This might suggest that, whatever products are formed in DNA structure after irradiation, their function is to direct incorporation of purines rather than of pyrimidines. I n this connection the data of Ono et al. (262) may be of interest. These authors observed that the effect of UV radiation on the priming activity of poly C may largely be reversed by addition of ATP, a noncomplementary nucleotide, to the incubation mixture. The priming ability resulted in the formation of poly (G,A) instead of poly G. This suggests that a t any site of injury on the DNA base, a noncomplementary base can be inserted in the structure of the product synthesized on that primer. This may indeed explain the changes in base ratios and in their nearest neighbors observed in the structure of RNA’s synthesized on irradiated DNA templates. experimental conditions are so Finally, it should be stressed that in vitro different from those prevailing in living systems, so loosely related to processes taking place in vivo,that one should be very cautious in extrapolating from one to the other. However, these data do suggest that radiation may, directly or indirectly, interfere with the processes of transcription and thus cause a severe unbalance in celI metabolism.

B. Radiation Effects on Genetic Transcription in Vivo The attempt to explain the radiation-induced inactivation and death of a living system in terms of a molecular damage in the genome DNA (or DNP) with resulting severe imbalance in metabolic processes has bene a major concern of modern radiobiology. Genetic transcription is a key process in the metabolism of all living systems. Consequently the problem of how radiation affect,s transcription may be central in molecular radiobiology. Information concerning this problem is very scanty, but such as may have a bearing on the processes of transcription in phage (virus)-hostbacteria, mammalian cells and some organs of higher organisms is summarized here.

EFFECTSON THE GENETIC TRANSCRIPTION OF VIRUSES Viruses are used effectively as a major tool to decipher the genetic code and to understand gene action and the function of mRNA. This system seems most appropriate for the study of radiation effects on DNA replication, but information on the radiation effects on viral transcription is very scanty. Experiments have been carried out with the DNA of phage alpha serving as a template for RNA polymerase. The experiments carried out by 1.

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Graziosi (263) were performed as follows. Cells multiply infected with phage alpha were allowed to start phage growth. At 5 minutes after infection they were rapidly cooled to stop phage growth and were irradiated at 0°C with doses ranging from 50,000 R to 200,000 R of X-rays at a dose rate of about 2000 R/min. The cells were then quickly warmed and kept 2-3 minutes at normal growth temperature. At this point a2Por ~ridine-~H was added, and after 5 minutes the cells were rapidly cooled, collected, and washed, and the pulse-labeled RNA was extracted. After purification on a Sephadex column, this RNA was used in annealing experiments with phage alpha DNA or with its purified “light strand” DNA. The results showed that X-irradiation caused the infected cells to synthesize smaller amounts of hybridisable, phage-specific RNA, as if this RNA seemed to lack some nucleotide sequences present in the RNA synthesized on nonirradiated DNA molecules. From these results, it could be concluded that the rate of synthesis of RNA, i.e., the transcription of the phage DNA, is also damaged invivoand that the structure of the newly synthesized mRNA is altered. The results are very similar to those observed in invitrosystems. Phage alpha is a double-stranded DNA virus. One could assume that this particular DNA contains a very large unit of transcription in which many cistrons are orderly aligned and orderly transcribed from a single initiation point. Hence, any radiation-induced lesion-single-strand scission or base damage-in this critical segment may slow down the initiation or the rate of movement of RNA polymerase along the DNA strand, resulting in a reduction in the rate of synthesis of phage mRNA. Alterations in RNA composition resulting from the absence of some nucleotide sequences, as revealed by the hybridization technique, may be explained by assuming that the population of templates changed because of the fragmentation of DNA induced by radiation. RNA polymerase is able to react with DNA molecules whether intact or fragmented. This idea seems to be supported by the observation of Hagen el al.(@a) that degradation with ultrasound and DNase does not prevent transcription. A distinct loss of template activity can be seen only in DNA molecules of molecular weight less than lo6(Fig. 17). A similar line of evidence indicating the synthesis of abnormal mRNA on irradiated DNA template was observed with UV-irradiated vaccinia virus (264), a double-stranded DNA-virus that multiplies in the cytoplasm of HeLa cells. It is thus possible, by pulse-labeling and preparation of cytoplasmic extracts, to examine the rate and duration of synthesis of the virus-specific RNA. Such experimentsshowed that UV-inactivated particles can function as templates in the process of transcription of “early” viral mRNA. The base composition of polysome-associated RNA in cells

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infected with UV-inactivated particles was dearly different from that of cells infected with nonirradiated virus particles (264). A significant increase in adenine and a decrease in uracil was observed. This resembles the changes observed in the base composition of RNA synthesized on irradiated DNA templates in RNA polymerase systems, described by Weiss and Wheeler (958). 2.THE EFFECTSO F

ItADIATION ON

GEKETIC

TRANSCRIPTION ITU'BACTERI 2 Bacteria appear to be appropriate for the study of the effects of ionizing radiation such as X-rays and y-rays on genetic transcription invivo. In many respects this is true, but it is also true that the processes through which bacterial cells are damaged are not understood. a. The Effects of Radiation on theSynthesis and Function of mRNA in Bacteria. Earlier experiments indicated that the synthesis of RNA is influenced to a much lesser degree by irradiation than is that of DNA (32, 59, 60, 267, 268), but these reports were based on estimates of the total amount of RNA, no importance being attached to the size and function of the several classes of RNA (32, 59, 60). Pollard and Barone (90) observed very little effect of gamma radiation (12,000 rads) on the function of preformed mRNA in E .coli15 T-L-. But it has been suggested that one important action of ionizing radiation is concerned with transcription processes (L69), and observations on the effect of ionizing radiation on the induced synthesis of formic hydrogen catalase (271, 272), and @-galactosidase(90, 269, 273)indicate lyase (270), an inhibition by low doses of radiation and gave experimental support to the view that in these cases transcription is affected. Pollard (269) showed that genetic transcription ceases if cells are irradiated shortly after induction. The production of @-galactosidasecontinues for a short time after irradiation and then ceases. The kinetics of cessation of transcription give information on both mRNA decay and the rate of transcription. Since the latter may be affected directly and indirectly, Pollard and Barone (90) attempted to determine which of the factors in genetic transcription are affected by peroxides and which by direct radiation effects. When bacteria were irradiated prior to induction in the presence of lW7M catalase (to prevent the indirect effects of radiation, i.e., the action of hydrogen peroxides formed in the medium), the rate of transcription of the galnctosidase operon (i.e., the synthesis of mRNA responsible for the synthesis of the enzyme) was reduced in a manner dependent on the dose. If under the same conditions a pulse induction is performed to elicit a short period of synthesis of mRNA priorto

174

b. T. KA??AZiR

irradiation, there is no effect on preformed mRNA, at least for doses up R. Thus it appears that there is no effect of radiation on the to 12,000 expression of preformed mRNA. At higher doses, transcription is almost completely prevented. Since one striking effect produced by direct irradiation is the degradation of DNA, the supression of transcription could be related to this degradation, although this cannot explain the complete inhibition of transcription. It cannot be excluded that the undegraded DNA may have a molecular structure that cannot be transcribed. It should be noted also that hydrogen peroxide generated in the medium after irradiation may temporarily inhibit the processes of transcription, but it may also destroy the preformed mRNA and prevent its expression (90). These results could mean that the rate of transcription (synthesis) of mRNA in bacteria and its structure may be damaged by radiation. The nature of the damage to DNA that prevents transcription is, however, not known. b. The Efects of Radiation on theSynthesis of Pulse-Labeled RNA, Ribosomal RNA,and Ribosomes inBacteria. Since all classes of RNA are transcribed on the DNA, one cauld expect radiation-induced damage to DNA to result in an altered rate of production and/or an altered structure either of pulse-labeled RNA, which contains mRNA, or the precursors of ribosomal RNA. However, in comparison with mRNA and tRNA, the rRNA forms a major portion of the total RNA of the bacterial cell. Up to now the only known function of rRNA seems to be a structural one, preserving ribosome conformation. Results from the studies of UV effects on bacteria strongly support the view that rRNA as well aa ribosome subunits may be damaged by radiation. It is known that UV irradiation effects temporarily the synthesis of DNA Salmonella typhimurium, and other bacteria (SZ, 158, 9 7 4 , in E. coli, whereas overall RNA synthesis is only slightly inhibited (%’,lS8, 274,278). However, the synthesis of the 50 S ribosome subunit and the 23 S RNA molecules is drastically inhibited in E .coli by U V irradiation (26’6, 276, 277’). The synthesis of 23S RNA seems more sensitive to W irradiation than that of 16 S RNA. According to the results of Drakuli6 etal.(665), the slowing down of 50 S ribosome synthesis takes place very shortly after irradiationmore exactly, during the period when the synthesis of DNA is still inhibited. This inhibition seems to be partial and temporary. After the repair of DNA damage, the synthesis of 50 S ribosomes is restored. The resynthesis of DNA and of 50S ribosomes appear to be simultaneous events after UV irradiation; they may be related processes. This idea is supported by the finding of DrakuliE etal.(265) that if the synthesis of DNA in bacteria is permanently inhibited, the specific activity of 23 S RNA remains lower than iq controls, It seems, therefore, that the repair of DNA and the

RADIATIOX-INDUCED

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175

resynthesis of 50 S ribosomes after UV irradiation are metabolically interrelated processes. One should thus expect that if ionizing radiation damages DNA templates, transcription of rRNA may be altered. As already mentioned, DNA in the bacterial cell is degraded upon exposure to X-radiation (57-62). However, little is known about the influence of this event on the subsequent synthesis of rRNA and ribosome subunits. The results discussed earlier showed that, despite the detrimental effects of X-rays on the synthesis and the stability of DNA in E. coli (5742) and Salmonella typhimuriurn (158)) a net synthesis of bulk RNA and proteins occurs for a period corresponding closely to the duration of postirradiation growth. However, Hudnik-Plevnik (138) showed that higher doses of X-rays may depress the synthesis of bulk RNA in a dosedependent manner. However, the resolution of rRNA into its components on sucrose density gradients and “chasing” experiments indicate that a decline in the rate of formation of larger RNP particles, the 50 S ribosome subunits, and changes in the synthesis of 23 S RNA take place after irradiation (278, 279). Since little is known about the effect of X-irradiation on the synthesis of pulse-labeled RNA, Hudnik-Plevnik etal.(278, 279) have been making typhimurium, strain LT2, was efforts to approach this problem. Salmonella irradiated during the logarithmic phase of growth and incubated at 37 C. Ten minutes later, the suspensions were “pulsed” for 3 minutes with [”PI inorganic phosphate and RNA was extracted by the method of Suzuki and Hayashi (280). The “step-down” culture was also used to The RNA was analyzed in ensure preferential synthesis of mRNA (281). sucrose density gradients. The sedimentation pattern of the RNA extracted from nonirradiated (normal) controls and from irradiated bacteria after R are presented in Fig. 18. There is no doses of 10,000R and 20,000 change in the absorbance at 260mp and in the relative amounts of 23S and 16 S RNA. This finding may indicate that there is no release of nucleases following secondary radiation damage to ribosomes. Although no degradation of bulk rRNA was observed, the pulse-labeled RNA from irradiated cells differed in two respects from t,he pulse RNA of controls in that there was an increase of label incorporated into all fractions of the RNA from the irradiated bacteria, but this increase was not uniform, being markedly higher in the region of 16 S to 23 S than in the region of 4 S to 16 S RNA. This difference became more evident when experiments were performed in “step-down” conditions. It should be pointed out that Klamerth (267) also found that RNA from irradiated E.colicultures showed consistently adenine. To decide whether structural higher incorporations of p4C] changes occur in the newly synthesized RNA, two kinds of approach have

176

D. T. KANAZIR

been made in our laboratory: (a) “chasing” experiments were performed and (b) base ratios of different RNA fractions were determined. For these purposes, suspensions of bacteria irradiated with 20 kR were3eft to grow for 3 minutes with 32Pi or [14C]uracil,washed, 10-15minutes, then pulsed and left to grow for another 30 minutes in an unlabeled medium (“chased”

--I

i-

0

%

-s 01

.

O n

b

+J

i

E

s

!

Tube m b e r

I

Tube number

I

I

FIG. 18. The sedimentation pattern of phenol-extracted RNA from cells of Salmonella t ~ p ~LTz ~ exposed ~ ~ to T a 3-min ~ u s2P043~ pulse. Gradient: 204% sucrose, Mga+ cone. 5 X lo- M ; 4.5 hours centrifugation at39,000 rpm. (A) and (C), controls for cells irradiated with 10 kR and 20 kR, respectively; (B)and (D), cells irradiated with 10kR and20 kR, respectively. From Hudiiik-Plevnik (d?‘8).

with uracil). Growth was then stopped, and the RNA was extracted and fractionated on sucrose density gradients. The results are presented in Fig.19.During the %base" period, the transition (transformation) of precursor to rRNA into stable rRNA was completed in the control cells,

177

RADIATION-INDUCED ALTERATION IN DNA

whereas in the irradiated bacteria this transition of precursor rRNA to stable 23 S rRNA appeared to be significantIy depressed; its specific activity was markedly lower than that of the 23 S component of RNA obtained from unirradiated bacteria. At the same time an accumulation of labeled RNA sedimenting in the region between 4 S and 16 S was observed

V

2

N

0

E

0

Q V

FRACTION

No

Pro. 19. Sedimentationpatternof phenol-extracted RNA from cells of S. typhzmurzum LT2 exposed to a 3 minute [14C]uracilpulse (A, B), followed by a 25-minute “chase” with cold uracil (C, D). Gradient: 2 0 4 % sucrose, Mgzf conc. 5 x 10-4 M ; sedimentation for 5 hours a t 39,000 rpm, SW 39 rotor, Spinco L Model ultracentrifuge. Graphs (A) and (C), unirradiated controls; (B) and (D), cells irradiated with 20 kR X-rays. 00, absorbancy at 260 mfi; @- - -@, radioactivity precipitable with cold trichloroacetic arid. From Hudnik-Plevnik (879).

in the irradiated bacteria. This is riot the case with the nonirradiated controls. Our results are in good agreement with the findings that the addition of either chloramphenicol or puromycin (276) results in an accumulation of 14 to 18 S RNA. Under such experimental conditions, the formation of ribosomes is prevented. Our results suggest t h t tlic syiiiliesis of the 23 8 rilmsomd RNA

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D. T. KANAZIR

component is altered in irradiated bacteria. The processes of synthesis of this rRNA fraction seem to be more sensitive to radiation than those of the 16S rRNA component. The accumulation of lower molecular-weight RNA's during "chasing" could suggest a degradation of precursor to rRNA during its transformation into the stable 23 S rRNA fraction. Furthermore, these data are also in good agreement with those of Frampton (268) and with the findings of Klamerth (267’). Frampton (268) studied the postirradiation synthesis of total RNA and ribosomes after exposure of E. coliB/r to X-rays (Fig. 20,A, B) and found that the net synthesis of RNA, measured by the orcinol reaction and by incorporation of [2-14C]uridine,is depressed in irradiated cells. Cells receiving doses of 40,000, 60,000, and 90,000 R all exhibited a leveling of

:3

16

15

14-1

A

I

Incubation time (mln)

FIG.20. (A)Effect of varying the dose of X-rays on incorporation of [Wluridine by Escherichia coliB/r.Cells were resuspended at one-half the concentration at harvesting in media containing 4 mMCi/ml of uridine-2J4C at a concentration of 16 pg/ml. After 45 minutes 4 mCi/ml of uridine-Z-"C were added at a concentration of 1 pg/ml to the preincubated irradiated cells (90kR). Samples (3ml) were removed at the times inon dicated, extracted with coldtrichloroacetic acid (10%) for 30 min arid roll~.ctrrl membrane filters for assay of radioactivity. From Frampton (268).

RADIATION-INDUCED

ALTERATION

179

IN DNA

net RNA synthesis between 30 and 40 minutes if cultured in a complete medium. A continued synthesis of RNA in cell suspensions exposecl to 20,000R was greatly reduced after this interval. Furthermore, when labeled uridine was added 45 minutes after 90,000It, i.e., at the time when RNA synthesis had ceased, a significant incorporation of uridine was

0.3

12

0.2

8

0.I

4

10 20

30 40 I0 20 Fraction number

30 40

FIG.20.(B)Density gradient sedimentation analysis of extracts prepared from whole B/r). After exposure to X-radiation (90kR,dose rate irradiated and control cells ( F .coli between 2700 and 2900R/min), the bacteria were resuspended i71 media at the original pCi/ml) a t B concentration concentration and again incubated a t 37 C.[3H]Uridine(0.5 of 0.5p g / m l was added after various periods of postirradiation incubation. After 1 minute, 500pg/ml of nonradioactive uridine was added, and the cells were incubated for an additional 15 minutes. The cell extracts were prepared as follows: The frozen cell pellets were resuspended in cold buffer, deoxyribonuclease was added (1 mg/ml), and the cells were disrupted in a French pressure cell at 8000-10,000 psi. The crude cell extracts were clarified by centrifugation a t 20,000 X g for 15 minutes. Immediately after clarification, 1.0 ml of each crude extract was applied to the top of the sucrose gradient (20-5 %). Centrifugation was carried out at 90,000 X g for5 hours. I n a Spinco Model L preparative ultracentrifuge (Spinco SW 25 swinging-bucket rotor), The various timea ofpostirradiation are indicated as follows: ( a ) control, pulse a t time zero; (b)irradiated cells, puke at time zero; ( c ) irradiated cells, pulse after 5 minutes (d) irradiated cells, pulse after 10 minutes; ( e )irradiated cells, pulse after 15 minutes; (f) same as (e) except that cell extracts were prepared in the presence of bentonite (1mg/ml). From Frampton (268).

180

T. KANAZIR

observed, indicating a probalde turiiover of somc fract,iori(s) of RNA. After a shiftdown in the rake of growth, irradiated hcteria were no longer able to synthesize RNA, but continued to incorporate radioactive uridine into acid-insoluble material. The RNA synthesis taking place under these conditions is presumably mRNA (281). Resolution of this RNA carried out on methylated albumin columns reveals alterations in the elution profile of radioactive material synthesized between 5 and 35 minutes after the shiftdown. When bacteria were irradiated (90,000 R), pulsed, and returned to “cold” medium (chased), an inhibition in the synthesis of 50 S ribosome subunits was observed whereas 30 S ribosomal material and smaller RNA components accumulated. The preparation of cell extracts in the presence of bentonite (1 mg/ml) to inhibit ribonuclease activity gave similar results. However, Klamerth found that irradiated bacteria (10,000 R) consistently incorporated more [’*C] adenine into the newly synthesized RNA. The total amount of RNA per cell remained unchanged for at least 25 ininutes after irradiation. Later, a slight decrease in mRNA content was observed, but differences in elution profiles were evident. The RNA preparations of nonirradiated controls showed no evidence of a fraction that in RNA preparations from irradiated bacteria, elutes at 0.3M NaCl, and the RNA fraction eluted at pH 9 was far less marked in the control than in irradiated cells. Although no differences in the profile of fractions eluting at 0.6 M NaCl, corresponding to tRNA (4 S to 6 S) was observed, the RNA samples from irradiated bacteria contained a series of smaller fractions corresponding to 12-16S fractions and eluting with 0.8M NaCl that usually were not observed in control RNA samples. These fractions are very slight in terms of quantity but show a high degree of incorporation, i.e., they have a high specific radioactivity. It should be noted that, after an accumulation of similar material was observed. It is “chasing” (279), possible that these fractions represent (a) incomplete mRNA of abnormal structure, not used for protein synthesis; (b) degradation products of precursors of rRNA; and/or (c) products of degraded mRNA. But the possibility cannot he excluded that radiation induces new sites of transcription resulting in the synthesis of new RNA’s of medium quantity, but with zt higher turnover instability than normal ItNA’s-i.e., RNA’s with abnormal structural and metabolic properties. It should be mentioned also that SaviC (282) found, in analyzing RNA from UV-irradiated S. typhimuI-ium on columns, some fractions of RNA with metabolic properties similar to those described by Klamerth. These fractions had altered base compositions. The changes may be attributed to radiation-induced damage to the DNA template resulting in faulty transcription. This assumption is supported by some observations from the i n vitrosystems. Thus, UV

RADIATION-INDUCED

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181

irradiation of poly U markedly stimulates the poly U-directed synthesis of poly A. Poly U is modified by UV irradiation (hydrates and/or dimers of uracil are formed) and this induces changes in the template behavior of irradiated poly U. In contrast t o the transcription reaction directed by unirradiated poly U, the rate of whivh decreases as the reaction proceeds, the irradiated poly U directs the synthesis of poly A by a “copy-slip” mechanism in which short template tracts of U residues terminated by a second type of residue (dimer or hydrate) are copied repeatedly. This is termed “reiteration.” It is postulated that transcription involves copying most of the template to produce an active template-product complex whereas reiteration involves copying of limited portions of the template repeatedly, so that an active coniplex is not formed. Furthermore it seem that RNA polymerase initiates copying randomly on these model templates and copies them sequentially in the 3 to 5’ direction (182). The biological significance of such reiteration reactions observed in vitro is not known, but similar reactions have been observed with several types of mammalian (283,284), avian (285), and bacterial extracts (286). I t may happen that, after irradiation in vivo, the “copying” is restricted to limited fragments of denatured DNA, resulting in a more rapid synthesis of some incomplete molecules of pulse-labeled RNA. The labeling of RNA thus formed may proceed more rapidly than that of RNA synthesized on the normal DNA template. The possibility of RNA digestion due to released RNase or other nuclease cannot be completely discounted, although this is udikely since within the interval during which the abnormal RNA fractions appeared, the total amount of RNA per cell remained unchanged. Nevertheless, one could assume an inherent instability of the highly-polymerized RNA synthesized de novo in irradiated bacteria that makes the pulse-labeled RNA more susceptible to the action of nucleases, leading to partial disintegration of polycistronic precursors to mRNA or rRNA. In order to elucidate this problem, new experimental approaches are needed. The results presented also suggest that the synthesis of the 50 S ribosome subunit is probably altered. It remains to be determined whether an inhibited synthesis of 23 S RNA causes immediate alterations in the synthesis of50 S ribosomes in X-irradiated bacteria. The problem arises as to why the specific cleavage of the precursor RNA into the stable 23 S RXA ribosomal component is altered. The solution to this problem cannot be given as the subsequent events in the formation of rRNA are not yet completely understood (see article by Perry in Vol. G of this series). In pulsechase experiments, including those in which continued incorporation from nonexchangeable nucleotide pools is prevented with actinomycin D, a sequential labeling of 45 S, followed by

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32-35S and 18 5 and finally by 28 and 18 S components (in mammalian cells) is observed. When part of the RNA guanine is replaced by 8-azaguanine, the early steps in this conversion can still occur, but later steps are greatly retarded (287). The later stage of conversion involves the transition of the 32-35S component to 18 S and 28 5 RNA. The niolecular events associated with the transition are still unknown. In any case, our results could suggest that after ionizing radiation this late step in conversion, i.e., the transition to 23 5 RNA, seems to be altered. If specificity toward cleavage enzymes is an inherent property of RNA precursor molecules, the effect of radiation may be regarded as arising from an altered affinity of the precursor RNA for its enzyme. This may be due to a change in base composition of the precursor RNA since it is established that the substitution of RNA guanine by azaguanine may influence the late steps of conversion of precursor RNA to stable rRNA. Alternatively, the necessary specificity may have its origin in the appropriate protein bound to RNA since it is known that puromycin can block the process (287-289). These conversion processes are not yet well understood. If it is assumed that the base composition of the precursor RNA may influence the activity of cleavage enzymes, it may follow that the base composition of the pulse-labeled RNA synthesized in irradiated bacteria is altered. A slight decrease of cytidine and the slight increase of adenosine observed in pulse-labeled RNA of irradiated bacteria may serve as evidence in favor of this idea. After a 30-minute “chase,” the base composition of labeled RNA should be the same as that of bulk RNA if transition of the precursor RNA to the stable form of RNA was completed. It can be seen from Table I that this seems to be the case in nonirradiated bacteria. However, in irradiated bacteria the base composition of “chased” RNA is slightly modified. The content of cytidine was slightly increased, whereas that of adenosine was decreased, with a slight decrease in the amount of guanosine. These data suggest, therefore, that base compositions-i.e., the structure of rRNA and, probably, that of the mRNA synthesized de novo in irradiated bacteria--are changed. This alteration could reflect the “errors” in DNA template transcribed into RNA’s during transcription processes. Further results indicate that both preexisting and newly synthesized RNA are broken down at virtually the same rate in irradiated bacteria. The breakdown of both types starts at the same time, at 30 minutes after irradiation. Since RNA synthesized 60 minutes after irradiation is not broken down, it seems that only RNA synthesized before or shortly after irradiation is subject to degradation (290, 291). These results indicate that the breakdown of RNA during the postirradiation growth of E ,coli B is not an immediate consequence of irradia-

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IN DN.4

TABLE I BASE COMPOSITION OF BULK RNA, DNA AND THE RIBONUCLEOTIDE COMPOSITIONS OF PULSE-LABELED AND CHASED RNA S FROM CONTROL AND IRRADIATED Salmonella typhimurium LT+b “Pulse” 3P, 3 min, 24 C NUCIMtides

CMP GMP AMP UMP a

Bulk RNA

DNA

23.4 32.0 26.1 18.6

26.8 27.0 24.0 22.8

Control

25.0 30.8 22.8 21.6

Irradiated, 20 kR

23.4 31.0 24.0 21.5

“Chase” SIP,30min, 37 C

Control

22.7 31.9 25.5 19.7

Irradiated, 20kR

26.0 30.0 22.6 21.5

From Hudnik-Plevnik (279).

* The base compositions of bulk DNA

and RNA were determined by quantitative paper chromatography. The symbols forbulk DNA and RNA correspond to the purine and pyrimidine bases. The symbols for “pulse-labeled” and “chased” RNA s correspond to the ribonucleotide composition, which was determined by Z*P distribution (279).

tion; there is always a lag of about 30 minutes before the appearance of breakdown products inside the cells and in the medium. Though delayed, this seems to be a radiation-induced process. In addition to delayed activation of nucleases, several complex mechanisms may be visualized in order to explain the observed effects. It may be that products of DNA degradation are transcribed, giving rise to RNA fractions more unstable or more susceptible to the action of nucleases. Moreover, the peroxides formed under action of radiation may prevent transcription and destroy mRNA and probably other classes of highly polymerized RNA, giving rise to intermediate smaller fragments that may be degraded to nucleotides. c. Effects of Radiation on T ransfer RNA in Microorganisms. Like mRNA and rRNA, tRNA is also transcribed on the DNA. This fraction accounts for 10-15% of the total RNA of the E .coli cell. Its size is relatively homogeneous, having a molecular weight of about 25,000(7Ck80 nucleotides). The role of tRNA in protein biosynthesis involves the acceptance of specific amino acids, the binding of the aminoacyl-tRNA’s to the messenger RNA-ribosome complex, and the release of the bound amino acid to the growing peptide chain. Change of secondary structure seems to be a general phenomenon in tRNA inactivation. UV irradiation produces specific alterations in particular bases of a polynucleotide chain, permitting the relation of these photochemical alterations to the biological action of Most work has been concerned with the inactivation radiation (292-295). of amino acid acceptance, but a number of experiments involving transfer From the evidence, and binding steps have also been reported (2924295). one may conclude that such inactivation implicates changes in the sec-

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ondary structure of tRNA, resulting at least in part from the formation of pyrimidine dimers and other photoproducts (296) ? Evidence concerning the action of ionizing radiation on tRNA synthesis and its function is very scanty although Yamamoto (297) recently showed that gamma irradiation may inactivate the function of the “pfI 5 fraction,” i.e., the function of the aminoacyl-tRNA synthetases.

3.RADIATION EFFECTS ON

MAMMALIAN CELLS Clear-cut demonstrations of the effects of ionizing radiation on the RNA metabolism of mammalian cells have been rather scarce in contrast to the many reports on the inhibitory effects of ionizing radiation on DNA structure and synthesis in such cells. In earlier studies, RNA was considered to be relatively resistant to radiation (76,298,699). The reason why the expected damage inflicted by radiation on RNA is not so clearly revealed by our tools of study as that on DNA is not clear. It is hardly conceivable that the structure of RNA itself is much more resistant to ionizing radiation than is that of DNA. From the biochemical point of view, the pathway of RNA synthesis has many processes in common with that of DNA, The synthesis of the precursors, their phosphorylation, and their polymerization are very similar. Since in the mammalian cell the synthesis of RNA is directed by DNA, which can be damaged b y radiation, one could expect that alterations in DNA structure will be transcribed to the RNA and thus affect RNA metabolism. Consequently, in the section that follows, the aim is to discuss the main changes produced by radiation in the metabolic patterns in mnmmalian cellular RNA. a. Radiation E$ectson theRNA Metabolism of Single Cells Cultured in Vitro. The experiments summarized here were carried out under different experimental conditions and with various cell strains so that close comparison of the results is neither justified nor possible. Ionizing radiation produces alterations in the reproductive integrity of cells owing to the damage to the DNA structure. The role of R NA in the expression of these alterations is not yet understood. Harbers et al.(300) were aniong the first to demonstrate an inhibition of synthesis of DNA4 and of nuclear RNA in X-irradiated Ehrlich ascites cells cultured i n vitro. After different doses (750R, 1500R, 3000R) of X-rays, the incorporation of labeled uracil int.0 nuclear RNA was significantly inhibited, while the total radioactivity of the acid-soluble fraction and the rate of uptake of labeled precursors in irradiated cells was almost the same as in the unirradiated ceIls and a slight increase in the specific radioactivity of cyto3

THE

RNA

OF

Seearticle by Setlow inV d . S ofthis series.

RADIATION-INDUCED

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185

plasmic RNA was observed. The inhibition of nuclear RNA synthesis paralleled that of the DNA. This suggests a close relationship between the biosynthetic pathways of these two macromoleculesin the nucleus. Furthermore, it reveals that the synthesis of nuclear RNA is more radiosensitive than that of bulk RNA. Irradiation of the cells (2000 R and 4000 R) causes a change in the “elution profile” of the “nucleolar” RNA, and a decrease in the incorporation of labeled precursors into this RNA fraction (300). However, the data of Myers and Skov (193)suggest that most of the RNA synthesis in thymocytes cultured in vitro depends on the integrity of deoxyribonucleoprotein.In an attempt to clarify the effect of radiation on nuclear RNA, Logan etal.(501) isolated nuclei from calf thymus and studied the effects of X-rays on the uptake of [14C]adenineand [Wlphenylalanine. The nuclei were exposed to 50 R, 300R, and 900 R prior to incubation with labeled adenine. Small doses, such as 50 R, inhibited the uptake into nuclear RNA, whereas DNA-bound RNA resistant to RNase contained little or no labeled adenine. It is generally accepted that the latter is the nascent, newly transcribed RNA. This finding therefore suggests that transcription processes are either very slow or completely stopped. Mori and Morita (502) made a similar approach to the problem, using nuclei isolated from the appendix of the rabbit. The control and irradiated nuclei (1000 R and 2000 R) were incubated with 32P(2 pCi/ml) for 30 minutes at 37 C and RNA was extracted by Kirby’s method and fractionated on DEAE-cellulose columns. Irradiation with 1000 R caused no significant change in the chromatographic pattern of the RNA, but higher doses (2000 R and 10,000 R) caused marked changes in the profiles, with the labeling of fractions eluting at higher salt concentrations (0.6-1 .O M NaCl) being markedly depressed. These experiments demonstrated that after X-irradiation RNA becomes reduced both in amount and in the degree of incorporation of 32P. Budnitskaya etal.(303) using autoradiography, studied the effects of radiation on the stability of RNA structure, the rate of synthesis and the transfer of RNA from nucleus to cytoplasm and found that the pulselabeled RNA is the most radiosensitive. The nucleolar pulse-labeled RNA seemed even more sensitive than the rest of nuclear RNA. The incorporation of [14C]cytidineinto nucleolar RNA was strikingly inhibited, especially a t short “pulses,” and this change was measurable even after only 100 R. Thus, it appears that radiation does affect both nuclear and nucIeolar RNA. The inhibition of the transfer of the label from nuclear to cytoplasmic RNA-i.e., the conversion to functional mRNA and stable rRNA (28S and 18S RNA)-seems also to be impaired by radiation. Better identification of radiation damage to the various types of RNA is certainly needed. I n this connection, Kim etal.(50.4) studied the effects of X-irradiation on the synthesis and turnover of rapidly labeled RNA in cultured human

186

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leukemic cells. The rate of RNA synthesis was measured by the extent of incorporation of tritiated uridine into RNA. Different classes of RNA were characterized by sucrose density gradient centrifugation and by DNA-RNA hybridization. Dose levels of 500-2000rads led t o an accelerated degradation of plelabeled RNA. DNA-like RNA was degraded more than was ribosomal RNA. From the results of Matsudaira etal.(506), it may be concluded that irradiation of hepatoma cells produces a slight inhibition in the rate of synthesis of rapidly labeled RNA. A depression of the synthesis of the pulse-labeled RNA fractions larger than 28 S was evident, though no changes in the UV absorption profiles were found. This interference of radiation with mRNA or rRNA formation should be damaging to the cell as RNA and protein synthesis are necessary for the initiation of DNA synthesis and for entry into mitosis (306). Negkovi6 etal.(207, 306), from morphological criteria, distinguished seven fairly typical development phases of growth during intermitosis of L strain cells. In chronological order, they are M, N, S, -4s, Lal, La2, O.M. The cells irradiated during interphase with y rays (776or 1552rads), and their normal controls were examined for their proliferative capacity. The number of nonirradiated control cells increased by 150% in 48 hours, whereas the cells irradiated with 776rads and 1552rads increased by only 2% and later died. Some of the cells that, at the moment of irradiation, were in the Laz and M phases of interphase divided in the 48 hours after irradiation; when irradiated in other phases, they failed to divide for 24 hours, the synthesis of RNA wa~smuch reduced, and that of DNA was almost stopped. The effect of actinomycin was similar to that of radiation. These results could mean that the initial effect of radiation is on the replication of DNA, the consequence of which is the blockage of mRNA synthesis, whence only those cells which at the moment of irradiation have accomplished DNA replication and mRNA synthesis can divide. Furthermore, blockage of the synthesis of mRNA and degradation of the mRNA already synthesized may explain the inhibition of mitosis in those cells that had not postulated that particular divided. From these results, Ne3kovi6 (207,306) mRNA’s are synthesized in each phase of interphase, and that are functional only during the phase in which they are required. From the action of agents that inhibit passage through the Gz stage (actinomycin D and puromycin), the synthesis of RNA and protein appears to be prerequisite to the formation of the mitotic machinery (207, 211, 212). Temporal differences between the action of the two inhibitors on cells passing from S stage to the Gzperiod of mitosis is consistent with the sequence of events involving first RNA and then protein formation. It is not unlikely that the formation of the RNA and protein essential for the subsequent mitotic division begins in the period of DNA replication or immediately after-

RADIATION-INDUCED ALTERATION I N DNA

187

ward. Thus, when exponentially growing mouse lymphoma ceIls in culture are irradiated with 200 R and then incubated with [3H]uridine,the location of radiation-induced G2 block in the life cycle is very close to the “switchingoff” time of the synthesis of nuclear RNA, possibly of the mRNA responsible for “division-protein” synthesis (307-$09). Hence, the radiation seems to affect the function of activated genes-i.e., their transcription and genetic translation. strongly suggest that the transcription, These studies with cells in vitro i.e., the synthesis of pulse-labeled RNA, is altered by radiation. The instability of pulse-labeled RNA synthesized de novo in irradiated cells seems to be an inherent property of such RNA fractions. This increased instability may be attributed to an abnormal RNA structure. Unfortunately, data on the base composition of RNA newly synthesized in are still lacking. irradiated cells cultured in vitro b. Radiation Action on R N A Synthesis inHigherOrganisms. The more recently developed ideas on the genetic code have stimulated investigations on the effects of radiation on RNA metabolism in a wide variety of organisms. Much of this work has been concerned with changes in the incorporation of precursors into the rapidly labeled RNA. There is general agreement that whole-body irradiation affects the synthesis (transcription) of “pulse-labeled” RNA in higher organisms. This is true even for resting liver, generally regarded as being a radioresistant organ, at leapt with respect to histological alterations (310). The rapid proliferation of cells that follows partial hepatectomy is a remarkable phenomenon in which cells of the adult liver with a low mitotic rate suddenly began mitosis, resulting in a rapid but temporary growth of the organ. “Regeneration” is the term commonly used to describe this compensatory hypertrophy and hyperplasia. Removal of two-thirds of the liver initiates an immediate rise in the RNA content of the nucleolus (311 ) and an overall increase in the rate of RNA synthesis (312, 313). The initial response seems to involve a rapid synthesis of a wide range of RNA molecules and ribosomes, an increase in the free polysome concentration, and a rise in the specific activity of hepatic RNA polymerases (238-2.40,314,3l5). Thus the liver in regeneration is used as a model system for the study of the studied effects of radiation on transcription processes. Uchiyama etal.(316) the effect of X-irradiation on the incorporation of [14C]oroticacid into rapidly labeled RNA of the regenerating liver by irradiating rats with 600 R, 1000 R, and 1500R a t 18 hours after partial hepatectomy. By 6 hours after irradiation, a drop in the specific radioactivity of the rapidly labeled RNA extracted from the regenerating liver of animals thus exposed was observed (Fig. 21A).The effect of X-radiation on the incorporation of [14C]orotic acid into the pulse-labeled (20min.) nuclear RNA of liver in

188

D. T. KANAZIR

I

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6 8 10 12 Dose in roentgens x

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FIG.21 A, B,and C .

RADIATION-INDUCED

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I N DNA

regeneration is reflected in the profiles obtained by centrifugation of RWA samples in sucrose gradients (Fig. 21 I3 and C) (316a). These profiles revealed that X-irradiation does not alter the sedimentation behavior of the bulk RNA. This is not degraded and thus therc is no release of nucleases as earlier assumed. The major change induced by irradiation is a clear reduction in the incorporation of [14C]oroticacid into the rapidly-labeled nuclear RNA fractions of the highest molecular weights (Fig. 21 C) prepared from 6-,12-, 18-and 24-hour-regenerating liver. The inhibition of the incorpora316a). tion was most significant 24 hours after partial hepotectomy (316, Similar results have been reported by Fausto et al.(83) and by Berg and Goutier ( S l y )indicating , an inhibition of the synthesis of rapidly labeled RNA in the nucleus of regenerating rat liver after whole-body X-irradiation. The reduction in incorporation of [TJorotate into rapidly labeled nuclear RNA could result from (a) changes in the permeability of the cell or nucIear membrane for the nucleic acids precursors; (b) activation of latent RNase, which could degrade newly synthesized RNA; (c) dilution of the labeled pool of precursors by unlabeled compounds derived from the breakdown products of nucleic acids in spleen, bone marrow, and lymph nodes; and (d) from a block in biosynthesis (transcription) of rapidly labeled nuclear RNA. FIG.21. Effects of total-body radiation on rapidly labeled RNA of rat liver. (A) Effects of X-irradiation on incorporation of [WJoroticacid in rapidly labeled RNA of regenerating liver. The ordinate refers to the cpm/mg of the RNA extracted (see ref. 316)from the pooled livers of three animals partially hepatectomized 24hours prior to sacrifice. The abscissa refers to the total-body doses of X-irradiation administered 6 hours prior to sacrifice a t 10 min after injection of[Wlorotic acid. A drop in specific activity of the rapidly labeled RNA is evident after exposure of animals to various doses of X-radiation with a marked reduction after 1500 R (316). (B)and (C) Effect of X-radiation on the incorporation of [14C]oroticacid into rapidly labeled nuclear RNA of regenerating liver. [14C]orotic acid (5pCi/100 g body weight) was given i.p. 20 min prior to killing. The livers were removed, the nuclei were fractioned and nuclear RNA was extracted with phevol and dodecyI sdfate by and purified (316u), the method of Di Girolomo et al.(J.Mol.Biol. 8,479,1964), with slight modifications (for details, see ref. S16a). The purified RNA was centrifuged in a Spinco SW-25 rotor rpm for 13hours a t 4 Cinlinear 5% to 20% sucrosegradients. Bamplesof RNA a t 20,000 (5 drops) were collected for determination of optical density and for radioactivity * ) in cpm X measurements. Fig. 21B shows the sedimentation of radioactivity ( 10-2and optical density a t 260 mp (-) obtained for the nuclear RNA fractions of nonirradiated animals killed 6,12,18 and 24 hours after partial hepatectomy and receiving [14C]orotic acid 20 min prior to killing (SL6a). C shows the sedimentation profiles of nuclear RNA from regenerating liver of irradiated animals. The experimental conditions were the same aa in 21 B, but the animals received 1500R of whole body X-radiation 6 hours before killing. The X-radiation produces a significant reduction in the rate of incorporation of [1*C]orotic acid into the nuclear RNA fractions of highest molecular weights. From Uchiyama et al.(3164.

-

190

D. T. KANAZIR

Large changes in the permeability of cell membraries to iiucleic acid precursors may be excluded. The incorporation rate of [‘4C](JrOtatCinto acid-soluble fractions and the specific radioactivities of “pool” precursors in the regenerating livers of normal and irradiated rats were almost identical (316). This also suggests that the pool of precursors is not diluted by unlabeled products derived from a breakdown of nucleic acids. The use of the RNase inhibitor, bentonite, during the extractions and the fact that no alterations in the UV profiles of RNA after irradiation were revealed (83, ,316) seem sufficient to exclude the possibility that the decrease in labeling of RNA is due to the action of latent RNase. These results therefore suggest that radiation does not affect the pool of RNA precursors or the activity of preformed enzymes involved in RNA synthesis (316, 316aJ$486823). The reduction of incorporation of orotic acid into pulselabeled RNA indicates rather that radiation interferes with the processes of DNA transcription and that the critical injury responsible for the depression and/or inhibition of nuclear RNA synthesis involves DNA, as is the case with microorganisms or in in vitro systems. Since the liver in regeneration contains a population of rapidly growing and proliferating cells that seem to be more sensitive to radiation, one can raise the question whether the irradiation impairs the function of resting liver composed of highly differentiated, mitotically inactive hepatocytes. An extensive study on the effects of radiation on different species of RNA and ribosomes in resting liver is in progress in our laboratory. PetroviE etal. (318) have demonstrated a stimulation in the labeling of the RNA of resting R). The rats rat liver induced by lethal total-body irradiation (800-1200 were irradiated with 850 R (119R f 9 R/min) and sacrificed a t 4 and 20 hours after irradiation and RNA metabolism was followed with [6-14C]oroti~ acid. The nuclei were separated from cytoplasm by the method of Chauveau (see ref. 318)the RNA from nuclei and cytoplasm was fractionated, and the specific activities of the pyrimidines of each RNA species was determined. The results are summarized in Table I1and show that the specific activities of pyrimidines as well as their ratio were marlredly altered in the irradiated animals. The type of change observed varies from one fraction to another and the extent of the alterations depends on the time following irradiation. Four hours after irradiation, the most important change in labeling of cytidine and uridine is detected in the NI fraction (probably containing mRNA). The uridine has a higher specific radioactivity than uridine from the N1 fraction of normal liver. The labeling of cytidine in the nuclear RNA fractions is significantly decreased, resulting in the increase of the ratio of U to C compared with that of the same RNA fraction from livers of nonirradiated animals. Within the same time interval (4hours) after irradiation, two types of

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I N DNA

TABLE I1 EFFECTSOF IRRADIATION ON RNA METABOLISM I N RAT LIVER^.^

Mi

Experimen t b Controls CP UP *P UP/CP UP/*P Irradiated 4 hours (1) CP UP \I.P UPhP UP/*P Irradiated 4 hours (2) CP UP *P UP/CP UP/*P Irradiated 20 hours CP UP *P UPKP UP/*P Irradiated 20 hours DNA CP UP *P UPKP UP/*P

+

Mz

S

NI

Nz

Na

2520 2520 4740 5410 5340 10500 - _ 5240 2.15 2.12 2.22 1.02 1.01 1 . 9 8

3330 22500 18600 6180 41600 32500 5290 1.75 1.85 1.86 1.17 -

c. 30000

Y34 0x3 2340 3100 2'275 13720 - _ 4240 3 32 3 59 5 87 0 78 0 57 3 40

1530 9600 6900 7550 51900 30400 3990 4.93 5 41 4 41 1 89 -

12900 49600 3 41

1330 1360 3320 3625 4150 20000 _ _ 4800 2.72 3 07 6.02 0.74 0.55 4 . 0 9

1710 8640 4880 5.05 1.77

462 1820 3.94 0.87

534 2140 2180 4.01 1.02

1370 6400 2680 4.67 3.05

974 7910 3920 29200 2100 4.03 3.70 1.87

1280 3280 2.56 1.07

1190 2920 2910 2.45 0.95

3GOO 7250 3460 2.02 2.36

2560 12700 8440 4680 23300 13050 3070 1.83 1.52

c. 40000

c.

1.4 -

-

No nuclei

-

-

-

-

12700 16800 -

-

From Petrovib et al.(318). The rats were exposed to total-body irradiation (850 R, 100% LDPo). The animals were sacrificed 4 and 20 hours after irradiation and 3.5 hours after injection of [14C]orotic acid (10.0 mCi/mmole). The livers were prefused in situ, pooled, washed, and homogenized. The nuclei were purified, and RNA fractions from nuclei and cytoplasm isolated as described in ref. 318. The symbols denote the origin of RNA fractions: NI,N,,Ns = nuclear RNA fractions; S = tRNA; MI and M2 = microsomal RNA fractions; Ms = postmicrosoma1 RNA fraction. The various RNA's were digested (0.30 N KOH, 19-20 hours at 37°C). The resulting mononucleotides were separated on Dowex 1 columns. The radioactivity of RNA pyrimidines was determined in a gas-flow (Nuclear-Chicago) counter. The specific activities of RNA pyrimidines are expressed as cpm/pmole of compound. The symbols Cp, Up, $p denote the mixtures of the 2'- and 3'-monophosphates of cytidine, uridine, and 5-ribosyluracil, respectively (SIB).

192

D. T. KANAZIR

changes were observed in cytoplasmic RNA fractions. The labeling of C was decreased in all cytoplasmic fractions as well as in the nuclear RNA, whereas that of U was slightly increased in Ma (postmicrosomal RNA fractions) and S (soluble RNA fractions), but significantly depressed in the MI and Mz fractions (microsomal and ribosomal RNA fractions containing about 85y0 of total cellular RNA). In all RNA fractions obtained 20 hours after irradiation, the specific activities of both C and U were Significantly decreased, but the values of TJ/C stilI remained very high.

(C)

(D)

FIG. 22.Effects of lethal X-irradmtion on the microsomal ItNA iuresting rat liver. Total-body X-irradiation was performed Rats were exposed to 850 R (100% LD,,-3,). in a Siemens-Stabilipan unit. The animals were sacrificed 2 hours after irradiation and iiijection of [l‘C]orotic acid. Liver microsomes were prepared as described by PetroviO etal.(391). Graphs (A-D)represent sedimentation patterns of Iabeled RNA s isolated from the “membrane” fraction and detached ribosomes of rat liver microsomes after 2 hours of in viuo labeling. Graphs (A)and (C) illustrate the sedimentation patterns of “membrane’] RNA from control and irradiated animals, respectively. Graphs (B) and (D) demonstrate the sedimentation profiles of RNA’s isolated from pyrophosphatereleased ribosomal subunits of control (B) and irradiated (D) rat liver. Sedimentation was performed in swrose grtitlients 5 4 0 % (w/v) by centrifugation for 3 hours a t 39,000 rpm, SW 39 rotor, in a preparrttive Spinc-o centrifuge, Model 1,. [H. I’etrovi6, personal comniunicat,it)ii(l%8)].

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FIG.23.Elution profiles ofnuclear RNA s from control and irradiated rat livers. The animals were irradiated and sacrificed under the same conditions as described for Fig. 22.The nuclei were isolated from livers as described by PetroviE (see ref. 318).The RNA fractions were fractionated on MAK-columns. Numbers over bars indicate NaCl molarity used for discontinuous elution of RNA fractions. The symbol D denotes tenaciously bound RNA eluted by dodecyl sulfate. The graph on the left represents elution pattern of nuclear RNA from liver of control animals. The graph on right illustrates elution profiles of nuclear RNA extracted from liver nnclei of irradiated animals. IS. PetroviE, personal communication (1968)j.

The results suggest that the changes in the biosynthesis and composition of various RNA species take place in the resting liver very early after irradiation though this organ is considered to be radioresistant. A high stimulation of RNA synthesis, with relative specific activities ranging between 150and 180% of the ControIs, is observed 3 hours after radiation and injection of the precursors in a RNA fraction strongly bound to microsomal membranes (Fig. 22,A, C). This fraction contains a significant portion of rapidly labeled RNA of the Iiver cell cytoplasm (320,321). At the same time, the increase in relative s p e c ~ cactivities of the nuclear RNA’s, including the “tenaciously bound” RNA is increased two- to threefold (Fig. 23).At the longer time intervals, progressively more radioactive surpIus RNA is found associated with cytoplasmic ribosomes. Thus, at the fifth hour, the relative specific activities of either 28 S or 18 S ribosomal RNA are some 50-700/, higher than those of corresponding controls (Fig. 22,C, D). The relative specific activities of “uniformly” labeled ribosomal RNA’s 24 hours after irradiation still exceed those of controls by 3040%. However, the increase in labeling of cytoplasmic RNA s never reaches the vdues observed for nuclear RNA’s. This fact suggests that most of the “surplus” nuclear RNA synthesized after irradiation is nevertransferred to the cytoplasm. Such nucleus-restricted

194

D. T. KANAZIR

RNA exists in a variety of animal cell nuclei (322). On the basis of our observations, the hypothesis may be advanced that X-irradiation stimulates the production of some RNA fractions in resting liver. A similar increase in transcription was observed in our experiments with livers from starved, fed, and adrenalectomized animals. Therefore it is unlikely that this increase represents an adrenal steroid-mediated change. Whether the surplus RNA that reaches the cytoplasm is physiologically useful and becomes translated is an open question. In favor of its translation, one may quote findings on the stimulation of protein formation in guinea pig liver ,stimulated aggregation of ribosomes and following X-irradiation (323,324 increased formation of polysomes in liver after irradiation, and also an enhanced uptake of amino acids by ribosomal oligopeptides inv i k owith That the RNA which reaches the ribosomes from irradiated liver (323,324), cytoplasm is functional in a certain sense may also be deduced from the fact that it persists undegraded for rather long intervals (24hours or more). Embryonic liver also provides a suitable model system for the studies of radiation action on the development of organs, i.e., on the macromolecular events taking place in the course of organogenesis. During the late period of fetal development, the mass of the liver tissues is tripled as the result Therefore, embryonic liver development mainly of cell proliferation (326). off ers experimental advantages at the cellular and the molecular level. During day 15of embryogenesis, new RNA molecules seem to appear in the liver since at that time the levels of many glycolytic enzymes of the liver are dramatically increased and a build-up of glycogen and fat commences (327). Information concerning radiation action on the embryogenesis is very scarce. Kafiani etal.(328, 329)studied the effects of radiation upon the fossiZis) synthesis of mRNA at the early stages of loach (Misprnus embryogenesis. The synthesis of mRNA species, their functions and quality change in the course of embryogenesis in a manner depending on the developmental stage. It is widely accepted that, up to the stage of gastrulation, there is no synthesis of new mRNA s. Beginning with gastrulation and during subsequent stages, various genes seem to become activated (“switched on”). Therefore the pool of the RNA molecules present in the cells at a given embryonic stage is representative of the phenotypic expression of gene activities regardless of whether or not they terminate in recognizable protein molecules. The authors have shown that in embryos of Misgurmus fossilis, the synthesis of RNA during the first 6 hours is very slow, but thereafter there is a dramatic enhancement. At that time the nuclei also become activated as their morphogenic functions commence. From hybridization experiments, it is also demonstrated that all the RNA synthesized during this first stage ofembryogenesis, up to the mid-gastrula

RADIATION-INDUCED

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stage, represents mRNA species. A dose of 40 kR was applied to inactivate the morphogenic function of nuclei without apparent damage to the cytoplasm. When fertilized eggs were irradiated 30 minutes after fertilization, the synthesis of presumable early mRNA’s seemed to be suppressed with consequent inhibition and elimination of the late stages of morphogenesis. Although this inhibition of &NA synthesis after irradiation was not complete, morphogenesis was completely arrested. Embryos irradiated at later stages (5-9hours, bIastula), moruIa and Iater, gradually and partidly recover their synthesizing activity, but only after a certain period of inhibition. Even with very high doses of radiation that damage the nuclei recovery of RNA synthesis was observed. Consequently the question was raised as to what may serve after irradiation as a template for this newly synthesized RNA, since the nuclear DNA was completely degraded. Cytoplasmic DNA may have served as a template for the synthesis of RNA although a repair of nuclear DNA damage could not be excluded. It was concluded that the radiation-induced block of embryogenesis was due to the inhibition of mRNA synthesis. The effect may be attributed either to quantitative changes in the amount of mRNA or to changes in its quality. The authors are inclined to believe that the change in quantity of mRNA is of primary importance for the processes of differentiation and embryogenesis since a similar block in embryogenesis may be produced by actinomycin D, which presumably affects only the rate of transcription but not the structure of the transcribed RNA. Furthermore, inhibition of embryogenesis may be due to faulty “switching on” and ‘(off” of coordinated genes resulting from radiation action on the regulatory machinery in the embryo. We extended our studies to the embryonic liver. As already mentioned, the day 15 of embryogenesis of rats is a critical period for organogenesis, growth, and metabolic functions. In preliminary work, SimiC et al.(SSO) observed that when embryos were irradiated on day 9 of embryogenesis, i .e., during the period of differentiation and very early organogenesis, and if the growth rate of liver was followed to between day 15and day 21,then the rate of synthesis of liver nucleic acid and protein was significantly depressed. Therefore, the objective of our studies was to follow the rate of synthesis of pulse-labeled RNA in embryonic liver and to analyze its composition under experimental conditions in which embryos were irradiated at day 9 of embryogenesis whereas RNA was analyzed at day 15 of embryogenesis, during intensive organogenesis. Rat embryos were irradiated on day 9 of embryogenesis with 100 R of X-rays either by exposing the pregnant mother to total body irradiation or by local irradiation of embryos in only one uterine horn. The RNA was “pulsed” with y2P, extracted by the method of Scherrer and Darnell ( S S I )and , analyzed on sucrose density

196

D. T. KANAZIR

B

A 26 S

1

90 A

70

50

.

X

x Y

30

L 20 30 10

Fractions

0

10

10

20 30 Fractions

FIQ.24. The sedimentation pattern of pulse-labeled ribonucleic acid (RNA)from liven of rat fetuses. The embryos were irradiated together with their mothers; the labeling of RNA with phosphorus-32 lasted for30 min. Gradient: 20% to 4% sucrose; 5 X 104 M magnesium ions; 4 hours of centrifugation at 39,000 rpm. (A)Control; (B) RNA from liver of irradiated fetuses. From Simi6 etal.($30).

B

A

26 S

1

0.121

10

20

1

30

Fractions

t . . . 10

20

30

Fractions

FIQ.25. The sedimentation pattern of pulse-labeled ribonucleic acid (RNA)from liver of rat fetuses. The embryos were locally irradiated; the labeling with phosphorus-32 4 hours lasted for 40 min. Gradient: 20% to 4% sucrose, 5 X 10- M magnesium ions; of centrifugation at 39,000rpm. (A) Control; (8)RN4 fromlwally irradiated liver fetuses. From SimiC et al.($30).

RADIATION-INDUCED

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gradients. The nucleotidc coniposit,ion of the pulse-la1)cled RNA was dctermined. In the experiments in whic.h the 9-day-old embryos were irradiated by exposing their mothers to radiation, the specific radioactivity of the pulse-labeled RNA extracted from the livers of the nonirradiated fetuses at day 15was 9600 cpm per milligram of RNA, whereas that from livers of irradiated fetuses was 22,000 cpm per milligram of RNA. Therefore, a stimulation of the RNA synthesis was observed 6 days after irradiation. Sucrose density gradients revealed also a faster incorporation of 32Pin all Thus the liver RNA fractions of the irradiated fetuses (Figs. 24 and 25). heavier RNA fractions (45-35 S) mcre more labeled in the liver of irradiated than in nonirradiated fetuses. When the effects of radiation transmitted from mothers to the liver RNA metabolism of their fetuses was bypassed by restricting the radiation to one uterine horn, enabling us to obtain from the same mother both irradiated and nonirradiated litters, again the heavier 45-35S RNA fractions in irradiated fetuses were significantly more labeled than in nonirradiated fetuses. On the other hand, the RNA fractions between 4 and 16S, presumably mRNA fractions, were more labeled in nonirradiated controls than in the irradiated fetuses (Fig. 25,A, B). This could mean that the conversion of pulse-labeled RNA into functional mRNA and stable rRNA fractions is altered by irradiation. The molecular events related to this transition are still unknown. Keeping in mind the fact that structural or configurational change may influence the rate of conversion of precursor RNA to functionally stable rRNA or mRNA, we determined the base composition of the pulse-labeled RNA. An alteration in nucleotide composition, as judged by the distribution of 32Pamong nucleotides in the pulse-labeled RNA, is manifested in the pulse-labeled RNA from liver of irradiation fetuses through an increase in the contents of cytidylic and guanylic acids, whereas the contents of adenylic and uridylic acids decreased compared with the control values. Since pulse-labeled RNA is composed of mRNA and precursors to ribosomal RNA, we still do not know which of the fractions is altered in composition. In any case, this phenomenon is very similar to those described by Petrovii: (318) in the resting liver and that described by Hudnik-Plevnik in bacteria (278). It is of interest to note also that Matsudaira et al.(306)observed a similar phenomenon in tumor cells of rats after X-irradiation. Yoshida =cites hepatoma cells (AH 130)were inoculated into the peritoneal cavity of the rat. On the fifth day after inoculation, one group of animals received whole-body irradiation (1000 R), the RNA of the tumorlcells was “pulsed” a t various times after irradiation, and the RNA was extracted and analyzed. The specific activities of the RNA fractions extracted by cold phenol and hot dodecyl sulfate-phenol were increased in dl but a few samples. This increase occurred at as early as 30 minutes and seemed to

198

D. T. KANAZIR

continue for about 20 hours after irradiation. The pool of precursors was slightly increased. This finding may indicate that possibly radiationinduced changes in cell permeability played no significant role in the impairment of RNA metabolism. Sucrose gradient analysis of the coldphenol RNA revealed that irradiation caused some increase in ~ridine-~H incorporation into RNA fraction although no change was found in the UV absorption profiles. The fractions extracted by hot phenol plus dodecyl sulfate revealed that RNA fractions heavier than 28 S RNA had been formed; these represented a major portion of the incorporated activity. Irradiation produced almost no change in UV-absorption profiles. Thus even in tumor cells an increased incorporation of [3H]uridinewas observed especially into peaks of the RNA of high molecular weight. The labeling of these cells is very similar to that observed in the RNA of fetal liver (330) or that of bacteria (268, 278). Barnabei et al.(332) also observed, in rat liver nuclei, an increase in RNA synthesis and in RNA polymerase activity after X-irradiation (600R), but a less obvious response in irradiated liver to cortisone. Similar results were obtained by Hidvdgi etal.(323) in their studies on RNA synthesis in guinea pig liver after X-irradiation. Lncorporation of [14C]orotateinto the microsomal and ribosomal RNA was almost doubled between 6 and 9 hours after irradiation with 2000 R i n uivo, and incorporation of the precursor into the rapidly labeled RNA of the nuclei continued with increased intensity for 6 hours after irradiation. The incorporation of the labeled precursor represents net synthesis since a 2040% increase of the RNA content was observed in the cell fractions between 12 and 24 hours. The authors claim that irradiation increases the DNA-dependent RNA synthesis since actinomycin D administered in uivo prior to irradiation inhibited the radiation-induced increase in the RNA synthesis. This may indicate that radiation induces a more rapid transcription ofactivated (“switched on”) genes. This actually appears to be the case since the increased RNA synthesis observed after X-irradiation was followed by an increase of tyrosine transaminase activity (332). Furthermore, Vartdresz et al.(3.24) reported that X-irradiation induces an increase in protein synthesis in the liver. The increase in protein synthesis appears to be due to an enrichment of polysomes, which seems to take place in liver after irradiation, the ribosomal fraction isolated from livers of unirradiated guinea pigs containing mostly monomers (50-60%). An increase of polysomes was observed in liver 2 hours after irradiation, and between 12 and 15 hours after irradiation the increase becomes even more evident. The enrichment in polysomes cannot be due to an artificial aggregation of ribosomes since they were as readily degraded as polysomes from normal liver by either endogenous or exogenous ribonuclease digestion. Parallel to that enrichment, the ribosomes display a higher amino acid incorporating ability whether the ribosomes were isolated from

RADIATION-INDUCED

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199

the mitochondria1 supernatant or from the microsomal fraction with deoxycholate. The result raises the question whether or not the phenomenon exists in vivo or is an artifact occurring in the course of isolation of ribosomes and polysomes. Two approaches have been made in order to solve this problem. First, 14C-labeled amino acids were injected shortly (60 minutes) after irradiation (1800R) and animals were sacrificed 11 hours later. More radioactivity was found in the polysomes isolated (in the presence of bentonite) from the livers of X-irradiated animals than in those prepared from livers of normal animals. The second approach, using poly U as an artificial messenger, demonstrat,ecl that, the irradiated microsome fraction contains more saturated endogenous messeriger ltNA than does t,he 11011irradiated one thus confirming the phenomenon. also reported that whole-body X-irradiation Cammarano elal.(333) (250, 550,and 850rads) elicits marked changes in the rat liver polysome organization. At 3,6,12,and 24 hours after irradiation, the liver ribosome population is characterized by a marked shift from monomers into the heavier polysomes. Sucrose density gradient centrifugation of the postmitochondrial fraction (without prior detergent t,reatment) revealed an enrichment in polysomes and a decrease in monosomes after irradiation. The relative amount of ribosomal subunits (45S and 60 S) remains essentially unchanged after irradiation. Pulse labeling (40 minutes, [14C]orotic acid) showed that the specific activity of the 18 S RNA derived from 45 S subunits is unchanged, suggesting that the radiation does not interfere with the rate at which ribosomal RNA is labeled (synthesized) and transferred to the cytoplasm. Unfortunately the authors did not present data on the labeling of 28 S RNA from 60 S ribosomal subunits, which could have been of interest since the results on irradiated bacteria indicate strongly that the conversion of precursor ribosomal RNA to 28 S stable RNA may be significantly affected by radiation. Furthermore, bilateral adrenalectomy failed to prevent the radiation induced shift from monomers into polysomes (as well as the increase in the heavy polysome fraction), so it cannot be argued that the observed effect is mediated through stimulation of adrenal activity that would, in turn, stimulate transcription processes in the liver. In our laboratory, Bebarevii: etal.(334) also observed in rat liver a marked shift from monosomes into heavy polysomes (Fig. 26,A-D) 24 (R), 48 (C) and 72 (D) hours after whole-body irradiation (800 R). Although no precise conclusion can be drawn at the moment on the mechanism responsible for the enrichment of polysomes in the liver after irradiation, it may be assumed, at least as L: working hypothesis, that irradiation enhances transcription (synthesis) of cither mRNA or precursor ribosomal RNA and that their transfer from the nucleus to the cytoplasm is enhanced. In connection with the observed polysome enrichment, one may assume

200

D. T. KANAZIR

also that radiation may esert an injurious effect on the membranes by reducing either their abilit,y to bind ribosomes or by producing a release of the bound ribosomes from the membrane which may form polysomes. Although many effects of radiation on cells appear t o have their origin in the nucleus, one cannot yet exclude the role that the cytoplasm may play in the production of radiation damage in mammalian cells and organs, since so little is known about radiation effects on the mechanisms control-

20 40 NUMBER OF FRACTIONS

FIU.26.Sedimentation profiles of ribosomes prepared from livers of unirradiated The rats were exposed to total-body X-irradiation and total-body irradiated rats. (850 R). At different time intervals after irradiation (24,48, and 72hours) the animals were sacrificed2.5hours after injection ofIW]orotic acid (20 mCi/mmole) and ribosomes were prepared from livers as follows. Ribosomes were released by treatment of the postmitochondrial supernatant with sodium deoxycholate ( h a 1 concentration 1.3%), and purified in a discontinuoussucrose gradient by centrifugation for 6 hours at 25,000 X g in a Spin00 ultracentrifuge. The purified ribosomes were suspended in 0.025M H, 0.001 M Mgp+, 0.1 M Trk-HC1,pH 7.4. Of this ribosome suspension, 0.3ml was layered on the sucrose gradients and centrifuged for 4 hours a t 39,000rpm in a SW 39rotor of a Spincocentrifuge. The graphs illustrate the profiles obtained for the radioactivity . and the . optical ) density at 260m p (0-0-0) foreach fraction collected (cpm, . after centrifugation of ribosomes in a sucrose gradient. A, Unirradiated control a n b l s ; B, irradiated animals sacrified 24hours after irradiation; C, irradiated animals sacrificed 48 hours after irradiation; D, irradiated animals sacrificed 72 hours after irradiation (334).

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201

IN DNA

ling cytoplasmic activity. Little is known about radiation effects OH nlitochondria and on their function, on the endoplasmic reticulum, or on lysosomes, ribosomes, soluble and bound enzymes, or on tRNA and 5 S RNA. It is very likely that radiation may. aIter the structure of ribosomes and polysomes directly or indirectly (via free radicals peroxides and (335) (Table 111)have shown that ribosome relensed enzymes). Metlag etal. preparation from the livers of irradiated animals (850 R), used in a poly U directed protein synthesizing system, exhibited a larger incorporation of phenylalanine (3040%)t,han did the ribosomes of nonirradiated animals during an incubation interval of 30 minutes. This may be attributed to a 03

D

7 2 hrs

500

01

#

48 hrs

C

03 500

\

N W

v)

t

W

p

._ E

0

01

D

03 500

0.1

20 4'0 NUMBER OF FHACTIONS

FIG.26 R, C, and D.

202

D . T. KANAZIH

TABLE: I11 EFFECTS OF in VivoX-IRRADIATION ON THE POLYPHENYLALANINE SYNTHESIZING ACTIVITY OF RAT LIVER RIBOSOMES~-~ Period after X-irradiation ~

Specific activity

~

~

Unirradiated 6 hours 24 hours ~

~~

Increased activity

6240 7189 9084 ~

0 15 32

~~

From Metla; et al.($36). * Reaction mixture: 172pmoles sucrose, 35 pmoles Tris-HC1 buffer; pH 7.6; 28pniole NH4Cl; 1 pmole KCl,5.7prnoles Mg acetate; 3 pmoles 0-mercaptoethanol; 0.48pmole ATP; 0.12 pmole GTP; 2.5pmoles phosphoenolpyruvate; 28 pg phosphoeiiolpyruvate kinase; 100pg polyU; 1.0mg pH 5 enzyme-protein, 0.5mg ribosomal protein; 1 MCi [aHIphenylalanine (1.5Ci/mmole, Amersham), all in a total volume of 0.6ml. The pH 5enzymeswere prepared according to M. Rosenbaum and R. A. Brown [Anal. Biochem. 2,15 (196l)l. The ribosomes were prepared according to Y. Tashiro and P. Siekevitz [J. MoZ.B i d .11,149(1965)]. After a 30-minute incubation at 37°C the reactionwas stopped with 5% CClaCOOH. The precipitate was washed twice with cold 5% acid, heated in 5% acid in a water bath at 90 Cfor 30 minutes plated on membrane filters, and counted in a nuclear-Chicago liquid scintillation system, Mark I. Corrections were made for endogenous messenger incorporation (536). a

direct or an indirect effect of radiation on preformed ribosomes. It seems that radiation may increase the number of ribosome binding sites. The mechanism leading to this increase is not known. In any case, the sucrose density gradient analyses of ribosomes or of ribosomal subunits were not synappreciably different. The normal ribosomal function in an in vitro thesizing system wshs apparently resumed around 72hours after irradiation, marked differences in their synthesizing properties being observed only up to 24 hours after irradiation. The fact that this change was observed only when ribosomes were prepared from microsomes with detergent treatment, but not if they were prepared from a post-mitochondria1 fraction suggests that the observed change may probably be attributed to tlhe action of some enzyme or enzymes released from tlhe microsonial membranes during the ribosome preparation. synthesizing system was Similar changes were observed when an in vitro found that the ability irradiated with y-rays. Maxwell and Nutwell (336) ofribosomes to synthesize protein in an invitro system is approximately a log function of the dose; 300,000 R produced no discernible change in bind 3&40% sucrose gradient profiles. Ribosomes so irradiated in vitro more [14C]polyU than uilirradiated systems. Irradiated ribosomes successfully compet,e with unirradint,ed ribosome in thc binding of poly IJ, but

RADIATION-INDUCED

ALTERATION

203

IN DNA

direct little protein synthesis. The results indicate no release of latent ribonuclease activity after such a high dose of radiation. These results suggest that some cytoplasmic factors reIated to the function of ribosomes and microsomes may be involved in the impairment of ribosome functions related to the processes of translation. Since tRNA is also involved in the translation processes an attempt has been made in our laboratory to study the effects of radiation on the acceptor capacity of tRNA, using resting liver as a system. Sein et aE. (337), using the total tRNA from the liver of irradiated animals, obtained results indicating that the phenyldanine acceptor activity decreased 1 hour after 850 R of total body X-irradiation (see Table IV). The tRNA’s isolated from livers 24,48, and 72hours after irradiation showed slight decreases in the acceptor activity for phenylalanine. In all the above studies the aminoacyl-tRNA synthetases from livers of nonirradiated rats were used. A significant decrease in the acceptor capacity of tRNA from liver was TABLE IV EFFECTS OF in Vivo X-IRRADIATION ON THE AMINO ACID ACCEPTOR ACTIVITYOF RAT LIVERtRNAasb Counts per minutee System

Control

Irradiated

Experiment IC [ 14C]Phe

[ I4C]Leu

Protein Hyd 1-01 yaa t e- ‘4C

Complete - CTP Complete - CTP Complei e - CTP

853 805 4,354 4,454 18,950 19,021

507 405 4,114 4,018 18,660 19,000

Experiment 2 [ WIPlie

[“CIIRU Protein Hydr~lyeate-~~C

Complete - CTP Complete - CTP Complete - CTP

677 691 3,449 3,121 14,458 14,678

350 378 3,019 3,005 14,873 15,438

From Sein etal.( S S 7 ) . The reaction mixture consisted of the following: 0.1 M Tris-HC1 buffer pH 7.4; 0.05 M KCI; 0.01 M MgAcr, 0.01 M ATP; 0.005 M CTP; 0.5 mg t,R.NA, 2 mg enzyme protein and [I4C]aminoacid all in a total volume of 1 ml. The [14C]aminoacids used were [14CC]leucine(8 mCi/mmole), [Wlphenylalanine (3.55 mCi/mmole), and [14C]protein hydrolyaate (600 pCi/mg) (Radiochemical Center, Amersham, England). Counts per minute of acid-precipitable material. Corrections were made for blanks in every experiment (3.37).

204

D. T. KANAZIR

found 24 hours after very high tloses (2500 R). This decrease was accompanied by changes in the melting profilc of bulk tRNA. The T, of bulk tRNA from irradiated livers seems to be shifted upward. This change may mean that some slight changes in secondary structure of tRNA in T,,, occur after irradiation and modify its function. B% These results suggest strongly that cytoplasmic factors may influence the rate of translation, which in turn may influence the processes of transcription and probably those of repair. Recently it has been found that the cytoplasm can exercise through some still unknown factors a positive action by stimulating both DNA and RNA synthesis in inactive nuclei of The chemical nature and mode of avian and mammalian origin (338). action on DNA replication and transcription of this factor(s) remains unknown.

VI. Biological Consequences Resulting from Radiation-Induced Damage to DNA It is not the purpose of this section to discuss the radiation-induced mutagenesis and the mechanisms of mutation expression. These subjects 93,9.6, All that can be are dealt with in recent review articles (23, done here is give a brief summary of recent observations on the effects of radiation upon genetic expression in highly differentiated mammalian tissues, keeping in mind the complexity of the system involved. Genetic expression in higher organisms is also mediated through transcription resulting in translation, i.e., in the synthesis of new protein. But, in addition to the external environmental influences, these processes are also subject to the internal neurohormonal control present in the multicellular organisms, and to the control of some factors capable of stimulating DNA replication and transcription present in the cytoplasm (338). It appears that the mechanisms concerned with the external environment, such as those expressed in hepatic tissue, are somewhat similar to those seen in microorganisms, including substrate induction and product The internal control of genetic expression may be mediated repression (339). through hormones influenchg the rate of genetic transcription and translation. One of the highly differentiated model systems, best studied i n vivo, is the regenerating and/or resting liver discussed above. It has been postulated that the rise in enzyme activity following partial hepatectomy depends on transcription, mRNA formation, and an increased synthesis of the enzyme proteins. Although the precise effect of radiation on the mammalian cellular enzyme systems is not as well defined as in bacteria, it is neverthe‘See articles by Pullman and Pullmttn and by Singer and Fraenkel-Conrat in this volume.

RADIATION-INDUCED

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IN DNA

205

less known that intracellular enzymes present a t time of radiation are not inhibited by doses that block their induction (251-256). Recent results (86,237-240,301-303, 516,317, 339-342) support this view. Radiation does not affect the activity of kinascs already synthesized, but even low doses prevent or delay biosynthesis of the enzyme. As already mentioned, irradiation soon after partial hepatectomy completely suppresses DNA replication, cellular multiplication (regeneration), and synthesis of induced enzymes. Here both functions of DNA, i.e., replication and transcription, seem to be impaired, but if irradiation is not applied until DNA replication has been initiated, DNA synthesis and liver regeneration occur relatively normally. In the resting liver, radiation damage in DNA impairs the processes of genetic transcription as followed by the synthesis of induced enzymes. The liver enzymes serine dehydrase and ornithine transaminase @do), which may be induced by the administration of the corresponding amino acids, are affected by whole-body radiation. A y-ray dose of 400 R to the whole organism inhibits enzyme induction by more than 60%. Initially, the induction of both enzymes is sensitive to the effects of actinomycin. This suggests that irradiation interferes with genetic transcription, i.e., with the The enzymes responsible elaboration of mRNA in the resting liver (840). for the catabolism of many foreign compounds such as cholinergic organic phosphate (341), are located in the liver, and they develop after the birth of animals reaching the adult level at about 6 weeks of age. The sublethal doses of X-irradiation cause a marked inhibition of development of this system. Doses as low as 100 R inhibit normal development of the liver enzymes that catalyze desulfuration of phosphorothioates to their corresponding oxygen analogs (341). Exposure of adult rats to X-irradiation has no effects on the activity. This suggests that radiation may exert its action on the formation and/or development of the enzyme-synthesizing system, i.e., on genetic expression in nondividing cells of the resting liver. In rat brain there are several isoenzymes of glycerophosphate dehydrogenase. Form I11 seems to be predominant in the mature brain, its development occurring between 18 and 45 days postpartum. The rise in activity occurs when cell division has ceased. Here again it is possible to study the effect of ionizing radiation on enzyme synthesis in the absence of cell division. Administration of 750 R to the head of 20day-old rats inhibited as much as 50% of the development of this form of the enzyme. This inhibition of a rise in activity is a delayed phenomenon probably due to the presence of long-lived mRNR is suggested by studies with actinomycin D, but no final recovery from the inhibition was observed. A more immediate phenomenon with no recovery was a 10% degradation of the DNA that takes place during the first 24 hours after-irradiation. The fully

206

D. T. KANAZIR

developed enzyme (form 111) was not affected by the irradiation. These results suggest an X-ray-induced nonreparable inhibition of the genetic transcription for enzyme I11 in nondividing brain cells (342). In sum (see Fig. 27),these observations suggest that the biological consequences of DNA damage in dividing cells is cell inactivation and, probably, reproductive cell death. In nondividing cells, the consequences of DNA damage are impaired genetic transcription and translation, leading to the suppression of synthesis of enzymes or of other specific proteins, causing altered metabolic functions that may in turn lead to shortening of the life span (aging), to malignancy, or to an inability of the organism to adapt itself to external environment. The “molecular genetic’’ theories of aging have proposed that aging of multicellular organisms is probably due to the increasing deterioration and ultimate failure of nonreplicating cells and systems. The rate of failure of such cells is also genetically programmed and is analogous to the DNA-programmed ontogenic sequence of early differentiation and growth (343, 344). Age-related changes in such organs as the brain, which contains nonreplicating cells, result primarily from a progressive accumulation of defects in the DNA. Ionizing radiation may increase the incidence of DNA defects and simulate some of the more specific processes of aging in cells as well as accelerate many of the more obvious structural and functional changes that normally accompany the Decreased life span after irradiation suggests an processes of aging (944). important role for DNA connected with somatic mutations. Consequently, mortality, longevity, and the rate of aging are completely genetically programmed and determined by the DNA structure at the time of fertilization. In addition, there is no satisfactory theory explaining the induction of carcinogenesis by radiation; ideas are numerous, often conflicting, and quite incapable of explaining clinical or experimental findings in detail. It is commonly believed that two steps or phases are involved; these are often referred t o as “initiation” and “promotion.” “Initiation” is thought of as sudden and irreversible, independent of time and dose, and envisaged as a “genetic change” in an individual cell, nowadays normally seen in terms of modifications of DNA structure. “Promotion” means a slow evolution, an expression of genetic damage demanding considerable time (346). Consequently, physicochemical events induced by irradiation that result in DNA structural damage may, unless repaired affect both of the main functions of DNA, replication and transcription. The biological consequences of DNA damage may be expressed as an inactivation of cell functions (cell death), as a variety of mutations and an increased frequency of their expression, as an alteration in the structure and function of enzymes resulting in loss of regulation of metabolic processes which again may cause

RADIATION-INDUCED

ALTERATION

IN DNA

207

a shortening of the life-span-aging, or finally, as cancer induction and, in a rather general way, a decrease in viability.

VII. A Working Hypothesis Today it is widely accepted that DNA is the carrier of genetic information in living systems, that genetic information is contained in the sequences of nucleotides comprising the DNA molecule, that the expression of the genetic information encoded in the DNA is achieved in the first instance by the synthesis of RNA whose nucleotide content and sequences are also prescribed by the DNA, that this RNA (messenger RNA) directs the synthesis of proteins by specifying one amino acid for each codon (triplet sequence), so that the sequence of amino acids in the protein molecule is determined by the sequences of codons in the mRNA’s determined by the original DNA (163, 346, 347). These concepts of molecular biology regarding the dominant function of DNA as the carrier of the genetic code have begun to influence and stimulate current research in the field of radiation biology. From these concepts it is evident that any damage produced by irradiation in the structure of the DNA may either (a)prevent replication, i.e., synthesis of new DNA, or (b) may be replicated as errors in the DNA structure of rapidly dividing cells and transferred to the progeny as definite changes in genetic constitution (Fig. 27).If this change occurs in a functional fragment of DNA, it will be transcribed by the RNA and transferred via translation processes as minute “errors” into the structure of the specific protein. In other words, the minute errors in the DNA structure may be phenotypically expressed by transcription and translation. A nonsense triplet in DNA transcribed to mRNA may cause premature termination of a peptide chain and result in the formation of nonfunctional protein (348). Radiation damage to DNA molecule may similarly be translated as changes in protein molecules, which under some physiological conditions may be lethal. It would appear, therefore, that studies on the incorporation of labeled amino acids into the nuclear and cytoplasmic proteins ofrapidly proliferating tissues after irradiation and their compositions (amino acid sequences) may well provide us with a clue as to a t least one of the possible type of codon alterations in the transcribing (functional) fragment of DNA. In this respect, the data of Van Lancker (348a,b) should be mentioned here. There is little effect of irradiation on the incorporation of [14C]leucineinto the proteins of regenerating liver, but there is a clear-cut inhibition of the incorporation of [‘4C]lysine.The low incorporation of [14C]lysineinto the

208

D. T. KANAZIR

Legend Filial generation

/

DNA’= RNA“= RS = INTER =

Expression M

--w--t Inhibition of the process

/

Damaged DNA Abnormal RNA Repair synthesis Interphase deoth

m

Reproductwe

f

Translation

f

mRNA*-Nonfunclionol

-

-

proteins +-Growl_h ribosomes

Chanse of metabolism-

w

h

h-o

t Fro.27.Diagrammatic representation of biological consequences resulting from radiation-induced damage to the DNA molecule (genome). Chemical alterations in the structure ofDNA may prevent, in dividing cells, DNA replication and cause immediate death (interphase death) or death after a few mitoses (reproductive). These alterations in the DNA structure block or induce abnormal DNA replication. Mutagenic alterations in the DNA structure consist of modifications of the bases, which do not prevent replication but produce “errors” in replication resulting in heritable changes in the base sequence of newly synthesized DNA. By this means, changes may be expressed as phenotypic (mutation), and appear either in germinal cells (progeny) or in somatic cells. The radiation-induced defects in the DNA structure may prevent DNA (genetic) transcription or may cause abnormal (incomplete) transcription. These alterations may give rise to changes in the metabolic functions of cells (organism), which in their turn may cause inhibition of growth or cell death. I n nondividing cells, radiation-induced damage to the DNA (genome) may prevent genetic transcription. This alteration may cause death. Transcription of the DNA may be incomplete (partial) and give rise to changes in the cell metabolism and thereby cause physiological and morphological alterations, cancer induction, and stimulation of the physiological processes of aging. Some of these changes in metabolism may also cause death. Radiation-induced damage to the DNA may be repaired by the repair mechanisms. This repair may result in normal replication and transcription of the DNA molecule, which can lead to normahation of the metabolic processes and assure the survival of the irradiated organisms.

RADIATION-INDUCED

ALTERATION

I N DNA

209

nuclear proteins after irradiation does not result from changes in the pool of amino acids, which could result from changes in permeability or a dilution of the lysine pool bg a degradation of proteins caused by radiation. This effect of X-radiation seems to result from an interference with the biosynthesis of newlysynthesized proteins, i.e., from damage to the structure of the corresponding DNA-codons. This damage may be expressed during translation, i.e., biosynthesis of newly-synthesized proteins. There are two codons for lysine, AAA and AAG, containing three and two adjacent adenine nucleotides. Lysine is the only amino acid whose codons are SO constituted. Consequently the scgment of the DNA molecule that codes the mRKA responsible for t,he biosynthesis of protcins rich in lysine must be rich in TT or TTT sequences. If such sequences were particularly sensitive to radiation or if radiation danixgc to such sequences were not rapidly repaired, the preferential interference of radiation with the incorporation of lysine could be better understood. Since the fragment of the DNA molecule that is damaged might be small, direct evidence for the alterations of TT sequences in codons after irradiation may not easily be detected by our methods. But if the low incorporation of [*4C]lysineinto proteins is due t o the effect of X-radiation on DNA structure, then it may be anticipated that this effect would be more evident in rapidly-proliferating tissues in which DNA replication and transcription are markedly more rapid than in resting tissues. The studies of Van Lancker (348a,b) on the incorporation of amino acids into the proteins of resting liver, kidney, brain, spleen, intestinal mucosa, and bone-marrow show that irradiation interferes with the incorporation of amino acid (lysine) in spleen, bone-marrow and intestinal mucosa, being most marked in the intestinal mucosa. No effect on the incorporation of [14C]lysineinto proteins of brain, resting liver and kidney was observed. However, it should be stressed that an alternative, very hypothetical explanation for the reported results might be offered, namely, that the changes in the incorporation might result from damage inflicted by X-radiation not t o the DNA but to the structure of the tRNA’s, which may play a role in the control of protein synthesis at the translation level. At any rate, it becomes evident that it is reasonable to assume that radiation damage to DNA may profoundly affect the mechanisms and fidelity of the repIication and transcription pattern of the DNA molecule. The biological consequences (Fig. 27)of the altered DNA replication and transcription are various; inhibition and/or delay of mitosis, mutations, loss of viability, temporary and/or lasting changes of biochemical processes and physiological functions, which in higher organisms may cause a radiation syndrome (mdi:ition sickiicss). This, fund:imentnl radiation biology is mpid1y becorning macromolccu 1: ~ r:idiohi r ology. I n view of all this, it is important t o iiridcrst~ntlthe physicochemical

210

D.

T. KANAZIR

nature of radiation damage to the DNA and the mechanisms by which DNA replication and transcription are affected by irradiation, because genetic stability and continuity depend on DNA replication and at least the first step in the expression of genetic information depends on transcription. It should be emphasized that we still do not know the intimate nature of DNA replication mechanisms. Jacob et al.(349) and Jacob (350) have proposed that replication of the bacterial chromosome DNA is initiated at a specific site, termed the replicator, attached at the mesosome and through it to the bacterial cell membrane. The attachment of the replicator to the cell membrane is an important part of the hypothesis as it provides a mechanism for the separation and segregation of the two daughter DNA molecules. Many reports give support to this coiicept (551, 362). Some suggest that DNA replication (synthesis) in the mammalian cell may also 354). In the model of the be initiated at the nuclear membrane (353, replicon proposed by Jacob et al.(349), the replicating site is fixed at the membrane and the DNA molecule passes through it as it is synthesized. Thus, both the initial and all subsequent replication steps should appear at the membrane. From this concept, one could assume that the initial lesion of ionizing radiation is equivalent to a break-up of the attachment of the DNA growing point (replicator) to the membrane. This may be followed by degradation of DNA, and by a drop in DNA transcription in that part of the genome (355). The radiation effects on the membrane may cause a disturbance in its permeability. The failure of transcription and leakage of nucleic acids precursors through the membrane may prevent synthesis of the cell wall and membrane constituents, which again may lead to cell death. This appears to be the case in radiosensitive organisms lacking repair enzymes, but in radioresistant cells the damage to DNA may be repaired and the structure of the cell wall and membrane restored. In such a case the cell is capable of surviving radiation damage. Although this assumption appears quite hypothetical, we still believe that the immediate and delayed radiation effects on the cell membrane, at least in bacteria, should be taken into consideration when discussing the primary site of radiation action. However, one should admit that a certain percentage of radiation-induced lethality may be attributed to “non-DNA’’ sites. But if DNA integrity and transcription are preserved and if DNA carries the repair information, then any damage on the rnembrane may be repaired and the cell should be capable of surviving otherwise lethal doses of radiation. Furthermore, the experimental evidence shows clearly that the biological consequence of radiation damage inflicted upon the DNA depends to a large extent on the genetic const,itutionof the living system at least in microorganisms (10-18). Thus, the quantity of radiation damage to tjhe DNA molecule of different, bacterial strains may

RADIATION-INDUCED

ALTERATION

IN DNA

211

be the same, but the let>halevciit. may not be the same. Therefore, large variations in sensitivity to ionizing radiation among different bacterial strains irradiated under the same conditions must be a result of differences in the DNA structure and of differences in the efficiency of the genedetermined multienzyme repair mechanisms. The existence of this DNA error-correcting-system” places a serious limitation on the use of the target theory in quantitating the cause-effect relationship and complicates the efforts to find an explanation of radiation action on living systems. Results consistent with the repair concept even in the cells of higher Repair may organisms can be provided from recent studies (226-224). be a universal phenomenon and of general biological significance in the maintenance of genetic stability. It should also be noted here that faulty transcription and translation may finally block repair synthesis and normal DNA synthesis due to a lack of repair enzymes and/or DNA polymerases. The importance of indirect radiation effects shouId also be taken into consideration. Irradiated media have an inhibiting effect on the synthesis principally from the of DNA, RNA, and protein in nonirradiated cells (go), 90).This must action of hydrogen peroxide and organic peroxides (19, mean that radiation-induced peroxides may act on DNA replication and transcription apart from the radiation itself. I n addition, hydrogen peroxide may destroy the capacity of mRNA to express itseIf. One may then ask which of these two classes of radiation action, direct or indirect, is the more important as a factor responsible for the disturbance of the DNA function. In early radiation biology there was no general agreement on the relative importance of direct and indirect action in living cells. The modification of radiation damage by oxygen or chemical protective agents has occasionally been interpreted as evidence in favor of the idea that the indirect action of radiation predominates. But to answer the question today, one must take into consideration the efficiency of mechanisms capable of repairing the damage to DNA induced by both direct and indirect actions of radiation. Radiation lethality is caused exclusively by the presence of unrepairecl defects in the DNA, regardless of their origin. Therefore, there is no necessity to cliscuss here the relative importance of direct and indirect radiation action on living systems. Instead, an oversimplified but nonetheless useful working hypothesis, which would unify all the available data in the frame of an overall picture on the radiation action on living systems, is presented. I n this working hypothesis, the attempt is made to summarize in a most general way the conclusions of many recent studies. The hypothesis is based on what seems to have become the central dogma of contemporary molecular radiation biology. (L

212

D. T. K A N A Z I R

The hypothesis assumes that the DNA macromolecule is the principal site ofradiation action in all living systems regardless of the complexity of their morphologicltl organization (Fig. 27).The physicochemical events after energy absorption give rise to subtle initial structural changes in the DNA that take place with great speed and are completed within milliseconds. These initial physicochemical changes lead to chemically stable structural defects, such as breakage of phosphodiester bonds, oxidation of the sugar moiety, deamination and dehydroxylation of bases and formation of peroxides (19-24). These defects may be induced by direct and/or indirect actions of radiation. The presence of these structural alterations may either (a) initiate repair enzyme activity, or (b) prevent DNA replication and transcription (synthesis of RNA),or (c) allow incomplete and partiaI replication and transcription. All these biochemical processes occur slowly during the first few hours after irradiation and they are crucial to survival. They can be influenced by the internal and external environments, that is, by the physiological state of the living system and by external factors. Thus, for example, the attenuation or failure of repair with caffeine increases the sensitivity to radiation. If the genome carries repair information, there is the possibility that repair mechanisms will eliminate all potentially lethal structural defects in the DNA, so that the informational “structure of the DNA may be reconstructed and restored.” This may be followed either by resumption of DNA replication and transcription, which could allow normal and cell replication (in dividing cell populations), or by normal transcription and genetic expression in nondividing cells resulting in normal cell functions. Either would lead to the survival of an irradiated system. Unrepaired radiation damage may have different biological consequences depending on the extent of the DNA damage and its topology (location). If the damage is located in a functional part of DNA, then faulty DNA replication may be either lethal or/and mutagenic. Faulty transcription, on the other hand, may result either in the lack of a functional protein or in the synthesis of a “nonsense” protein, causing early and/or late physiological alterations (phenotypic changes) or death of the living systems. The attempt to describe changes induced by radiation in the frame of a unitied mechanism is thus based on the concept that the main radiation effects-inhibition and delay of cell division, and loss of viability and mutagenesis-are dependent on the structural and functional integrity of DNA, i.e., ofgenetic information that it carries. Any radiation damage to the DNA, the expression of which is mediated through replication and transcription, may result in physiological changes or in death of the living system if the DNA does not carry the repair information. Consequently,

RADIATION-INDUCED ALTERATION I N DNA

213

minute and subtle defects in DNA may be amplified by transcription and expressed as significant physiological changes, which may be lethal. The question may be asked : Is there any direct evidence to support the ideas in this tentative hypothesis? My answer would be that there is no direct evidence for all the postulates of the hypothesis to make it universally valid, but recent experimental data strongly support some of the ideas. Some of the experimental data that give support to the main postulates of the hypothesis may be summarized as follows. (a) There is evidence that DNA is an important target for lethal radiation effects produced in viruses (48-52, 98-100, l a g )bacteria , (10-18, 25-33,54-73), and mammalian cells (74-80, 204-205,215-224). (b) The physicochemical nature of (c) There is some structural DNA damage is already explained (19-64). evidence indicating a relationship between the nature of structural changes in the DNA and lethality in viruses (129) and bacteria (I@-18). (d) Certain DNA structural defects may block DNA replication invitro (84, 86,88, 88a) and inVivo (84, 86,87), and suppress DNA transcription in vitro and invivo(84-88a). (e) Faulty DNA replication, giving rise to abnormal DNA and abnormal pulse-labeled RNA, may occur in irradiated living systems (71, 73,154,159, 160,245, 267, 278, 318). (f) There is evidence indicating that irradiation inhibits the synthesis of induced enzymes in bacteria (251-255, 269-27.4) and mammals (937-939, 339-3.@), whereas the preformed enzymes are not affected by the same doses of radiation, suggesting that irradiation prevents genetic expression and that the phenomenon is universal, i.e., it is the same in bacteria (251-255,269-274), in hepatocytes of rat liver (638, 340,341)or in brain cells (342). (g) The phenomenon of repair appears to be of general occurrence, having been demonstrated in and in mammalian cells (d15-625), although different bacteria (11-17) biochemical mechanisms may be involved in the repair of mammalian cells. It is my belief that as the situation stands today, there is considerabIe evidence in support of the proposed working hypothesis, which must, however, be subjected to further tests to prove in a more direct way how far it is applicable to higher living systems. Finally, it is my hope that the hypothesis, although frankly oversimplified, may stimulate basic research in the field of molecular radiation biology and thus lead to new ideas and original approaches to the problem. ACKNOWLEDGMENTS The author would like to express his gratitude to Dr.P. Martinovi6 for his helpful criticism and for reading the manuscript. Thanks are also due to colleagues, Dr.A. Beharevid, Dr.S. Petrovib, Dr. T. Hudnik-Plevnik, Dr. Marta Sirnib, Dr.B. KrajinEaniE, atid Dr. K. Sein, fortheir help on the preparation of the manuscript. The author also owes thanks to Mrs. Danka Filipovid and Miss Vera Bogosavljevi6 for their enthusiastic technical assistance.

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The presented research was supported by grants of the Yugoslav Federal Research Council and the Yugoslav Federal Nuclear Energy Commission.

REFERENCES 1. Z. M. Bacq and P. Alexander, “Fundamentals of Radiobiology.” Pergamon Press, Oxford, 1961. 2. M. Errera and A. Forssberg, eds., “Mechaniums in Radiobiology.” Academic Press, New York, 1961. 3.A. M. Iiusin, “Osnovi Radiacionoy Biologii.” Nauka, Moscow, 1964.

4.E. F. Rornancev, V. D. Blohina, N. N. Koscheenko, and I.V. Filipovich, “Ranii RadiacionobiohernicheskiiReakcii.” Atornizdat, Moscow, 1966. 6. D. E. Lea, “Actions of Radiation on LivingCells.” Cambridge Univ. Press, London and New York, 1947. 6 .D. E. Lea, R. B. Haines, and C. A. Coulson, Proc.Roy.SOC.B120, 47 (1936). 7. N. W. Timofieff-Ressowsky and K. G. Zimmer, “Biophysik I.Das Trefferprinaip in der Biologie.” Hirael, Leipzig, 1947. 8. E. C. Pollard, W. R. Guild, F. Hutchinson, and R. B. Setlow, Progr. Biophys. Chem.6,72 (1955). 9, K. 2. Zimmer, “Studies on Quantitative Radiation Biology,” p. 1.Oliver &Boyd, Edinburgh, 1961. 10. W. Szybalski and Z. Opara-KLlbinska, “Cellular Radiation Biology,” p. 223. Williams & Wilkins, Baltimore, Maryland, 1965. 11.D. Pettijohn and P. C. Hanawalt, J .Mol.Biol. 9,395(1964). 12.R. B. Setlow and W. L.Carrier, Proc. Natl. Acad.Sci.U.S.61,226 (1964). 13.R. P. Boyce and P. Howard-Flanders, Proc. Natl. Acad.Sci. U.S.61,293 (1964). 14.B. E.B. Moseley and H. Laser, Proc.Roy.Soc.B162, 210 (1965). 16.R. H. Haynes, Radiation Res.Suppl. 6,1 (1966). 16.R. A. McGrath and R. W. Williams, Nature212, 534(1966). 17 .H. S.Kaplan, Proc. Natl. Acad.Sci. U.S. 66,1442(1966). 18.C. J. Dean, P. Feldschreiber, and J. T. Lett, Nature209, 49 (1966). 19. R. Latarjet, B. Ekert, S. Apelgot, and N. Reybeyrotte, J .Chim.Phys.68,1046 (1961). 20. B. Ekert, Nature194, 278 (1962). 21.G. %holes, Progr. Biophys. Mol.Bwl. 13, 59 (1963). 22. A. Wacker, This series, 1, 369 (1963). 23.J.J.Weiss, This series, 3,103(1964). 24.L. E. Orgel, Advan.Eptzymol. 27,289(1965). 26.M. Demerec and R. Latarjet, ColdSpring HarborSymp.Quant. Biol. 9,38 1949). 26.R. Latarjet, Compt.Rend.Acad.Sci. 227, 1354(1949). 27. S. S. Cohen and A. Arbogost, J .Ezptl. Med.91,637(1950). 28.L. W. Labaw, V. M. Mosley, and R. R. Wyckoff, J .Bacteriol. 66,330 1953). 29.A. Kelner, J .Bacteriol. 66,252 (1953). 30. D. Kanaair and M. Errera, Biochim. Biophys. Acta14, 62 (1954). 31. S. S. Cohen and H. D. Barner, Proc. Natl. Acad.Sci.U.8.40, 885(1954). 32. D.Kanazir and M. Errera, ColdSpringHarborSymp.Quant. Biol. 21, 19 (1956). 33.D.Billen and E. Volkin, J .Bacteriol. 67,191(1954). 34.K. Paigen and B. N. Iiaufrnan, J .Cellular Comp.Physiol. 42,163(1953). 36.G. Klein and A. Forssberg, Exptl. CelERes.6, 211 (1954). 36. A. Forssberg and G. Klein, Exptl. CellRes.7 ,480 (1954).

RADIATION-INDUCED

ALTERATION

IN DNA

215

37.H. Harrington and P.Lavik, Arch.Biociiem. Biophys. 64,6 (1955). 38. D. F. Petersen, F. W. Fitch, and I<. B. DuBois, Proc. SOC. Ezptl. Biol. Med.88, 394(1955). 39. A. Howard, CibaFound. Symp.Irmising Radiation CellMetab., p.196(1956). 40. M. G. Ord and L. A. Stocken, Biochim. Bioph9s. Acta29,201 (1958). 41. L. S.Kelly, Progr. Biophys. Biophys. Chem.8,143(1958). 42. L. G. Lajtha, R. Oliver, R. Berry, and W. D. Noyes, Nature182,1788(1958). 43.G. A. Kritskii, Biokhimiya 20,158 (1955). 44. A.Bebarevic?, K. KaEanski, D. ManEid, and Lj. Hadiib, Experientia 12,23 (1956). 45. A. M. Kmin and V. I.Tokarskaya-Merenova, BioJizika 4,446 (1959). 46. L. G. Lajtha, i n“The Nucleic Acids” (E. Chargaff and J. N. Davidson, eds.), Vol. 3,p. 527.Academic Press, New York,1960. 47. P. Mandel and P. Chambon, i n“Immediate and Low Level Effects of Ionizing Radiation,” p.71.Taylor and Francis, London, 1960. 48. G. Bertani, J .Bacteriol. 79,387 (1960). 49.F. W. Stahl, J. M. Grasemann, L. Okun, E. Fox,and C. Laird, ViToZogy13,98 (1961). 60. H. Tanooka, Radiation Res.21,26 (1964). 51. D. Freifelder, Proc. Natl. Acad.Sci. U.S.64,128 (1965). 52. D. Freifelder and D. R. Freifelder, Mutation Res.3, 177(1966). 55. V. I.Tokarskaya, R ~ i o ~ ~ o 1, l 193 o g (1963). ~~a 54.H. S. Kaplan and P. A. Tomlin, Radiution Res.12,447 (1960). 55. H. S. Kaplan, K. C. Smith, and P. Tomlin, Nature 190, 794(1961). 56. H. S. Kaplan, K. C. Smith, and P. Tomlin, Radiation Res.16, 98 (1962). 57. J. H. Stuy, J .Bacteriol. 79,707 (1960). 68. J.H. Stuy, Radiation Res.14,56 (1961). 59. B. MiletiE, Z. KuEan, M. DrakuliE and Lj. Zajec, Bioehem. Biophys. Res.Commzan. 4,348 (1961). 60. B. MiletiE, KuEan, and Lj.Zajec, Biochem. Biophys. Res.Commun.4, 343 (1961). 61. B. MiletiE, Z. KuEan, and D. Novak, Nature 202,106(1964). 62. E. C. Pollard and C. Vogler, Radiation Res.16,109(1961). 63.H. S. Kaplan and R. Zsvarine, Bioehem. Biophys. Res.Gommun.8,432 (1962). 64.E. C. Pollard and P. M. Achey, Science 146, 71 (1964). 65. R. H. Haynes, M. H. Patrick, and J.E. Baptist, Radiation Res.22,194(1964). 66. D. Billen, R. Hewitt, and G. Jorgenson, Biocliim. Biophys. Acta103, 440(1965). 66a. R. Hewitt, J.C. Suit, and D. BiHen, J .Bacteriol. 93, 86 (1967). 67.W. Saybalski, J .Arkansas Med.SOC. 62, 486 (1966). 68. H, S.Kaplan, Radiation Res. Sz~ppl. 6, 185 (1966). 69. R. A. McGrath, R. W. Williams, and C. S. Swartzendruber, B i o p h pJ .6, 113 (1966). 70.2. TrgovdeviE and 5. KuEan, Intern. J .Radiation Biol. 12,193(1967). 71. E. W. Frampton and D. Billen, Radiation Res.28,109(1966). 78. D. Billen, K. Hen-itt, T. Lapthisophon, and P. M. Achey, J .Bacteriol. 94,1538 (1967). 73.P. M. Achey and E. C. Pollard, Radiation Res.31, 47 (1966). 74. B. Djordjevid and W. Szybalski, .I. Ezptl. Med. 112,509 (1960). 76. G. F. Whitmore, C. P. Stanners, J. E. Till, and S. Gulyas, Bioehim. Biophys. Aetn 47, 66 (1961).

5.

216

D. T. KANAZIR

76. R. B. Painter, Radiddon Res.16, 846(1962). 77. R. B. Painter and R. E. Rasmussen, Nature201, 162,409 (1964). 78. J.T.Lett, G .Parkins, P. Alexander, and M. G. Ormerod, Nature203,593 (1964). 79.T.P. Brent, J. A. V. Butler, and A. R. Crathorn, Nature210,393(1966). 80.J. T. Lett, I. Caldwell, C. J.Dean, and P. Alexander, Nature214, 790 (1967). 81. 0. F. Nygaard and R. L.Potter, Radiation Res.10, 462 (1959). 82. Y. Sugino, E. P. Frenkel, and R. 1,.Potter, Radiation Res.19, 682 (1963). 83. N. Fausto, A. 0. Smoot, and J. L. Van Lancker, Radiation Res.22, 288 (1964). 84. H. Harrington, Proc.Natl. Acad.Sn’. U .S. 61, 59 (1964). 86. H. Harrington, Arch.Biochem. 117, 615 (1966). 86. H. Eckstein, V. Paduch, and H. Hiz, EuropeanJ .Biochem. 3,224 (1967). 87. F. T.Hatch, J.L. Moore, and J.J. Koranda, Radiation Res.31,605 (1967). 88. F. Zimmermann, H. Kroger, U. Hagen, and K. Keck, Biochim. Biophys. Acta 87,160(1964). 88a. U. Hagen, H. Kroger, and E. Petersen, Studiu Biophys. 6 ,62 (1967). 89. E. C. Pollard, Science 146, 927 (1964). 90. E. C. Pollard and T. F. Barone, Radiation Res.Suppl. 6, 194(1966). 91. P. Howard-Flanders, R. P. Boyce, E. Simpson, and L. Theriot, PTOC. Natl. Acad. Sci. U.S. 48,2109(1962). 92. R. H. Haynes, Photochem. Photobiol. 5, 429 (1964). 93. N. P. Dubinin, “Molekdarnaya Genetika i Destiviye Izlutchenii na Nasledstvenost.” Gosatoddat, MOSCOW, 1963. 94.E. Freese and E. B. Freese, Radiation Res.Suppl. 6, 97 (1966). 96. E. M. Withm, Radiation Res.Suppl. 6, 30 (1966). 96.R. B. Setlow, J. Cellular Comp.Physiol. 64, Suppl. 1, 51 (1964). 97. P. Howard-Flanders and R. P.Boyce, Radiation Res.Suppl. 6, 156 (1966). 98. W. Harm, Virology 19, 66 (1963). 99. W. Harm, Mutation Res.1,344(1964). 100. G. E. Fradkin, in “Nucleinnoviye Kisloti i Biologicheskoye Deystivie Yoniziruyuchey Radiacii,” p.161Nauka, Moscow, 1967. 101. W. C. Summers and W. Szybalski, J. Mol.Biol. 26, 227 (1967). 101a. D. T. Denhardt and R. L.Siheimer, J .Mol.Biol. 12, 674 (1965) 102. G. Hotz and H. Reuschl, MoZ. Gen.Genet. 99, 5 (1967). 102a.G. Hotz, Mol.Gen.Genet. 102, 44 (1968). 103,B. Straws, H. Reiter, and T. Searashi, Radiation Res.Sup& 6,201(1966). 104. B. E. B. Moseley, Intern. J .Radiution Biol. 6, 489 (1963). 106. P.Alexander, C. J. Dean, and .T. T. Lett, 3 u X Boris Kidrich Inst., SuppE.17, 1 (1967). 106.T. Hudnik-Plevnik, N. Djordjevib, D. Savib, B. KrajinEani6, and D. Kanazir, in preparation, 1968. 107.J. Hillova, BuEl.Inst. BorisKidrich Suppl. 17,11 (1967). 108. W. SzybaIski and 2. Larkiewics, Abhandl.Deut.Akad.Wiss. Berlin, Kl.Med. p.63 (1962). 109. S. Greer, J. Mol.BioE.22, 618(1960). 110. H. S.Kaplan, J. D. Earle, and F. L. Howsden, J .Cellular Comp.Physiol. 64, Suppl. 1, 69 (1964). 111. H. S. Kaplan and F. L. Howsden, Proc.Natl. Acad.Sci.U.S. 61, 181(1964). 11.9. H. S. Kaplan, Radiation Res.Suppl. 6, 185 (1966). 115. R. P. Boyce, Nature208, 688 (1966). 114. H. D. Mennigmann and W. Szybalski, Biochem. Biophys. Res.Commun. 9, 398 (1963).

RADIATION-INDUCED

ALTERATION

IN DNA

217

116.1).Kanazir, Bioclbim. Biophys. Acln30, 20 (1958). 116.It. H. Haynes, in“Physical Processes in Radiation Biology,” ( L .Augenstein, R. Mason, and B. Rosenberg, eds.), p. 51.Academic Press, New York, 1964. 117. J. Marmur, W. F. Anderson, L. Matthews, K. Berm, E. Gayevska, D. Lane, and P. Doty, J .Cellular Comp.Physiol. 68, Suppl. 1, 33 (1961). 118.E. M. Witkin, Science 162,1345(1966). 119. R. F. Kirnball, Radiation Res., Suppl. 6, 51 (1966). 120.G. Hevesy, in“Radioactive Indicators.” Interscience, New York, 1948. 191.A. M. Kuzin and A. A. Wainson, Nature 212, 820 (1966). 122.H. J. Burki and S. Okada, Radiation Res.31, 645 (1967). 1.83. A. H. Sparrow and H. G. Evans, Brookhaven Symp.Biol. 13,76 (1961). 18.4. J. 5. Bedford and E. J. Hall, Radiation Res. 31, 679 (1967). 19.5. J. Storrer, Radiation Res.31, 699 (1967). 196. E. D. Wills and A. E. Wilkinson, Biochem. J .99, 657 (1966). 187. E. D.Wills and A. E. Wilkinson, Radiation Res.31, 732 (1967). 198.J. B.Clark, European J .Biochcm. 2, 19 (1967). 129.D. Freifelder, Radiation Res. Suppl. 6,80 (1966). 129u. D. Freifelder, personal oommunication, 1965. 130.G. Scholes and J. Weiss, Exptl. Cell Res.Suppl. 2, 219 (1952). 131. G. Scholes and J.Weiss, Biochem. J .63, 567 (1953). 152. G. Scholes and J. Weks, Biochem. J .66,65 (1954). 153.A. H. Sparrow and F. M. Fhenfeld, Science 104, 245 (1946). 154. J. A. V. Butler, Radiation Re8.Suppl. 1, 403 (1959). 136.M. Errew, ColdSpringHarborSymp.Quant. Biol. 12, 60 (1947). 136.L.J. Cole and M. E. Ellis, Radiation Res.6,252 (1956). 137. P. Alexander and K. A. Stacey, in“Progress in Radiobiology,” p. 105.Oliver I% Boyd, Edinburgh and London, 1956. 138.T. Hudnik-Plevnik, Doctoral Thesis, University of Belgrade (1963). 139.G. A. Kritskii, Biolchimiya 29, 701 (1964). 140.V. V. Konstantinova, Radiobiologiya 3, 336 (1963). 141. G. P. Toropova and N. V. Ermolaeva, Med.Radiol. 30, 24 (1956). 148.L. S. Isupova and V. S. Balabuha, Med. Radiol. 8,36 (1961). 63, 681 (1950). 143.G. Limperos and W. 13. Mosher, Am. J .Roentgenol. 144.V. A. Struchkov and A. M. Kuzin, Radiobiologiya 1, 153 (1961). 145.P. F. Davidson, D. Freifelder, and B. W. Holoway, J .Mol.Biol. 8,1 (1964). 146. U .Szybalski, Radiation Res. Suppl. 7, 147 (1967). 147. P. Alexander and J. T.Lett, Natwe187, 933 (1960). 148. R. A. Cox, Nature 176, 919 (1955). 149. A. R. Peacocke and B. N. Preston, J .Polymer. Sci. 31, 1 (1958). 150. U. Hagen, Nature 194, 554 (1962). 151.D. L.Dewey, Intern. J .Radiation Biol. 12, 497 (1967). 152.D. C. Huston and E. C.Pollard, Biophys. J .7, 555 (1967). 153. A. Kuzin, Intern. J.Radiation Biol. 6 ,201 (1963). 164. E. C. Pollard and P. M. Achey, Radiation Res. 27, 419 (1966). 155.J.D. Chapman and E. C.Pollard, Radiation Res. 31,607 (1967). 156.L. J. Grady and E. C. Pollard, Biochim. Biophys. Acta146, 837 (1967). 167. C. R. Shaffer and R. A. McGrath, Exptl. CellRes. 39, 604 (1965). 158.T. Hudnii-Plevnik, personal communication. f59. E. W. Frampton and D. Billen, Radiation Res.26, 190(1965). 160.P. M. Achey and D. Billen, Radiation Res. 31, 606 (1967). 161. T. Hudnik-Plevnik and A. Stoclren, Nature 192, 554 (1961).

218

D. T. KANAZIR

162. J. H. Taylor, ed., “Molecular Genetics,” Vol. 1, p. 65. Academic Press, New York,

1963. 16s. Ph. E. Hartman and 5. R. Suakind, in“Gene Action,” (S. R. Suskind and Ph. E. Hartman, eds.), p.1. Prentice-Hall, New Jersey, 1965. 164. J. Cairns, J .MoZ. Biol. 6,208 (1963). 165. A. Kornberg, I. R. Lehman, M. J. Bessman, and E. S. Simms, Biochim. Biophys.

Acta21, 197 (1956). 166. ,J.N. Davidson, R. M. S. Smellie, H. M. Keir, and H. H. McArdle, Nature 182,589

(1958). 167. F. J. Bollum, J .Am. Chem.Soc. 80, 1766 (1958). 168. R. Mantsavinos and E. S. Cannelakis, Biochim. Biophys. Acta27, 661 (1958). 169. A. Kornberg, “Enzymatic Synthesis of DNA.” Wdey, New York, 1962. 170. M. J. Bessman, I. K. Lehman, E. S.Simms, and A.Kornberg, J .Biol. Chem.233,

171 (1958).

C. L. Schildkraut, C. C. Richardson, and A. Kornberg, J .Mol.Biol. 9,24 (1964). R. B. Inman, C. L. Schildkraut, and A. Kornberg, J .Mol.Biol. 11, 285 (1965). C. C. Richardson, R. B. Inman, and A. Kornberg, J .M o t Biol. 9, 46 (1964). F. J. Bollum, J .Biol. Chem.236, PC 18 (1960). K. Hory, H. Fujiki, and Y. Takagi, Nature210, 604 (1966). A. G. Lezius, S. B. Hennig, C. Menzel, and E. Metz, European J .Biochem. 2, 10 (1967). 177. F. S. Bollum and R. B. Setlow, Biochim. Biophys. Acta68, 599 (1963). 178. K. A. Stacey, Biochim. Baophys. Res.Commun.6, 486 (1961). 179. E. R. Walwick and R. K. Main, Biochim. Biophys. Acta66,255 (1962). 180. C. M. Wheeler and S. Okada, Intern. J .Radiation Biol. 3,23 (1961). 181. P. Cerutti, H. T. Miles, and J.Frazier, Biochem. Biophys. Res.Commun.22, 466

171. 172. 173. 174. 175. 176.

(1966). 182. R. Adman and L. Grossman, J .Mol.Biol. 23, 417 (1967). 183. S. Robev and Z.Marinova, Nature 213, 935 (1967). 184. W. Ginosa and A. Norman, Nature 179, 520 (1957). 185. I. Tessman, E. S.Tessman, and G.8. Stent, ViTOlOglJ4, 209 (1957). 186. I. Tessman, Virolog~/ 4, 7, 263 (1959). 187. W. Sauerbier, Virology 16, 465 (1961). 188. D. J. Fluke and E. C. Pollard, Ann.N . 1’. Acad.Sci. 69,484 (1955). 189. L. W. Labaw, V. M. Mosley, and W. G. Wyckoff, J .Bacteriol. 66, 330 (1953). 190. R. B. Setlow, J .Cellular Comp.Physiol. Suppl. 64, 51 (1964). 19Oa. R. B. Setlow and J. K. Setlow, Proc. Natl. Acad.Sci. U.S.48, 1250 (1963). 191. A. Phil and T. Sanner, Radiation Res.28, 96 (1966). 192. M. Delbriick, Radiation Res.Suppl. 6, 234 (1966). 19s. D. K. Meyers and K. Skov, Can.J .Biochem. 44, 839 (1966). 194. J. E. Cleaver, Biochim. Biophys. Acta108, 42 (1965). 195. W. A. Creasey and L. A. Stocken, Biochem. J .72, 519 (1959). 196. D. W. Van Bekum and 0. Vos, Brit.J .Ezptl. Pathol. 36, 4 (1955). 197. A. M. Kuain and K. S. Trincher, Bioph~siku 6, 533 (1960). 198. G. P. Toropova, Dokl.Akud.NaulcS S S R117,266 (1957). 199. K. P. Hanson, Radiobwlogiya 6, 44 (1965). 200. H. M. IUouwen and i\. W. M. Appelman, CancerRes.27, 255 (1967). 201. I. Betel, Intern. J .Radiation Biol. 12, 459 (1967).

202.A. Howard and S. R. Pelc, Heredity Suppt. 6,261 (1952). 203.L. G. Lajth, R. Olner, and F. Ellis, Brit. J .Cancer8,367 (1954).

RADIATION-INDUCED

ALTERATION

IN DNA

219

204.J. E. Till, Ann.N . Y .Acad.Sci. 96,911 (1961). 206.S. Mak and J. E. Till, Radiution lies. 20, 600 (1963). 206.T. Terasima and L. J . Tolmach, Biophys. J .3, 11 (1963). 207.B. Negkovid, Nature 206, 837 (1965). 208.J.S.Bedford and E. J. Hall, Radiation Res.31, 679 (1967). 209.W. C. Dewey and It. M. Humphrey, Radiation Res.16, 503 (1962). 210.F.J.Bollum, J. W. Anderegg, A. 13. McFlya, and V. R. Potter, CancerRes.20, 138 (1960). $11. M. Fujioka, M. Koga, and I. Lieberman, J .Biol. Chem.238, 3401(1963). 212.S.Kishimoto and I. Lieberman, Ezptl. CeEL Res.36, 92 (1964). 213.E.W. Taylor, Exptl. Cell Res.40, 316 (1965). 214.B. Jacobson and T. Lee, Radiation Res.31, 368(1967). 215.R. E.Rasmussen and R.. B. Painter, J .CellBiol. 29,11 (1966). 216.M. M. Elkind and H. Sutton, Radiation Res.13, 556 (1960). 217.M. M. Elkind, J. Genetics (Tokyo) Suppl. 40,176 (1965). 218.M. M. Elkind, Bull. Boris Kidrich Inst Suppl. 17,25 (1967). 819.A. Han, B. Mileti6, D. Petrovid, and D. Jovid, Intern. J .Rdiation Biol. 8,201 (1964). 220.A. Han and D. PetroviE, Bull. Boris Kidrich Inst. Suppl. 17, 33 (1967). 221.R.A. McGrath, Intern. J .Radiation Biol. 2,177 (1960). 222.N. K. Das and M. Alfert, Proc.Natl. Acad.Sci. U .S.47,1 (1961). 223.0. F.Nygard and S. Guttes, Intern. J .Radiation Biol. 6,23 (1962). 22.4. R. A. McGrath, W. M. Leach, and J. G. Carlson, Exptl. CellRes.37, 39 (1965). 226.S.Damjanovich, T .Sanner, and A. Phil, European J .Biochem. 1, 347 (1967). 226.G. Hevesy, Advan.Biol. Med.Phys.1, 409 (1948). 227.R. Abrams, Arch. Biochem. 30, 90 (1951). 228. E.L.Bennet, L. S. Kelly, and B. Kreuckel, Federation Proc.13, 181 (1954). 229.0. F.Nygaard and R. L. Potter, Radiation Res.12, 120(1960). 2SO. 0. F.Nygaard and R. L. Potter, Iiadiation Res.16, 243 (1962). 231.J.L.Van Lancker, Biochim. Riophys. Acta46,57 (1960). 232.J.L.Van Lancker, Biochim. Biophys. Acta66,61 (1960). 2%. W. B.Looney, L. 0. Chang, M. L. Pardue, and F. W. Banghart, Rudiation Res. 19,203 (1963). 234.C. W .Bishop and J. N. Daviclson, Brit. J .Radiot. 30,367 (1957). 236.F. J.Bollum and V. K. Potter, J .Biol. Chem.233, 478(1958). 236.R. E.Beltz, Biochim. Biophys. Res.Commun.9,78 (1962). 237.N. Faust,o, T. Uchiyama, and J. L. Van Lancker, Arch. Biochem. Bioplrys. 106,447 (1964). 258.L. Baugnet-Mahieu, R.Goutier, and M. Semttl, Radiation Res.31, 808 (1967). 239.J. S.Roth, Life Sci. 3, 1145(1964). 240. E.V. Budilova, Radiobiologya 7, 323 (1067). 2.41. D. W. Van Bekum, Biochim. Biophys. Acta26,487 (1957). 2.42. G. P.Toropova, Doklady Akud.Nauk SSSE If7, 266(1957). 243.J.W. Whitfield and T. Youdale, Exptl. CeltRes.43, 153(1966). 244. J.W. Whitfield and T. Youdale, Nature 211, 212 (1966). 246.N. Voiculetz, J. R. Maisin, and I,. Ledoux, Proc.2nd Intern. Biophys. Congr., Vienna, 1968Abstr. 662. 246.V. A.Struchlcov, i n“Nukleiriovie Kialoli i Hiologicheskoye Deystvie Yoniziruyuschei Radiacii,” p. 56,Kauka, Moskva, 1967.

220

D. T. KANAZIR

247.L.A.Stocken and M. G. Ord, i n CellNucleus Metabolism and Radiosensitivity,” p. 141. Taylor & Francis, London, 1966. 247a.M. G. Ord and L. A. Stocken, Proc. Roy.SOC.Edinburgh (B)70,117(1968). 248. M. Berenbom and E. Peters, Radiation Res.6,515 (1956). 249.A.Bebarevib, S.Petrovib, V. Jankovib, D. T.Kanasir, and G. Jovicki, i n“Biological Effects of Ionizing Radiation at the Molecular Level,” p. 421.Intern. Stomic Energy Agency, Vienna, 1962. 250.V. I.Tokarskaya, Radiobiologyn 2,161 (1962). ,951. P. A. Swenson and R. B. Setlow, Science 146,791(1964). 252.G.W . Ruqhisky, M. Riley, L. S. Prestidge, and A. B. Pardee, Biochim. Biophys. Acta46,70 (1960). ,953. D. Nakada and B. Magasanik, J .Mol.Biol. 8, 105(1964). ,954. T .Kameyama and G. D. Novelli, Biochim. Biophys. Actu97,529(1962). ,955. A. B.Pardee and L. S. Prestidge, J .Bucteriol. 95, 1210(1967). 266. J. J. Weiss and C. M. Wheeler, Nature 203,291 (1964). 257.R.C. Huang and J. Bonner, Proc. Natl. Acad. Sci. U.S.48, 1216(1962). 257a.G. P. Georgiev, This series 6 ,259 (1967). 2.58. J. J. We& and C. M. Wheeler, Biochim. Biophys. Actu146,68 (1967). 259.W .B. Wood and P. Berg, J. Mol.Biol. 9,452 (1964). 260.J. Hurwits, J. J. Furth, M. Anders, and A. Evans, J .Biol. Chem.237,2752 (1962). 261. A. Ishihama and T. Kameyama, Biochim. Biophys. Actu138, 480 (1967). 26.2. J. Ono, R. G. Wilson, and L. Grossman, J .Mol.Biol. 11,600 (1965). 263. F. Grariosi, in“Radiation Research (G.Silini, ed.), p.438.North-Holland Publ., Amsterdam, 1967. 264.B. Woodson, Biochim. Biophys. Res.Commun.27, 169(1967). 265.B. Brdar, M . Drakulib, and E. Kos, Biochirn. Biophys. Acta119,362 (1966). 266.E. Kos, M. Drakulib, and A. Brdar, Nature 208, 1125(1965). 6/37. 0.Klamerth, Intern. J .Radiation Biol. 8, 291(1964). 268.E. W. Frampton, J .Bacteriol. 87,1369(1964). ,969. E.Pollard, Science 146,927 (1964). 270.D. Billen and H. C. Lichstein, J.Bucteriol. 63,522(1952). g 7 f .H. Chantrenne and S.Devrew, Biochim. Biophys. Acta. 31,134(19.59). 272.M. M. Wagle and G. B. Nadkarni, Radiation Res.33,174(1968). 873.G. D. Novelli, T. Kameyama, and J. M. Eisenstadt, J .Cell. Comp.Physiol. Suppl. 68, 225 (1961). ,974. P. A. Swenson and R. B. Setlow, 6.Mol.Biol. 16,201 (1966). 275.P. C. Hanawalt and R. B. Setlow, Biochim. Biophys. Actu41,283 (1960). 276.A. Riirsch, A. Edelman, and J. A. Cohen, Biochim. Biophys. Acta68,271(1963). 277.H. H. Kroes, A. M. Schepman, and A. Rorsch, Biochim. Biophys. Acta76, 201 (1963). ,978. T. H u d n i k - P l e d , Biochim. Biophys. Acta103, 515(1965). dr9.T. Hudnik-Plevnik, 6thAnn.Meeting European SOC. Radiation Biol. Interlaken, Switzerland, 1968. Abstr. 280. H. Suzuki and M. Hayashi, Biochdm. Biophys. Acta87, 610 (1964). 681. S.Spiegelman, ColdSpring HarborSynip. Quant. Biol. 26,75 (1961). 282.D. Savib, personal communication. 283.J. R. Wakes and R. M. S. Smellie, Biochim. J. 99,347 (1967). 284.M. Edmonds, J .Biol. Cheni. 240, 4621 (1965). 285.P. R.. Venkataraman and H. R. Mahler, J .Biol. Chem.238,1058(1963).

RADIATION-INDUCED

ALTERATION

I N DNA

221

T. T. Trujillo, D. L. Williams, and F. N. Hayes, J .Biol. Chem.241, 1915(1966). 287. R. P. Perry, Natl. CancerZnst. Monograph. 18, 325 (1965). 5 88.T. Tarnaoki and G. C. Mueller, Biochim. Biophys. Acta108, 73 (1965). 289. J. J. IIoland, Proc. Natl. Acad.Sci. U.S.50, 436 (1963). 5’90. I. PeEevski and B. Miletib, Nature 210, 323 (1966). 291. I.PeEevski and 2. ICuEan, Biochim. Biophys. Acta145, 310 (1967). 292. P. A. Swenson and S. Nishirnura, Photochem. Photobiol. 3,85 (1964). 29.5’. H. G. Zachau, Z. Physiol. Chem.336, 176(1984). 294. M. H. Buc and J. F. Scott, Biochim. Biophys. Res.Commun.22, 459 (1966). 295. Kin-Ichiro Miura, This series, 6, 68 (1967). Biophys. Res.Commun.27, 579(1967). 5 96. 11.Adam and P. Doty, Biochzm. 297. 0. Yamamoto, Intern. J .Radiation Biol. 12, 467(1967). 298. J. F. Scaife, Can.J .Biochem. 42, 623(1964). 299. J.F. Scaife, Can.J .Biochem. 43, 119(1965). 500. E. Harbera and Ch. Heidelberger, J. Biol. Chem.234, 1249(1959). 301.R. Logan, M. Errera, and A. Ficq, Biochim. Biophys. Acta32,147 (1959). 302.J. F. Mori and T. Morita, Nature200,1323(1963). 303. E. V. Boudnitskaya, M. Brunfaut, and M. Errera, Biochim. Biophys. Acta80, 567 (1964). 304. J. H. Kim, S. H. Kim, and A. S. Gelbard, Radiation Res.31, 610(1967). 305. 11. Matsudaira, T. Sekiguchi, Ch. Nakamura, and T. Hishizowa, Intern. 6. Radiation B i d .13, 21 (1967). 306. B. A. Negkovi6, A. Milosavljevi6, M. Pantelid, M. Bugarski, 2. AjdaEi6, R. Tomin, and S.Nikolib, Neoplasma 14, 6 (1967). 507. Y. Doida and S. Okada, Radiation Res.31, 607 (1967). 508. R. A. Tobey, E. C. Anderson, and D. F. Petersen, Proc. Natl. Acad. Sn’. U.S.56, 1320 (1966). 309. S. Kishimoto and 1. Lieberman, Exptl. CellRes.36, 92 (1964). 310. W. A. Creasey, Biochem. J .77, 5 (1960). 311. M. Swift, L. Rebhum, E. Rash, and L. Woodard, in“Cellular Mechanisms in Differentiation and Growth,” (D. Rudnick, ed.), p.107,Princeton Univ. Press, Princeton, New Jersey, 1956. 312. M. Fujioka, M. Kaga, and I.Lieberman, J. Biol. Chem.238, 3401(1963). 513. N. L. Bucher and M. N. Swaffield, CancerRes.24, 1611(1964). $14. K. Tsukada and I.Lieberman, J .Biol. Chem.239, 2952(1964). 316. I<. Tsukada and I. Lieberman, J .Biol. Chem.240, 1731(1965). 516. T.Uchiyama, N. Fausto, and J. L. Van Lancker, Arch. Biochem. Biophys. 110, 191(1965). 3 1 6 ~T. . Uchiyama, N. Fausto, and J. L. Van Lancker, J .Biol. Chem.241, 991(1966). 317.T. L. Berg and R. Goutier, Arch. Intern. Physiol. Biochim. 74, 909 (1966). 518. S. P. Petrovib, D.T. Kanazir and A. BeCarevib, in“Biological Effects of Ionizing Radiation at the MolecuIar Level,” p- 429.Intern. Atomic Energy Agency, Vienna, 1962. 320. S. PetroviO, Biochim. Biophys. Acta61,842 (1962). 521. S. Petrovib, A.Bebarevib, and J. Petrovib, Biochim. Biophys. Acta95, 518(1965). 32g. K. W. Sherrer and B. J. McCarthy, Biochemistry 6,283(1967). 523. E. J.HidvBgi, J. Holland, E. Boroni, and V. VbtBresz, ActaBioehim. Biophys. Acud.Sci. (Hungary) 1, 107 (1966). 324.V. VartBresz, E. J. Hidvkgi, J. Holland, L. D. Ssabo and F. Antoni, Proc.5th 886. D. A. Smith, R. L. Ratliff,

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D. T. KANAZIR

Meeting Europenn Soc. Radiatkrn Biol., Spindlerov Mlin, Czechostovaicia, Studia Biophys. 6, 213 (1967). 8.26. F. Daljanski, Intern. Rev.CytoE.10,217 (1960). 397.V. B. Petters, G. W. Kelly, and H. M. Dembitzer, Ann.N . Y .Acad.Sci. 111,87 (1963). 328.K.A. Kafiani, M. Timofeeva, A. A. Neifan, J. A. Rackus and H. A. Meljnikova, Biohimiya (USSR) 81, 365(1966). 329.M. Timofeeva, A. A. Neifan, and K, A. Kafiani, in “Nukleinovie Kisloti i Biologicheskoye Deystvie Yoniziruyuschei Radiacii,” p. 175,Nauka, Moskva, r967. 330.M. M. Sirnib, T. Hudnik-Plevnik, and D. T. Kanazir, Nature 214, 489(1967). 331.K. Scherrer and J. E. Danell, B
Optical Rotatory Dispersion and Circular Dichroism of Nucleic Acids’ JEN TSI YANG Cardiovaseular Reseurch Institute and of Biochemistry, and Department of California San Francisco University MedicalCenter, Sun Francisco, California AND

TATSUYASAMEJIMA Department of Chemistr:y, ofScience and Engineering, College Aoyama GakuinUniversity, Tokyo,J a p a n

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I. Introduction . , . . . . . 11.Optical Rotatory Dispersion and Circular Dichroism . . A. Origin of Optical Activity , . . . . . . B. Expressions forORD and CD . , , . . . . C. Correlation between ORD and CD . . . . . D. DrudeEquation. . . , , , . . , . E. Calibration of the Instrnmetits . , , . . F. Difference ORD . . . . , . . . . . 111.Mononncleosides and Monotiricletrtides . , . , . A. Comparison of the Purine and Pyrimidine Derivatives . B. The a- versus p-Linkages of the Sugars, . . . . IV. Oligonucleotides . . , . . . . . . . A. Dinucleoside Phosphates . . . , . . . . B. Trinucleoside Diphosphates and Other Oligomers . . V. Synthetic Polynucleotides . . . . . . . . A. Poly A, Poly U, and Their Complexes , . . . . B. Poly C, Poly G , Poly 0.Poly C, Poly I, Poly I.Poly C, and P01y A.2Poly I . . . . . . . . . . C. Poly d(A-T), Poly dA.I-’olydT, I’oly dC, Poly dA, and I’oly dT . . . . . , . . . . . I). Hybrids of Polyriboiiucleot ides aiid Polydeoxyriboiiucleotides 1 1. DNA aiid RNA . . . , . , . . . A. Similarities arid Differences bet,weeii DNA and RNA . B. Single- versus Double-Stranded Structure . . , . C. BaseTilting. . . . . . . . . . . I). Calculstioti of t,he ORD of 1lNA . , . , , . ,

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224 225 225 227 228 229 229 230 231 231 233 238 238 244 249 249

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257 261 262 262 269 271 277

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VII. Complexes ofNucleic Acids

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AND TATSUYA

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A. Ribosomes, Viruses, and Deoxyribonucleoproteins B. Complexes withSmall Molecules . . . . VIII. Visible Rotatory Dispersion . . . . . IX. Concluding Remarks . . . . . . References . . . . . . . .

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1. Introduction Three classes of biopolymers, nucleic acids, proteins, and polysaccharides, are optically active; they all contain asymmetric carbon atoms. Our current interest in the optical properties of these polymers is not in the rotations arising from the asymmetric configuration, but in correlating the optical activities with the conformations of these polymers (the so-called secondary, tertiary, and quaternary structures). The constituent pentoses of DNA and RNA, deoxyribose and ribose, are optically active. But it is the induced optical activity of the base chromophores attached to the pentoses that provides information concerning the overall molecular architecture of the nucleic acids. Although the history of optical activity goes back more than one s we hundred and fifty years (I), it was only in the middle 1950 that witnessed a renaissance of ORD? the change in optical rotation with wavelength. This technique has yielded many fruitful results in studies of conformation of proteins and polypeptides, and it has become a standard 3). It is therefore natural that experimental tool in protein chemistry (2, the same technique should be used widely in the study of the conformation of nucleic acids. This has been reinforced by introduction of R comple mentary technique, CD2 (4,5 ) .More data are now available from ORD than fromCD, but it is almost certain that the latter will rapidly accumulate. accurate ORD measurements of nucleic acids were conPrior to 1963, fined to wavelengths above 350 nll.r (for a brief review, see ref. 6). But the ultraviolet region enables us to observe directly the circular dirhroic bands that are responsible for the characteristic CD and ORD of nucleic acids. first managed, with a manual spectropolarimeter, Simmons and Blout (7) Section I1 for definitions) for the to detect a positive Cotton effect (see RNA isolated from tobacco mosaic virus with an inflection point around 260 mp, which shifted to 272 mp in 8 M urea. Fresco et al.(8) also attempted to push the ORD of calf thymus DNA and calf liver RNA toward 220mk and reported that both displayed multiple Cotton effects. However, the appearance of artifacts due to the use of solutions having an absorbance greater thnn 2 (i.e., the tranemittctl light intensity was much less than 1%) 2 Abbreviatiaris used in (his review: ORD, optical rotittory dkpersion; 0, rircidttr tlicliroism.

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interfered with the interpretation of t,heir data. Yang and Sarnejirno (9) reinvestigated the ORD of calf thymus DNA and found the multiple Cotton effects to have two peaks and one trough around the 260-mp absorption band: their results differed from those reported by Fresco etal.(8).This general ORD spectrum for nucleic acids was soon confirmed by inore precise measurements with recording spectropolarimeters (10). Since the introduction of the latter, accurate ORD results on nucleic acids, polynucleotides, and their constituent components have accumulated rapidly. I n 1963,Brahms and his collaborators (11-14) began a detailed exanination ofthe CD of poly- and oligonucleotides and nucleic acids; their measurements did not go below 220-230mp, but recent results from other laboratories extend down to about 200 mp or lower. Thanks largely to Tinoco and his co-workers (15-17 we ),have a better understanding of the origin of optical activity of these polymers and an explanation of the experimental observations in terms of the conformation ofthese polymers. Much, however, remains to be done before we can interpret such data in quantitative terms. This review begins with a brief description of the principles of ORD and CD. Following this, we present the optical activity of mononucleosides and mononucleotides, which are in a class by themselves, and discuss the similarities in the ORD and CD between oligonucleotides and polynucleotides and the differences between DNA and RNA. In essence, this is a review of work on the ORD and CD of nucleic acids of the past five years. The t(heoretica1interpretations are left in the competent hands of theorists, and it is hoped that in a few years it will be time for a review on the theoretical advances in this field.

II. Optical Rotatory Dispersion and Circular Dichroism (1,4, 18-.%?0) A. Origin of Optical Activity Optical activity is one ofthe many phenomena grouped under the term “spectroscopy,” which deals with the interaction of electromagnetic fields with matter. Polarized light is employed in the measurements of optical activity. Linearly poIarized light may be regarded as the resultant of right and left circularly polarized light of the same frequency. The two components are of equal magnitude but rotate in opposite directions; to an observer facing the light source the right-hand component rotates clockwise and the left-hand one counterclockwise. (This is not merely a convenient fiction, since experimentally a linearly polarized wave can be separated into

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its circularly polarized component,s or, conversely, these components can be superimposed to form a linearly polarized wave.) In an optically active medium, the right and left components would have unequal speeds causing a phase difference upon recombination, and the plane of polarization of the emergent light would be rotated through an angle, a,the optical rotation. Since the refractive index of the medium is related to the speed of light in the medium, optical rotation is simply a measure of circular birefringence, although this term is rarely used in the literature. Fresnel in 1825showed that a

(inradians)

= d ( n L-

~R)/X

(1)

where a is the angle of rotation at wavelength A, 1 is the light path of the medium, and nL and n R are the refractive indices of the left and right circularly polarized components. By convention, a clockwise rotation (similar to the right circularly polarized light) is termed dextrorotatory or positive (+) and a counterclockwise one (similar to the left circularly polarized light) is designated levorotatory or negative (-). When linearly polarized light is passed through an absorption band, the intensity of both circularly polarized components is reduced. If they are absorbed unequally, the absorption band is said to be optically active and we have CD, the difference in absorbance between the two circular polarizations. Since the incident light in this case is converted into elliptically polarized light upon emerging from the medium, we can also express the CD in terms of the ellipticity, \E, n7hich is simply related to the difference in absorbance by a proportionality constant (see below). Figure 1 illustrates an idealized CD spectrum and its corresponding ORD through an optically active absorption band. They are commonly called Cotton effects after their discoverer. In the idealized case, the CD extreme is located at the wavelength of maximum absorption, but unlike an absorption band, the CD can be positive or negative, depending on whether the absorbance of the left circularly polarized component is greater or smaller than that of the right one. Also by convention, the Cotton effect is positive (or negative) if the ORD maximum (or minimum) is on the long wavelength side. The maximum and minimum of the ORD have customarily been called the peak and trough, and the absolute magnitude of the difference in rotation between the two extremes (peak and trough) is defined as the amplitude of the Cotton effect. The wavelength of the crossover (zero rotation) coincides with t,hat of the absorption maximum in as distinthe idealized case. The Cotton effect shown in Fig. 1 is single, Cotton effects with two or more optically active guished from the multiple bands.

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ORD AND CD O F NUCLEIC ACIDS

CD

Negative

t x

0 Positive

n /Peak

FIG.1. Idealized Cotton effect of an isolated, optically active absorption band with its maximum at xi.The left-half is negative and the right-half positive Cottoneffect. From Yarig(20).

B.

Expressions forORD and CD

The optical rotatory power is expressed in terms of its specific rotation, by [].A = Cu/Z’C (2) where a is the observed rotation in degrees at wavelength A, 1‘ is the light path in decimeters, and C is the concentration in grams per milliliter. Its unit is degrees X cm2/decagram. For simple compounds of molecular weight M )the molar rotation [MI,is reIated to the specific rotation by [MI = (M/100)[a]. For polymers, the mean residue rotation, [m],provides a better representation than [MI.This is done merely by replacing M by the monomer residue molecular weight Mo. The late Moffitt (21)introduced rotation, [m ], which converts [m]to that the term reducedmean residue under vacuum by a Lorentz correction, i.e., [a], defined

[m’l

=

-+

13/(n2 -f 2>1[ml = 13/(nP 2)1(Mo/l00)bl

(3)

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where n is the refractive index of the solvent, which is also wavelength is widely used for ORD of proteins and dependent. The quantity [m ] polypeptides, but theoreticians still disagree on the validity of this Lorentz correction for polymers in solution. It is rarely used for the ORD of nucleic acids. The dimension for [MI,in?],and [?a is] degree X cm2/decimole. or m,and others use R and Some workers prefer to use the symbol @ for ill R instead of m and m . The CD can be expressed simply as the difference in molar absorptivity EL - eR,which equals (AL - AR)/mZ. Here the subscripts L and R refer to the left and right circularly polarized components, the A sare the absorbance, m is the molar concentration, and 1 is the light path in centimeters. The dimension of EL - ER is cm2/mole. As in the case of optical rotation, CD can also be expressed in terms of its ellipticity. Thus, we have the specific ellipticity, [*I, molar ellipticity, [el,and mean residue ellipticity, [el, simply by replacing a. M , and m in Eq. (3)by \k, 8, 0. Their dimensions are identical with the corresponding quantities for optical rotation. It can be shown (80) that EL - eB is related to its ellipticity by

[*I

=

3 3 0 ( A-~ ApJ/ZC

(4)

and

Here again the light path, Z, is in centimeters, and the concentrations, C and m, are in grams per milliliter and moles per liter. The A sfor [el, of course, refer t o the absorptivity based on mean residue rather than the entire polymer molecule.

C. Correlation between ORD and CD ORD, a dispersion property, and CD, an absorption property, share a common origin, the optical activity, and are thus closely related. Theorems of very general validity niake it possible to calculate the dispersion characteristics of a molecular system from its absorption properties orvice versa. Indeed, Moscowitz, using the Kronig-Kramers transform, has given explicit formulas that correlate ORD and CD (18,2% :)

imd(~>l =

(z/T)

hm

[ei(x‘)1[h’/(x2 - x’z)] ~JX’

(6a)

[m;(A )][X 2/(X2 - X’7] dX

(6b)

and [&(A)]

=

- (2/rX)

Equation (6a, b) can easily be solved with a computer program. For a

ORD AND

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CD OF NUCLEIC ACIDS

Gaussian CD, Eq. (62)would give an ORD profile similar to that shown in Fig. 1. An important theoretical quaiitity of optical activity is the rotational strength, Ri, which is related to the experimental ellipticitv, I P j ,by (18):

or

Here h is Planck's constant, c is tthe speed of light in vacuum, N I is the number of molecules per milliliter, and \Eiis in radians. The dimension of Ri is erg X cm3 X radian. The rotational strength determines the sign and magnitude of the partial CD and the corresponding ORD.

0. Drude Equation Drude, in 1896, first introduced a general expression for rotations in the region farfrom any of the optically active absorption bands:

where X i is the wavelength of the ith electronic transition and k; is a constant proportional to the rotational strength, Ri. Equation (8) was It later derived correctly from quantum mechanical considerations (23). can also be shown that for an idealized Cotton effect Eq. (6a) is reduced to one Drude term in a wavelength range distant from the absorption band (18). Equation (8) can be approximated as [m]A =

k / ( x2 A:)

(9)

if - << 1X2 - Xfi. This, however, is only a mathematical approximation and lacks any physical meaning. The graphical solution of Eq. (9) is done by plotting X2[cu]~against [.]A, which yields a straight line with X l and k as the slope and the intercept on the ordinate (Z.4). The ORD of all nucleic acids and polynucleotides thus far studied obey the one-term Drude equation over a certain range of wavelengths in the visible region.

E.

Calibration of the Instruments

Several recording spectropolarimeters and circular dichrometers are now commercially available; an excellent discussion of the instrumentation is The calibration of a polarimeter is given in the review by Holzwarth (25). commonly done by measuring the ORD of a known standard such as a freshly prepared sucrose solution (U.S. National Bureau of Standards

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grade) or the quartz control plates. The calibration of a circular dichrometer was, until recently, very uncertain because there were no universally accepted standards. However, Velluz el nl.(4) have compiled a list of numerical values for the CD of many organic compounds. Recently, Thibry (25u; private communication) developed a computer program for CD calibrations. By measuring the ORD and CD of a simple compound having a well-defined CD band and utilizing the Kronig-Kramers transform, we can derive the following expression:

+ a k X V ( X z - A);

[74cx*tl = fblcalc

(10)

Here the [mIealc is computed from the experimental CD using Eq.(6a), and the Drude term on the right-hand side of Eq. (10) represents the background rotations of all other CD bands distant from the CD band under consideration [see Eqs. (8)and (Y)]. The factor f is introduced to compensate for any imperfect adjustment of the instrument. The computer solves the three parameters, j , ak,and A h so that the computed rotations on the right side of Eq. (10) matches the experimental rotations. With the Thi6ry program, Cassim and Yang (26) found that d-10-camphorsulfonic acid in water has an EL - CR of f2.20 at 291 mji (the corresponding molar rotation at the 306-mp peak is +4120) and d-camphor in dioxane has +l.69 at 300 m p . The latter value compares fairly well with the value of +l.6 reported by Velluz etaE. (4). Thus, the CD data in the literature might be subject to revision when some common standards for calibration are accepted. Note that the absolute magnitude of the CD and ORD of d-10-camphorsulfonic acid or any other compound depends on the purity of the sample used, but it will not affect the factor f for a particular instrument, since both ORD and CD are measured on the same sample. On the other hand, the method of CD calibration based on Eq.(10) assumes that the spectropolarimeter for ORD is perfectly calibrated.

F. Difference ORD As in difference absorption spectra, difference ORD has the advantage of measuring small changes in rotation when a sample is compared to a reference material. In one commercial instrument that has a folded beam by using a mirrored Faraday cell (25), the coordinate system of the light beam is inverted after reflection from the mirror. Thus, if two identical solutions in cells of equal path lengths are placed in the incident and reflected beams, the rotation produced by the former is exactly canceled by that of the latter. This technique can then detect any small difference in rotation between two solutions, since the bulk of rotation of the polymer proper is canceled. The difference rotation can further be increased by increasing the concentration of the solution without exceeding the limits of

ORD A N D CD OF NUCLEIC

ACIDS

231

rotation in the present commcrcial instruments. However, the difference ORD method has not yet been applied to the study of nucleic acids. One problem inherent in this technique is the appearance of artifacts when an absorbing material, be it optically active or inactive, is placed in the reflected beam. Details of the correction measure have been described recently (27). Because nucleic acids absorb strongIy in the ultraviolet region, precise measurements of CD as well as ORD are very difficult. As a rule of thumb, the total absorbance of the solution including the cell must be kept well below 2 to avoid possible artifacts ( 2 )Extreme . caution in experimental measurements, especially when the degree of rotation is very small, cannot be overemphasized.

111. Mononucleosides and Mononucleotides A. Comparison of the Purine and Pyrimidine Derivatives All nucleosides and nucleotides have an absorption maximum near 260 mp; the optically inactive base chromophores are induced to produce Cotton effects when they are attached to the optically active sugars. The ORD in the ultraviolet absorption region of these compounds was not measurable prior to 1963, mainly because of instrument limitations. The small rotations in magnitude throughout most of the ultraviolet region are compounded by the high absorptivity of these compounds, thus making precise ORD measurements extremely difficult. Yang etal.(9, 28)have since examined 16 mononucleosides and 5’-mononucleotides in neutral, acidic and, in a few cases, alkaline solutions. Figure 2 shows the Cotton effects of 5’-deoxyribonucleotides and Table I lists the pertinent numerical have also reported the ORD of values of the ORD data. [Lin etnl.(30) dAMP, Fasman et ul.(31) of clCMP, Lamborg etat. (32) of 2 ,3 -AMP and 2’,3’-UMP, and Miles etal.(33) of uridine, thymidine, and cytidine.] The ORD of these compounds in all cases displays a single Cotton effect above 220-240 mp with a crossover near 260 mp. The troughs of the cytosine derivatives, however, suggest the overlapping of more than one Cotton effect. This is in accord with their absorption spectra, which show a shoulder at 230 mp at neutral p H in addition to a inaxiinum near 270 mp (29, 34). But the most important finding is that the purine derivatives Cotton effect and the pyrimidine derivatives a always have a negative positive one near the 260-mp absorption band. This generalization applies t o all mononucleosides and 5’-moaonuclcotides of P-D-pentose, be it, ribose or deoxyribose. The OltD of the cytosirie and thymicline derivatives was extended down to 200 inp (Fig. 2), and revealed another peak much larger

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2 0

-2

c

4

\

0 -4 -8

u 200

250

300

u 350

X (mp)

FIG. 2 .ORD of 5'-deoxyribomononucleotides. Note the difference in the ordinate scale between the upperand lower halves. From Yang et al.(88).

in magnitude than the first peak or trough. This is undoubtedly due to the intense absorption band below 210 mp (29). The ORD profiles of the derivatives of each purine or pyrimidine base are very similar, although the magnitude of the Cotton effects can vary. The attachment of the phosphate group to C-5’of the pentoses does not change the rotations significantly. Neither does the presence or absence of the 2'-hydroxyl group on the pentoses seem to cause much difference in rotations. Protonation of the bases can change significantly the absorption spectrum and therehy the ORD of nucleositles nnd nucleotides. Voet etal. (29) ohscrved changes in absorption spectra of cytosine derivatives and, to a lesser extent, adenine derivatives when the pII of the solution was

ORD

AND

CD O F NUCLEIC

233

ACIDS

TABLE I THE COTTON EFFECTSO F MONONUCLEOSIDES A N D 5'-MOiVONUCLEOTIDES" Peak Substance

dA A dAMP AMP dG G dGMP GMP dC C dCMP CMP dT dTMP U UMP

Crosuover

Trough

Amplitude Absorption (mp) [m]x 10-3 i(mp)i(mp) [nz]x 10-3 [m] x 10-8 x (rnpIb 260 275 268 272 258 268 270 28 1 288 288 288 288 290 260 283 283

-1.55 -3.30 -2.49 -3.00 -2.40 (-2.90). -3.35 (-1.61)' +5.85 $6.80 +6.30 $7.70 +a. 20 $1.50 $4.00 +3.50

258 256 258 257 250 249 248 247 273 273 273 272 277 278 271 272

245 245 253 246 231 2.70 233 237 250 242 245 24 0 260 257 253 255

-3.80 $2.25 $1.60 -4.90 +1 30 -3.79 $1.80 -4.80 -6.05 +3.65 (f 1 .(35)" ( -4. 8 5 ) C --6.45 +3.10 ( + O , 60)c ( -2.2 1)" $13.1 -7.20 -13.6 +20.4 -9.10 +lt5.4 -13 0 +20.7 -4 30 +6.50 -6 20 $7.70 -9.80 +13.8 +13.2 -9.70

259.5 260 260 260 252.5 253 252.5 253 272 271 272 270 267 268 261 262

For solutions in 0.1 M phosphate buffer (pH 7.5-7.9). Data taken from Yang etal.($8). * Data taken from Voeteta2. ($9). The figures in parentheses are only rough estimates.

adjusted to 2.5or lower. Similarly, the cytosine derivatives, for instance, showed a reduction in the amplitude of the Cotton effects accompanied by a red shift of the ORD spectrum. The derivatives of thymine and uracil are not protonated and therefore their ORD is unchanged in acid solution. But the guanine derivatives show a marked drop in the magnitude of their Cotton effects at pH 2,even though their absorption spectra are little The derivatives of guanine and thymine, affected in acid solution (29). having alkaline pK's between pH 7 and 12,also showed ORD spectra at pH 12 different from those at neutral pH, whereas the derivatives of adenine and cytosine remained essentially unaffected in alkaline pH's. Among the five bases studied, the magnitude of the Cotton effects is largest for the cytosine derivatives.

B. The

Q-

versus /?-Linkages of the Sugars

The sign for the Cotton effects of the purine and pyrimidine derivatives is highly dependent upon the anomcric configuration at C-1’of the pentoses. I ) synthesized and measured the ORD Ulbricht, and his co-workers (35-4 spectra of a series of mononucleosides and mononucleotides. The most

234

JEN TSI YANG

AND

TATSUYA

SAMEJIMA

interesting conclusion is that replacing the P-ribose (or ,&deoxyribose) by a-ribose (or a-deoxyribose) immediately reverses the sign of the Cotton effects; that is, it is positive for the a-nucleosides of purines and negative for those of pyrimidines, just the opposite of those described in the preceding section. Table I1 presents the Cotton effects of some of these anomers. This inversion must reflect primarily the orientation of the transition dipole of the base with respect to the sugar, which is different for the a- and p-anomeric nucleosides. Whether the sugar moiety is ribose or deoxyribose makes no major difference in the ORD spectrum. Variations of the suhstituents in the C-3 and C-5‘ positions appear to have little effect 011 the magnitude of the ORD curves. The 2’-epimers, however, show different magnitudes; for instance, the 2’-hydroxyl group in a cis configuration with respect to the pyrimidine base, as in P-D-arabinosy1s, have a larger magnitude than those in trans configurations, as in 8-D-ribosyls. The sign of the Cotton effects is also not affected by the nature of the substituents in the purine or pyrimidine rings. [Hudson’s isorotation rule, based on the [(Y]D, predicts that the a-u-configuration of a pair of anomeric glycosides is more dextrorotatory than the corresponding 8-configuration (42). It has now been shown that this rule does not apply to pyrimidine ribo- and deoxyribonucleosides (43-46). Needless to say, assignment of anomeric configurations based on measurements at a single wavelength is very arbitrary (41, 45). The Cotton effects, on the other hand, provide a simple and sensitive method for determining such configurations.] The study of models indicates that in the nucleosides of uracil, thymine, and cytosine, the proximity of the carbonyl group at C-2of the pyrimidine ring to the groups at C-2’ and C-5 of the sugar interferes with free rotation about the glycosyl bond; hence, the pyrimidine ring, e.g., uridine (structure I), although not rigidly held, has a preferred anticonformation so that the oxygen atom on C-2 of the pyrimidine ring is directed away from the furanose ring. Clark and Tinoco’s classification of spectral bands in pyrimidines shows that the Btuband (near 260mp) in uracil is polarized along the line joining the two keto groups (46). On this basis, Ulbricht ctat.(40) and proposed the following rule: The sign of the Cotton Emerson et al.(41) effects of pyrimidine pentofuranosyls will be positive if (a) the nucleoside has a preferred conformation owing to restricted rotation about the glycosyl bond and (b) a line from the C G O (or C4-NHJ group passing through the C2=0 group aIso passes from above to beIow the plane of the furanose ring provided that the chromophore is not twisted to such an extent that the line passes through C-5’. (“Above” is defined here as the same side of the furanose ring as C-5’.) For instance, spongouridine, like uridine, fulfills the above conditions and therefore has a positive Cotton effect [see also FriE et at.(47)]. On the other hand, 8-pseudouridine hns the line

ORD AND

CD OF NUCLEIC

ACIDS

235

Uridine (1)

passing from below to above the plane of the furanose ring; C-2,5’-cyclouriand 3-ribosyluracil has no preferred dine has the line passing through C-5 ; conformation (since both keto groups are orthoto the glycosylic nitrogen atom). The Cotton effects in these cases are therefore negative. However, /3-6-azauridines and P-6-azacytidines also have negative Cotton effects, even though their conformations are not expected to differ significantly , from those of uridine and cytidine. According to Emerson etal.( d l )the direction of polarization of the Bzuband in these compounds might not be the same as in uridine. The proposed rule also does not cover compounds containing chromophores other than those in the bases. As in pyrimidine nucleosides, the orientation of the purine nucleotides, e.g., AMP (structure 11),is also believed to favor the anti form with (3-2 of the purine ring directed away from C-5’ of the furanose ring, even though there may be considerable variation in the degree to which the base form. Indeed, Emerson et al.(39) found approximates completely an anti that the sign of the Cotton effects of 6-8,5‘-cycloadenosine, which has a fixed anticonformation, is the same as that of adenosine. alee and Mudd (CS), however, reached a different conclusion when they investigated a number of adenosine derivatives including some containing sulfur substituents on the (2-5 The . introduction of a sulfur atom of either the t,hioether or sulfonium type changes the sign of the Cotton effects around the 260-mp absorption band. These workers thus suggested that t,he nucleosides of purines may favor the synform (that is, the C-2 of the purine ring is directed toward the furanose ring), which was altered into the antiform in the case of the sulfur derivatives mentioned because of steric hindrance. But quite possibly there may be some electronic interactions between the sulfur atom and the purine ring that could alter the sign of the Cotton effects, even though such an interaction is not obvious from the absorption spectra. On the other hand, Lin etal.(50) proposed a hydrogen bond between the 5‘-phosphate hydrogen and the N-3 of the purine to account for the pH effect on the ORD of AMP. This intramolecular

COMPARISON

OF THE COTTON

EFFECTSOF

SOME

TABLE I1 ANOMERIC NUCLEOSIDES AND NUCLEOTIDES IN

Peak Substance

Configuration a t C-1’

(mp)

A4QUEOUS SOLUTION

Trough

[m]X 10-2

Crossover A (mp)

(w)

[m]X 10-3

Amplitude [m]X 10-8

rlbsorption , A, (mp) ~

Thymidine

a

B Ribopyranosylthymine

a

B 5-Hydroxymethyl2’-deoxyuridine 5-Fluoro-2 -deoxyuridine

ff

B a

P 3’-CMP

ff

B 2‘-Deoxycytidine

CY

B

286 282 281 287 279 279 286 286 283 289

-3.66 +1.89 -7.45 +1.06 -5.68 +5.90 -6.30 $4.35 -5.90 +5.10

283 290

-7.40 -4.63

252 255 248 260 249 249 252 253 254 250 -231 249 266

+6.78 -7.59 +12.7 -3.64 +8.10 -2.02 +11.55 -6.60 +2.46 -10.3 ---13.0 +13.6 -6.50

~~

-10.4 +9.48 -20.1 +0.47 -13.8 +7.92 -17.9 +11.0 -8.36 f15.4

267 267 265 265 266 266 263 269 271 271

-21.0 +11.1

271 280

0

2’-Deoxy-B-aeacy tidine @-Uridine @-Spongouridine P-Pseudouridine j3-3-Ribosylurrtcil D-Ribosyluracil

a

B B P P

P a

P D-Arabinosyluracil

a

P

I,-UMP

a

P

L-Thl P

a

B

z-CMP D-~~PVIP

a B a

B a

270 292 279 275 278 280

$2.03 -4.58 $4.10 4-11.5 -3.10 -3.30

-

283 284 281 280 282 283 288 290 289 290 2i5 279

-14.1 $5.1 -3.8 $14.7 $9.9 -3.5 $6.9 -2.3 $13.5 -8.4 $2.6 -5.3

269 273 271 266 270 273 274 279 273 274 26 1 262

-

-

252 233 247 245 258 256 252 253 250 249 252 254 254 258 235 250 235 250 245 243

--

-0.78 0 -7.6 -17.9 -0.8 +3.90

+2.78 -4.58 +11.7 +29.4 -2.3 -7.20

266 266 262 263 262 261

+23.3 -10.1 $12.0 -19.7 -18.4 +12.3 -15.6 $5.9 -15.7 +10.8 -3.6 +5.9

-37.4 +15.2 -15.8 +34.4 $28.3 -15.8 +22.5 -8.2 +29.2 -19.2 +6.2 -11.2

264 262 263 263 263.5 262 269 267.5 271.5 271 259 259.5

Dat,a were taken from Emerson etal.(41) (upper part) and Nishimura etal.(34) (lower part).

2!U c1

u

0

r Z

2r M

i; b3

z

238

JEN TSI YANG

AND TATSUYA

SAMEJIMA

hydrogen bonding requires AMP to have the syn conformation. However, Schweizer etal.(49) were unable to confirm the observations reported by Lin et al.(30). Finally, we note that all the X-ray studies and stereoindicate that most purine chemical analyses based on X-ray data (50-5.2) have the and pyrimidine nucleosides and nucleotides, including AMP (53), anticonformation in the solid state. Although the conformation of these compounds in solution is less clear, the X-ray studies show a substantial rotational barrier between the syn and antiforms, which makes a rapid conversion between them unlikely. In addition, the nucleoside bases in double-stranded DNA are known to have the anti form. Recent nuclear magnetic resonance studies also indicate that the nucleosides and 5’nucleotides in solution must be in the anti conformation (49, 54). Since the stereochemistry of a sugar at C-1’ is the same in the a - ~ configuration as in P-L-configuration, the purine and pyrimidine derivatives of the @+kind are expected to have positive and negative Cotton effects just as do the corresponding ones of the a-n-kind. Emerson et al.(39) reported the first measurement of a 8-L-nucleoside, namely, /%-adenosine, which has a positive Cotton effect. This is now confirmed by the work of Nishimura and his co-workers, who synthesized (55)and measured the ORD (34) of about 40 anomeric nucleosides and their 5’-phosphates Invariably, the ORD of @-L- and a-D-pyrimidine derivatives (see Table 11). show negative Cotton effects, and a-L- and @)-pyrimidine derivatives positive ones. Likewise, these workers confirmed that a-D-purine derivatives display positive Cotton effects as compared with the negative ones for 0-D-purine derivatives. Nishimura et al.(34) also observed that the configuration at C-2’ affects the magnitude of the Cotton effects, but those at C-3 and G4 do not. No systematic studies of CD of these mononucleosides and mononucleotides have been reported. There are few available data that can be quoted, except that CMP and UMP show one positive band at 272 and 265mp (56) and that AMP has a weak negative band near 255mp (57). Recently, 4-thiouridine and its phosphate display complex CD bands: two positive ones near 320-330and 260-270mp and a negative one at 230mp (58).

IV. Oligonucleotides A. Dinucleoside Phosphates Warshaw etal.(59) and Warshaw and Tinoco (60, 61)examined the ORD of all 16 dinucleoside phosphates of adenine, uracil, guanine, and cytosine at neutral, acidic, and alkaline pH’s. Figure 3 shows some of the

ORD AND

CD OF NUCLEIC

239

ACIDS

FIG.3. Rotation and absorption per base of six dinucleoside phosphates at pH 6.9 and ionic strength 0.1(0.01 M phosphate buffer plus 0.08 M KClO,) (SO).The absorption of ApA constituents (A PA) represents that of AMP alone.

+

representative ORD results together with the absorption spectra. The absorption spectrum of each compound is essentially characterized by that of its component monomers. The salient small differences are the hypochromicity near the absorption maximum and the hyperchromi~ity~ near 3 Thepercent hypochromicity (or hyperchromicity) is defined as [I - E(A)D/E(A)M]~OO, where E(X)D and E(A)M are tjhe molar absorptivities of the dimer and the equimolar mixture of its component monomers. The percent hypochromism (or hyperchromism) is defined as (1 - f~/faa)lOO, where f’s are the oscillator strength of the dimer and the equimolar mixture of the component monomers. The f’s can be calculated from the

x:

absorption spectrum and equal 4.32X 10-0

[e(h)/Xa] dX where Al and XB are the short

and long wavelength cutoffs. See Mahler etal.(62).

240

J E N TSI YANG AND TATSUYA SAMEJIMA

the long wavelength tail (except GpU at pH 1 and UpU at pH 11.5). In cont,rast, the ORD spectrum of each dinucleoside phosphate differs niarkedly from that of its component monomers. The general ORD profile is characterized by a long wavelength peak, a deep trough that occurs near the wavelength of the absorption maximum, and another peak occurring above 225 mp. This two-peak-and-one-trough profile was also observed for dinucleoside phosphates having deoxyribose instead ofribose (28), and it closely resembles that of polynucleotides and nucleic acids, discussed in Sections V and VI. Warshaw and Tinoco’s work reveals a striking sequence dependence of the ORD of dinucleoside phosphates. For instance, ApG and GpA at pH 7 have very different ORD’s with respect to the positions of the peak and trough (60, 01). Even those compounds that have similar ORD profiles, such as CpU and UpC, show dierent magnitudes of rotations. Thus, the ORD method can be utilized to distinguish between sequential isomers, even though their absorption spectra are, for all practical purposes, superimposable. Much evidence indicates that the bases in the dinucleoside phosphates interact strongly and tend to “stack” on top of one another. Among the nuclear physical methods used are absorption spectra (60, 61, 63,64), magnetic resonance (65-68), osmotic pressure measurements (69,70), and also ORD and CD. The multiple Cotton effects near the 260-mp absorption band as shown in Fig. 3 can be explained in t e r m of the base stacking (71, 72)and have been predicted by the theory of Tinoco (73). Warshaw and Tinoco (61) further classified dinueleoside phosphates into two groups by designating those with hypochromism less than or equal to 3yoas “unstacked” and those with more than 3% as “stacked.” (Of course, even the “unst,acked” ones possess some degree of stacking; thus their ORD differs from that of the component monomers.) Table 111 reveals two interesting features. First, at neutral pH A, G, and C stack, but U does not. This conclusion is consistent with the self-association studies of Ts’o etal. (69), who showed that uracil has the least tendency to stack and that stacking seems to decrease in the order: purine . purine > purine . pyrimidine > pyrimidine pyrimidine. Second, charges of the same sign on both bases cause unstacking, but a charge on one base does not. Exceptions to the above generalization are that at pH 7, UpG and CpU stack, at pH 11.5, CpU- stacks and ApG- does not, and a t pH 1, G+pU, UpG+, C+pG+, and G+pC+ stack. (Because of lack of titration data on these compounds, there is uncertainty about the last two compounds, which may actually be charged singly). Bush and Tinoco (74)also calculated the ORD ofthese dinucleosidephosphates by considering exciton interactions of the electronic transition moments above 220 mp as well as the directions and magnitudes

-

ORD

AND

CD OF NUCLEIC

THE

a

241

ACIDS

CLASSIFICATION

TABLE 111 DINUCLEOSIDE PHOSPHATES

OF

(61)

Borderline.

of these moments from the absorption spectra and found the position and relative magnitude of the first peak and trough (on the long wavelength side) in good agreement with experiment. That base-stacking interactions give rise to the observed multiple Cotton effects is also confirmed by the ORD study of GMP gel (76). In concentrated solution (10 mg/ml) and at low temperature (27, the monomers of this compound are stacked to form a left-handed helical structure (76). Accordingly, the Cotton effects of the gel are multiple instead of a single one around the 260-mp absorption band for the monomer. However, the ORD profile is inverted from that shown in Fig. 3 and resembles that of the helical poly I (see Section V, B), that is, it displays two troughs and one peak near the 260-mp absorption band. Raising the temperature of the solution causes a reversible transition from the helical aggregates to the monomer with a T,,, around 15". Brahms et al.(66) studied the CD and also absorption spectrum of n variety of 3 ’ 35' and 2'- 5' dinucleoside phosphates in a range of temperature between -20 and 80". Their results were classified into two categories. One is represented by essentially two adjacent positive and negative CD bands of almost equal rotational strength in magnitude witsorption mnximuin. Since the sum ofthe rotnt8ionnIstrengths in the spectrnl region untlcr investigation is approximately zero, the Cn is therefore designated "con-

242

J E N TSI YANG AND TATSUYA SAMEJIMA

servative,” for instance, ApA. The second category is composed of one or two positive CD bands, characteristic of the guanine and cytosine derivatives (Fig. 4). Since the sum of rotational strengths in the range of wavelengths studies is not zero in this case, the CD is termed “nonconservative” (for instance, CpC). The conservative type is in accord with the that is, the interactions among the prediction of the exciton theory (73), chromophores in a dimer give rise to a splitting of the monomer into two bands. The nonconservative type is attributed to contributions other than the exciton splitting, that is, t,o those arising from the interactions between near and far ult,raviolet transitions (77; see Section VI, C). The difference 10

8 6 4% 2 ;

0 -2

-4

220

240

260

200

300 220 X (mp)

240

260

280

300

FIG.4. CD and absorption per base of four dinucleoside phosphates at neutral pH in 4.7 M KF plus 0.01 M Tris and at -18 to -20°C (66). The absorption spectra are: left, UpG ( - - - ) and GpU (- . -); right. CpA ( - - - ) and ApC (- -). The dots for CD are the average of the monomer constituents.

.

between these two kinds of CD cannot be readily seen from their corresponding ORD. On the other hand, the results in Fig. 4 also reveal the sequence dependence of the CD of dimers just as ORD does (see Fig. 3).

EFFECT OF TEMPERATURE The rotation of the stacked dinucleoside phosphates decreases in magnitude with increasing temperature. This can be interpreted as the unstacking of the bases at high temperature, although the possibility that increasing torsional fluct,uation rather than base unst,acking causes the decrease in rotation cannot be completely ruled out. Similarly, the intensity of the CD of dimers decreases markedly with increasing temperature. With the linear van? Hoff plots, Brahms et nl.(56) estimated the thermodynamic parameters, AH , AS”, and AF , of the stack-unstack processes and could find no fundamental diflerenre among the dinucleoside phosphates studied warranting a division into L‘stxked”and “unstacked” groups. This is contrary to the widespread acceptance of such a division, as shown in

ORD AND CD O F NUCLEIC ACIDS

243

Table 111. However, it is not too difficult to reconcile this viewpoint with that deduced from optical properties. For instance, if the two stacked bases rotate relative to each other, the Cotton effects are markedly reduced, but the thermodynamic quantities may not be affected significantly. Note that these thermodynamic quantities are determined on the basis of a two-state model, which assunies that a dinucleoside phosphate molecule can have only two conformations, one stacked and the other unstacked. The stacked form is favored at low temperatures, and the equilibrium between the two states is shifted toward the unstacked form with increasing temperature. This model could be oversimplified, since the dinucleoside phosphate molecule is expected to have a continuous spectrum of the stacked conformations and it is difficult to visualize a sharp division on the molecular level between the two states. Davis and Tinoco (78)further pointed out that the calculated thermodynamic quantities of, for instance, ApA are not self-consistent, since optical rotation and hypochromicity methods gave very different results. Davis and Tinoco (78)and Glaubiger et aE.(79) thus proposed as an alternative a torsional oscillator model in which the parallel bases oscillate with respect to each other. This model predicts that the ORD of dinucleoside phosphates should show an exponential decrease at higher temperatures (as contrasted with the van't Hoff equation) and that the hypochromism should be almost independent of temperature. While the model fits the ORD data as well as the two-state model, it still does not predict the correct temperature dependence of the hypochroniism. David and Tinoco (78)concluded that both the two-state model and the oscillating dimer model fit some of the experimental data, but neither is completely satisfactory. Needless to say, we still cannot draw any definite conclusion about a specific conformation or conformations of dinucleoside phosphates in solution in spite of many physical studies of these and related compounds (free bases, nucleosides, and polynucleotides). True, the bases tend to stack at low temperatures in aqueous solutions and unstack at high temperatures and in denaturing solvents. But we cannot speculate much beyond this statement. Brahms et al.(56) also reported the CD of 2' --f 5' dinucleoside phosphates such as CpC, ApC, and ApA. The general profile is vcry similar to that of the 3’ 5' isomers, but the intensity in this case is relatively small and also the band position is slightly displaced. The changes in intensity of the 2 ’ - 5' dimers were some 104070 from 80" to -20 ,whereas the intensity of the 3 ’ 4 5 ’dimers can change by a factor of three. These findings suggest that the 2’ + 5’ dinucleoside phosphates also have a dissymmetric structure with weakly interacting bases and that their predominant conformation is probably close to the unstacked form in the region of temperature used. Brahms et al.(56) postulated that the 2’--f

244

J E N TSI YANC AND TATSUYA SAMEJIMA

hydroxyl group of ribose might form an intramolecular hydrogen bond, probably with an oxygen atom, which in turn favors the base stacking. This, they conclude, is supported by the study of deoxyribonucleoside phosphates such as dGpdG, which lacks the 2’-hyclroxyl group and, therefore, yields an exceedingly weak CD. We note that Ts’o et at. (80) aIso measured the ORD of dApdA, which shows Cotton effects similar to those of ApA except that their magnitude appears to be smaller than that of the ribosyl dimer.

6. Trinucleoside Diphosphaies and Other Oligomers Cantor and Tinoco (81) measured the ORD and hypochromism at pH 1, 7, and 11.5 of seven trinucleoside diphosphates: A-A-A, A-A-C, A-A-U, G-A-U, A-G-U, G-G-U, and G-G-C (Fig. 5). Again, by designating those with hypochromism less than 3% as “umtacked,” greater than 6% as “stacked,” and from 3 and 6% as “partially stacked,” these authors found that the first five compounds are stacked at pH 7,and A-A-C and A-A-U also stacked at pH 11.5. G-G-C is the only compound to show much stacking at pH 1. More recently, Cantor and Tinoco (82) reported the ORD of four additional trimers, A-C-U, G-C-C, A-G-Cp, and G-A-Cp at pH 7. The last two compounds contain 3’-terminal phosphates, which

220

260

300

340

220

260 300 X (rnp)

340

220

260

300

340

FIG.5. Molar rot.ation per base of seventrinucleoside diphosphates at pH 7 and ionic st,rength 0.1 (0.01 M ph0sphat.ebuffer plus 0.08 M KCIO,) (81).Solid line, experirnent,al; broken line, calculated from Eq. (14).

ORD

AND

CD OF NUCLEIC

245

ACIDS

seem not to affect greatly the optical properties of the trimers. This is also in accord with the finding of Vournakis etal.(83) that the ORD profiles of A-A-Cp and A-A-C are in excellent agreement. Inoue et al.(8%) also reported the ORD and hypochromism at pH 1, 7, and 11 of seven trinucleotides all ending with Gp : A-A-Gp, A-C-Gp, C-A-Gp, C-C-Gp, U-A-Gp, A-U-Gp, and U-U-Gp. The first five compounds are stacked at pH 7. Trimers having one or no protolytic base (uridylyl or guanylyl residue) are still stacked at pH 11, whereas those having two or three protolytic bases are unstacked at t,his pH. C-C-Gp appears to be the only compound to exhibit appreciable stacking at, pH 1. Inoue et al.(83a), however, believe that the S’-terminal phosphate is a significant factor in the formation of ordered conformations in oligonucleotides, and consequently to their ORD and hypochromicity, although its importance may diminish with increasing chain length as the ratio of base to phosphate approaches unity in polynucleotides. Like dinucleoside phosphates, sequential isomers of trinucleoside diphosphates show very different ORD profiles, for example, G-A-U and A-G-U at pH 7 (Fig. 5 ) .Ideally, all 64 trimers should be prepared and their ORD measured, but this would be a formidithle task for any single laboratory. Enthusiasm for studying higher oligomers becomes even less when one realizes that there are 4N possible oligomers of chain length N . Thus, Cantor and Tinoco (82) have developed a simple semiempirical approach for calculating and predicting the optical properties of these 64 trimers from those of their constituent dinucleoside phosphates. This is based on the assumption that only nearest-neighbor interactions between nucleoside bases contribute to the optical properties (cf. Felsenfeld and Hirschman, 8%).

NEAREST-NEIGHBOR INTERACTIONS The molar rotation, [MI,at any wavelength of a dinucleoside phosphate, NpN’, can be written as [MNN’] = [ M N ]

+

f [MN ] INN!

(11)

where I N N is the contribution of the nearest-neighbor interactioIi to the rotation. Since the molar rotation of a nucleoside and its corresponding nucleotide are usually very close and small compared to the rotation of a dinucleoside phosphate, one can take the average of the molar rotation of pN’ and N for [n/!,,], for instance. Likewise, for a trinucleoside diphosphate, we have [ M N N N J t ]= [ M N I

f

[MNt]

+

[2vN’t1

+

JNN~

f JN’N”

+

JNS

,

(12)

Here again, J , and J N f N , ,are the terms for nearest-neighbor interactions,

246

JEN TSI YANG

AND

TATSWA

SAMEJIMA

but J ,,= 0, since the effect of the next-nearest-neighbor interaction is ignored in this treatment. If the bases in the dimer and trimer have the same relative conformation, J N N ~ = I ,and J N 1 = I N t N . Combining we have Eqs. (11)and (12), [hf

N ]

= [.$fNN ]f LiTfNtNtC]-

[nrNr]

(13)

or, in terms of mean residue rotation, [m], [ ? I t N N N ~= l

(a[m,iY~]

+

2[rnN,N,tI

- [,nxrI l/3

(14)

Ignoring the end effects, Eq. (14)can be extended to higher oligomers and polymers :

where X N N ~ is the mole fraction of NpN’ and XN of nucleoside N. If the sequence of a polynucleotide is random, we can further assume that XNN’ = X N . X N ~ ,and Eq. (15) becomes A

A

For a homopolymer, Eq. (16)is greatly simplified as [mpolgrner] = a[mNN]

-

[mX1.

(17)

The calculated ORD, using Eq. (14)for the trinucleoside diphosphates, agreed very well with the experimental values (Fig. 5 , broken lines), thus supporting the theory of the nearest-neighbor interactions between bases. It also gives a strong argument for similar conformation of bases in dimers and trimers and makes it possible to predict the ORD of oligomers from their constituent dinucleoside phosphates. While this practical approach promises to become a new tool in determining the sequence of oligonucleotides, its usefulness may be limited to trimers and perhaps tetramers because differences among the ORD of longer sequence isomers will probably become very small or vanish. It should also be pointed out that the work of Inoue et al.(8%)on seven trinucleotides shows a rather poor agreement between experimental ORD and that based on Eq. (14), assuming a negligible effect of the 3’-terminal phosphate. These workers cannot draw any definite conclusion as to whether the disagreement is greater in trinucleotides than in trinucleoside diphosphates (cf. Fig. 5). They further suggest that the dinucleotide rather than dinucleoside phosphate should be used as a unit structure in the nearest-neighbor calculations, ifthe effect of the 3’-terminal phosphate on the ORD is not small in

ORD AND CD O F NUCLEIC ACIDS

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oligonucleotides. Equations similar to Eq. (14)can easily be obtained for the mean residue rotation of a trinucleotide and a trinucleoside diphosphate. The importance of this terminal phosphate effect must be clarified by future investigations. In the case of polymers, the use of Eqs. (15), (IS), and (17) requires that the rotatory contributions of the nearest-neighbor bases, -NpN'-, can be approximated by those of the corresponding dinucleoside phosphates, NpN'. This again must await more experimentation. measured the ORD of a series of oligoadenylic Michelson et al.(84) acids with a degree of polymerization varying from N = 2 to 12 and also one polymer with N = 200. At pH 6.75 the amplitude between the first peak (at 277 mp) and trough (at 255 mp) increased gradually with chain length and more than doubled for N = 200 over N = 2. Note that Eq. (17) actually predicts that the mean residue rotation of a homopolymer is about twice that of its dimer. At pH 4.86, the increase in the amplitude was also gradual between N = 2 and 12, but a drastic difference in the amplitude was observed for the polymer, which was about &fold that for N = 2. This is undoubtedly due to a transition of the protonated double-stranded, helical form of poly A, known to exist below pH 5 (see Section V, B). The ORD of di-, tetra-, and hexamers of adenylic acid, PA)^, PA)^, and (pA)e, at neutral pH over the temperature range of 5 to 85" were also reported by Poland et al.(85). Their results can be accounted for by a theory of cooperative stacking of bases for the single-stranded oligomers, which shows that the stacking in this case is only slightly cooperative. 86) corroborated well the The CD studies of oligoadenylic acids (57, ORD results of Michelson etal.(84). The general spectrum is the same for all the oligomers snd high polymers, as illustrated by the trimer, heptamer, and polymer at neutral pH in Fig. Ga. The intensity of the CD bands are an order of magnitude higher than, and entirely different in profile from, that of the monomer AMP. As the degree of polymerization increases, the positive band becomes broader arid increases in intensity more rapidly than does the negative band. There is also a general shift toward lower wavelengths with increasing degree of polymerization. As expected, the intensity reduces at elevated temperatures as a result of unstacking of the bases. The mean residue rotational strength of the positive band is also gradually enhanced with increasing number of residues (Fig. 6b). In contrast, the positive rotational strength in acid solution (pH 4.5) shows a sudden increase a t about the level of the heptamer and the sharp increase approaches a constant plateau at about 15 residues. These results support the conclusion that oligoadenylic acids and poly A at neutral pH adopt single-stranded conformations stabilized by base stacking, whereas in acid solution a double-stranded, helical conformation can be assumed only by oligomers larger than the heptamer. Brahms etal.(57) also determined the

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FIG.6. (a) CD per base of ApApA, A ~ ( A P ) ~and A , poly A at p H 7.4in 0.1 M NaCl plus 0.01 M Tris and at various temperatures (57). Left, curves 1-6‘: -2,0.5, 4.5, 18,25, and 47 C.Center, curves i4 : -2,0.5,8, 18.5,32, and 40 C.Right, curves 1-6: - 2 t o 6, - 17,34,42,57, and 74 C;curve 7,quaternary ammonium salt of p l yA in 98% ethanol at 0 C;and curve 8, AMP at 0°C. (b) Rotational strength of the positive band of oligo- and poly A as a function of the Open circles, p H 4.5; filled circles, pH 7.4. Temperature: degree of polymerization, N (67). about 0 C.

thermodynamic parameters of these compounds and concluded that thermal ‘Ldenaturation”of these single-stranded structures is largely a noncooperative process, whereas that of the double-stranded ones in acid solution is cooperative.

ORD

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249

Adler et al,(8’7) studied iu detail the ORD and absorption of both oligomers and polymers of CMP and dCMP at neutral pH over a wide temperature range. They concluded that ribosyl oligo- and polymers display greater rotatory power, hypochromicity, and heat stability than do the comparable deoxyribosyl compounds. Comparison of the two oligomer series further showed that asymmetric macromolecular structure was attained more readily, as a function of chain length, in the oligoribonucleotides. Detailed CD studies of oligocytidylic acids and poly C have also ) . these compounds at neutral pH possess a singlebeen reported ( 8 7 ~Again stranded, stacked-base conformation, but in acid solution (pH 4.0) the double-stranded, helical structures begin with the heptamer. In contrast with the above natural 3’-+ 5’ oligocytidylic acids, a series of 2' 4 5 ’ oligocytidylic acids at neutral pH showed no marked temperature dependence for the CD spectra, suggesting the lack of significant stacking between the bases in this case ( 8 7 ~ ) . Simpkins and Richards (87b) reported the spectrophotometric titration, hypochromism, and ORD ofoligouridylic acids (up to the undecamer) and also of poly U and concluded that these compounds are devoid of temperature-dependent base-stacking interactions at 20 Cin 0.1 M NaCl. The Cotton effects of the oligomers (from dimer up) and polymer are closely similar; their shape and the magnitude of the mean residue rotation show little dependence on chain length.

V. Synthetic Polynucleotides A. Poly A, Poly U, and Their Complexes Studying synthetic polynucleotides will help us understand the more complicated nucleic acids which contain a multiplicity of bases and base sequences. (For reviews on polynucleotides, see refs. 88-95.) Like the oligomers, the general ORD profile of all polynucleotides (except poly I) consists of two peaks and one trough centered around the 260-mp absorption band, and their CD has a positive band (on long wavelength side) and a negative one above 22&230 m p . Additional Cotton effects are also observed below 220-230mp. and Lamborg etal.(32) first reported the ORD's Sarkar and Yang (94) of poly A and poly U above 200 mp, and, more recently, Ts'o et al.(80) have extended the measurements to about 190mp. Holcomb and Tinoco (95) also presented the ORD of poly A in neutral and acidic solutions at various temperatures. Figures 7 and 8a illustrate the ORD spectra of these polymers and their complexes ; Table IV lists the pertinent numerical values of the ORD of various polynucleotides. The multiple Cotton effects

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TABLE I V THE COTTOX EFFECTS OF POLYNUCLEOTIDES Peak 1 Polymer and solvent

Poly A (0.15 M KF, pH 7.5) Poly U (0.15 M KF, pH 7.5) Poly A . poly U (0.15 M KF, pH 7.5) Poly C (0.1 M NaCl plus 0.01 M cacodylate, pH 7.0) Poly Go (0.1 M NaC104 0.001 M cacodylate, pH 7.0) Poly G . poly C (0.1 M Tris, pH 7.5) poiy ~ ( A - T )(0.15 n l NaCl 0.015 M sodium citrate)

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all have a large peak at 282-286mp and trough at 252-260mp, followed by a small peak near 230-240mp. Their magnitude is reduced by the disruption of the secondary structure ofthe polymers at elevated temperature. Poly A at neutral pH is known to be single-stranded; small-angle X-ray scattering study indicates a molecule consisting of rodlike segments with a linear dimension of 3.4A per nucleotide residue (98). Thus, the melting curve in this case is rather broad (Fig. 7a, inset). The ORD profile of poly A in acid solution is similar to that at neutral pH, but the magnitude of the Cotton effects is larger at pH 4.55than at pH 7.5and a sharp transition occurs in acidic buffer (for instance, 0.1 M sodium acetate plus 0.1 M NaC1) with a T, of about 60"(94, see also 96). This agrees with the X-ray diffraction study of oriented fibers (99) and the small-angle X-ray scattering study (98), which show that poly A possesses a double-stranded helical conformation in acidic solution. The Cotton effects of poly U have a much smaller magnitude (except at 1-2’)than those of poly A. There is no significant change in rotation with heating at temperatures above 20",but a sharp increase in the magnitude of the Cotton effects occurs below 20"(Fig. 7b,inset). These results are in accord with conclusions drawn from optical, hydrodynamic and titration studies; poly U lacks any regular, cooperative structure at room temperature and thereby exhibits little stacking of bases (100). Such helical conformation at low temperature, however, is rather unstable, as witnessed by its low T,. The conclusion of Richards etal.(100) has been challenged by Michelson whose various optical studies of single-stranded poly U and Monny (IOOa), indicate a significant amount of helical structure arising from interplanar interactions. Simpkins and Richards (87’b), however, point out that the existence of a Cotton effect per se cannot be taken as evidence of stacking interaction as implied by Michelson and Monny (IOOa),since UMP also shows a Cotton effect. Simpkins and Richards (87’b, 100b)also attempt to reconcile this disagreement by suggesting that base-stacking in poly U is favored only in high salt solutions, e.g., above 0.5 M NaCI. This is also consistent with the CD evidence of stacking interaction of UpU in 4.7M KF even at 20 C(56). Poly A and poly U are known t.0 form two complexes, polyA poly U and poly A 2 poly U,when appropriate proportions of poly A and poly U respectively (IOI-lO4). are mixed at an A:U molar ratio of 1:1 and 1:2, The Cotton effects of poly A . poly U (Fig. 8a) and poly A . 2 poly U (not shown here) have a magnitude (per base) between t.hat of poly A and Both complexes show a sharp melting curve in agreepoly U (Table 111). ment with their double- and triple-stranded helical conformation, and the variation of the first peak (on the long wavelength side), parallels the

ORD

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253

ACIDS

hyperchromicity of t.liese complexes at, eicvated temperatures. The first trough also undergoes a shift t,oward the red with increasing temperat>ure. Comparison of the experimental ORD of poly A . poly IJ with that calpoly U indicates that the latter is located at longer culated from poly A wavelengths than the experimental curve (94).It implies that base pairing in a double-stranded helical conformation causes a shift toward the blue of the ORD of ribosyl polymers (see Section VI, B). Brahms (106, 106)and, more recently, Hashizume and Imahori (107) reported the CD of poly A, poly U,arid their complexes, some of which are also found forpoly A shown in Fig. 8b. Hashizume and Imahori (107) another large positive band at 220mp with a shoulder around 235mp and a negative one near 208mp. They also reported that the melting curves of poly A, poly U, and poly A .poly U were very close to the ORD results found that at pH 4.9, where poly A shown in Figs. 7 and 8.Brahms (105) is double-stranded, the positive band is much enhanced and the negative one reduced; this is accompanied by a blue shift of the CD spectrum. Poly U at neutral pH shows a CD profile very similar to that of poly A, except that its intensity is much less than that of poly A and its variation with temperature is also small (106). The CD of the complex of homopolymers, poly A poly U (Fig. Sb, curve .2), and the random copolymer, poly (A,U)(curve S), are very similar; the slight difference between these two curves may indicate a less ordered structure for the copolymer due to some random distribution of Poly (A,U)also shows a broader melting bases according to Brahms (106). curve (based on hyperchromicity) than poly A . poly U. The CD spectra of these two complexes have the same features, that is, a large positive band at about 262-265mp and a very reduced negative one at about 240-244 mp, whereas the CD bands of their components, poly A poly U, are located on the longer wavelength side (curves i and 4). This again indicates that the formation of the double-stranded helical conformation is accompanied by a blue shift of the spectrum. The CD of the alternating copolymer, poly (A-U),appears to be quite different from that of poly A . poly U (K. Imahori, private communication). One interesting feature in Fig. 8b is that the CD of poly A poly U and poly (A,U)is markedly reduced in intensity and the positive band is poly U (curves 1 and 4).[The broadened as compared with poly A and 4 (at 2’)occurs because poly U difference between curves 1 (at 14-15’) adopts an ordered structure at 2" and its CD is more intense than at Brahms (106) suggested that the antiparallel temperatures above lo0.] arrangement in the double-stranded poly A - poly U and poly (A,U) could account for their low CD intensities. Since the only difference between a parallel and an antiparallel double-stranded helix in this case is

+

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J E N TSI YANG A N D TATSUTA SAMEJIMA

that the orientation of the ribose residucs in the two strands is in the same direction in one case and opposite iii the other, it is hard to believe that this can be the cause of the observed reduction in intensities of the CD spectra. It seems more likely that interactions among bases in adjacent strands in addition to those among stacked bases of the same strands could be responsible for tfhemarked change in optical properties of these polymers.

B. Poly C, Poly G, Poly G Poly C, Poly I, Poly I Poly C, and Poly A 2 Poly 1 Fasman etal.(31) found that the ORD of poly C has a single Cotton Cantor etal. effect between 255and 350 mp (see also Ulbricht etal.(108);

FIG. 9. (a)ORD of poly C at pH 8.4 in 0.05 M sodium phosphate. From Ts o etul.(80). (bj CD and absorption of poly C at pH 4.0 (in 0.1 M NaCl plus 0.05 M acetate) and 7.5(in 0.1 M K F plus 0.01 Af Trisj (Vaj. Solid lines, CD at about 0°C; broken lines, absorption a t 25 C.

(108a)J; Sarkar and Yang (96) extended the measurements to 225 mp and observed another shoulder around 250 mp, thus indicating that poly C also has muItipIe Cotton effects with two peaks a t 293 and 250 mp and one trough at 267mp; more recently, Ts’o et at. (80) reexamined the ORD of poly C to about 200 m p (Fig. 9a). The magnitude of the first peak and trough (on the long wavelength side) for poly C is much larger than that of other polynucleotides, although the melting curve a t neutral p H was very broad and indicative of a structure for the single-stranded polynucleotide not highly ordered. In acid solution, e.g., p H 4.0-4.4, where poly C is believed to be a double-stranded helix (88, 109, 110); the magnitude of the first peak dropped and that of the first trough was enhanced. This is

ORD

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CD O F NUCLEIC ACIDS

quite different from the p H effect on poly A (Section V, A), where the magnitude of the peak and trough was larger a t p H 4.85 than a t neutral Since protonation of AMP does not alter its absorption spectrum pH (94). very much, although the spectrum of CMP is significantly changed in acid solution, protonation as well as conformational change (to a doublestranded helix) must be responsible for the difference. Sarkar and Yang (111)concluded that protonation of the cytosine base was a t least partially responsible for the reduction in magnitude of ORD for poly C in acid solution, since the same phenomenon has also been observed for DNA's and RNA's even in the absence of any conformational change. The Cottoil effects of poly C are shifted to the longer wavelength upon acidification instead of the blue shift characteristic of the formation of double-stranded helices of other polynucleotides such as poly A in acid solution. This must be attributed t o the drastic change in optical properties when cytosine is protonated. The C D and absorption spectrum of poly C at p H 7.5and 4.0 (Fig. 9b) fully substantiate this conclusion. At neutral pH, poly C has only one main positive CD band a t 276 mp, but in acid solution it has one positive band a t 287 mp and a negative one at 266 mp (86, 87a). Poly G was until recently very difficult to prepare; it aggregates easily in aqueous solution. Ulbricht elal.(108) reported that a t neutral pH and a total ionic strength of 0.15, poly G has an ORD with two peaks a t 266 and 222 mp and one trough at about 243 mp (the amplitude of the first Cotton effect on the long wavelength side was +39,400). The ORD changed little as the pH of the solution was lowered to 1,suggesting that the conformation of the polymer survived complete protonation. On the other hand, the ORD at pH 12.2 showed only a single, negative Cotton effect typical of the purine p-mononucleotides, indicating a complete loss of secondary strucobserved similar ORD for poly G a t p H 7.0(in ture. S. K. Arya (96a) 0.1 M NaC1O4 0.001 M cacodylate), except that the two peaks and trough were located at 271,224, and 249 mp (see Table IV), which were The several millimicrons longer than those reported by Ulbricht elal.(108). corresponding C D showed a large positive and a small negative band with extremes a t 260 and 237 mp and another extremely small positive one near 288 mp (9th). Fig'ure 10 shows the ORD of poly C poly C (prepared by synthesizing poly G on a poly C template), which has two peaks a t 276 and 22&230 mp and one trough at 245 mp (96). This polymer complex is believed to be a double-stranded helix and therefore has a very sharp, but irreversible, melting curve with a T, near 90"as contrasted with about 60" for poly A poly U (Fig. Sa). At elevated temperatures, both the peak and trough of the ORD reveal a large red shift together with a reduction in their magnitude. No CD of poly G . poly C is as yet reported. I<. Imahori (private

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communication) now observes a large positive CD band at 270mp and a small negative one near 235m p , but he is unable to detect any small band around 300mp (cf. Fig. 10). The ORD of an equimolar mixture of (G), and (C), differs from that shown in Fig. 10for the positions of the peaks and troughs. Ulbricht etal.(108) found that the first Cotton effect had a peak and trough at 287and 260mp, which were at longer wavelengths than those also found that increasing the ionic shown in Fig. 10. S. K. Arya (96~) strength of the solution resulted in a blue shift of the Cotton effects of the equimolar mixture of(G), and (CIn.We are tempted to suspect that there were fewer base pairings in the eyuimolar mixture of poly G and poly C than in polyG . poly C (see Section VI, B). The question of complex formation has been a controversial one. Fresco (118) indicated that there could be two complexes, 2 poly G * poly C and poly G . 2 poly C, whereas Pochon and Michelson (113) found only a stable 1 :1 complex, except that degraded polyC’ and poly G rapidly formed

ORD AND CD O F NUCLEIC ACIDS

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a complex of 2 poly G . poly C. Since the state of aggregation for poly G in aqueous solution is still not clear, this problem must be clarified before we can resolve the question of its complexes with poly C. Poly I, an analog of poly G, distinguishes itself from other polynucleotides by displaying two troughs and one peak between 240 and 300 mp instead of the familiar profile of two peaks and one trough around the 260-mp absorption band (96); it resembles the Cotton effects of the GMP gel (Section IV, A). The conformation of poly I, and thereby its Cotton effects, are known to be very dependent on the ionic strength of the 115). In low salt solution, e.g., 0.1 M NaCl, the Cotton solution (114, effects of poly I are relatively small and its melting curve is broad and indicative of little secondary structure. On the other hand, in high salt solution, e.g., 1 A l NaCI, where poly I assumes the triple-stranded helical the ), rotation of the peak near 260 mp is increased by conformation ( l l j an order of magnitude as compared with that in 0.1 M NaC1. Furthermore, its melting curve showed a sharp transition between 40 and 45",which reflects the cooperative melting of a highly ordered structure. Complex formation of poly I with poly C and poly A producing poly I . poly C and poly A . 2 poly I (116, 117)immediately inverts the ORD profile of poly I to one characteristic of other polynucleotides with two The peaks and one trough around t>he260 mp absorption band (96). melting of poly I . poly C occurs near 60" and is accompanied by a red shift. Poly A . 2 poly I melts around 40"with a blue shift; furthermore, unlike other polyribonucleotides it has n serond peak larger than its first one at room temperature.

C. Poly d(A-TI, Poly d,A Poly dT, Poly dC, Poly dA, and Poly dT Figure 11 (97) shows the ORD of poly d(A-T), a copolymer of alternating sequence that forms a double-stranded helix, which may be represented as poly d(A-T) . poly d(A-T), through the base-pairing between dA and dT in the opposing strands. Note that the second peak is about twice as large as the first one. This appears t o be true for all double-stranded polydeoxyribonucleotides as well as DNA's (see Section VI,A), whereas the polyribonucleotides always have a second peak close to zero rotat,ion. Poly d(A-T) has a reversible transition with a T, at about 60". Recently, Ts'o etal.(80) examined the ORD of poly d(A-T) from two sources: one synthetic as that used in Fig. 11 and one natural, isolated from Cancerantennarius. The Cotton effects for the two poly d(A-T)'s are very close. Those ofthe synthetic one are essentially the same as those shown in Fig. 11, except for some small differences in magnitude of the peaks and

258

JEN T S I YANG

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TATSUYA

SAMEJIMA

FIG.11.ORD of poly d(A-T) at pH 7.4in0.15M KF.Curves: 1,27-60 %’C; J 65 C; and ,!IJ81i"C. From Samejima and Yang (97).

troughs and in the fine splitting of the peaks. These workers also made measurements to about 200 mp and observed another trough and peak below 225 m p . The subtle difference between the natural and synthetic copolymers was the effect of temperature on the magnitude of the first,peak just before the onset of the melting. The synthetic copolymer had a smaller peak a t 282mp than the natural one, but its magnitude increased with t,emperature (between 20’and 50") up to the level of the natural one, whereas the natura1 one was not so sensitive to heating in this temperature range. This phenomenon is probably due to some difference in the secondary struct.ure of these two polymers at room temperature. The natural d(A-T)7L has been shown to be a straight-chain, double helix without branching or hairpin-like structures (118); it also contains about 3% guanine plus cytosine in its base composition, which is unlikely to have a serious effect

ORD AND CD O F NUCLEIC ACIDS

259

on conformation and ORD. The synthetic d(A-T),, however, has been known to have branches and hairpin-like structures through intrachain 120). Ts'o et al.(80) attributed the difference in tembase-pairing (119, perature response of the peak at 282inp prior to melting to this dissimilarity in structure of the two copolymers. The ORD of poly dA . poly d T (see Fig. 13,Section V, D), which like its counterpart poly rA . poly rU forms a double-stranded helix ( l d l ) , shows more complicated Cotton effects than that of poly d(A-T) (Fig. 11). At room temperature, the ORD curves of the two kinds of polymers are distinctly different, but above the T , they show much similarity. The T, of poly d(A-T) is also 5" lower than that of poly dA . poly dT (80). Recent X-ray study on fibers indicated that the structure of poly d(A-T), but not 123). Thus, poly dA - poly dT, is similar to that of the B-form of DNA (122, 1 I I I I I I

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AND TATSUYA

SAMEJIMA

the difference in ORD may be due to this difference in helical structure. An alternate interpretation is that the sequential difference in the constituent polynucleotide chains can give rise to a dissimilarity of the exciton splitting of the absorption bands and thereby the difference in ORD. The ORD profile of poly d(G-C) is reported to be more complicated than that of poly d(A-T) (97). The copolymer used, however, was suspected to contain more dG than dC; a reinvestigation of it seems warranted. Ts'o etal.(80) measured the ORD of poly dC, poly dA, and poly dT, and Adler ef al.(87) the ORD of poly dC. The shape of the curve for poly dC (Fig. 12) is quite similar to that of poly C (see Fig. 9),but the magnitude of its Cotton effects is only about 3040% that of the latter. This implies much less stacking interaction for poly dC than for poly C, a conclusion also supported by the finding of less than 1% hyperchromicity when the solution of poly dC is heated from 23 to 90".In acid solution, both the shape and magnitude of the ORD curves for poly dC and poly C are very similar, except that the second peak is positive for poly dC and negative for poly C. The pH of the single-to-double-strand transition at room temperature is 7.2 for poly dC and 5.7for poly C in 0.05 M sodium ions. Thus, the absence of the 2'-hydroxyl group seems to stabilize the helical structure of poly dC in arid solution. Ts'o et al.(80) suggest that the intramolecular hydrogen bonding of the 2'-hydroxyl group to the 2-carbonyl group greatly reduces rotational freedom around the glycosyl bond, thus enhancing the stacking of bases of poly C. The lower stability of the double-stranded helix ofpoly C in acid solution can be explained as follows : (a) the intramolecular hydrogen bonds greatly hinder participation of the 2-carbonyl group in interchain hydrogen bonding; and (b) the dissociation constant of the ribosyl cytosine group is lowered by intramolecular hydrogen bonding. Both effects will tend to lower the transition pH ofpoly C as compared with poly dC. The difference in ORD between poly dA and poly A at neutral pH is very striking (cf. Fig. 7a). The prominent first peak and trough for poly A between 250and 320mp are much reduced in the case ofpoly dA. But in acid solution their ORD curves are very similar, except that the magnitude of the Cotton effects is smaller for poly dA and its first peak shows a blue shift of about 10 mp. The transition pH for poly dA was about 1.5units lower than that required by poly A, suggesting that the poly dA helix in acid solution is less stable than that of poly A (80). This may be ascribed to the intramolecular hydrogen bonding of the 2'-hydroxyl group to the base, which enhances the base stacking of poly A as compared with poly dA. The ORD's of poly dT and poly U are much the same at room teniperature and in the absence of Mg*+ (cf. Fig. 7b). But unlike poly U, the ORD ofpol9dT in solutions containing Mg2+ was found to be insensitive

ORD

AND

CD OF NUCLEIC

26 1

ACIDS

to temperatux siid to lack :my tr:nwition at low tempertbture, thus indicating little stacking interaction arid secondary structure (80).

D. Hybrids of Polyribonucleotides and Polydeoxyribonucleotides The observation that the ORD of double-stranded polydeoxyribonucleotides always has a much larger second peak than the first one and the has led Ts’o el aE.(80) t o opposite is true for polyribonucleotides (10) examine the hybrids of polyribonucleotides and polydeoxyribonucleotides. Figure 13 includes the ORD of all four permutative pairs of 1:1 mixtures of poly r h. poly dT, poIy dA ’ poly dU, poly rA poly rU and poly dA poly dT (the last two are included for the sake of comparison). The positions of the peaks and troughs of these curves are roughly the same, but their magnitude is quite different (poly dA . poly d T also has more than one peak in the 260-290mp region). The two hybrids aIso have sharp 1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

1

I

I

1

I.,

20

.

I

15

10

5

0

5

-5

x

T

E u

-10

-15

-20

- 25 1

1

1

1

1

1

I

A (mp)

FIG.13. ORD of ( 1 )polyrA . polyrU,(2)polydA . polyrU,(31polyrA . polydT, (4)poly dA * poly dT (enzymatic) in0.05 M NaCI04at 20°C. FromTs’o etul. (SO).

and

262

JEN TSI YANG

AND

TATSUYA

SAMEJIMA

melting curves with T,’sof 41.5" for poly dA . poly rU and 59"for poly rA . poly d T in 0.05 M NaC104 (pH 7.0), as compared with 51"for poly rA poly rU and 61.5" for poly dA . poly dT under the same conditions. We have already mentioned that the ORD of double-stranded deoxyribosyl complexes has a second peak much larger than the first one. But the formation of the hybrids brings an increase in the first peak as compared to the original deoxyribosyl complex and a decrease in the second peak as compared to the original ribosyl complex, resulting in an ORD profile approaching that of the double-stranded rihosyl complex. This phenomenon cannot be attributed to the rotatory cont,ributions of ribose and deoxyribose and these differences must reflect different types of helical structure (see Section VI, C for further details).

-

VI. DNA and RNA A. Similarities and Differences between DNA and RNA Accurate ORD results of nucleic acids have been accumulating since 124-127). DNA and RNA from vari1963 (see, for example, 10,32,59,97, ous sources have similar ORD profiles of two peaks and one trough centered around the 260-mp absorption band, as typified by the examples shown in Fig. 14.Table V lists the pertinent numerical values of several DNA's and RNA's. ForDNA, the first peak on the long wavelength side occurs at 290mp, the first trough near the absorption maximum, and the second peak around 230 mp. These positions may vary a few millimicrons among the DNA's studied, but the general shape has been predicted by the theory of Tinoco (see Section VI, C). When measurements are extended below 220mp, a second trough is observed near 215mp and a third peak, much larger than the first one, can be located near 200mp. The ORD of RNA also has three peaks and two troughs between 190 and 300 mp, but its first peak is centered near 280 mp, about 10 mp less than that of DNA. The first trough again can be found near the 260-mp absorption maximum. The most striking difference between DNA and RNA is the relative magnitude of the first two peaks; invariably, DNA has a second peak that is about twice as large as the first one (on the long wavelength side), the reverse being true for RNA, whose second peak is always very small and has a rotation very near zero. An explanation for this dissimilarity between DNA and RNA is given in Section VI, C. Brahms and his co-workers (11, 14,86,106,106) reported in detail the CD of nucleic acids and polynucleotides. Their measurements were, however, limited to wavelengths above 220-230mp. They found a positive band on the long wavelength side and a negative one immediately following

OltD AND

CD OF NUCLEIC ACIDS

263

it on the short side with a crossover near the absorption maximum. This is true for a wide variety of nucleic acids, irrespective of their origin. However, DNA has two almost equally large (in magnitude) CD bands with a maximum a t about 272 mp and a minimum near 245mp. In contrast, RNA has a very large positive band centered at 265 mp, several millimicrons shorter than that for DNA, and an extremely small negative one around 252mp, a few millimicrons longer than that for DNA. It should be noted also that the intensit,y of the first positive band is stronger in RNA than in DNA. When measurements are extended below 220 mp, both DNA and RNA ?how additional negative arid positive CD hands (Fig. 14), which are undoubtedly related to the absorption h:Lnd near 200mp.

TABLE V THE COTTON EFFECTS OF NUCLEICACIDS".* T,

Substance or source DNA 1. Aerobacter aerogenes 2. Bacillus megaterium 3. Esckichia coli 4. M~cobacterium tuberculosis

5. Serratia marceseens

6. T2 phage 7. Calfthymus 8. Salmon sperm

9. Poly d(A-T) 10. Poly dG * poly dC

Peak 1 Temperature ("C) X (mp) [a]

27 96 27 87 27 90 27 96 27 96 27 90 27 96 27 96 27 85 27 90

290 292 290 292 290 292 290 292 290 292 290 292 290 292 290 292 283 287 310 320

+2310 t-1200 +1820 +790 +2200 1080 4-2610 +I260 4-2410 +1170 +1380 +770 +1910

27 90 27 90 27 90

280 292 283 290 280 290

+3140

+

+850

+1930

+850 +2450 +900 +250 -130

Trough 1

Peak 2

h (m#)

Iul

X (mp)

[ul

257 260 258 260 257 260 256 260 258 260 258 262 257 260 256 262 255 263 285 295

-2920 -2770 -860 -1180 -2230 -2590 -3690 -1380 -2540 -1420 -2410 -2120 -1920 -1980 -2000 -1900 -5780 -3730 -830 -600

225 225 225 225 225 228 225 228 228 228 235 230 228 230 228 228 236 235 264 270

+4650 +615 +4480 +2070 +4200 +1640 +4360 +4130 +1450 +3900 +lo60 +3730 +660 +3920 +950 +5480 +2770 +2520 +I610

251 258 253 258 252 260

-2770 -2040 -3220 1790 -3500 -2580

227 227 225 225 228 228

-160 -750 +240 -120 +120 -750

+870

Trough 2 X

(mr) 217

[PI

Peak 3 A (mp)

[a]

From 290 Literature mp peak value ("C) ("C)

+2980

-

-

215

f3.550

-

-

215

+3090

91

93.5

82

85

-

-

86

90.5

215 215 217

+3390 -1160 +2680

92

97

90

93.5

80

83

84

87

85

87.5

63.

65

-

-

218

+3150

216

f2830

-

-

-

-

215

+2940

-

-

-

-

247 242

-4590 -2190

217 215 215 210 217 210

-1360 -1030 -1430 -1380 -1150 -3100

-

RNB 11. Yeast tRNA 12. Tetrahynena rRNA 13. Rat liver

+800 +3060 +730 +3400 +980

-

Data taken from ref. (87). M K F (pH 7.4), except substance 1, 0.01 M NaC1; substance 3, 0.13 M KF; substance 9, 0.15 M NaCl 10, 0.02 M NaCl 0.01 M phosphate buffer (pH 7.1); substance 12, 0.15 M KF 0.01 M phosphate buffer (pH 7.0). c From 283 m# peak. a

6 Solvents used: 0.15

+

+

+ 0.015 M sodium citrate; substance

ORD AND

265

CD O F N U C L E I C ACIDS

A new negative CD band around 300 mp that had escaped detect>ion by previous workers because of its extremely low intensity (128, 129)has recently been reported. The minimum of this band for DNA is located near 310 mp and its magnitude is about 1/500 of the first maximum; the minimum for RN A is found between 295 and 298 mp, and its magnitude is 1/10 to 1/30 of the first maximum. None of the synthetic polynucleotides or their complexes, however, shows such a negative C D band near 300 mp, with the exception of poly I, which seems to have a very small one. Since the intensity of the first positive band is much stronger than that for nucleic acids, it could be that an adjacent small negative band is submerged and escapes detection. K. Imahori (private communication) has informed us that the CD of poly (A-U) and poly (G,C), but not poly G poly C, did show a small negative band between 290 and 300 mp.

-

1. TEMPERATURE DEPENDENCE

The Cotton effects of the nucleic acids are conformation dependent (Fig. 15). The first and second peaks of the ORD for DNA are obviously temperature-dependent; so is the first peak of RNA (the first trough in both cases seems not very sensitive to temperature). Since the denaturation of DNA is known to be a cooperative phenomenon, the Cotton effects of DNA remain little changed below its T ,and thereafter fall drastically with further increase in temperature. A plot of the rotation at 290 mp, for instance, versus temperature would show a sharp helix-coil transition, which parallels closely the hyperchromicity of the 260-mp absorption band ; indeed, the T ,as determined from both methods agrees very well. RNA, on the other hand, is known to have a broad melting curve; accordingly, its Cotton effects also fall gradually with increasing temperature. It should be noted also that a t elevated temperatures the peak and trough of RNA undergo a red shift of about 2-12 mp. Since the base chromophores in nucleic acids give rise to the observed Cotton effects, the interactions among various bases are expected t o contribute different rotations. Empirically, a simple linear relationship has been found between the rotation at the 290-mp peak and the base composition of DNA in terms of the content of guanine plus cytosine (Fig. 16) (97) : [a]290 =

26.5 X mole percent (G

+ C) + 550

(18)

15.4 X mole percent (G

+ C) + 220

(19)

for native DNA’s and [a1290=

for heat-denatured DNA’s. No comparable C D data on DNA’s are available, hut sudi linear relntionships shoultl cxist . Two notahlc exreptioris in arid poly (l(A-T). The formrr contains hylroxyFig. 10 arcthe T2 phtigc

I

I

I

I

Salmon sperm DNA

I

I

200

250

,

A

I

300

1

Yeast iRNA

,

I

I

350

200

250

I

300 X (mp)

(a) I

I

I

I

I

1

I

I

I

I

I

I

I

42-

o -2

\

-

DNA

- -4I

I

I

2

I I

J

I

I

I

I

6I

4-

2-

0

I

I

I

I

I A (mp)

(b) F1c:. 15a a11db.

I

A

I

350

(

ORD AKD CD O F NUCLEIC

ACIDS

267

mcthylcgtosine which may not necessarily contribute the same rotation as that of cytosine. Poly d(A-T) has a, much larger rotation than that read from the straight line; unlike thc irregular sequence of bases in DNA, the regvlarity of the alternate sequence in poly d(A-T) may have enhanced the Cotton effects and thus the rotation at 290 mp (cf. Fig. 11). Also, poly d(A-T) is an exception in the melting curve reported by Marniur and Doty (130). Equation (18) provides a simple means for determining the base composition of DNA having standard bases, adenine, thymine, guanine, and cytosine. It is complementary to other analytical methods such as chromatography (131), CsCl density gradient (232, I$$), absorbance ratio of 260 t o 280 mp a t pH 3 (134), bromination (135), and melting temperature (130). It has the advantage over the T,-technique in that only a single rotation a t room temperature is measured, thus avoiding the difficulties associated with heating (e.g., turbidity) that often make measurement difficult. OF PROTONATION 2. EFFECT

Although the Cotton effects of nucleic acids as well as polynucleotides have been demonstrated to be conformation dependent, protonation of the nucleoside bases can also change the magnitude of these Cotton effects without disruptbig the secondary structure of the polymers (111). For instance, the rotation at the 290-mp peak for calf thymus and sperm whale DNA's is reduced about 10% when the p H of the solution is lowered from 7 to 4,whereas the second peak near 230 mp and the trough near 260 m p are almost identical a t the two pH's. Further lowering of the p H of the solution t o 3,however, did reduce both the peaks and the trough; the rotation at the 290-mp peak a t p H 3 is about half that a t pH 7.In addition, another small Cotton effect appears near 270 mp, its peak and trough being levorotatory; the rotation of the peak is close to zero. The effect of p H on the multiple Cotton effects is more drastic for RNA than for DNA. For instance, the rotation a t the 280-282 mp peak for several RNA's may decrease about one-half from p H 7 to 4; this is accompanied by a red shift of this peak by several millimicrons. Unlike DNA, however, RNA does not show a new Cotton effect between 265 and 275 mp, FIG.15. (a) ORD ofsalmon sperm D N A and yeast tRNA at pH 7.4in 0.15 AT KF and a t various temperatures. Left, curves 1-4: 27",80", 85", and 96°C. Right, curves 1-4: 27",55",go", and 27"after slow cooling from 90°C. From Samejima and Yang (97). (b) CD ofcalf thymus DNA and tRNA a t p H 7.4and a t various temperatures (14). Upper, DNA in 0.01 M NaCI, 0.01 M Tris, and 0.001 M EDTA: curves 1 ,20 C;2, 45 C;3,heated to boiling temperature for 10-15min and cooled rapidly; and 4,80 C. Lower, RNA in 0.15 M NaCl, 0.01 M Tris, and 0.001 M EDTA: curves 1-4,22",45", 70", and 80°C.

268

JEN ‘PSI YANG

AND

TATSUYA

SAMEJIMA

FIG.16.The relationship between therotation atthe290-mppeakandthe (guanine cytosine) content ofDNA. Symbols: 1, Mycobacterium tuberculo sis; 2, Serratia mrcescens; 3,Aerobacter aerogenes; 4,Escherichia coli; 6,salmon sperm; 6, calfthymus;7, Bacillus ,o megaterium; 8,T2 phage;and 9,poly d(A-T).From SamejimaandYang (97).

+

G + C (mol%)

even at a pH as low as 2.5(32). Difference absorption spectra ofnucleic acids at pH 4 with a pH 7 solution as reference show almost no change at the 260-mp absorption maximum, but a hyperchromic effect occurs around 280-290mp. A similar study of poly d(A-T) shows no such pH effect,but the Cotton effects of poly I . poly C are markedly changed at low pH (111). In Section 111,A we have already mentioned that the Cotton effects ofthe adenine derivatives show little changes even at pH 2,but marked displacement (red shift) of the Cotton effects are observed for the cytosine derivatives upon protonation. It is thus stipulated that protonation of the base cytosine, not adenine, is responsible for the observed changes in ORD. Since DNA is known to be stable at pH 4 (in the presence of salt), this effect of protonation cannot be associated with any gross conformational change of the polymer. The same may be true for RNA’s such as yeast tRNA (111). Similar conclusions have now been reached by Zimmer et al.(136), who

269

ORD AND CD O F NUCLEIC ACIDS

studied the protonation of Xtreptonzyces clwysomallus DNA and found that the new Cotton effect for this (G C)-rich DNA began to appear at pH below 4, when the cytosine base is 3040% protonated; the peak at 260262 mp has a positive rotation, and the two troughs are located at 276-279 and 245-247 mp.

+

B.

Single- versus Double-Stranded Structure

The differences in the ORD arid CD of DNA and RNA are an intriguing question, and their origin is still not fully elucidated. Two possibilities may be considered. First, the major difference in chemical composition of the nucleic acids is the constituent sugar, deoxyribose in DNA and ribose in RNA. This, we believe, plays an important role in determining the geometric arrangement of the polynucleotide chains (see Section VI, C). Second, most DNA molecules form double-stranded helices, whereas the RNA molecules are usually single-stranded and they may adopt hairpinlike or loop-like models. Thus, there is a difference in the conformation of nucleic acids that could account for the observed differences in the ORTI and CD profiles. This question can be clarified by studying RNA’s having rice the double-stranded helical conformation such as reovirus R,NA (137), dwarf virus RNA (RDV-RNA) (138, 139), cytoplasmic polyhedral virus RNA ( I @ ) , and the replicative form of MS2 bacteriophage RNA (141). Figure 17illustrates the ORD and CD of RDV-RNA. Evidently, this RNA possesses the same profiles as a single-stranded RNA; the second peak of the ORD is much smaller than the first one (on the long wavelength side) and the negative CD band is extremely small, unlike the ORD and CD of DNA. Thus, we can reasonably rule out the second possibility that singleversus double-stranded conformation of the nucleic acids is responsible for the described differences in the ORD and CD profiles. The intensity of the positive CD band for RDV-RNA in Fig. 17 is much stronger than that of any ot,her nucleic acids studied. This undoubtedly can be attributed to the high degree of base stacking in this polymer molecule. Raising the temperature of the RDV-RNA solution also causes a sharp, irreversible helix-coil transition just as does the “melting” of the double-helical DNA (139). One interesting finding in Fig. 17 is that the extremes of the positive and negative bands are located at 261 and 235 mp, several millimicrons shorter than the corresponding ones for a single-stranded RNA. I t strongly suggests that the formation of a doublestranded helix of RNA results in a blue shift, albeit small, of the CD bands and the corresponding ORD profile. In Section V, A,we showed that the formation of poly A . poly U from the two homopolymer components also results in a blue shift of the CD bands (Fig, Pb). The ORD in this case

270

JEN TSI YANG

I I

I

I

200

250

300

350 200 X (mp)

AND

TATSUYA

I

I

250

300

SAMEJIMA

FIG.17.ORD and CD ofricedwarfvirus RNA in0.01 M SSC solutions at various temperatures (139).Curves: I ,24"; 2,93"; 3,’24 C after slowcooling from93 C.

(Fig. Sa), however, was less clear-cut; the first trough of poly A . poly U was located at a shorter wavelength than either poly A or poly U, but the first peak did not show a blue shift. On the other hand, the ORD of the double-helical poly G . poly C is shifted toward the long wavelength side when the complex is disrupted upon "melting" (see Fig. 10). These results together with the recent theoretical calculations of the ORD of RNA (see Section VI,D) have led Hashizume and Imahori (107) to suggest that the location of the CD band at 261 mp reflects the presence of a perfect double-stranded helical conformation of the RNA molecule. (Note, however, that the crossovers of both poly A . poly U and poly G . poly C are located near 265mp. It remains to be seen whether the exact position of the crossover is an accurate and sensitive indicator for detecting such helical conformation.) These workers further stipulated that the separation of paired bases through the breaking of hydrogen bonds causes a red shift of the CD bands, whereas the unstacking of the bases markedly reduces the int,ensity of the CD bands. Similar conclusions were also reached by Adams et al.( l 4 d who ) , found that the crossover of the ORD of yeast tRNALeuwas shifted from 260.5to 263.3mg and the CD maximum from 260.3to 262.3 m p on going from the biologically active to the inactive form. [The biologically inactive form is considered to have nearly the same amount, of

ORD

AND

CD O F NUCLEIC

ACIDS

271

secondary structure as the active form; for further details, see Fresco et al.(143), Muench (144), Sueoka et al.(145).] ddams et al.(14% at) tributed the observed small shifts to the disruption of several base pairs in the RNA molecule. These shifts are also accompanied by a very small decrease in the rotational strength of the Cotton effects, indicating some degree of unstacking during the conversion of an active molecule to its inactive form. It is, however, not possible at present to decide whether the unpaired bases entered into single-stranded stacking or unstacking conformations. Neither can we speculate how the conformation of other stacked bases is affected by the breaking of a few paired bases. We simply do not have the detailed knowledge about the effect of base composition and sequence of RNA on the position, shape, and magnitude of its CD and ORD. But it is clear that both techniques are potentially very powerful tools for studying the fine details of the RNA conformation in solution.

C. Base

Tilting

The Cotton effects of the derivatives of each purine or pyrimidine are very similar, whether the constituent sugar is ribose or deoxyribose (Section 111,A). But data on polynucleotides as well as nucleic acids clearly demonstrate the pronounced effect of the 2’-hydroxyl group of the pentose on the ORD and CD of these polymers. We are led to believe that differences in conformation of the polyribonucleotides and polydeoxyribonucleotides rather than the configuration of the 2’-carbon atom of the pentose give rise to the striking differences in the Cotton effects of ribosyl and deoxyribosyl polymers. Since we can eliminate the explanation based on single- versus double-stranded structure, the alternate possibility is then the difference in geometry of the stacked bases between DNA and RNA. It may be instructive to consider the known structures of the double-stranded nucleic acids. Figure 18 illustrates the helical models of the A- and B-form of DNA, RNA, Table VI lists the pertinent parameters and the hybrid DNA.RNA (146). for all four models. Clearly, the mode of stacking is quite different for the A- and B-form of DNA. Furthermore, the sugar rings in the A-form are incorporated into the main helical chain, but those in the B-form are situated radially from the helical axis. Another salient feature is that the pIanes of the base pairs are almost perpendicular to the helical axis for the B-form, but tilted by about 20 degrees for the A-form. The distance between the helical axis and the paired bases also differ in the two forms (see Table VI).Of particular interest is the finding that the structure of double-stranded RNA or of DNA.RNA hybrids is very similar to that of the A-form of DNA. The replacment of thymine in DNA by uracil in RNA has no influence on the molecular structure, since the DNA from PBS

272

JEN TSI YANG

AND

TATSUYA

SAMEJIMA

. ,

I

I IA)

(8)

(Cl

IDl

FIG. 18.Doublestranded helical modelof nucleic acids. (A) A-form of DNA; (B)B-form of DNA; (C) RNA; (D)hybridof DNA and RNA. Symbols:0 , oxygen; 0 , phosphorus; 0 , nitrogen; 0, basepair. From Tsuboiand Higuchi(146).

2 bacteriophage, which contains uracil has the same structure as normal DNA’s (151). Indeed, model building of the RNA molecule shows the interference of the Z'-hydroxyl group of the ribose resulting in formation of a double helix similar to the B-form of DNA. The successive bases in this case are tilted or twisted (that is, the base pairs are not coplanar) to accommodate such steric hindrance. We therefore suggest that the mode of base stackmgs, the orientation of the sugar rings, and the orientation of the base pairs with respect to the helical axis are responsible for the observed differences in ORD and CD between DNA and RNA. [Atpresent the

ORD AND

CD OF NUCLEIC

273

ACIDS

TABLE VI THEHELICALSTRUCTUREOF DOUBLE-STRAKDED NUCLEIC ACIDS* DNA Measurement Pitch ofhelix Number of residues per turn Inclination of base pair from horizontal line Distance between axis of helix and base pair Orientation of PO;: (a) Angle between axis ofhelix and 0-- - -0 line (b) Angle between lhe helical axis and the bisector of angle L OPO

A-forms 28.2A 11 ’LO"

4 i

55"

B-ford

3 3 . 7 A 30.5A 10 or 11 10 2"

15"

0

4-5 K

.55"

3 65"

3 65"

li S&

DNAdlKA hybridsd 28 A 11

20"

4A

70"

6606(in DSA

40"

chain, 65"; in RNA chain, 70") 55"e(in DNA chain, 70"; in RS.4 chain, 40")

____

*From Tsuboi and Higuchi (146). a Fuller etat. (147); Sutherland and Tsuboi (14Ta). Langridge etal.(148, 1.68~). Sat0 etaf (158); Fuller P t al.(148); Arnott etal.( 1 4 9 ~ ) . Milman etal.(150); Higuchi eta/.( 1 6 0 ~ ) . e These values are the effective orientation by superimposing the orientation ofboth the DNA and RNA chains of the hybrids.

exact dimension of the double-stranded RNA remains somewhat uncertain, that is, whether it, has 10 or 11 base pairs per turn. The RNA helix in solution is even Iikely to have 12 base pairs per turn and, therefore, less tilting, probably 10 degrees, than that listed in Table VI (P. 0. P. Ts'o, private communication). This will not alter our working hypotheses (see below) .I We further speculate that the same rules apply to single-stranded as well as double-stranded polymers. Our working hypotheses for nucleic acids in aqueous solution can be summarized as follows (poly I is excluded in this discussion since it has a n inverse profiIe of the Cotton effects; see Section V, B). 1. The relative magnitudes of the two peaks of ORD on the long wavelength side and of the corresponding positive and negative CD bands of polynucleotides as well as nucleic acids are primarily determined by the geometry of the stacked bases, which in turn is influenced by the presence or absence of the 2'-hydroxyl group on the pentose. 2.T he stacking of bases perpendicular to the helical axis would lead to a higher second peak (near 225-230nip) of O R D and a large negative CD h n d as in DNA. Tilting of the Imes woultl reduce the second peak nntl

274

JEN TSI YANG AND TATSUYA SAMEJIMA

increase the first one, and decrease the negative and increase the positive CD band as in RNA. 3.The deoxyribosyl polymers can have stacked bases perpendicular to the helical axis or tilted, whereas the stacked bases of the ribosyl polymers or DNA.RNA hybrids are always tilted. 4. For any polynucleotide or nucleic acid, the magnitude of the Cotton effects decreases with unstacking of the bases (e.g., at elevated temperature). A large change in the magnitude indicates a stacking-unstacking process, but a small change does not necessarily preclude the presence of stacked bases. 5 . Protonation or deprotoriation of the bascs can change the magnitude of the Cotton effects, even though the stacking of bases may remain unchanged. 6. For RNA and ribosyl polymers, the formation of base-pairs leads to a significant blue shift of the ORD and CD. In contrast, only a very small blue shift could be detected for the formation of a double helix of DNA (see Table V). Brahms and Mommnerts (14 reported ) an intermediate form of DNA in 80% ethanol at low salt concentration and between 40 and 55 Cwhich differs from both native and completely denatured DNA. The CD in this case resembles that of RNA; the positive rotational strength of this intermediate form was about three times that of native DNA and the negative CD band almost completely disappeared. It is highly tempting to suggest that this intermediate form has tilted bases instead of bases perpendicular to the helical axis. Similarly, we found that poly dT and poly dC can have an ORD similar to that of a ribosyl polymer, but no ribosyl polymer has been found to have an ORD profile of the B-form of DNA. Ts’o et al.(80) have also discussed in detail the large influence of the 2’-hydroxyl group of ribose on the conformation and helix-coil transition of polydeoxyribonucleotides and polyribonucleotides. They explain the differences in ORD between these two classes of polymers on the hypothesis that the 2 hydroxyl group forms hydrogen bonds with the 2 keto group of cytosine and uracil and with the N-3of the adenine in the polymer, even though the predominance of such hydrogen-bonded structure of ribosyl polymers in solution has not yet been demonstrated experimentally. Ts’o et al.(80) further stipulated that the stack of the bases of the ribosyl helix is morc oblique and that of the deoxyribosyl helix more parallel. Ultimately, our understanding of the dissimilaritiesof the Cotton effects of nucleic acids must depend on advances in theoretical calculations. But a qualitative description here may suffice for our purpose. The exciton t,heory of optical activity (15-17, 73)prellicts that the m r * transitions in a helical molecule such as DNA produces many overlapping 0hands arid the

ORD AND

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275

ACIDS

resultant spectrum shows two nearly equal CD bands of opposite signs with :I crossover at a wavelength of maximum absorption. The opposite rotat,ionalstrengths sum to zero and the spectrum is therefore conservative. The corresponding ORD as calculated from the Kronig-Kramers relationship [Eq. (Sa)] shows two peaks and one trough (or two troughs and one peak) with the trough (or peak) located near the wavelength of maximum absorption. Qualitatively, this is in good agreement, with the experimental CD and ORD ofDNA (Fig. 14). In contrast, the CD of RNA does not appear in pairs of nearly equal positive and negative bands and the spectrum is t,herefore nonconservative over the range of wavelength studied (above 220 mp). Bush and Brahms (77)examined Tinoco’s general theory of and concluded that. interactions of the CD bands in optical activity (152) the region of 200 and 300mp with far-ultraviolet transitions can give rise to a CD of t,he nonconservative type such as found in RNA. They further suggested that base tilting cannot be expected to give large nonconservative CD bands, which of course is contrary to our speculations mentioned above. Bush and Brahms considered the striking difference between the CD of DNA and RNA to be due t,o some geometrical factors and they hypothesized that formation of a double-stranded structure would be reflected in changes of spectra and in t,heappearance of a conservative type of CD such as in DNA. This explmation, however, cannot, be applied to the CD of double-helical RNA, for instance. Thus, the matter is still an open question and must await more detailed theoretical calculations. has now refined his exciton theory by taking into conTinoco (253) sideration both the conservative and nonconservative types of spectra. The theory can be qualitatively described as follows. For a polymer array of N identical monomer units the singlc electronic transition of each monomer will be combined into N polymer transitions, that is, N absorption bands at N frequencies for each band in the monomer. If the bands in the polymer are not too shifted relative to the monomer band, the optical properties of the polymer can be represented as t~ weighted sum of the monomer absorpt,ion band plus its first derivative, second derivative, ete. For simplicity the second derivat,ive and higher derivatives will be ignored. Thus, the CD of the polymer at any given frequency, can be written as CL

-

+ bz(d~i\l/du).

CR = ( X ~ C ~ I

(20)

Here EM represents the molar absorptivity of a single electronic transition of the monomer at frequency u. The coefficient al characterizes the interactions among different absorption bands of the monomers (nonconservative type) and the coefficient b2of the same band in all identical monomers of the polymer molecule (conservative type). The corresponding molar rotation at a.ny wavelength can he obtained from Eq. (20) by using the

270

JEN TSI YANG

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Kronig-Kramers transform. Figure 19 illustrates the CD 011 the left-hand side and ORD on the right for a modified Gaussian band (curves 1 and l a ) and its negative derivative (curves 2 and Sa). The Cotton effects of the polymer can have the shape of these characteristic curves with either sign In or any combination of these curves (curve 1 f 2 ; curve l af.%’a>. general, both the conservative and nonconservative types will contribute to the CD and ORD and their contributions are very dependent on the geometry of the molecule. Our speculation is simply that the nonconservative contributions are small for a helix having bases perpendicular t o the helical axis such as the B-form of DNA. The above general idea can be applied to real polymers; heteropolynucleotides can be repreeented by an average effective monomer, and double-stranded polymers can be treated as a single-stranded polymer of monomeric units having two transitions (for a paired base). Native DNA's are represented mainly by curves d and 2u,and the RNA's characterized mostly by curve 1 2 and curves l a 2u.

+

+

t 0 c

0 - aa l -

U

> L

0 C

c

+

+ -e

.-

0 5

- a 0

+ 0

-

1(mpl

Fro.19. Theoretical CD (left) and ORD (right) cwves (155). The top two curves are the characteristic curves for a modified Gaussian band and its first derivative which combine to give all others. Each curve can be positive or negative.

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277

ACIDS

The exciton theory deals with the T-T* transitions; the existence of an n-r* transition in nucleic acids remains unsettled. We merely wish to suggest that the small negative CD band found near 300 mp (Fig. 14) might be attributed to this n-r* transition.

D. Calculation of the ORD of RNA Cantor e2 al.(10812) recently extended the nearest-neighbor formalism (Section IV,B) to take into account the double-stranded conformation, using an equat,ion analogous to Eq. (15): (21)

The summatioris are carried over the base sequence of oxily one of the two strands, and the choice of the strand is irrelevant. Here [m&J is the molar rotation per base of the double-stranded dinier containing the sequence NpN' and its antiparallel complement and [m:] the molar rotation per base of the double-stranded monomer. With the approximations of random nearest-neighbor frequencies, identical base composition of both strands, and separable effects of the single- arid double-stranded interactions, Eq. (21)can instead be written as (cf. 836): [WLRNA]

+

= [?n1]/2 f [ w . L ] / ~

+

X ~ ~ ~ A T ~ Z - A UX&’~A?~-GC

+

XAUXGC2Am--Au/GC

(22)

Here [ml] arid [ m ~are ] the mean residue rotations of the two single strands; the 2Am's are the additional contributions of two sequential A U pairs and G C pairs, and the average interactions of all the remaining possible sequential base-pairing combinations; the X A U and XGC are the mole fractions ofthe A . U and G . C pairs. In other words, the three 2Am terms are the increments in rotation caused by the interstrand interactions, due to the G . C and A U hydrogen bondings, with respect to the curves calculated from the single-stranded base-stacking model. They can be roughly estimated from the data on double-helical polynucleotides (94,96'), that is,

-

-

2 0-AU~ = ~ [mvOiy A . poiy u] -

and 2Arn-cc =

[mpoiy G

-

poty

cl -

([?))poiy A]

-b

[mPoiy u])/2

GI + [ m p o lc~I ) / ~

([mpoiy

(23)

(24)

Cantor etal.(10812)calculated the [ m p o l yaccording ~] to Eq. (17), since no experimental data were available for single-stranded poly G. The term Am-AUlGC is assumed to be the average of AW~-AU and A m - G C , since no experimental estimate is :tvailshle. This of course is a very rough estimate and is the best we can do at present.

278

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Using the known sequence of yeast alanine tRNA (154), Cantor etal. (208a) constructed a model of this polymer with a maximum number of antiparallel A . U and G . C pairs but without unstacking any bases in the single-stranded polynucleotide chain. They were able to calculate and almost reproduce the peak, trough, and crossover of the experimental ORD, but the magnitude of the calculated rotations was much larger than the observed ORD, due undoubtedly to the approximations and assumptions used in such computation. Vournakis and Scheraga (127) also used the ) calculated the ORD for the possible method of Cantor et al.( 1 0 8 ~ and conformation of both alanine tRNA and tyrosine tRNA previously proand Madison et al.(155). Their results were posed by Holley etal.(254) . two slightly different very similar to those of Cantor ef al.( 1 0 8 ~ )That models of alanine tRNA gave equally satisfactory agreement between experimental and calculated curves suggests a degree of uncertainty in predicting the correct model for the RNA molecule. It seems therefore premature to conclude that the crude calculations support the conformation also ) of any particular chosen model for the tRNA. Cantor el al.( 2 0 8 ~ found that the calculated ORD of the base-pairing model for alanine tRNA is shifted to the shorter wavelength side when compared with that based on a single-stranded RNA without base pairings, a finding in agreement with experimental observations (see Section VI, B). McMullen et al.(156) have also applied the method of matrix rank analysis to a large body of experimental ORD data to determine and identify the minimum number of independent components contained within it. They found the ORD of tobacco mosaic virus RNA to be a superposition of only two basic spectra of the single- and double-stranded helical forms of the molecule. That leads to a direct calculation of the percent composition of the double strands at any of the conditions considered, if a structural model in terms of an equilibrium between the two forms is postulated. This method of analysis seems to be powerful and of wide applicability, since it is independent of the source of data. But it has the disadvantage of yielding only the shape of the spectral components and variations in amplitude are specifically ignored in the reduction process. It is therefore necessary to find the molar rotation at any given set of conditions for the two forms. The method of caIcuIation described in this section should be equally applicable to the CD of RNA, but no comparable detailed experimental data on the 16 dinucleoside phosphates and poly G . poly C exist at present. Theoretical calculations of the ORD and CD of DNA have also not yet been tried. Comparison of the ORD results in Sections V and VI shows that the magnitude of the Cotton effects of synthetic polyribonucleotides is much larger than that of RNA. This cannot be attributed to the imperfect base

ORD AKD

CD O F NUCLEIC ACIDS

279

pairing in RNA as compared with poly A . poly U, for instance, since a t elevated temperature where the secondary structure of the polymers is disrupted the first peak of the ORD of polynucleotides is still higher than have suggested that the two that of RNA. Brahms and Mommaerts (14) intertwined double-stranded helical structure of poIynucleotides such as poly A . poly U and poly A in acid solution may be parallel as contrasted with the antiparallel base pairings in RNA. Accordingly, the rotational strengths of the base pairs are assumed to he additive in the former case and subtractive in the case of RNA. This suggestion, however, fails to account for the findings that poly U, for instance, is known to have little ordered structure above 10” and it still has a higher first peak of its ORD than do most RNA’s. Rather, we suspect that the difference arises from the regularities in base sequences. It is quite possible that the interactions among an array of identical bases or bases of alternating sequence differ significantly from those of mixed bases of irregular sequence. This hypothesis seems also to apply to polydeoxyribonucleotides; for instance, poly d(A-T) has larger Cotton effects than those expected for DNA (see Figs. 11 and 15a). It is contended that the regularity of the base sequence would enhance the Cotton effects as contrasted wit,h the irregular sequence found in nucleic acids. I n view of this difference in rotation, estimation of the amount of secondary structure in RNA using synthetic polynucleotides as model compounds seems very uncertain. Vournakis and Scheraga (I%?),however, suggested that the amount of G . C hydrogen bondings in a RNA molecule can be made by comparing the mean residue rotation at 276mp with that of the 276mp peak of poly G . poly C, neglecting the small contributions due to the A . U base-pairings. They recognized that the lack of adequate model compound data and calculation techniques made possible only crude ) a different approach to the estimates at best. Cotter etal.( 1 5 6 ~suggest calculation of ORD for RNA. These workers question the correctness of Eqs. 23 and 24,because they are based implicitly on the single-stranded states of homopolymers, which are unlike the RNA molecule with its “random” sequence of bases. Instead, Cotter etal.derive the composition of the base-paired part from thermal difference absorption spectra. The rotations for the unpaired part are then calculated from Eq. (15) using shndard dinucleoside phosphate data and assuming a random sequence of the appropriate base composition of RNA, as applied by Cantor et al. (108a). However, the rotations for the double-stranded part are still obtained from the data of Sarkar and Yang for poly A . poly U and poly G poly C (94,96),which differ from the double-stranded RNA of a “random” sequence, thus making such calculations somewhat uncertain. Suffice it to say, any quantitative interpretation of the ORD of RNA should presently

280

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be viewed with reservations. In this respect),the use of a double-helical RNA is perhaps a more realistic as a model [see Hashizume and Imahori (lor)] approach and worth further investigation. Even here we have to bear in mind that structural possibilities with many base-pairing arrangements other than those proposed by Watson and Crick cannot be overlooked;4 furthermore, structures with more than two strands are also possible.* These are the challenges that must be met if ORD can be used for such calculations. The very nature of any empirical analysis cautions us against overenthusiasm toward its quantitative interpretations, at. least at this stage of development,.

VII. Complexes of Nucleic Acids A. Ribosomes, Viruses, and Deoxyribonucleoproteins Nucleic acids are known to form complexes with a variety of substances, from small molecules, such as metallic ions, to macromolecules, such aa proteins. The study of the optical properties of these complexes might enable us to elucidate the interactions and the conformational changes, if any, that occur when these constituents are incorporated into the complexes. Blake and Peacocke (157)initiated the ORD study (above 230mp) of rabbit reticulocyte ribosomes, and their isolated constituents, rRNA, and soluble proteins. Examination of the Cotton effects and also the rotation of RNA at a given wavelength led to the conclusion that the secondary structure of RNA in mammalian ribosomes is the same as the one that the isolated RNA molecule possesses in solution. McPhie and see also Gratzer, 1,5959) studied the ORD of ribosomes from Gratzer (158; coli as well as rabbit reticulocyte. Quantitatively, the yeast and Escherichia results of the three species were very similar. Dissociation of the yeast ribosomes and their disorganization by high concentrations of chelating agent essentially had no effect on the ORD results. Sarkar and Yang ( l a gsee ; also Sarkar et al., 160) investigated both the ORD and CD of Eseherichia coliribosomes (70S ) ,their two basic subunits (the 30 S and 50 S particles), and the corresponding 16 S and 23 S RNA's and found no notable difference in the mixtures of 70 S, 50 S, and 30 S particles and also no difference in mixtures of 50 S and 30 S particles. This implies the absence of any conformational change when the 30 S and 50 S particles associate to form the active 70 S ribosomes. Figure 20 summarizes the results for the subunits of Escherichia coliand their corresponding RNA moieties. Since the ribosomal proteins have no CD bands above 250mp, the portion of the spectrum in this wavelength range approximates that of 4

See article by Pullman arid Pullman in this volume.

ORD

AND

CD OF NUCLEIC

281

ACIDS

A (rnpL1

(rnFLJ 250

300

I

I

200

250

300

I

I

3

1 P)

+ '0

3 2

-25 4

4

2 P)

+ 'Q

0 2

-9

2

X (mp)

A (mp)

FIG.20. CD (upperpart)and ORD (lower part) of 30 S arid 50 S Escherichia coli ribosomal subunittr and their corresponding RNA moieties (139).Allsolvents are made up of 0.005 M Tris, 0.0005 M MgZC,and 0.1 M KC1.

the corresponding RNA's, which is about 63% of the ribosomes. Significant difference in the spectrum between ribosomal subunits and the RNA moieties, however, occurred below 250mp, where the contributions of the peptide chromophores become prominent. While the 23S RNA, for instance, displays a small positive CD band at about 230mp, the ribosomal subunit actually shows a shoulder of a negative CD band and also a negative shoulder in ORD near 230 and 235mp. It is now well recognized that the a-helical (right-hand) polypeptides show three CD bands between 185and 250mp, one positive with a maximum at 191 mp and two negative with a and the corresponding ORD double minimum at 210and 222mp (161,162), has a trough at 233mp and a peak at 198m p (16.2, 163). Thus, the results in Fig. 20 strongly suggest the presence of helical segment,sin the ribosomal

282

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protein moieties. Analyses ofthe difference ORD and CD spectra between ribosomal subunits and their RNA moieties indicated that the ribosomal proteins, at least in thecase of 50 S particles, are partially a-helical (about 20-25%).The portion of the isolated ribosomal proteins of Escherichia coli that could be solubiIized was also found to have about the same amount of helical content. On the other hand, the protein moiety of the yeast ribosomal proteins contained about 30% a-helix, but once isolated from the RNA moiety the sdubfe portion of the proteins had no detectable a-helical content and was presumably denatured (158). THE COTTON Peak 1

TABLE VII EFFECTSOF VIRUSES(166) -

Trough 1

Peak 2

Trough 2

Class 10

T2 T2(gt) T4 T6 T6(gt)

x,

290 290 290 290 290 290

270 270 27 1 272 270 270

-3960 -3000 -4050 -4650 -3650 -3620

240

240 240

2280 2060 1680 2350 2430 2240

620 771 627 825 640

255 265 260 260 258

-1820 -2360 -2580 -2240 -2200

230 240 237.5 225 240

3200 630 238 1320 180

520 4840 4030 3450

260 252 255 257

-3660 -7860 -8000 -5800

245 -

-1180 -200 -1500 -1700 -1370 -480

237.5 240 241

Class I1

T5 Tli B3 X(C)

MK)

290 290 295 290 290

Class 111 292.5 +X1?4 290 MS2 285 f2 285 R17 ViralDNA and RNA 290 a T2 DNA 290 X(C) 290 DNAb 290 DNAb ax174 290 DNA T7DNAb 290 R17RNA 285 0

-2350 235 - 235 - 232 235

2780 258 1280 260 2230 (2140) 258

-1910 236 -2100 230 -1920 228

2730 4480 4380

2210 (2140) 258

-1540

228

4290

1920

-2180

230

2870

260

4050 -2690 225 2090 (20%) 258 4070 252.5 -4100 227.5 -1070 217

-4290 -8860 -11800 -10100

- 1780

These viruses have no peak at 290 mp. yt = nonglucosylated. The values in parentheses we the rotations at 290 m,u computed from Eq. (18).

ORD

AND

CD OF NUCLEIC

283

ACIDS

Maestre and Tinoco (164, 165)attempted to elucidate the internal structure of the nucleic acids and their relation to the protein coats in Each virus has viruses; they measured the ORD of 1 G viruses (Table VII). a characteristic ORD curve, by which it can be easily distinguished even in the cases of very closely related phages. The general profiles (Fig. 21)can be classified into three types: (I) t,he T-even class has levorotations above 250 mp with a trough at about 270 mp, although its ORD retains the two-peak-and-one-trough shape of nucleic acids; (11)the T-odd class has an ORD very similar to that of nucleic acids (Section VI) with a peak near 290 mp and a trough near 260mp; (111) the third class shows a characteristic deep trough near 235 m p in ORD caused by a high percentage of ;i3 5

2

c 0

0

1

41 0 E -1

-$ - 2 T)

no- - 3 2-4 U

Y

1

'

"

'

MS2

-

\

-

-

l

220

i

l

240

,

l

L

[

280

260

,

I

300

,

I

,

320

XCmP) CC)

FIG.21.ORD of (a) T-evenphages, (13) T-odd phages, and (c)viruses containing a large contribution from theprotein coat(166).

284

JEN TSI YANG

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protein. The rotations of the nucleic acid and protein moieties were found to be nonadditive, that is, the ORD of the intact phage cannot be explained as the simple sum of the ORD of isolated protein coat (virus ghost) and nucleic acid. On the other hand, the ORD of osmotically shocked T2 phage can be identical, within experimental errors, with that of the sum of the purified components. Since osmotic shock releases the nucleic acid from the head of the phage, but leaves the protein coat essentially intact, the difference in ORD between intact and osmotically shocked phage must be the result of the packing of the DNA molecule inside the head of the phage. + relemedDNA, Furthermore, this difference in rotations, [aIphage- [aIgho8t shows qualitative similarities among all the DNA phages tested; all displayed a negative Cotton effecttwith a trough between 280 and 290 mp (the low wavelength regions, however, were obscured by the rotation of the protein). This implies that the packing of the DNA molecule inside the suggested, as virus has a common characteristic. Maestre and Tinoco (166) did Pollard (166) and Tikchonenko et al.(167), that the DNA molecule inside a virus is in a dehydrated state that altered the stacking structure of the bases with the resultant change in optical properties. In support of this contention, lowering the activity of water by using a high LiCl concentration does alter the ORD of the T2 and T7 phages, for instance, and the difference ORD between normal DNA and that in high salt concentration is very similar t o that between intact and osmotically shocked phages. We note, however, that the ORD of T2 phage in 24.4% LiCl (pH 7.4) (165) still retains a larger second peak than the first one characteristic of all normal DNA’s. If our hypothesis about the base tilting versus base perpendicular to the helical axis (Section VI, C) is valid, then the DNA molecule inside the viruses may still have a structure similar to the B-form. It could be that other types of geometrical change that have escaped our detection are involved in the condensation of the DNA molecule into the compact package of internal DNA. Maestre and Tinoco (164) also found a linear relationship between the specific rotation at the 290-mp peak (based on the concentration of DNA) and the percentage by weight of DNA in each phage after corrections were made for the rotations due to the glucosylation of the 5-hydroxymethyl cytosine of the T-even phage family (Fig. 22). The deviation A[LY]~~,, for T-even phages from the straight line (Fig. 22) was corrected by assuming that T2, T4, and T6 were 70, 100, and 147% glucosylated (168). The correlation shown in Fig. 22 is also consistent with the hypothesis that the main influence on the ORD of a polymer is the iriteractioii among the polymer units, and the differences in rotations for the different phages mirror differences in conformation of the DNA molecule inside the phage (164). Thus, a high DNA concentration in the phage should lead to a

285

ORD AND CD O F NUCLEIC ACIDS I

I T 7

1

T-6

I

30

I

I

* I

40 50 60 Weight % DNA in bacteriophage

I

I

70

a0

FIG.22.The relationship between the rotation at the 2 9 0 - upeak (based on the concentration ofthe DNA moiety) and the weight percent of DNA in the phages (164).

conformation of DNA different from that in solution. However, an entirely satisfactory explanation for these results is still lacking, because extrapolation of the straight line in Fig. 22 to zero DNA content (presumably approximating free DNA in solut,ion) leads to an [ c Y ] ~ ~higher o than any measured in solution or expected from Eq. (18) (ef. Fig. 16). Whatever interpretation for these results is finally established, the ORD is apparently very sensitive to the state of DNA inside the phage and will be very useful in the study of bacteriophages. Maestre and Tinoco (165) have also analyzed the ORD of the T2 ghost (protein coat) using both the bo and methods see refs. 2 and 20) and 233-mp trough methods for proteins (for concluded that the helical content of the phage ghost is quite small (5205Z0). They also found the influence of the protein coat on the rotations above 233 mp (away from the Cotton effects of the protein) due to the whole phage was small compared to that of the nucleic acids. Oriel (169) has studied the ORD of calf thymus deoxyribonucleo-

286

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protein (DNP). Both the particulate fraction and the soluble DNP showed the same ORD [the particulate fraction is the gellike fraction and amounted to about 15% of the final preparation (170)l. It had a trough at 235 mp in addition to one near 260 mp, typical of the DNA molecule; this second trough can be readily recognized as the characteristic 233-mp trough foran a-helix of proteins. Using the 233-mp trough method, Oriel estimated that particulate DNP, soluble DNP, and the corresponding histones (separated from the DNA molecule in 2 M NaC1) contained about 20% a-helix. This result agreed well with the estimated helical content from the bomethod for acid-extracted histones in 1 M NaCl (171, 272). Thermal denaturation studies indicated that histones associated with the DNA molecule are more stable than those dissociated from DNA, suggesting a strong interaction between the proteins and DNA in DNP (169).

B. Complexes with Small Molecules The binding of dyes to nueleic acids has been studied for two decades with respect to the staining properties of the dyes and their mutagenic properties (88, 90,173-176). This interaction is known to cause a shift in the wavelength ofthe absorption maximum of the bound dyes (177). The binding of acridine orange to DNA (178-182) and of proflavine to DNA and RNA (179,183, 184)induces Cotton effects in the absorption band of studied the ORD and absorption the dyes. Blake and Peacocke (279) spectra of acridine orange bound to native DNA (Fig. 23a). At low DNA

400

500 X (mp)

600

460

500

540

X (mp)

( al (bl FIG. 23.(a)ORD andabsorption spectra ofacridine orange boundto native DNA at, pH 7.0in 0.009 M NaCl plus0.001 M sodiumphosphate at20 C. DNA (phosphorus)/ dye ratios: low ( - - - 1, 28.48 p M to15.68 high(-), 85.44 p M to15.68 pM. Theuppercurveineachpairistheabsorption spectrum. FromBlakeandPeacocke (179). (b)CD and absorption spectra ofacridine orangeboundtonative DNA atpH 6.6. DNA (phosphorus)/dye ratios: (-), 3 and ( - - -), 9 in 0.001 M buffer, and ( . . . ), 15 at0.1ionic strength. The uppercurves areabsorption spectra; (- - -) absorption spectrum of acridine orangein ethanol. Dye concentration: 3 x 10-6If. From Gardnerand Mason (182).

a;

ORD AND

CD OF NUCLEIC

ACIDS

287

(phosphorus) :dye ratios (about 1.8), the intensity of the 292-mp absorption maximum became smaller as compared with the free dye but that of the shoulder near 470 mp was enhanced. The ORD of the complex showed a small negative Cotton effect with a trough near 475 mp and a peak at however, about 440 mp. At high DNA (phosphorus) :dye ratios (about 5.4), the intensity of the absorption maximum near 492 mp was enhanced; this was accompanied by a gradual red shift of the band. Two Cotton effects were induced in this case, a relatively large positive one centered at the band on the long wavelength side and a negative one superimposed on it. The magnitude of the positive Cotton effect first increased gradually with increasing DNA concentrations, reached a limiting level, and thereafter decreased with further increase in the DNA:dye ratios. A similar study has been reported by Yamaoka and Resiiik (180), but comparison of the results from different laboratories under various experimental conditions is difficult, since the Cotton effects of the DNA-dye complexes are very sensitive to the solvent composition (ionic strength and pH) and also to temperature. Figure 23b shows the CD and absorption spectra of the The CD consisted of three DNA-acridine orange complexes (181, 182). bands, one positive at 505 mp and two negative at 488 and 465 mp. At DNA:dye ratios less than 3 and at low ionic strength, only the 505- and 465-mp bands could be observed; they were, however, preferentially suppressed with increasing DNA :dye ratios. In all cases, lowering the ionic strength of the solution increased the intensity of these two bands but reduced slightly that of the 488-mp band. Gardner and Mason (182) attributed the 488- and 465-mp negative CD bands t o the interactions between bound dye monomers and between bound dye dimers, whereas the 505-mp positive band is largely due to the dye dimers. On the other hand, Yamaoka and Resnik (180) analyzed their ORD data of the DNAacridine orange complexes, using the Kronig-Kramers transform (see Eqs. 6a, b), and concluded that at least 4 CD bands were required to fit the experimental ORD curves. Several laboratories have also attempted theoretical calculations to explain the observed ORD results. The problem is a complicated one; for instance, we are still unable to differentiate between the fractions of monomers and poIymers of the total dye molecules bound to DNA, or between the degrees of aggregation in the polymeric dyes. However, it is the current belief t!hat the monomeric dye molecules are intercalated between neighboring base pairs and that the dye aggregates may be bound to the phosphate groups of the DNA molecule. Blake and Peacocke (179, 183)found that the binding of proflavine to DNA produced a positive Cotton effect centered at the 443-mp absorption band of the dye. Its magnitude depended on the ionic strength, tempernratio. Lowering thr ionic strength of the solution from ture, and DNA :(lye

288

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AND TATSUYA

SAMEJIMA

0.1 to 0.001, for instance, doubled the magnitude of the Cotton effect. A t a constant concentration of proflavine, the magnitude of the Cotton effect increased linearly with the DNA concentration until the molar ratio of the DNA (phosphorus) to proflavine was about 4 or 5, and then decreased with further increase in the DNA concentration. These findings suggest that the bound dye molecules contribute less to the rotation when they are sparsely distributed over the DNA molecule than when they are near-neighbors. In other words, a group of such ligands bound to neighboring sites is required for induced opticaI activity. Yamaoka and Resnik (184) restudied the binding of DNA with proflavine and found the first Cotton effect to have a peak and trough at 482 and 457 mp. They also observed another broad peak near 410 mp; the rotations approached zero between 420and 400 mp, but became increasingly negative below 400mp. These authors were able to fit their ORD data with two Gaussian CD bands, one positive and the other negative. More recently, Yamaoka and Ziffer (185)reported the ORD and CD of DNA-actinomycin D complex (see also Permogorov and Lazurkin, 186). Free actinomycin D shows a major absorption band near 440 mp, which is weakly optically active (187).However, the absorption maximum shifted to about 460 mp with the appearance of a strong negative CD band when actinomycin D was mixed with DNA, thus indicating the participation of the actinomycin chromophore in the complex formation. Mahler et al.(188) studied the interactions at low ionic strength between DNA and steroidal diamines such as irehdiamine A (pregn-5-ene3P,2Oa-diamine) and malouetine [5a-pregnan-3~,20a-ylenebis(trimethyliodide)] by means of various physical methods. They found that the positive CD band of DNA and the corresponding 290-mp peak of its ORD is first enhanced by the addition of steroids, reaches a maximum effect at a steroid :DNA(phosphorus) ratio of about 0.2 to 0.3, and then decreases at still higher steroid :DNA(phosphorus) ratios. These results suggest the formation of two types of complexes, depending on the molar ratios of steroids to DNA. The first complex is characterized by enhanced thermal stability as compared with native DNA. The second complex is less stable thermally, and at room temperature its optical parameters are those characteristic ofpartially disoriented DNA. The interactions between nucleic acids and metallic ions have been studied with a variety of physicochemical methods, but only recently did Cheng (189) report the ORD of caIf thymus DNA in the presence of divalent ions such as Ca2f, 2n2+,Mg2+,Mn2+, Cu2+,and Hg2+ (Fig. 24). In all cases the ORD spectra were altered. While manganese, magnesium, calcium, and zinc ions did not significantly shift the positions of the peaks and troughs, cupric ion showed a characteristic reduction of the magnitude ofall cxtmmes and eliiniimtion of tJhrseconil peak. Mercuric ion produced

ORD

289

AND CD O F NUCLEIC ACIDS

X (mp)

FIG. 24.ORD of calf thymus DNA in the presence of divalent cations at26 C.A , Ca2+;C,ZnZ+; D,Mg2+;E,Mn2+;F ,none; G, C U ~ +and ; H, Hgz+.Concentrations: DNA, 1.0 X W 4M (phosphorus); divalent ions, 2 0 x 10-4M ; buffer, 5 X 10-3M NaCIOl M Tris-HC1 (pH 7.5). From Cheng (189). plus 5 X

an even more remarkable change in the spectrum; the Cotton effect turned negative on the long wavelength side. These changes were attributed to the specific action of the divalent ions on the DNA molecule. The less sensitive absorption spectrum could not, detect such interactions between DNA and these metallic ions.

VIII. Visible Rotatory Dispersion Prior to 1963,almost all the ORD studies of nucleic acids, polynucleotides, and their constituents were confined to the visible and near ultraviolet regions (109,110, 190-200). Although we have shown that CD and ORD in the ultraviolet region can provide a wealth of information about the conformation of these polymers, it seems appropriate to conclude this review by mentioning briefly their rotatory dispersion in visible light. The nucleic acids and their constituents all show featureless and monotonous ORD in the visible region. With the exception of TMP and UMP (28), the ORD usually obeys a one-term Drude equation and the constants of k and A, in Eq. (9) are conformation dependent. The range of wavelength over which the Drude equation applies varies widely with the compounds studied. Table VIII summarizes the pertinent numerical values for some of the nucleic acids and polynucleotides. The lists are not intended to be complete, but they serve to illustrate the lack of any genera1 trends. The

290

JEN TSI YANG

AND

TABLE VIII THEONE-TERMD R U D E EQUATION OF NUCLEIC ACIDS AND

Substance

+

0 15 M NaCl 0.015 M Nn citrate, pH 7-7.5 Salmon sperm DNA 0.15 M KF, pH 7.4 (a) 27" (b) 80" (c)goo 0.1 M NaCl 0.1 M Na TMV RNh citrate 0.001 21.1 MgC12, pH 5.50 0.1 M KF, pH 7.4 Yeast tRNA (a) 27" (b) 55" (c)90" 0.02 M phosphate buffer 0.18 M NaC1, pH 7, 27" 0.02 M phosphate buffer Yeast rRNA 0.18 &f NaCl, pH 7 (a) 27" (b) 80" Turnip yellow mosaic 0.02 M phosphate buffer 0 18 M NaCI, p H 7 virus RNA (a) 27" (b) 80" 0.02 M phosphate buffer R17 viral RNA 0.18 M NaC1, p H 7 (a) 27" (b) soo 0.1 M Na acetate 0.1 M PolyA NaCI, pH 4.85 (a) 20" (b) 27" 0.05 M Tris + 0.1 M NaC1, pH 7.8 ( 4 6" (b) 27" (c) 75" 0.1 M N a acetate 0.1 M Poly NaCl, pH 4.85 (a) 20" (b) 80" 0.05 M Tris 0.1 M NaCl, pH 7.8

+ +

38.0

230 220

36.0 38.5 4.7 46.5

227 228 286 256

34.5 18 .O 1.2

245 267 316

28.0

248

45.0 3.8

245 296

50.0 8.5

255 292

55.0 8.0

239 334

+

+

+

+

+

+

u

+

100 94

50

37 1.0

4.2 1.0

SAMEJIMA

POLYNUCLEOTIDEd

k X lo8, deg cm' decagram-1 Xe (mp)

Solvent

Calf thymus DNA

TATSUYA

279 278

275 276 314

302 307

Reference

ORD AND

CD OF NUCLEIC

291

ACIDS

TABLE VIII (Continiied) 6

Substance ~

Poly

c

Solvent ~

~

x

106,

degcm* decagram-’A. (mp) Reference ~

(a)6-7" (b)27" (e) 75" 0.1 M Na acetate 0.1i l l NaCl, pH 4.85 (a) 20" (b) 80"

+

9 .,5 5.0

20.5

290 298 316 (109)

4.2

1.6

302 307

only generalization for the data on these polymers is that X, increases and k decreases when their secondary structure is disrupted. Unlike the measurements of the Cotton effects in the ultraviolet region, study of the visible rotatory dispersion usually requires a large

quantity of samples to ensure precise measurements. Since not much information is derived from these studies as compared with those on Cotton effects, the ORD in the visible region is not usually used for studies on nucleic acids and their constituents. On the other hand, in solvents that absorb strongly in the ultraviolet region, e.g., many organic compounds, chloride ions, we must use measurements in the visible region 200). (see, for example, refs. fgr,

IX. ConcludingRemarks Accurate measurements of the Cotton effects of nucleic acids, polynucleotides, and their constituents began to accumulate only five years ago. The purine and pyrimidine base chromophores themselves are optically inactive, but once attached to the optically active sugars they are induced to produce Cotton effects. These are conformation dependent and provide us with a new means for probing the molecular structure of these biopolymers in solution and for following their conformational changes when exposed to different environments. Significant advances have also been made in theoretical treatments that help us to understand and interpret the experimental observations, although quantitative calculations are still a formidable task at present. The mononucleosides and mononucleotides are in a class by themselves as far as optical activity is concerned. A11 display a single Cotton effect around the 260-mgabsorption band. For natural compounds having the 0-D-furanese f i g , the purine derivatives have negative signs and tha

292

JEN TSI YANG AND

TATSUYA

SAMEJIU

pyrimidine ones have positives ones. Replacing the ,&glycosyl bond by an a-linkage reverses the sign. The purine and pyrimidine base rings are considered to favor the anti rather than the syn form with respect to the sugar ring as they do in the nucleic acids. Furthermore, since the a-D-and p-L-anomers have the same configuration at C-l', the Cotton effects of these purine or pyrimidine derivatives are of the same sign. The ORD and CD of dinucleoside phosphates and trinucleoside diphosphates are sequence-dependent ; the positions of the extremes and their magnitudes vary among the isomers, even though the general spectra are very similar. Thus, optical activity can become a new means for determining the base sequence in oligonucleotides. There will probably be applications for the study of the primary structure of nucleic acids when they are degraded and separated into various fractions of oligomers. The very simplicity and the small quantity of samples required for the optical method make it very attractive. But it remains to be seen whether this approach has definite advantages over other physicochemical methods such as chromatography. The oligonucleotides and polynucleotides as well as nucleic acids display multiple Cotton effects around the 260-mp absorption band. The general ORD profile shows two peaks and one trough between 230 and 300 mp, with the trough centered near the wavelength of absorption maximum (poly I and GMP gel are the exceptions, their Cotton effects having two troughs and one peak over the same wavelength region). The corresponding CD has a positive band (or two bands as in the case of some dinucleoside phosphates) on the long wavelengt,h side and a negative band following it. These results can be interpreted in terms of the stacking interactions among the bases, which cause a splitting of the mr*transitions and thereby the appearance of the positive and negative bands around the 260-mp absorption band. The most important contributions to the optical activity of a polynucleotide chain are the nearest-neighbor interactions, which are present from dimers to polymers. These base interactions are largely responsible for the stability of helical polynucleotides, be it singleor double-stranded. Both DNA and RNA also show a small negative CD band near 300 mp, which is probably attributable to an n?r* transition. Our concept of the molecular forces that stabilize the helical conformation of polynucleotides and nucleic acids has modified remarkably in the past several years. Originally, the hydrogen bonds between the base pairs in the Watson-Crick model were thought to be the major factor holding the DNA double helix together in aqueous solution as well as in solid state. The linear variation of the T, of DNA’s with the G C content had been attributed to the formation of three hydrogen bonds in the G . C pair and only two in the A . T pair. We now believe that the hydrophobic interac-

+

293

ORD AND CD O F NTJCLEIC ACIDS

tions giving rise to base stacking best explain the stability of the helical polynucleotides. The importance of such hydrophobic interactions is fully supported by recent theoretical treatments of helical polynucleotides. I n aqueous solutions, the base planes tend to stack on top of each other rather than to be exposed to the water molecules. The fact that organic solvents break up the helical structure can be interpreted in terms of the weakening of the hydrophobic interactions among the stacked bases in the presence of nonaqueous solvents. We have already mentioned that oligo- and polynucleotides can exist in single-stranded helical conformations stabilized by base stackings without the benefit of any hydrogen bondings. Unstacking of the bases would disrupt the helical conformation. This is manifested in the reduction of their Cotton effects and is accompanied by a hyperchromic effect, just as in the denaturation of double-helical DNA. Study of the dinucleoside phosphates clearly indicates that, of the four RNA bases, uracil stacks the least with neighboring bases, be it adenine, guanine, cytosine, or another uracil. Since thymine is very similar to uracil, DNA T content would therefore be expected to be the most with the lowest A stable. Thus, the dependence of the T ,of DNA on the G C content can be explained in terms of stacking interactions without resort to hydrogen bondings. The same is true for the linear correlation between the G C content and the [m]290 of DNA [see Eq. (18)]. Of course, hydrogen bondings among the paired bases would enhance the stability of the helical conformation of polynucleotides, as reflected by the sharp T, of doublestranded helical polynucleotides. Although DNA and RNA show the same two-peak-and-one-trough ORD profile around the 260-mp absorption band, the relative magnitude of the two peaks is in marked contrast. Invariably, the first peak (on the long wavelength side) is smaller than the second for DNA, whereas the second peak is close to zero rotation for RNA. This rule applies irrespective of whether the nucleic acid is single- or double-stranded. Undoubtedly, the 2’-hydroxyl group plays an important role in the geometric arrangement of the polynucleotide chains. Madel building indicates that a double-stranded polyribonucleotide can only have paired bases tilted to the helical axis, unlike the B-form of DNA that has the bases nearly perpendicular to the helical axis. It is therefore suggested that bases perpendicular to the helical axis lead to a second peak larger than the fist, whereas tilted bases drastically reduce the magnitude of the second peak. It is further speculated that the same rule applies even to single-stranded ribosyl and deoxyribosyl po1ynucleotides, and that the double-stranded hybrids will resemble the ribosyl polymers. This difference in the mode of stacking between ribosyl and deoxyribosyl polymers is believed t o affect the interactions between near and far ultraviolet interactions of the bases, thus accounting for the

+

+

+

294

JEN TSI YANG

AND TATSUYA

SAMEXMA

observed “conservative” and “noncoriservative” OED and CD spectra. Ultimately, our hypothesis must be proved or disproved by future experimental results as well as theoretical calculations. The theory of nearest-neighbor interactions explains clearly the enhancement of the Cotton effects with base stacking in a single-stranded polynucleotide chain. For double-stranded helical conformation, however, we must also consider additional interactions among the paired bases and diagonal interactions among the nearest-neighbor bases in the two separate strands. We have shown that base pairing between ribosyl polymers, and to a lesser extent between deoxyribosyl polymers, causes a blue shift of the Cotton effects. This blue shift is also substantiated by empirical calculations of the ORD of RNA based on results of synthetic polyribonucleotides. Thus, in principle we can estimate, albeit very crudely, the extent of base pairing in a RNA molecule from the position of the peak and trough of its ORD. We have also mentioned that the magnitude of the Cotton effects of homopoIynucleotidesor polynucleotides of regular sequence is much larger than that found in DNA and RNA. It is for this reason that any quantitative calculations of the extent of base stacking in a natural polynucleotide based on model synthetic polynucleotides should be viewed with reservation; much depends on future modifications and refinements. The ORD and CD studies of complexes of nucleic acids with other compounds, as in ribosomes, viruses, and nucleohistones, would enable us to detect any change in conformation when the constituents are incorporated to form the complexes. With the newly developed technique of difference ORD, it will be possible to measure small changes in rotations with more confidence. Introduction of chromophores into nucleic acids could induce new Cotton effects as in the case of the binding of dyes, which in turn would provide additional information concerning the conformation of the nucleic acids. Even the binding of small ions such as Hg2+could drastically change the optical properties and thus indicate an alteration of the conformation of the nucleic acids. In this review we have briefly described only a few examples. The vast field of interactions between nucleic acids and many other compounds remains to be explored, as far as ORD and CD are concerned. There is every reason to believe that such investigations will be accelerated in the next few years. In this review we have not listed any numerical values of the CD of nucleic acids, partly because this technique has not yet been utilized as extensively rn ORD and partly because the numerical values reported in the literature might be subject to further refinement due to the lack of some universally accepted standard for the calibration of the CD instrument (see Section 11, E). We are confident, however, that both CD and

ORD

AND

CD OF NUCLEJC

ACIDS

295

ORD will play an even more important role than they do now in advancing our knowledge of the structures ofnucleic acids. ACKNOWLEDGMENTS We thank Professor Masamichi Tsuboi formany valuable discussions and for calling to our attention the models of double-stranded nucleic acids (Fig. 18),J. T. Y. is indebted to Professor I. Tinom, Jr., for several stimulating discussions. Doctors H. Drucker, W. B. Gratzer, I(.Imahori, P. I<.Sarkar, and P. 0. P. Ts’o read and commented on the manuscript. Thanks are also due Mrs. M. Arrieta and M. Kita for their technical assistance.

REFERENCES 1 . T. M. Lowry, “Optical Rotatory Power.” Longmans, Green, London, 1935; Dover, New York, 1964. 2.P. J. Urnes and P. Doty, Aduan.Protein Chem.16,401 (1961). 3.J.T. Yang, i n “Poly-a-amino Acids” (G. D. Fasman, ed.), Chapter 6.Dekker, New York, 1967.

4.L. Velluz, M. Legrand, and M. Grosjean, “Optical Circular Dichroism.” Academic Press, New York, 1965. 6. S. Beychok, i n “Poly-a-amino Acids” (G. D. Fasman, ed.), Chapter 7.Dekker, New York, 1967. 6. J. T. Yang, i n“Newer Methods ofPolymer Characterization” (B. Ke, ed.), p.103. Wiley (Interscience), New York, 1964. 7 .N. S.Simmons and E. R. Blout, Biophys. J .1,55 (1960). 8. J. R.Fresco, A.M. Lesk, R. Gorn, and P. Doty, J .Am. C h m .SOC.83, 3155(1961). 9.J. T.Yang and T.Samejima, J .Am. Chem.SOC.86, 4039(1963). 10.T. Samejima and J. T. Yang, Biochemistry 3,613 (1964). 11.J. Brahms, J .Am. Chem.SOC.86, 3298(1963). 12.J.Brahms, W. F. H. M. Bilommaerts, and C. Sadron, Compt.Rend.Acad.Sci. 268, 2203(1964). 13.W. F. H. M. Mommaerts, J.Brahms, J. H. Weill, and J.P. Ebel, Compt.Rend. dead.Sci. 268, 2687(1964). 14.J. Brahms and W. F. H. M. Mommaerts, J .Mol.Bio2. 10,73 (1964). 16.I.Thoco, Jr., R. W. Woody, and D.F. Bradley, J. Chem.Phys.38, 1317(1963). 16.D. F. Bradley, I. Tinoco, Jr., and R. W. Woody, Biopolymers 1,239 (1963). 17.I. Tinoco, Jr., Radiation Res.20,133(1963). 18.A. Moscowits, in “Optical Rotatory Dispersion” (C. Djerassi, ed.), p. 150. McGraw-Hill, New York, 1960. 19.P. CrabbB, “Optical Rotat,ory Dispersion and Circular Dichroism in Organic Chemistry.” Holden-Day, San Francisco, 1965, 20. J. T.Yang, i n A Laboratory Manual of Analytical Methods in Protein ChemAlexander and H. P. Lundgren, eds.), Vol.5,Chapter 2. Pergamon Press, ktry” (P. Oxford, 1968. 21.W. M o 5 t t and J.T.Ymg, Proc. Natl. Acad.Sci. U .S.42, 596 (1956). 22.W. Moffitt and A. Moscowita, J. Chem.Phys.30, 648 (1959). 23.V. L.Rosenfeld, 2.Physik62, 161(1928). 24. J.T.Yang and P. Doty, J .Am. Chem.Soc.79,761 (1957).

296

JEN TSI YANG

AND TATSUYA

SAMEJIMA

25.G. M. Holswarth, i n A Laboratory Manual of Analytical Methods in Protein Chemistry” (P. Alexander and H. P. Lundgren, eds.), Vol. 5,p.34. Pergamon Press, Oxford, 1968. 25n.J. ThiBry, J .Chim.Phys. 66,98 (1968). 16. J. Y. Cassim and J. T. Yang, to be published. 27.B. J. Adkinsand J. T. Yang, Biochemistry 7, 266 (1968). 28. J. T.Yang, T. Samejima, and P. K. Sarkar, BwpoZymers4, 623 (1966). 29.D. Voet, W. B. Gratzer, R. A. Cox, and P.Doty, Biopolymers 1 , 193 (1963). 30. C. Y. Lin, D. W. Urry, and H. Eyring, Biochem. Biophys. Res.Commun. 17, 642 (1964). 31. G. D. Fasman, C. Lindblow, and L. Grossman, Biochemistry 3,1015 (1964). 39.M. R. Lamborg, P. C. Zamecnik, T. K. Li, J. Kagi, and B. L. Vallee, Biochemistry 4,63 (1965). 33.D. W. Miles, R. K. Robins, and H. Eyring, Proc. Natl. Acad.Sci. U .S. 67,1138 (1967). 34. T. Nishimura, B. Shimisu, and I.Iwai, Biochim. Biophys. Actu167, 221 (1968). 35. T. L. V. Ulbricht, J. P. Jennings, P. M. Scope, and W. Klyne, Tetrahedron Letters p.695 (1964). 36.T. L. V. Ulbricht, Ann.Rept. Chem.Soc.61,446 (1964). 37.T.L. V. Ulbricht, Ann.Rept. Chm. SOC.62,402 (1965). 38. T.L. V. Ulbricht, T. R. Emerson, andR. J. Swan, Biochem. Biophys. Res.Commun.

19, 643(1965). 39. T. R. Emerson, R. J. Swan, and T. L. V. Ulbricht, Biochem. Biophys. Res. Commun.

22, 505 (1966). 40. T. L. V. Ulbricht, T.R. Emerson, and R. J. Swan, Tetrahedron Letters, p.1561

(1966). 41. T.R. Emerson, R. J. Swan, and T. L. V. Ulbricht, Biochemistry 6,843 (1967). 42. C. S. Hudson, J. Am. Chem.SOC. 31, 66 (1909). 43. R. V. Lemieux and M. Hoffer, Can.J. Chem.39,110(1961). 44. J. J. Fox, N. C. Yung, I. Wempen, and M. Hoffer, J. Am. Chem.Soc. 83,4066 (1961). 45.T. R. Emerson and T. L. V. Ulbricht, Chem.Ind.(London) p.2129(1964). 46.L. B. Clark and I. Tinoco, Jr., J*. Am. Chem.SOC.87, 11 (1965). 47.I.FriE, J. gmekal, and J. Farkas, Tetrahedron Letters p. 75 (1966). 48. W. A. Klee and S. H. Mudd, Biochemistry 6 ,988 (1967). 49. M. P. Schweizer, A. D. Broom, P. 0. P. Ts’o, and D. P. Hollis, J. Am. Chem.Soc. 90,1042(1968). 60. J. Kraut, Ann.Rev.Biochem. 34, 247 (1965). 61. V. Sasisekharan, A. V. Lakshminarayanan, and G .N. Ramachandran, in“Conformation of Biopolymers” (G. N. Ramachandran, ed.), Vol. 2,p. 641.Academic Press, New York, 1967. 52. A. E. V. Haschemeyer and A. Rich, J. Mol.B i d .27,369 (1967). 63. J. Iiraut and L. H. Jensen, ActaCryst. 16,79 (1963). 64. R. J. Cushley, K. A. Watanabe, and J. J.Fox,J. Am. Chem.Soc.89,394 (1967); Chem.Commun.p. 598(1966). 66. T. Nishimura, B. Shimisu, and M. Futai, Biochim. Biophys. A d a 129,654 (1966). 66.J. Brahms, J. C. Maurieot, and A. M. Michelson, J. MoE. Biol. 26,481 (1967). 57.J.Brahms, A. M. Michelson, and K. E. Van Holde, J. Mol.Biol. 16,467 (1966). 58.T. Samejima, M. Kita, and F. Srtwada, to be published. 59. M. M. Warshaw, C. A. Rush, and I. Tinocw, Jr., Riorhrni. Rzophys. Rus.Commim. 18,G33 (1965).

ORD

AND

CD O F NUCLEIC

ACIDS

297

60. M. M. Warshaw and I. Tinoco, Jr., J .11401. Biol. 13,54 (1965). 61.M. M. Warshaw and I. Tinoco, Jr., J .MoZ. Biot. 20,29 (1966). 62. H. R. Mahler, B. Kline, and B. D. Mehrotra, J .MoE. Bwl.9, 801(1964). 63. M. Leng and G. Felsenfeld, J .Mol.BioE. 16,455 (1966). 64.J.Applequist and V. Damle, J .Am. Chem.Soc. 88, 3895(1966). 65. S. I.Chan, M. P. Schweizer, P. 0. P. Ts’o, and G. K. Helmkamp, J. Am. Chem. SOC.86, 4182(1964). 66. M. P.Schweirer, S. I. Chan, and P. 0. P. Ts’o, J .Am. Chem.Soe. 87,5241(1965). 67.S.I. Chan, B. W. Bangerter, and H. H. Peter, Proc.Natl. Acad.Sci. U.S .66, 720(1966). 68. C. C. McDonald, W. D. Phillips, and J.Lazar, J .Am. Chem.Soc. 89, 4166(1967). 69.P. 0. P. Ts’o, I. S. Melvin, and A. C. Olson, J .Am. Chem.Soc. 86, 1289(1963). 70.P. 0. P. Ts’o and S. 1.Chan,f . Am. Chem.SOC.86,4176(1964). 71.I. Tinoco, Jr., in“Molecular Biophysics” (B. Pullman and M. Weissbluth, eds.), p.269.Academic Press, New York, 1965. 72.H. De Voe, f . Chem.Phys.43, 3199 (1965). 73.I. Tinoco, Jr., .I. Am. Chem.Sac. 86, 297 (1964). 74.C. A. Bush and I. Tinoco, Jr., J .Mol.Biol. 23,601(1967). 76.P.K. Sarkar and J.T. Yang, Biorhrm. Biophys. Res.Commun.20,346 (1965). 76.M. GeIIert, M. N. Lipsett, and D. R. Davies, Proc. Natl. Acad.Sci. ti. S.48,2013 (1962). 77.C. A. Bush and J. Brahms, J .Chem.Phgs.46, 79 (1967). 78.R. C. Davis and I. Tinoco, Jr., Biopolymws 6, 223 (1968). 79.D. Glaubiger, D. A. Lloyd, and I. Tinoco, Jr., Biopolymers 6,409(1968). 80.P. 0. P. Ts’o, S. A. Rapaport, and F. J.Bollum, Biochemistry 6,4153(1966). 81.C. R. Cantor and I. Tinoco, Jr., J .Mol.Biol. 13, 65 (1965). 82. C. R. Cantor and I. Tinoco, Jr., BiopoZymers 6,821(1967). 83. J. N.Vournakkis, H. A. Scheraga, G. W. Rushizky, and H. A. Sober, Biopolymers 4,33 (1966). 8Sa. Y . Inoue, 6.Aoyagi, and K. Nakanishi, J .Am. Chem.SOC.89,5701(1967). 8%. G. Felsenfeld and S. Z. Hirschman, J .Mol.Biol. 13,407(1965). 84.A. M. Michelson, T. L. V. Ulbricht, T. R. Emerson and R. J. Swan, Nature 209, 873 (1966). 86. D. Poland, J. N. Vournakis, and H. A. Scheraga, Biopolgmers 4,223(1966). 86.J. Brahms, Proc. Roy.SOC.8397, 150 (1967). 87.A. Adler, L. Grossman, and G. D. Fasman, Proc.Natl. Acad.Sci. U .S.67,423 (1967). 87a.J. Brahms, J.C. Maurizot, and A. M. Michelson, J .Mol.Biol. 26,465 (1967). 87b.H. Simpkins and E. G. Richards, J .MoE. Biol. 29, 349 (1967). 88. R. F.Steiner and R. F. Beers, “Polynucleotides.” Elsevier, Amsterdam, 1961. 89.A. Rich and D. W. Green, Ann.Rev.Biochem. 30, 93 (1961). 90.A. M. Michelson, “The Chemistry of Nucleosides and Nucleotides.” Academic Press, New York,1963. 91.I. Tinoco, Jr.and D. N. Holcomb, Ann.Rev.Phys, Chem.16,371 (1964). 98.A. M. Michelson, J.MassouliB, and W. Guschlbauer, Proffr. Nucleic AcidRes. MoZ. Biol. 6, 84 (1967). 93.G. FelsenfeId and H. T.Miles, Ann.Rev.Biochem. 36, 407 (1967). 94.P. K. Sarkar and J. T. Yang, J .Biol. Chem.240,2088(1965). 96. D. N. Holcomb and 1. Tinoco, Jr., Biopolymers 3, 121(1965). 96.P. If. Sarkar and J.T. Yang, Rioehrmistq/ 4, 1238(1965). 96a.S. K. Arya, unpnhlishetl work.

298

JEN TSI YANG

AND TATSUYA

SAMEJIMA

97.T.Samejima and J. T.Yang, J .Biol. Chem.240, 2094(1965). 98.V. Luzzati, A. Mathis, F.Masson, and J.Witz, J .Mol.Biol. 10, 28 (1964). 99.A. Rich, D. R. Davies, F. H. C. Crick, and J. D. Watson, J .Mol.Biol. S,71(1961). 100.E. G. Richards, C. P. Flessel, and J. R. Fresco, Biopolymers 1, 431 (1963). 100a.A. M. Michelson and C. Monny, Proc.Natl. Acad.Sci.U .S .66,1528(1966). 100b. H. Simpkms and E.G. Richards, Biopolymers 6,551(1967). 101.A. Rich and D. R. Davies, J .Am. Chem.Soe. 78, 3548(1956). 102.G. Felsenfeld, D. R. Davies, and A. Rich, J .Am. C k m . Soc. 79, 2023(1957). 103.G. Felsenfeld and A. Rich, Biochim. Biophys. Acta26,457 (1957). 104. G. Felsenfeld, Biochim. Biophys. Acta29,133(1958). 106.J.Brahms, Nature202,797 (1964). 106. J. Brahms, J .Mol.Bid.11, 785(1965). 107. H. Hashizume and K. Imahori, J .Biochem. (Tokyo) 61, 738 (1967). 108.T.L. V.Ulbricht, R. J. Swan, and A. M. Michelson, Chem.Commun. p.63 (1966). 208a. C. R. Cantor, S. R. Jaskunas, and I. Tinoco, Jr., J .Mol.Biol. 20,39 (1966). 109.P. 0. P. Ts’o, G. K. Helmkamp, and C. Sander, Biochim. Biophys. Actu66,584 (1962). 110.G.K. Helmkamp and P. 0. P. Ts o, J .Am. Chem.Soc.83, 138(1961); Biochim. Biophus. Acta66,601(1962). 111. P. K. Sarkar and J. T. Yang, Arch.Biochem. Biophys. 112, 512(1965). 112.J. R. Fresco, in “Informational Macromolecules” (H. J. Vogel, V. Bryson, and J. 0. Lampen, eds.), p. 121.Academic Press, New York, 1963. 113.F. Pochon and A. M. Michelson, Proc.Natl.Acad. Sci. U.5. 53, 1425(1965). 114. A. Rich, Biochim. Biophys. Acta29,502(1958). 116.R. Haselkorn, Ph.D. dissertation, Harvard University, 1959. 116.A. Rich, Nature181, 521(1958). 117.D. R. Davies and A. Rich, J .Am. Chem.Soe. 80,1003(1958). 118.N. Davidson, J. Wilholm, U. S. Nandi, R. Jensen, B. Olivera, and J. Wang, Proc. Natl. Acad.Sci.U.S.68,111(1965). 129. R. B. Inman and R. L.Baldwin, J.Mol.Biol. 6,185(1962). 120.R. B. Inman, C. L. Schildkraut, and A. Kornberg, J .Mol.Biol. 11, 285 (1965). 121.M. J. Chamberlin, Federation Proc.24,144(1965). 122.D. R. Davies and R. L. Baldwin, J .Mol.Biol. 6 ,251 (1963). 123.R. Langridge, Abstr. WBl410thNatl. Meeting Biophys. SOC.,Boston, Massachusetts, 1966. 124.G. D. Fasman, C. Lindblow, and E. Seaman, J .Mol.Biol. 12, 630(1965). 126.M. R. Lamborg and P.C. Zameenik, Biochim. Biophys. Res.Cmmun. 20, 328 (1965). 126. C. M. Kay and K. Oikawa, Biochemistry 6 ,213(1966). 127.J. N. Vournakis and H. A. Scheraga, Biochemistry 6,2997(1966). 128.P.K. Sarkar, B. Wells, and J. T. Yang, J .Mol.Biol. 26,563(1967). 129.P. K. Sarkar and J. T. Yang, in“Conformation of Biopolymers” (G. N. Ramachandran, ed.), Vol.1,p.197.Academic Press, New York, 1967. 130.J. Marmur and P. Doty, Nature183, 1427(1959); J .Mol.Biol. 6 ,109 (1962). 131.B. Magasanik, in“The Nucleic Acids” (E. Chargaff and J. N. Davidson, eds.), p. 373.Academic Press, New York,1955. 132.R. Roue and M. Meselson, Proc. NatZ.Acad.Sci.U.S. 46,1039(1959). 133.N. Sueoka, J. Marmur, and P.Doty, Nature183, 1429(1959). 134. E.Fredericq, A. Oth, and F.Fontaine, J .Mol.Biol. 3,11 (1961). 136.S. Y. Wang and J. M. Hachagen, J .Mol.Biol. 8,33 (1964).

ORD AND CD OF NUCLEIC ACIDS

299

136.C. Zimmer, G, Luck, 11. Venncr, and J. I+%, Biopotymers 6,563 (1968). 137.R. Langridge and P. ,J. Gomatos, Sciancc 141,694 (1963). 138.T.Sato, Y. Kyogoku, S. Higurhi, Y. Milsui, Y. Litaka, M. Tsuboi, and K. Miura, J .MoZ. Biol. 16,180(1966). 139.T. Samejima, H. Hashizume, K. Imahori, I. Fujii, and K. Miura, J. Mol.Biol. 34, 39 (1968). 140.K. Miura, I. Fujii, T. Sakaki, M. Fuke, and S. Kawase, J .Virol. In press. 141.P. J.Oriel and J. Wilder, Nature214, 702 (1967). 142.A. Adamx, T .Lindahl, and J. R. Fresco, Proc.Natl. Acad.Sci. U.S.67, 1684 (1967). 143.J.R. Fresco, A. Adams, R. Ascione, D. Henley, and T. Lindahl, Cold Spring HarborSymp.Quunt. Biol. 31, 527 (1966). HarborSymp.Quant. Biol. 31, 539 (1967). 144.K. Muench, ColdSpring 145.N. Sueoka, T .Kano-Sueoka, and W. J. Gartland, ColdSpringHarborSymp. Quant. Biol. 31, 571 (1967). NucZeicAcids, Enzymes(in Japanese) 13, 146.M. Tsuboi and S. Higuchi, Proteinq 533 (1968). 147.W. Fuller, M, H. F. Wilkins, H. R. Wilson, and L. D. Hamilton, J. Mol.Biol. 12, 60 (1965). 147a.G.B. B. M. Sutherland and M. Tsnboi, Proc.Roy. SOC.8239,446 (1967). 148.R. Langridge, D. A. Marvin, W. E. Seeds, H. R. Wilson, C. W. Hooper, M. H. F. Wilkins, and L. D. Hamilton, J .Mol.Biol. 2,38 (1960). 1.48~. R. Langridge, H. K. Wilson, C. W. Hooper, M. H. F. Wilkins, and L. D.Hamilton, J .MoZ. BioZ.2, 19 (1960). 149.W. Fuller, F. Hutchinson, M. Spencer, M. H. F. Wilkins, J .Mol.Biol. 27, 507 (1967). 149a.S. Arnott, M. H. F. Wilkins, W. Fuller, and R. Langridge, J. Mol.Biol. 27, 535 (1967). 150.G.Milman, R. Langridge, and M. J. Chamberlin, Proc. Natl. Acad.Sci. U .S. 67, 1804(1967). 150a.S. Higuchi, M. Tsuboi, and Y. Litaka, to be published. 161. R. Langridge and J. Marrnur, Science 143, 1450(1964). 152.I. Tinoco, Jr., Advan.Chem.Phya.4,113 (1962). 153.I.Tinoco, Jr., J .Chim.Phys.66,91 (1968). 164.R. W. Holley, J.Apgar, G. A. Everett, J. T. Madison, M. Marquise, S. H. Merrill, 3. R. Penswick, and A. Zamir, Science 147,1462(1965). 155.J.T. Madison, G. A. Everett, and 13. Kung, Science 163, 631 (1966). 156.D. W. McMullen, S. R. Jasknnas, and I.Tinoco, Jr., Biopolymers 6, 589 (1967). 256a.R. I.Cotter, P. McPhie, and W. B. Gratzer, Nature 216, 864 (1967). 157.A. Blake and A. R. Peacocke, Nature208,1319(1965). 158.1 .McPhie and W. B. Gratzer, Biochemistry 6,1310(1966). 159. W. B. Gratzer, Biochim. Biophys. Acta123,431 (1966). 160. P. I<. Sarkar, J. T. Yang, and P.Doty, Biopolymer 6, 1 (1967). 161. G. M. Holzwarth and P.Doty, J .A m . Chem.SOC.87, 218 (1965). 162. J. T. Yang, in “Conformation of Biopolymers” (G. N. Ramachandran, ed.), Vol.1,p.157.Academic Press, New York, 1967. 163.E. R. Blout, I. Schmier, and N. S. Simmons, J .Am. Chem.Soc. 84,3193(1962). 164.M. F. Maestre and I. Tinoco, Jr., J. MoZ. Biol. 12, 287 (1965). 165.M. F. Maestre and I. Tinoco, Jr., J.Mol.Biol. 23, 323 (1967). 166.E. C. Pollard, “The Physics of Viruses,” p. 15.Academic Press, New York, 1953.

300

LEN TSI YANG AND

TATSUYA

SAMEJIMA

167.T. I. Tikchonenko, E. N. Dobrov, G. A. Velikodvorskaya, a d N. P. Kiseleva, J .Mol.Biol. 18, 58 (1966). 168.G. S. Stent, “Molecular Biology of Bacterial Viruses.” Freeman, San Francisco, 1963. 169.P. J. Oriel, Arch.Biochem. Biophys. 116, 577 (1966). 17 0. G. Zubay and P. Doty, J .Mol.BioZ.1, 1 (1959). 171.B.Jirgensons and L. Hnilica, Biochim. Biophys. Aeta109, 241 (1965). 172.E. M. Bradbury, C. CraneRobinson, D. M. P. Phillips, E. W. Johns, and K. Murray, Nature206, 1315(1965). 173.D. F. Bradley and M. K. Wolf, i n “The Neurochemistry of Nucleotides and Amino Acids’, (R. 0. Brady and D. B. Tower, eds.), p.80.Wiley, New York, 1960. 174.S. Brenner, L. Barnett, F. H. C. Crick, and A. Orgel, J .MoZ. Biol. 3, 121(1961). 176.A. Orgel and S. Brenner, J .Mol.Biol. 3, 762(1961). 176.L. S.Lerman, J .Cellular Comp.PhysioE. 64, Suppl. 1, 1 (1964). 177.L. Michaelis and S. Granick, J .Am. Chem.SOC.67,1212(1945). 17 8.D. M. Neville and D. F. Bradley, Biochim. Biophys. Aeta60, 397 (1961). l78a. V. 1. Permogorov, Y. S. Lazurkin, and S. Z. Shmurak, Dokl.Biophgs. (English Transl.) 166, 71 (1964). 179. A. Blake and A. R. Peacocke, Biopolymers 4, 1091(1966). 180.K. Yamaoka and R. A. Resnik, J .Phys.Chem.70,4051 (1966). 181.S. F. Mason and A. J. McCaffery, Nature204, 468 (1964). 182.B. J. Gardner and S. F. Mason, Biopolymers 6, 79 (1967). 185.A. Blake and A. R. Peacocke, Nature206, 1009(1965). 184. K. Yamaoka and R. A. Resnik, Nature213, 1031(1967). 186. K. Yamaoka and H. Ziffer, Biochemistry 7 ,1001(1968). 186.V. I.Permogorov and Y. S. Lazurkin, Biojizika 10, 17 (1965). 187.H. Ziffer, K. Yamaoka, and A. B. Mauger, Biochemistry 7 , 996 (1968). 188. H. R. Mahler, G. Green, R. Goutarel, and Q. Khuong-Huu, Biochemistry 7, 1568(1968). 189.Ping-Yao Cheng, Biochirn. Biophgs. Aeta102, 314 (1965). 190.P. A. Levene and E. S. London, J .Biol. Chem.83, 793 (1929). 191.B. H. Levedahl and T. W. James, Biochim. Biophys. Acta17, 453(1955). 192.B. H. Levedahl and T. W. James, Biochim. Biophys. Acta26,89(1957). 193.P. Doty, H. Boedtker, J. R. Fresco, B. D. Hall, and R. Haselkorn, Ann.N . Y . Acad.Sci. 81, 693(1959). 194. P. Doty, H. Boedtker, J. R. Fresco, R. Ha~elkorn,and M. Litt, Proc. Natl. Acad. Sci.U.S. 46, 482 (1959). 196.J. R. Fresco, Trans. N . Y .Acad.Sci. 21, 653 (1959). 196.M. N.Lipsett, Proc.Natl. Acad.Sci.U.S.46, 445 (1960). 197.G. Helmkamp and P. 0. P.Ts’o, J .Am. Chem.SOC.83, 138(1961). 198.J. R. Fresco, Tetrahedron 13, 185(1961). 199.P.0. P. Ts’o, Tetrahedron 13, 198(1961). ,%XI. P. 0. P. Ts o, G . Helmkamp, C. Sander, and F. W. Studier, Biochim. Biophys.

Acta76,54 (1963).

The Specificity of Molecular Hybridization in Relation to Studies on Higher Organisms

I. Introduction

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A. The Specificity of Hybridixdiori . . . . 11. Factors Affecting the Stability of the Formed Duplex A. Artificial Polymers . . . . . . . B. Natural Nucleic Acids . . . . . . . C. Effect ofNoncornplernentary Regions . , D. Discrimination andSequenceHomology . . . 111. Rates of Reassociation . . . . . . . A. Factors Affecting Rate . . . . . . B. DNA Families in Higher Organisnis . . . . IV. Practical Implications of DNA with DiBerent Repetition A. D N A . DNA Interactions . . . . . . B. D N A . RNA Interactions . . . . . V. Conclusions . . . . . . . . . . References . . . . . . . . . . ,

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1. Introduction It is seven years since Schildkraut, Marmur, and Doty (1) showed that hybrid molecules could be formed from the DNA of different bacterial species. It is therefore not inappropriate to look at what has become, particularly since the advent of the simple filter techniques (2, S), a popular but highly empirical art. Molecular hybridization has shown that mice and rats are more closely related to each other than either are to Escherichia coli, and we are prepared therefore to use the method in fields where no such comforting controls are available. It is not intended here to consider the details of the many techniques available, but to concentrate on two aspects of the method of prime importance in its extension to the DNA and RNA of higher organisms. Both result from the thousandfold greater complexity of the genome of higher organisms compared with that?of a bncterium. This causes a correspondingly greater chance ofthere being similar but not identical cistrolls in their DNA, so that thc limits of the spcrificity of molecular hybridization, important enough in microorganisms, becomes crucial in interpreting results from animals and plants. It is not that we are likely, except in very 301

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special conditions, to be able to identify a single cistron, but that the presence of families of cistrons, of a known degree of resemblance, niay grossly color any deductions we make about DNA-DNA or DNA-RNA homologies. This brings us to our second main consideration. It was apparent early in our investigations of the DNA of mice and rats that only part of the DNA reacts (4)in the agar-gel method and that this fraction (about a third) can be isolated from the nonreactive DNA. Britten and Kohne (5) showed that this was expIained by the presence in the DNA of higher organisms of related families of sequences, whose relative abundance could be measured by their rates of reassociation. Because fanlilies are defined substantially in the same way as the techniques used to detect DNA homology between species and RNA-DNA homologies between RNA from different functional classes or different tissues, a very complicated situation can occur. An incubation mixture may contain RNA molecules at widely different concentrations, whose interactions are to be studied with a DNA itself enormously heterogeneous. In this DNA, a cistron transcribing RNA may be unique in a genome with no cistrons of sufficient sequence similarity to enable them to react with these RNA molecules. On the other hand, another RNA may come from a cistron that is a member of a family whose members are sufficiently alike that all will react with this RNA. To avoid confusion, we define three related terms that will be used with refers to the general ability of special meanings in this chapter. Speci$cizty the hybridization method to distinguish between sequences of nucleotides refers to the even if they come from the same organism. Discrimination special case of specificity in which DNA s from different organisms are compared. Degreeof homologyrefers to the similarities in terms of base substitutions between two polynucleotide chains.

A. The Specificity of Hybridization The basic problem revolves around the specificity of the hybridization reaction; put qualitatively, we need to know how different polynucleotide sequences have to be before they fail to form a duplex or hybrid molecule. Quantitative definition is more difficult since it, involves a number of interacting parameters, the most important of which is the stable minimum length of n nucleotides. That is the shortJestsequence which, when bound to its exact complement, will st,abilizetwo polynucleotide chains to form a partially duplex molecule (6). The lengt’h of this sequence depends on the

following faotm:

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1. The ionic environment during the incubation. 2. The temperature of incubation (T,). 3.Whether the polymers involve deoxyribose, ribose, or a mixture. 4. The base composition of the sequence. 5. The nucleotide sequence itself; for example, whether purines and pyrimidines alternate or not. 6.The length of any single-stranded ends attached to the sequence. 7. The presence and length of other sequences separated from the first sequence by one or a few mismatched bases. Of these, the first two are under experimental contrd, and the effects of the remainder can be inferred from the numerous studies on the temperature of the midpoint of the thermal transition (T,)ofoligo- and polynucleotides. T,,the temperature ofrenaturation or reassociation, is related somewhat first showed, it has a broad loosely to T,, since 5ts Marmur and Doty (7a) peak with a maximum some 2 5 O less than the T,. In most hybridization experiments, the quality of the hybrid-that is, the value of n-is determined by T,and the ionic environment, although the hybrids once formed can be further studied by thermal chromatography. The latter allows the product to be separated into different fractions dependent on their stability at different temperatures-that is, on their TnL. For a general review of denaturation and renaturation in DNA, Marmur et al.(7b)should be consulted. So far, we have mentioned only factors affecting specificity, but since hybridization is a bimolecular reaction, the concentration of the reactants and the time of incubation are also important in governing the amount of hybrid formed. It is convenient, however, to consider these two aspects separately. In the following section we first consider the more general effects of salt concentration and polynucleotide structure on helix stability, before discussing the relation of chain length to T,,in synthetic and natural polynucleotides. This is followed by more circumstantial evidence on the ability of the method to discriminate between different DNA’s or RNA’s. I n Section 11,D the relation between the stable minimum length and the homology, in terms of base substitutions, between two natural polymer strands is considered. Section I11 considers the significance of rates of reassociation and the definition of a family of DNA cistrons in the light of our earlier section on the specificity of the hybridization technique. This part also presents a paradox and the biological implications thereof. I n Section IV, some of the practical limitations of the method are discussed. This article is therefore an attempt to define in more quantitative terms the evolutionary differences in DNA sequences detected by hybridization techniques (discussed also by McCarthy in Volume 4 of this series).

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II. Factors Affecting the Stability of the Formed Duplex A. Artificial Polymers 1.SALT CONCENTRATION

One of the main difficulties in extracting meaningful comparisons from published data is the wide range of counterion concentrations that have been used. A number of results (7-10) show that there is a linear relation between T, and log Na+ below 0.1M Na+, and that above 1 M Na+ the T, tends to level off. Mg2+ ions are about 100 times more effective in raising T, than are Na+ ions, and particular care may be necessary in eliminating all Mg2+ in annealing experiments. We would predict from this that for a given temperature of incubation during hybridization, the higher the salt concentration the shorter the stable minimum length.

2.BASE SEQUENCEAND

SUGARBACKBONE The stability of the duplex is affected both by the base composition (7u) and by the sequences of bases along the individual strands, in particular in THE

the mixed ribose-deoxyribose polymers (11-13). The general rule appears to be that for identical sequences (U # T), the ribose helix is more stable than the deoxyribose, but U-containing polymers are less stable than their T analogs so that poly rA . poly rU melts 12°C lower than poly dA poly dT in 0.1 M Na+. In the mixed ribose-deoxyribose polymers, helices containing purine deoxynucleosides are markedly less stable than those with purine ribonucleosides. This is true for adenine, hypoxanthine, and guanine (1.6) and causes poly dA . poly rU to have a T ,at least 19" lower than poly rA poly dT. There is also a curious relationship between the stability of the mixed compared to the all-deoxyribose polymers. Thus, while poly dA poly rU is less stabIe than poly dA . poly dT, poly dG . poly rC is more stable than poly dC . poly dG, so that both the mixed polymers in the C . G series are interniediat,e in stability between poly dC . poly dG and poly rC . poly rG. The greater range of stabilities of the RNA-DNA compared to the DNA-like polymers should, as Chamberlin and Patterson (12)pointed out, cause a greater breadth of thermal transition in a DNA RNA hybrid compared to its parent DNA. This was found when +X RNA. DNA hybrid was compared with duplex 4X DNA (15). These interrelations between polynucleotides of different base composition result from the relative contributions of stacking and hydrogen bonding for the different base pairs. Other results with more complex polymers have,

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presumably similar explanations; Wells etal.(16) showed that poly d(T-C) poly (A-G) has a T ,5" less than poly d(T-G) . poly (A-C); and Riley etal. (13) found that the alternating poly d(A-T) has its T , 7.5"lower than poly dA poly dT. For mixtures containing ribose, an addit,ional factor affecting stability is the 2'-OH group.

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3. CHAIN LEKGTHS

Several studies have been made on the effect of chain length of artificial and by Michelson and Monny polymers on T,,chiefly by Lipsett etal.(17) (18) for the poly A . poly U series. Comparable information for the poly Cr poly C series has been more difficult to obtain owing to the high stability of G G interact,ions, which gives poly G a highly stable intrastrand secondary structure and reduces interactions with C. However, Lipsett (19) has investigated oligo G . poly C stahilities, which are more relevarit to hybridization reactions. She finds that in 0.2M Na+, ( G P ) ~has a T ,of 58",

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FIG. 1. Therelationbetweenlength of oligomer and T ,for various invztro synthesized A, Oligo rG poly rC, from Lipsett (19) in 0.2 M NaCl polynucleotides. A0.002 M Na cacodylate, pH 6.2.00 , Oligo rA . 2 poly rU from Michelson and Monny (18) in 0.1 M NaCl 0.01 MgCla 0.05M Na cacodylate. AA, Oligo rA . 2 poly rU,from Lipsett etal.( 17)in 0.001 M MgClz 1M NaCl 0.002 M K2P0,,pH 7.4 - - - 0 , Oligo rU . poly rA, as above. U--B or 0, Oligo dA poly dT,from Cassnni and Bollurn (20) in 0.04 M H phosphate pH 7.0 0.008 M MgC12. Open syrnhol vnliie was ralciilatcxl from the rel:iiiotr given in thereference.

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and predicts that ( G P ) ~would ~ have a T , of 83°C. Cassani and Bollum (20) have made similar studies of the interactions of (dA), and (dT)%and find a linear relation when the reciprocals of chain length and T, (OK) are plotted together. Some of these results for oligomer . polymer stability are plotted in Fig. 1. There are certain discrepancies in the values for the long-chain polymers between these results and those from Chamberlin’s group, perhaps due to differences in salt concentration. The main finding is that, in general, oligomers of chain length 2 2 0 will form duplexes very little less stable than the polymers. The exceptions to this are the oligo dA poly dT series, where longer sequences may be needed for stability, and we would predict from Section 2 above, that even longer chain lengths will be needed for oligo rU poly dA. The C . G data are less complete, but the steepness of the curve for oligo rG poly rC indicates that the much less stable deoxypolymers and their mixed analogs, will still form duplexes with a very high stability when n 2 10. This evidence suggests [see also the discussion by Thomas (6)] that for DNA . DNA duplexes of average composition, the “stable minimum For RNA DNA hybrids, this length’’ may be in the region of 10-20. sequence may be the same unless there are considerable stretches of A in the DNA.

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B. Natural Nucleic Acids 1.COMPLEX SEQUENCES

One natural DNA sequence, now largely known, has been studied; this is the single-stranded ends of phage A. Wang and Davidson (61), have studied its melting by band sedimentation, which distinguishes closed and open circles. They find T, sof 50 and 63 in 0.1M and 2.0M NaCl, which compare well with those found by infectivity assay (22). These ends have a sequence of 20nucleotides on each strand; of these, 15are G and C, with 12 clustered as a run at one end of the sequence. By infectivity assay in 0.01M Mg2+:the T, is 61 ,but this is reduced to 42 if four guanine nucleotides are added with DNA polymerase to an end adjacent to the double-stranded part, thus reducing the length of the effective binding site by probably more than 4 nucleotides. Before the publication of this sequence, Kallenbach and Crothers (23), predicted from thermodynamic considerations that the shape of the melting curve, if only G . C base pairs were involved, would best fit 11 base pairs if some stacking interaction is allowed. On the other hand, 25 base pairs would fit an all A . T sequence best, but with considerable base stticking. This gives some measure of the range of size to he expected from extreme compositional bias in thc oligonucleotides.

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Since oligonucleotides of defined sequence in the critical range of 10-20 bases are now becoming available (24), it should not be long before we know the precise T ,for these defined sequences of known length. 2.HYBRIDIZATION EXPERIMENTS WITH NATURAL POLYNUCLEOTIDES OF DEFINED LENGTH

Four such experiments are known to the author, which give somewhat contradictory results. These four experiments are essentially concerned with T,, the temperature of renaturation, although McConaughy and McCarthy (25)released the duplexes from the filters by increasing the temperature. They broke mouse DNA into short chain lengths by acid depurinatian and subsequent hydrolysis and then separated the oligomers on DEAE-Sephadex. The oligomers and denatured DNA were heated to 60°C in 0 . 5 M KC1 and then slowly cooled to 20" over 10 hours before trapping on filters. The trapped duplexes were then released from the filters by increasing temperature. This is not, as they acknowledge, a very sensitive test since many of the duplexes trapped may be stabilized by very short regions. Nevertheless, they found increasing stability as the chain length was increased from 20 to 200,and specificity of interaction between different DNA's with chain lengths of 33.In another experiment, they used oligomers from T4 DNA and found that, although the amount of oligomer retained increased from a length of 17 to one of 28nucleotides, the T , did not. In comparable experiments, GiIIespie and Spiegelman (26)partially separated the RNA into size classes digested ribosomal RNA from E.coli, and incubated oligomers of known chain length with DNA at various M Na+ a t 67",50 nucleotides were temperatures. They found that in 0.33 needed to give a stable duplex, while at 55" and 44",only 32and 17 nucleatides were required. On the other hand, Niyogi and Thomas (27’) syntheon T4 or T7 DNA primers and digested it with T1 sized RNA in vitro ribonuclease before separating the oligomers on DEAE-Sephadex. Below the decamer, no binding to the homologous DNA immobilized on filters couId be detected at any temperature of incubation in 0.825 M Na+ after 6 hours. Oligomers containing 11 or 12 nucleotides showed a dramatic increase in the amount bound, which began tjolevel off after 14nucleotides. They conclude that only 12 perfectly complementary nucleotides are using essentially needed to form a stable complex. Riiger and Bautz (282, the same methods, confirmed that 12 nucleotides are sufficient to distinguish T4 DNA from various deletion strains. They report a lower optimum temperature of incubation for T4 than do Niyogi and Thomas for T7,which is consistent with the different base compositions of T4 and T7 DNA.

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These figures are very much snialler than those of Gillespie and Spiegelman, and it does not seem likely that differences in salt concentration in the two experiments account for the difference. Another explanation is suggested by the much simpler genome used by In their experiments, both the ability of even 12 Niyogi and Thomas (27). nucleotides to discriminate between T7 DNA and T4 DNA and the small by the largest oligomer tested suggest that fraction of DNA covered (3.3y0) there is very little cross-reaction between similar DNA sequences and that the RNA is annealing with the same cistron that transcribed it. In higher organisms such as the mouse, studied by McConaughy and McCarthy (25), there may be many related sequences, each differing slightly from the others (see Section 111, B, 1 for a fuller discussion of this point). For any particular 12-nucleotide stretch, the chances of finding its exact complement will be fairly small, because many of the related sequences will have differing bases somewhere within that sequence. If the length of the oligomer being tested is increased, the chance that one stretch of twelve anywhere along the oligomer can find its exact complement is increased n - 12times. For this reason, lengthening the oligomer does not affect the stable minimum length or the degree of sequence homology derived from it in Section 11,D, 3. This discussion raises two more questions. So far we have assumed that a perfect complementary sequence of known length is necessary before binding occurs. What is the effect of mismatching of bases, and can a longer sequence compensate for less than perfect complementarity? Will it be necessary to increase the length of an oligomer if it has to support a length of unbound single strand?

C.

Effect of Noncomplementary Regions

1.MISMATCHED BASES

The effect of mismatching introduced into poly A . poly U has been studied by Bautz and Bautz (99).From their results, we can predict that in oligomers longer than 10, the presence of an additional mismatched base at the end of the sequence does not have a large effect on its stability. They also tested a poly r(A,N). 2 poly rU system in which N varied from 0 to 30% of A N and could be C, U, G or I. G and I had the least effect, and U and C a progressively greater one. On average, 30% of mismatch lowered by 20" in 0.5M NaCl and 0.01 M Trig. The T, of the pure poly A the T,,, 2 poly U (77 C) is the same as in Fig. 1, and it is tempting to read off the length of the oligomer corresponding to the lower T, of the polymer with 30% mismatched bases. It is 9 nucleotides, which does not correspond to the size ofaverage runs of A's in the 30% case. Despite the inadequacy of

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this model for the DNA DNA or DNA RNA situation, we can conclude that, while there is extensive destabilization, purine sequences probably contribute by hydrophobic interactions to the stability of a neighboring sequence separated by a nonpairing base. It would be instructive to know whether purines introduced into pyrimidine runs produce a greater effect owing to the greater dislocation caused by purine-purine interaction. An interesting and relevant result is provided by a comparison of the renaturation behavior of the artificial poly d(A-T) and the naturally occurring A-T rich satellite DNA of crab. Poly d(A-T) has the same T , on a second heating unlike the renatured crab DNA which has a T , 5" lower than in the native state (30). The latter has also a small percentage of G C (<3%),which nearest-neighbor studies show are not conIt is therefore probable that the C's and G's are centrated into runs (31). rarely in register in the renatured molecules, and that this is sufficient t o make the polymer behave as though it were composed of oligomers of about 30 nucleotides (see Fig. 1). This fits quite well with the expected distribution of average intact chain lengths, particularly if G A interactions are more disruptive than are C . T.

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2.STABILIZATION OF SINGLE-STRANDELI

REGIONSBY

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DUPLEXSEQUENCE

The often large amount of labeled RNA removed from filters by RNase digestion shows that long lengths of single-stranded RNA can be held by a duplex sequence a t high temperatures of incubation. A similar process best explains certain results with reassociated DNA, such as those on minor the components of mouse satellite DNA reported by Flamm et al.(32), higher T, obtained by thermal chromatography compared to optical spectroscopy, and the simplificationof band patterns after enzyme digestion reported in the original DNA DNA homology experiment (I). Zimmerman et al.(33)in designing an assay for the E. coliDNA ligase, made use of this property and were able to show that single-stranded DNA could be enzymatically joined to cross-linked and duplex DNA immobilized on hydroxylapatite. In these experiments} therefore, duplex regions with long single-stranded ends behave like duplex DNA. Crothers et al.(3.4) have shown that DNA with less than a thousand base pairs normally melts from the ends of the molecules while longer lengths may also support internal looping-out. The relative rarity of internal loops is due to the greater energy required to break the considerable interactions a t both sides of the initial base-pair compared to the single position a t an opening end. Similar considerations favor one large internal loopcompared to several smali adjacent ones. Cooperative melting may be aided by the

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increasing contribution made by the diffusional and repulsive energies of the growing free ends to the breaking of the hydrogen bonds and hydrophobic interactions of the adjacent bases. The same arguments can perhaps be applied to the rather different situation presented by the reassociation of polynucleotide strands that are only partially complementary. Here free ends or internal loops are prevented from closing as the temperature is lowered by mismatched bases. These loops and ends will make an energy contribution tending to disrupt the duplex region. The minimum sequence that will stabilize an oligomer to a polymer must therefore be increased in order to compensate for this.

D. Discrimination and Sequence Homology 1. DISCRIMINATION

So far we have considered onlithe stability of sequences in a homologous incubation and have ignored the problem of discrimination-that is, the ability of a given sequence of RNA or DNA to distinguish a DNA of diff erent origin. Surprisingly, even the 12-nucleotide sequence derived from phage shows discrimination (27, 28). This again may be a function of the relatively small size of these phage genomes, but discrimination is also detectable in the McConaughy and McCarthy experiments with DNA chain lengths of 33.Nucleotide sequences of about this length from mouse DNA gave much less stable products when incubated with hamster or E. coZiDNA than when incubated with mouse DNA. We can summarize the foregoing sections by saying that for DNA RNA interactions, a minimum of 12 nucleotides is needed to form a stable duplex at relatively low temperatures of incubation in high salt. This length is dependent on base composition and must be longer if the temperature of renaturation is increased or the salt concentration decreased. The DNA . DNA situation is not so clear; while DNA . DNA duplexes are less stable than RNA RNA (11, 35,36)) optical melting studies (15) and elution of fragments from agar columns (37)show that DNA RNA hybrids are less stable than DNA. DNA duplexes, owing, in part at least, to the low stability of the rU dA pair. This favors a minimum stable length of less than 12 for DNA * DNA hybrids, but the rather low stability of the cohesive ends of X (22) contradicts t,his. It therefore seems probable, taking all the data into consideration, that DNA DNA hybrids of average composition require about the same minimum stable length as DNA . RNA hybrids. In both cases, the sequences must be longer if they are partially mismatched and if a short sequence has to stabilize single-stranded ends. Even so, it does not seem likely that much more than 20 nucleotides are needed.

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Factors increasing the minimum stable length are shown diagrammatically in Fig. 2 .In practice, single-stranded ends and bops may not be very large since all DNA sequences are related, however distantly, by the relatively simple steps of base substitution, inversion, and translocation. It is therefore highly probable that any DNA sequences having 20nucleotides in common will also share others along their length. This brings us first to a technical point regarding the interpretation of dat.a on RNA . DNA hybrids after RNase treatment and then to a discussion of sequence homology. 2.REMOV-iL

OF UNBOUND

P A R T S OF THE

MOLECULE

It is implicit in this discussion that loops of single-stranded DNA or RNA must join duplex regions of various degrees of stability, and it is the practice to remove these loops and ends with RNase in DNA/RNA experiments, but to neglect this step in DNA/DNA work because no suitable enzyme is readily available. McCarthy (58) has questioned the RNase procedure, and it does appear that in many instances the method gives results difficult to interpret. In phage, possibly in bacteria, and in special systems in higher organisms (such as ribosomal cistrons and highly repetitive satellite sequences), RNase treatment is probably justifiable since in such “simple” DNA, the presence of 12 nucleotides complementary to the RNA carries the high probability that the remainder of the DNA is also complementary. RNase treatment will remove the small fraction of loops and free ends present as a result of the vagaries of chain-scission during preparation. It will also remove unbound RNA that might otherwise be stabilized by short (<12) regions formed on cooling the filter after incubation. A very different situation exists in higher organisms for DNA not in those categories mentioned above; here the chances of a messenger RNA binding to its parent cistron and forming a perfect duplex are remote. Nearly all of it will bind to regions with sequences more or less similar to the parent cistron. I n such a situation, the effeet of RNase is to leave the DNA covered by short segments of RNA, whose distribution and quantity depends on the resemblance of the DNA to the various RNA species added (Fig. 3).If, instead, the hybrids are eluted from the support medium at higher temperatures, then only those molecules with the degree of homology It is then possible to defined by the temperature will be measured (39). measure the amount of DNA having cistrons like the RNA or DNA with which it is challenged rather than in terms of the number of short sequences that have reacted.

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During annea I ing Cool I

(C)

During annealing

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FIQ.2. Factors effecting the minimum stable length ( M S L ) .(A) The oligomer polymer situation, where the MSL is drawn with 7 nucleotides for convenience (B). The oligomer iY longer but can bind over only part of its length. The binding region a t

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( A ) In phage

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r , 1 PIG.3. Differences in the effect of RNase on RNA . DNA hybrids in (A) microorganisms and (B) more complex organisms. I n microorganisms, sequences that find complements probably do so over most oftheir length, because few, if any, ditTerent long sequences have a “minimum stable length” in common. The enzyme will remove only unbound ends, as shown. I n higher organisms the hybrids have internal regions that cannot bind. These will be excised by the enzyme, leaving short bound regions that give little quantitative measure of the RNA similar to the DNA. In this figure, t2helengths ofthe bound and unbound regions aredrawn shorter than in the real situation. Thin and thick lines indicate RNA and DNA, respectively.

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the annealing temperature is increased to MSL 2 to compensate for the unpaired ends. On cooling, other regions (y), too short to form duplex regions a t high temperature, can now bind. (C) Very long mismatched polymer . polymer interactions. A long matched region (MSL z )is needed t o give II high probability of other regions longer than the M S L ( M S L a and M S L b)occurring a t other regions along the length. On cooling other short regions (c) reform as described in (B) above.

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If this argument is accepted, the molecular weight of the reactants becomes important, since the amount of RNA or DNA bound to DNA by a given nucleotide sequence will increase with the size of the fragment. On the other hand, as the fragment size increases, so does the chance, given random base substitutions, of other sequences appearing that can also bind to the molecule. But because there is no reason to believe that, in polycistronic messengers, the order of related cistrons shouId be the same in different organisms, it would seem reasonabk to keep the fragment size to about that of the cistron. 3. CALCULATIONSON SEQUENCEHOMOLOGY

Let us consider two partially complementary single strands of DNA, each 500 nucleotides in length. If n is the minimum stable length, how many base substitutions can occur in either strand before no such sequence of length n remains? At one extreme there could be cistrons coding for two enzymes with identical active sites of n/3 amino acids. All other amino acids would be different and there would therefore be not more than 4 complementary nucleotides at equivalent positions on the chains, although by looping-out it might be possible to obtain positions with a longer sequence in register at nonequivalent positions. At the other extreme, the base substitutions could be spaced n - I nucleotides apart to give the minimum number of substitutions (500/n),which would destroy all homology between the strands. Because proteins and their cistrons have evolved from common ancestors, a more realistic model is one in which base substitutions occur randomly along the chain. It is possible to simulate this situation with a random-number generator and a suhable computor program (designed by H. Watkins), and to calculate the number of hits required to leave intact different percentages of the chains with various values of n. The results for a large number of runs at each value of n are shown in Fig. 4. The results are, in principle, directly applicable to homology experiments, since if we assume n = 40,then 25% homology between two strands 500nucleotides long implies that they differ by 66/500or 13% base substitutions. These are probably minimum figures since, except in the case mentioned earlier where the substitutions are equally spaced, other nonrandom configurations should allow more substitutions before the sequence, n, is extinguished. Mismatching, even when compensated by a longer sequence that is otherwise complementary, still further increases the number of substitutions allowed, since the effective complementary sequence must always be shorter than the product of n and the number of mismatched pairs, if no sequence n is to be found.

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I N HItiHER ORGANISMS % ALTERED BASES (-

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FIG.4. Calculation of the percentage of sequences of a minimum stable length (n), which are left after a given number of “base substitutions” in a 500nucleotide sequence length. Either 500 (n = 30-70)or 250 (n < 20)separate runs were m d e , in which random numbers (1-500) were generated to give the position of the base substitutions, and the number of hits needed to extinguish the sequence length (n) were noted. The percentage intact sequences refers to the fraction of runs at which a given number of hits per 500bases extinguished the sequence (- - -). The dotted lines therefore represent the number of “mutational events” that must occur, but, particularly for smaller values of n, a proportion of the hits will occur a t thesame position and some ofthose will revert to the original base. The full lines -( ) allow for multiple hits and give approximately the percentage of altered bases. Three hits a t the same position have been neglected together with reversions adjacent to the last intact sequence.

A major result of this calculation is the large number of base substitutions that can occw before any minimum complementary sequences of, say, 20 are lost. Substitutions up to 14% can be toIerated before hybridization methods can begin to discriminate between the DNA from two species. It is also important to note that thereafter the percentage of intact sequences falls relatively rapidly, and that all homology is extinguished in this example after 42% of the bases have changed. These results have farreaching implications for the interpretation of both DNA DNA homology experiments and the reassociation data from the DNA of higher organisms. In the next section we shall therefore consider the significance ofrates of reassociation of DNA, before returning to the implications of these calculations.

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12.0

.2

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2I 0.365 a 0.375

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FIG.6. (T o p )The relation betweenanalytical complexity or molecular weightand thekinetic complexity obtained fromrenaturation data. (Bottom) Rateconstants forthe renaturation of ratascites tumor DNA. The rateconstants are6.4l/mole-sec and 2.6X 10-8l/mole-sec for theintermediate andslow fractions ofthis DNA. From Wetmur (10). and Davidson

111. Rates of Reassociation A. Factors Affecting Rate In their first paper on DNA renaturation, Marmur and Doty (40) showed that phage DNA renatures more rapidly than that of bacteria in conditions in which calf thymus DNA does not renature at all. It has been generally accepted that these differences are due to the increasing complexity of these genomes, since if the fragments of DNA are all broken to the same size during preparation, the chances of a single strand finding a complement,would be the greater, the smaller the genome. This relation has

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been extensively studied by Rritten and Kohne ( 5 )and Wetmur and Davidson (10). The latter have also extended the study (8, 41-43)of the effect of various factors on the rate of reassociation of DNA. Their findings can be summarized as follows : the renaturation rate is1. inversely proportional to relative viscosity. 2. proportional to the square root of the single-stranded molecular weight ofthe DNA. 3.slightly faster for DNA rich in G and C. 4. independent of pH between 5 and 9 at high salt concentrations. 5.highly dependent on salt concentration. 6.inversely proportional to the complexity of the DNA-that is, to the size of the genome-if repetitive fractions are neglected. The latter relation is illustrated in Fig. 5,compiled from Wetmur and Davidson’s data. This also shows the renaturation kinetics of two fractions of rat DNA, the slower of which requires incubation at 70 m g / d for 1 hour before it would be half-renatured. This explains the failure of Marmur and Doty to find a significant degree of renaturation in the similar DNA from calf thymus. Why then did Hoyer et al.(44) obtain reassociation in DNA from higher organisms, using their DNA-agar technique? One difference is that they used DNA of low molecular weight as one component of the incubation mixture. This releases minor fractions that could react more rapidly than most of the DNA. But they also published evidence that they interpreted as showing that all the DNA could, in their experiments, form duplexes because at high inputs of lowmolecular weight DNA, nearly as much of the latter was bound as there was high-molecular weight DNA trapped in the agar. Walker and McLaren (4), who obtained a similar result, also showed, as mentioned in the introduction, that the DNA fragments could be separated into two fractions, a minor one, which bound to the DNA in agar, and a major one, which did not. Presumably at high input concentrations, some of the low-molecular weight DNA can bind to other such fragments, which are in turn bound to the DNA trapped in the agar. This would result in branched, bushlike structures and a considerable increase in the apparent length of the trapped DNA covered. We now know from Britten’s work on the rates of reassociation of DNA that the minor component contains “repetitive” DNA organized into families, while the major fraction is nonrepetitive and probably contains one or a few copies per haploid genome. In addition to these two fractions, we have found it convenient to separate from the repetitive fraction an even more highly repeated fraction with a G C content that sometimes allow it to be separated by CsCl gradient centrifugation as a satellite band. We can therefore, describe three broad classes of DNA in higher organisms.

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1.Fast. This may not be present in all such DNA s but can sometimes be isolated as a satellite band. It is repeated from perhaps lo5to 5 X lo6 times per genome and may comprise as much as 15% of the DNA in a nucleus. This may be up to 70% of the DNA, but is usually 2.Intermediate. l0-20%.It occurs in all the higher organisms studied by Britten and Kohne (5)and probably contains a number of families repeated lo2to lo5 times. 3. Slow. The remainder of the DNA, which appears to have only one or a few repeats per genome. The degree of repetition can be determined from the rate of reassociation and the fraction of the genome. Intermediate DNA, because of the size of the fraction of the genome it occupies and the comparatively small amount of repetition, must contain a number of different classes of molecules that reassociate, which Britten has called families. The reassociation rates, which define families, have been measured by ultraviolet hypochromicity, optical rotatory dispersion, and hydroxylapatite fractionation-that is, by techniques essentially similar to those used in hybridization experiments. The same restrictions on specificity and sequence length must therefore also apply to the definition of a family.

B. DNA Families in Higher Organisms 1.WHAT DEFINES A FAMILY? If we accept the burden of the earlier discussion, we must accept that low molecular weight DNA from the same organism will reassociate if they have a complementary sequence of about 20 nucleotides. If the length of the DNA strands is, perhaps, 200 nucleotides, then there will also be other complementary sequences contributing to the stability and hypochromicity of the duplex. It is also evident that strands can make use of different sets of sequences in their reactions with other strands; consequently one molecule can be a member of several different families. Consider an enzyme with s short amino acid sequence in, let us say, a region of an a-helix, and with another sequence at its active site. There may be many other proteins with the same peptide sequence in their helieaI regions, and these proteins will have DNA cistrons that may reassociate to form a numerous family. The active-site peptides may also be common to a smaller number of other enzymes. Their cistrons may constitute another family with a lower repetition rate, which need not overlap the larger to any great extent.

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2.THE CENTRAL YARADOX

This brings us to a paradox central to any discussion of annealing procedures: if the sequences are so short that many base substitutions can be tolerated, why is all the DNA of higher organisms not in families instead of the 10-207&generally found, and why is it possible to detect large differences in DNA homology in even such closely related species as the laboratory mouse and rat? On the face ofit, if all amino acid sequences have changed as slowly as those few whose sequences have been determined in a wide range of animals, we should have difficulty in discriminating between DNA from quite distantly related animals. We have recently considered this problem from an evolutionary viewpoirit (45)and can summarize the argument as follows: When mouse intermediate DNA is hybridized with whole mouse or rat DNA, we C ~ R detect only 8% as much reassociation with DNA from the rat, compared to DNA from the mouse, if rigorous procedures for detecting DNA. DNA homology are used. Taking a liberal interpretation of the length of the intact sequence to be 40 nucleotides in a strand of 500,the computor simulation results (Fig. 4)show that an average of 80 out of 500could have changed during the divergence of rats and mice. There is evidence that slowDNA isat least as different as intermediate DNA in the few comparisons made (5), so we can extend this proportion of base substitutions to the whole genome (neglecting the satellite DNA), and so obtain a figure of about 8 X loa base substitutions that must have occurred since mice and rats diverged from common stock. This may have been 5 X lo6 years ago, but Kimura (46) has calculated that one base substitution per 2 years might be expected by extrapolating data obtained from the rate of sequence evolution in the few proteins studied to the whole genome. Even allowing for the shorter generation time of rodents, there is a considerable discrepancy between the peptide substitutions expected and the nucleotide substitutions found, which seems unreasonable in any view of the rate of evolution of peptide sequences. There seem to be three ways of resolving the paradox: 1. The technique is really much more precise than all our previous discussion indicates. It is worth noting that even if we increased the stable minimum length to 80,the discrepancy is not much reduced. No evidence suggests that perfect matching is essential. 2.The DNA in different organisms is arranged differently. Translocation of cistrons is probably common, but since, in our experiments at least, all components are less than cistjronsize, it is difficult to see how the method can be sensitjiveto cistron arrangements.

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3.The proteins so far studied are atypical. 4. The differences are due to degeneracy in the code. This seems a possible explanation. If the wobble hypothesis (47) is true, there may be little or no selective advantage in using some of the alternative third code letters in synonymous codons. Mutations at these third positions will then be much more easily preserved than in the other two. The question then resolves itself into a consideration of whether a random selection of alternative bases in cistrons coding for similar proteins would allow these cistrons to hybridize. There is evidence (48)that vertebrates preferentially use some of the E. coliassigned codons, and fail to use others, notably the codons for alanine and serine containing C and G. Despite this, there are still many alternative codons available to these animals, which for most proteins ).a minimum complementary averages about 2 codons per amino acid (46 If sequence of n nucleotides is necessary for binding, the number of alternative sequences in proteins of average composition, is approximately 2 . This implies that for n = 20 a set of proteins using only one of the available alternatives for each amino acid would have its DNA organized as a family with a reassociation rate 400 times faster than the same set using instead a random selection of code letters. These proteins could therefore be transcribed from either the intermediate or slowclasses of DNA, depending on the selection pressure in favor of specific codons during the evolution of the species. If there was no selection for one rather than another of the alternative codons, all the DNA would reassociate a t the slow rate and there would be no families of cistrons. On the other hand, if the presence of only one codon a t matching positions in related proteins was advantageous, then it is possible, given the rather small degree of similarity that allows DNA to reassociate, that all the DNA would be in families. If all DNA codes for proteins, this might still allow, for example, about lo* sets of related proteins each with lo3 members. The absence of selective advantage for specific codons could also result in two related organisms having DNA without detectable homology. Homology wiIl be shown only if two related organisms use at least some of the alternative codons in common. We can detect some homology between the DNA of, for example, rats and mice, and since their DNA is organized in families, they do not use a random selection of the available codons for all proteins. But is the majority of their DNA cslowor l“single copy” because the majority of their proteins do not have 7 amino acids in common with any of the other lo7proteins that, may he made a t some time in the life of n rat or a mousc?

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A hypothesis, that the 70% of rodent DNA in the slowfraction is simply that part of the genome for which there has been no selective advantage in using specific codons, accounts rather well for a number of anomalous results difficult to explain in other ways. First, it explains how 49). closely related organisms can have demonstrably different DNA (4.5, Second, it provides an explanation for a number of Britten and Kohne’s (5) important results. They have shown that the size of the intermediate fraction varies considerably; the ox has 40% in this fraction compared to the mouse with 30%,of which at least 10% is satellite DNA, and to salmon and Amphiuma with up t o 70%. There is surely no fundamental reason why the proteins of these vertebrates should be so different, and it is much more reasonable to invoke a trivial difference involving, for example, the availability of certain transfer RNA’s or the need to prevent secondary structure in messenger RNA (50) as the cause of the bias of the DNA toward a more ordered selection of codons. As stated earlier, if different organisms have DNA in which there has been no selection for specific codons, we would not expect much homology between the DNA. This is exactly the situation Britten and K o b e found when they incubated labeled slow (single copy) DNA from the mouse with high concentrations of whole DNA from rat or ox.Those sequences that did reassociate in the r a t . mouse case were of much lower stability than those in the mouse * mouse control. They have also shown a lack of DNA with degrees of repetition between 2 and 100 (see also Fig. 5),that is between slowand intermediate, which is diffcult to explain on a plain evolutionary hypothesis, since one would expect some recent members of the 100-1000 class to be now in the 10-100 class. They explain it hy supposing that some sequences are preferent,idly and repeatedly duplicated during evolution. It is argued here that all or most proteins might share 7 amino acids with at least 100 other proteins, and therefore qualify as members of a DNA family. For a proportion of these proteins (which varies in different organisms), there is, however, strong selection in favor of retaining the use of specific codons at corresponding positions in their cistrons. For the remaining proteins, there is no such selective advantage in uniformity, and any alternative codon may be used. It is not of course essential to this hypothesis that there be no “unique” polypeptide sequences in this relaxed sense, only that the differences in the proportion ofDNA organized in families and the lack of homology between related DNA should be accounted for by a rapid evolution of the DNA not reflected in changes in the protein. Somewhat unexpectedly, molecular hybridization may be useful in distinguishing species or even subspecies and

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races, not because the technique is so sensitive, but because their DNA s are so different.

IV. Practical Implications of DNA with Different Repetition Rates Without related families in the DNA of higher organisms, molecular hybridization would, except in bacteria and viruses, have been unrewarding. With them, we are certainly restricted, but a knowledge of the restrictions might at least prevent naive and optimistic conclusions.

A. DNA DNA Interactions We can meaningfully compare DNA from different species if we know the rates of repetition of the various fractions. The lower limit to the degree ofrepetition is set by the incubation conditions, but anomalous results may be obtained because of the presence of a highly repetitive fraction in the DNA. For example, asymmetrical results were obtained by McLaren and in their comparisons of rat and mouse DNA, which are due to Walker (51) the size of the fast-reassociating DNA in the mouse. A problem raised by the random selection of codons and its effect on the DNA is whether this fraction ofDNA has size and nature ofintermediate the same functional significance in related organisms. It is not certain that it should code forthe same spectrum of proteins in different animals, or, put another way, whether it is possible that the hemoglobin cistron may be one of a number of related cistrons in one animal, but in the single-copy DNA ofanother. Further work is clearly necessary in studying the organization of the genome; relationships between DNA families in related organisms reflect their evolutionary history ; the relationships between the families of the same organisms may also have a functional significance in differentiation and speciation.

B. DNA * RNA

Interactions

This is a much more complex situation, since the fraction ofthe genome transcribed and the number of copies of RNA made from any one site must vary considerably in different tissues and at different times. We presume all this occurs against a background of DNA, invariant except for the multiplication ofribosomal cistrons during oogenesis (52). One of the consequences of the varying rates of reassociation of the DNA of higher organisms is that hybridization methods, which do not allow the easy collection of kinetic data, may conceal important information. This is the main reason forpreferring the solution methods, in which

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filters or hydroxylapatite colums (53)are used to separate dupIex from single-stranded molecules of isotopically labeled DNA. The main advantage is that reassociation of this DNA is of immobiliaing DNA on f3ters (3) prevented, although very highly repetitive sequences may reassociate as the DNA is concentrated onto the filter during charging. However, it is possible to arrange the concentrations in solution experiments so that DNA RNA interactions are favored at the expense of the self-renaturation of the DNA. The form ofthe association curves of an RNA fraction incubated with a well-defined fraction of, for example, intermediate DNA should show whether the RNA contains an equal number of copies of all the DNA cistrons present or whether some of the DNA has been transcribed in excess of others. The easy recovery of DNA hybridized with RNA by means of the hydroxylapatite method could be a very powerful method. In principle, the isolation of cistrons functioning in different tissues is possible, although attempts in this laboratory (54) along these lines have shown that only a small proportion of the more abundant families react even in the presence of a large excess of RNA. This is likely to be a serious limitation since its extension to rarer families may involve high and unacceptable RNA concentrations. Is there a way out? It can only lie, I think, in further purification of, primarily, the RNA. Since most of the RNA in these experiments may be ribosomal, which cannot react because the ribosomal cistrons are few and have a base composition that allows their removal by density-gradient centrifugation, its elimination would allow a higher concentration of the effective sequences. Nevertheless, it may be difficult to penetrate into incognita of “single-copy” DNA. the terra The conditions for finding a rare sequence in a complex genome are clear; the challenging RNA or DNA should have as short a sequence as possible; it shouId be single-stranded to avoid self-renaturation, and it should have a high specific activity, since this will determine the concentration of the complex genome needed in order to allow a detectable amount of labeled DNA or RNA to be bound. RNA transcribed invitro from a simple viral DNA can fulfill these conditions. So will single strands isolated from mouse satellite. Experiments have been published (32)in which the presence of sequences like mouse satellite DNA in the DNA of other rodents has been investigated. Here very sensitive t>ests,capable of detecting as little as 10-6 of the heterologous DNA, can be made, since a single strand of mouse DNA mg/ml will only renature in the presence of a t a concentration of 2 X its complement, which may be w few sequences in a large excess (x20,OOO50,000) of low-molecular weight heterologous DNA.

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In experiments of this kind, the simplicity of satellite DNA allows it to remsociate at concentrations sufficiently low that a large excess of DNA can be added without increasing the viscosity excessively. A more complex DNA or RNA would require a higher concentration to renature in the presence of its complement.

V. Conclusions Hybridization techniques appear to require a rather short complementary sequence (<20) for annealing to occur. This minimum stable length may have to be increased to some extent to compensate for mismatching of bases and for longer noncomplementary regions. An attempt is made to relate the minimum stable length to the degree of difference between two polynucleotide strands expressed as the proportion of base substitutions. Such calculations show that DNA svery different genetically should not be distinguishable by hybridization methods, but in fact they are. This paradox can be resolved by allowing a rate of evolution much faster for DNA than for proteins. It is possible that both the size of DNA families and the differences in the DNA of related organisms can be explained by the divergence in the third base of many codons, which can occur without alteration in the protein sequence due to the degeneracy of the genetic code. Finally, the limitations imposed by the presence of these DNA families on the DNA . DNA and DNA . RNA hybridization are discussed. It is difficult to see how any RNA transcribed by ‘Lsingle-copy”DNA can be easily detected by the method, and the importance of understanding the kinetics of the reactions has been stressed.

ACKNOWLEDGMENTS During the preparation of a somewhat speculative essay, I havehad many helpful discussions with Drs. W. G. Ramm, Anne McLaren, and E. Southern. I would also like to thank them and Dr.M. Birnstiel forcomments of the manuscript. This work has been generously supported by the Medical Research Council.

REFERENCE^ 1 . C. L. Schildkraut,

J. Marmur, aridP. Doty, J. MoZ. Biol. 3,595 (1961)

2. A. P. Nygaard arid B. D. Hall, J .Mol.Biol. 9,125 (1964).

3. D. Gillespie and S. Spiegelman, J. MoZ. Rid.12,829 (1965). 4 .P.M. B. Walker and A. McLaren, J .Mol.B i d .12,394 (1965). 5.R. J.Britten and D.E. Kohne, Currzegie Inst. Yearbook 66, 78 (1967). 6. C. A. Thomas, This series, 6,315 (1966).

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ORGANISMS

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7n. J. Marinmir and P. Doty, J .Mot.Biol. 6, 109 (19tU).

7b.J. Marmur, R,. Rownd, and C. L.Schildkrant, Thisseries I,231 (1963). 8. W. F. Dove and N. Davidson, J .Mol.Biol. 6,467 (1962). 9. R. Thomas, Biochim. Biophys. Acta14,231 (1954). 10. J.G. Wetmur and N. Davidson, J .Mol.Bid. 31, 349 (1967). 11.M. Chamberlin, R. L. Baldwin, and P. Berg, J .MoZ. Biol. 7, 334, (1963). 12.M. J. Chamberlin and D. L.Patterson, J .MoZ. Bid. 12, 410 (1965). 13.M. Riley, B. Maling, and M. J. Chamberlin, J .Mol.Biol. 20, 359, (19613). 24. M. Chamberlin, Federation Proc.24, No. 6,1446 (1965). 16.M. Chaniberlin and P. Berg, J .Mol.Bid.8, 297 (1964). 16.R. D. Wells, E. Ohtsuka, and H. G. Khoraiin, J .Mol.Bid.14, 221 (1965). 17.M. Lipsett, L. Heppel, and D. F. Bradley, 6.Biol. C hem. 236, 857 (1961). 18. A. M. Michelson and C. Monny, Biochim. Biophgs. Acta149, 107 (1967). 2.9.M. N. Lipsett, J .Biol. Ch,em. 239, 1256 (1964). 20. G.Cassani and F.J. Bollum, J .Am. Chem.SOC. 89, 4798 (1967). 21. J. C. Wang and N. Davidson, J. Mol.B i d .19, 469 (1966). 22.R. Wu and A. D. Kaiser, J .Mol.Biot. 36, 523 (1968). $3. N. R. Kallenbach and D. M. Crothers, Proc. Natl. h a d .Sci. U.S.66, 1018 (1966). 24.H. G. Khorana, Biochem. J .109, 709 (1968). 95. B. L. McConaughy and B. J. McCarthy, Biochim. Biophys. Acta149, 180 (1967). 96.D. Gillespie and S. Spiegelman, BacterioE. Proc. LosAngeles, 1966G-112. 27.S. K. Niyogi and C. A. Thomas, Biochem. Biophys. Res.Conimun. 26, 51 (1967). 88. W. Riiger and E. K. F. Bautz, J .Mol.Biol. 31, 83 (1968). 89. E. K. F. Bautz and F. A. Bautz, Proc.Natl. Acad.Sci. U .S.62, 1476 (1964). 30. N. Davidson, J. Widholm, U. S. Nandi, R.. Jensen, B. M. Olivera, and J. C. Wang, Proc.Natl. Acad.Sci.U .S.63, 111 (1965). 3 1 . M. N. Swartz, T. A. Trautner, and A. Iiornberg, J .B i d .Chem.237, 1961 (1962). 3% . W. G. Ffamm, P. M. €3. Walker, and M. McCallum, J. Moh. Biol. in press (1969).

33. S. B. Zimmerman, J. W. Little, C. I<. Oshinsky, and M. Gellert, Proc.Natl. Acad. Sci. U .S. 67, 1841 (1967). 34.D. M. Crothers, N. It. Kallenbach, and B. H. Zimm, J .Mol.Bid.11, 802 (1965). 36. J.Fresco, i n“Informational Macromolecules” (H. J. Vogel, V. Bryson, and J. 0. Lampen, eds.), p. 121. Academic Press, New York, 1963. 36. P. J.Gomatos and I. Tamm, Proc. Natl. Acad.Sci. U .S.49, 707 (1963). 37.E. T. Bolton and B. J. McCarthy, J .MoZ. Bid.8, 201 (1964). 38. B.J.McCarthy, General aspects of homology studies with nucleic acids. In “First International Symposium on Tumor Viruses: Subviral Carcinogenesis” (Y. Ito, ed.), pp. 43-60. Sponsored by Res. Inst., Aichi Cancer Center and the Japan. Cancer ASYOC.,1966. 39. H. Wallace and M. Birnstiel, Biochim. Biophys. Ada 114, 296 (1966). 40. J. Marmur and P. Doty, J .Mol.B i d . 3, 585 (1961). 4 1 .J.Marmur and D. Lane, Proc. Natl. Acad.Sci. U.S.46, 453 (1960). 42.J.A. Subirana and P. Doty, Biopolymers 4, 171 (1966). 4 3 .K. J. Thrower and A. R. Peacocke, Biochim. Biophys. Acta119, 652 (1966). 44.B. H. Hoyer, B. J. McCarthy, and E. T. Bolton, Science 144, 959 (1964). 46.P. M. B. Walker, Nature219, 228 (1968). 46.M. Kimura, Nature 217, 624 (1968). 47.F. H. C. Crick, J .Mol.Bid.19, 548 (1966). $8. R. E. Marshall, C. T. Caskey, and M. Nirenberg, Science 166, 820 (1967).

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49. F.W. llobertson and M. Chipchase, Bioclwnz. J .108,36P (1968). 50. F.H. C. Crick, Personal communication.

61. A. McLarenand P.M. B.Walker,Genet. Res.6,230(1965). 6.2. D. D.Brown and I.B.Dawid,Scieme160, 272 (1968). 63. M. McCallumand P.M. B.Walker,Biochem. J .106,163(1967).

54.D.Malcolm,in preparation.

Quantum-Mechanical Investigations of the Electronic Structure of Nucleic Acids and Their Constituents' BERNARDPULLMANAND ALBERTE PULLMAN Universit3 of Paris, Ilzstitut de Biologk

P~.~s~co-C~~rniq~~, Paris, France

. 328 I.Introduction . . . . . , . . . . . 329 11. Types of Calculation . . . . . . . . , 332 . 111. Schematic Description of the Met.liods of Calculativii . 333 A. The Self-consistent Field Method . . . . . . 336 B. The Huckel Approximation . . , . . . . 337 C. The Representation of the u-Bonds , . . . , 338 D. The Extended Huckel Theory . . , . . . , 339 E. The Iterative Extended Huckel Theory . . F. The SCF Procedures for AllValence Electrons 340 (the C N D 0 / 2 Method) . . . . . . . . 240 IV. Problems Investigated . . . . . . . . , 340 V. Electronic Properties of the Purine and Pyrimidine Bases . 340 A. Electron Distribution and Dipole Moments . . . B. Energies of Molecular Orbitals. Ionization Potentials a ~ i d Electron hffinities . . . , . . . . . . 347 . . . . . . . . . 349 C. Electronic Transitions VI. Interbase Interact,ions . . . . , . . . . . 354 A. The Forces Involved . . . . , . . . . . 354 B. In-Plane van derWaals-London Int,eractions and the . . . 357 Configuration of Hydrogen-Bonded Base Pairs . C. Vertical Interactions between Stacked Bases and the Interaction of the Base Pairs in the DNA Helix . . . . 369 D. Related Problems of Molecular Associations Involving Purines and Pyrimidines , , . . , . . . . . . 373 . . . . . . 375 VII. Problems in Radio- and Phutobiology . A. Spin Denait,ies in Free Radicals Derived from Nucleic Acid Bases 375 B. The Mechanism of Thymine Photodimerization . . . . 377 . . . . . . . 383 VIII. Electronic Factors in Mutagenesis IX. Conclusion , . . . . , . . . . . . 393 References . . . . . . . . . . . . . 395

.

.

The work was supported by Grant No. 67-00-532 of the Dblbgation Gknbrale A la Recherche Scientifique et Techniqlte (Cornit6 de Biologie Molbculaire), Grant No. of the Institut National des Qcierlceset de Recherche MBdicale (Intergroupe CR-66-236 Cancer Leucbmie), and Grant No. GM 12289Of the U.S. Public Health Service (National Institute of General Medical Sciences). 327

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1. Introdudion The establishment of the main features of the molecular structure of DNA by Watson and Crick in 1953,besides producing the tremendous development of modern molecular biology, had also the probably less predictable effect of stimulating the rise of another new branch of science, The designation refers to the quantitative namely quantumbiochemistry. application of the concepts and methods of wave mechanics, as developed previously, e.g., in the field of quantum chemistry, to the investigation of the electronic structure of biological molecules in relation to their behavior as substrates of life and tlo their involvement in the biochemical and biophysical processes characteristic of living matter. Although isolated attempts had been made occasionally earlier in this field, they had been very rare and generally of a qualitative and speculative nature. It was the sudden understanding by quantum chemists of the central role played in the phenomena of life by the purines and pyrimidines of the nucleic acids, i.e., by molecules very similar to those t,hat they had been familiar with in their studies for a long time, that more than anything else pointed to the possibility of applying the ideas and procedures of quantum chemistry to the problems of molecular biology. Together with this stimulus came immediately the understanding of the potentialities and enormous advantages of extending this type of study to other fields of biochemistry as well. A review of such a general application may be found in our book “Quantum Biochemistry” ( I ) In . the present paper, we restrict ourselves to the primary field of the contribution of quantum-mechanical calculations to studies on nucleic acids and their constituents. As a preliminary it must, of course, be realized that since the nucleic acids are extremely large, aperiodical polymers, there is no possibility, at present, of calculating the electronic structure of a real molecule of a nucleic acid. The calculations have therefore been restricted mostly to the evaluation of the electronic characteristics of the essential components of these macromolecules, namely, the purine and pyrimidine bases and the purine-pyrimidine complementary pairs. The first theoretical calculation on the fundamental purines and pyrimidines was carried out in 1956(2). The first theoretical calculations on the purine-pyrimidine complementary . papers and the refinements of base pairs were published in 1959( 3 )These these original calculations quickly demonstrated the possibility of approaching a number of the electronic features of the nucleic acids themselves by interactions between the introducing appropriately the intermolecular base-pairs. We shall consider at least some aspects of this problem in Section VI. The quantum-mechanical calculations so far performed on purines and

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pyrimidines or on the nucleic acids may be considered in two ways: first, they may be classified with respect to the method of calculation used; second, they may be classified in connection with the problems that they have dealt with. We shall first consider the methodology of the calculations.

II. Types of Calculation Allthe calculations so far published have been carried out within the general frame of the molecular orbital (MO) method. In the course of their development, they have, however, involved different technical procedures, corresponding to different degrees of refinement of the method. A very brief outline of these different techniques is given in the next section. In this section we shall simply name them and indicate their most general characteristics. From the technical point of view, five stages may be distinguished in the development of the calculations on the electronic structure of the bases: 1. The first period, initiated with the publications mentioned above and , to embracing a large number of subsequent papers ( C I S )corresponds calculations carried out within the classical Huckel approximation of the molecular orbital method (HMO) and limited to the ?r-electronicsystem of the compounds studied. In spite of the well-known shortcomings of the Huckel approximation, these calculations yielded a great mass of extremely useful information concerning the structure and the properties of purines and pyrimidines, which enabled a general interpretation to be made of a large body of experimental data concerning the chemical, physicochemical, and biochemical properties of these molecules and of the nucleic acids themselves and led to a number of predictions that have since been largely substantiated by more refined treatments and, what is more important, by subsequent experimentation. The success of the procedure was due mainly to the careful manipulation of the method, which was used essentiaIly for study of the electronic characteristics of the compounds the comparative scale of investigated, in other words for the classification on a relative compounds or molecular regions or their constituent atoms with respect to the electronic properties under investigation. The results of this work have already been summarized ( I ) It . may be added that in spite of the utilization in more recent years of more refined procedures (see below) Hiickeltype molecular orbital calculations are still being carried out in connection with different problems concerning the nucleic acids or their constituents. When properly handled, they are of very great significance. 2. The second stage corresponds to the advent of more refined calculations going beyond the HMO prorcrliirp. The first step in thcsr refinements

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consisted of adopt,ing the self-consistent jield molecular orbital procedure (SCF MO), generally in one of its semiempirical versions (in particular the so-called Pariser-Purr-Pople approximation, PPP-SCF-MO), some of the calculations including a further refinement in the form of allowing configurational interaction or mixing (SCF CI MO). The calculations were still limited to the ?r-electronic system of the compounds studied. The first calculation utilizing the PPP-SCF-MO method is due to Veillard and Pullman (14, 15). It was followed by a number of other similar publications (16-21). As is well known, the self-consistent field methods yield, inprinciple, more reliable absolute values for a number of indices of electronic structure and should therefore be suitable for a direct comparison of theoretical and experimental results. I npracfice, although the whole scheme of these procedures is more satisfactory than that of the HMO method, they nevertheless sufTer, in calculations concerning heteromolecules, from many of the same drawbacks. Generally, they do introduce in one way or another some semiempirical parameters into their scheme, in particular in connection with the different integrals related to heteroatoms. As a consequence of this situation, the absolute values of the quantities they evaluate may be appreciably in error. A very careful scrutiny of the choice of the integrals appearing in the method and their optimization on reference compounds are indispensable for a successful calculation. This, unfortunately, has not always been done, leading in some cases to serious mistakes. It has, however, been accomplished recently (22-25), yielding much more reliable results t)hat may be considered as very satisfactory for a number of problems. 3.The calculations referred to above were limited, as stated, to the *-electronic system of the molecules considered. This is, as is well known, a very classical and usual method of procedure in studying quantummechanically the electronic structure of conjugated molecules, especially in connection with their chemical properties. I t cannot, of course, be considered as completely satisfactory, especially for investigating certain physicochemical properties of molecules (e.g., their dipole moments) in which the distribution of the u electrons must also play an important role. For this reason, concomitantly with the late period of stage 1 and with stage 2,calculations have been performed, essentially by Berthod and Pullman (26, 27)[see also (28)] on the characteristics of the underlying u skeleton of the purine and pyrimidine bases. These calculations have been carried out by a refinement, due to these authors, ofthe method of DeI Re (29, SO) for studying u bonds, which may be considered as the counterpart for u electrons of t3heHiickel method for a electrons. The results of this type of calculation for the u electrons have then been combined with the results

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of the calculations carried out for the x electrons, whether by the Huckel approximation or the self-consistent field molecular orbital method, leading thus to a general picture of thc electronic distribution. 4. This way of calculating separately and then adding together the distribution of the u and R electrons suffers from the obvious drawback of neglecting the fine aspects of their mutual interactions. To remedy this electrom calculations, dealing simultaneously with the u defect, all-valemce and R systems, have been carried out recently for the purines and pyrimiby three procedures developed in recent years. These are the dines (31-33) extended Hiickel theory (EHT), the iteratiue extended Hiickel theory (IEHT), and the so-called CNDO method (the abbreviation standing for “complete neglect of differential overlap”), A general comparison of the results While the first two of obtained by the three procedures is given in (34). these procedures are extensions to all-valence electrons of the basic Hiickel procedure for x systems, the CNDO method is based on the self-consistent field molecular orbital scheme. Generally speaking, the utilization of these all-valence electrons methods has confirmed the validity, in many respects, of the previous additive procedures for the treatments of the CT x systems, when these treatments are appropriately parametrized, but has also brought into evidence some new aspects of results. Quantitatively, these methods are still far from being completely adequate and each of them seems to be, at present, particularly suited for the study of some aspects of the electronic structure. 5.Finally, the last of the five stages in the computational refinements of purines and pyrimidines is represented by calculations ab initio for all electrons (not only all-valence electrons). This type of computation is in progress presently and is not discussed here. This work utilizes the approach 36) and uses a developed primarily by Clementi and his associates (35, Gaussian atomic basis-set for the construction of the molecular orbitals. I n principle, it is the less empirical of the techniques used. In practice, the quantitative significance of its results is sometimes not completely satisfactory, because ofthe limitation in computational possibilities. It is thus obvious t,httt we have available at present for the basic constituents of the nucleic acids, the purine and pyrimidine bases, a large number of calculations carried out with a wide variety of computational techniques, starting from the simple procedures limited to x electrons and culminating in refined all-electrons calculations. It is essential to realize that all these techniques are approximation methods as there is no possibility at present of solving rigorously the Schrodinger equation for large molecular systems. All of them involve at one stage or another, although to a different extent, some technical approximations, the validity of which can

+

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be best ascertained only through the comparison of the theoretical results with experimental data. Moreover, because the different technical approximations mentioned above have sometimes more drastic influences in the refined techniques than in the simpler ones and have therefore to be treated carefully, the mere label of a “refined” procedure is not necessarily by itself a guarantee of a higher precision in the results. It is therefore essential to know both the advantages and the limitations of each procedure and to judge appropriately the significance that may be attached to the results obtained in each case. On the other hand, when properly used, all the methods are useful, and in many respects yield qualitatively and even quantitatively similar results. When these elementary data and precautions are kept in mind, the methods of quantum-mechanics appear as an exceedingly useful tool for the investigation of the structure and properties of the nucleic acids at a very fundamental level, whether for interpreting the observed phenomena in the appropriate terms, relating them to the appropriate electronic indices or predicting unknown properties and establishing new correlations. It is not possible, of course, to present in this paper the detailed results of all the aforementioned calculations or to carry out a detailed comparison between the results obtained by the different techniques of the molecular orbital method. We shall limit ourselves to the presentation of a selected series of results and will rather try to show their general significance for some problems of the molecular biology of the nucleic acids.

111. Schematic Description of the Methods of Calculation These methods, as already stated, all lie within the general framework orbital method. Only an extremely brief outline of the very of the molecular general principles of this method and of its different approximations can be given here. The details of the different procedures may be found in the technical literature : e.g., a detailed presentation of the Hiickel approximaa presentation of the self-consistent field method in tion in (1, 37-39), (40-45). The all-valence electrons and all-electrons procedures, because very recent, must be looked for in the original papers indicated below [for a brief review see (&)I. The basic idea that lies at the foundations of themethodof molecular orbitals is a very general one, first used in the quantum-mechanical descripthewavefunction of tion of polyelectronic atoms. It consists of constructing a polyeleetronic systemas a suitable combination of individual one-electron wavejunctions. The most “suitable” combination has been shown t o be of the general form

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a(1) a(2)

b(1) b(2)

c(1). . . 4 2 ). . .

a(n)

b(n)

~ ( n. ). .

a notation standing for the determinant built on the n individual wave part functions a,b, G, etc. Since each of these is a product of an orbital7 cp(lf,Y,Z)

and a spin function a orp, the total wave function for an even number of electrons is written as: cp1(1)41) cpl(lM(1) c p z ( 1 ) d ) cp1(2)42) cp1(2)P(2) cp2(2)42) P2(2)P(2) . . .

Such a “Slater determinant,” as it is often called, would in fact be the correct wave function for a system of noninteracting electrons. However, electrons do interact in real niolecular systems. Thus in order to obtain a satisfactory representation, theindividual orbituls cp aredetermined so as to takeintoaccount thepresence of theotherelectrons.

A. The Self-consistent Field Method The best procedure, which allows the determination of the individual molecular orbitals, is the “self-consistent field” method, whose main features are shs follows: a. One writes the exact total Hamiltonian for the system with explicit inclusion of electron interactions:

H = C H ( . ) + Z r, L y

Y

*
where H ( Y )is the Hamiltonian foroneelectron Y in the field of allthe b are nuclei. b. One expresses the total energy of the system by the standard quantum mechanical expression :

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in terms of the individual orbitals ‘p, by using the determinantal expression of 9. c. One satisfies the variation principle for the energy. This is a standard procedure that rests 011a fundamental theorem of quantum mechanics, namely, that the energy calculated by the above expression using an approximate wave function lies always higher than the exact energy; thus if one uses an approximate wave function expressed in terms of certain parameters, minimization of the energy with respect to these parameters will yield the best possible energy value attainable with this form of \k. Carrying out this program yields the general “Fock” equations, one for each individual orbital, cp :

Fv, = E N ; where F is an operator playing the role of an individual Hamiltonian and eiis the individual energy of one electron occupying the orbital ‘pz An essential characteristic of the Fock equations resides in the fact that each individual operator F depends on all the orbitals occupied in the system (on account of the explicit inclusion of the interaction terms) : thus each ‘p is given by an equation that depends on all the ‘p’s. The way out of this difficulty is to choose arbitrarily a starting set of ‘p’s, calculate the F(v) s, solve the series of equations for a new set of ‘p%, and go over the same series of operations again and again until the pth set of ‘p’s reproduces the ( p - 1)th set to a good accuracy. Hence the name “self-consistent” given to the procedure. The orbitals obtained in this fashion are in principle the best possible orbitals compatible with a determinantal 9. One restriction must be made, however, about this last statement: a choice must be made of a starting set of ‘p’s. Since it is impossible to guess ab initio the appropriate analytical form of a molecular orbital, one must rely on a “reasonable” possibility. Thus the final orbitals are the best possible orbitals oftheformchosen. The classical choice of the starting orbitals is based on the following idea: suppose that we deal with a chemical bond formed between two monovalent atoms A and B by the pairing of their valence electrons, one on A, the other on B. It is natural to assume that when one electron in the molecule is close to nucleus A, its molecular orbital will resemble the atomic orbital that it would occupy in A, and a similar situation would lar occur in the vicinity of B. This leads to the idea that them o l e ~ ~ orbital may be approximated by a Zinear co~b~nu~~on: ‘p

= ClXA

+

c2XB

where the x sare the atomic orbitals. The idea can of course be extended to a polyatomic molecule and

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generalized so that, eachmolecular orbital i na,molecule i sa linear combination of alltheatomicorbitals occupied by theelectrons intheconstituent atoms:

This is the classical and general LCAO approximation (linear cornbination of atomicorbitals) of the molecular orbital method. Given this convention, the Fock equations yield the unknown coefficients crby an iterative procedure provided that a starting set c,O is chosen. The practical calculat>ion,however, involves the tedious evaluation of a large number of integrals, a number that increases so rapidly with the number of electrons that for large molecules, complete self-consistent field calculations are presently out of question. molecules, which are usually defined as When dealing with conjugated molecules containing double bonds separated from each other by not more than one single bond, another classical and general simplification consisted until very recently, and still consists frequently, in treating their system of P electrons alone in the field created by the nuclei andthe so-called “u-core.” For this reason, one writes a Haniiltonian SOTtheP electrons:

defining a corethat includes everything but the ?r electrons. Provided H is so defined, self-consistent procedure can be applied to the ?r system alone thus he computed using the Fock equations apand the bestn orbitals propriate for the system. When the niolecular orbitals are taken as linear combination of the atomic p , orbitals only, the form taken by the equations amounts to solving a determinantal equation :

- CAY,,\

=

0

where S,,= ~ x p xdr, is the overlap integral and F,, = Jx$ x,dr the p q matrix element of the Fock operator. Solving the equation in e yields the possible values of the individual energies and for each ~i the corresponding coefficients G+ of the atomic orbitals xrin the molecular orbital pi are obtained by a system of linear equations 9

The resolution of the equation yielding t can be achieved by an iterative

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procedure of a “self-consistent” character, since all the quantities involved in-F,, can be, in principle, calculated when the u-core is assumed to have a definite configuration. In practice the u-core is admittedly taken a8 the sum of the atomic cores with the u electrons occupying atomic valence-state orbitals. This purely theoretical scheme yields, in principle, the best possible LCAO r-molecular orbitals for a given problem. However, the practice of the calculation forces in a number of errors, one of which is the use of a valence-state u core instead ofan exact u core, rarely available. Another is the choice and the limitation of the basis of atomic orbitals on which one builds the molecular orbitals. Last, but not least, there is the intrinsic error contained in theusual concept ofdoubly occupied orbitals, which permits two electrons of opposite spins to be a t the same time at the same place, thus ignoring at least in part what is called their “correlation.” In order to make up for these defects of the self-consistent field method, some empirical or semiempirical corrections must be introduced, the calibration of which are obtained by a close comparison of the theoretical results with experimental data for some fundamental compounds. The resulting procedure in its current form is the so-called Pariser-Purr-Pople approximationof theself-consistent jield method.One of its essential features is the hypothesis of zero differential overlap : XpXn

=0

which greatly simplifies the calculations. The results obtained can still be improved by conjiguration i,nteraction ormixing, correcting for the residual correlation error.

B. The HuckelApproximation There is another approach to the determination of individual molecular orbitals, which is different in its spirit from the preceding one in that it forgets about the apparent rigor of the self-consistent formalism. Thus, in this approach, instead of trying to determine the bestmolecular orbitals of an LCAO form that minimize the energy corresponding to a determinental wave function, one looks for approximate molecular orbitals (always of an LCAO form) in the following way: one considers that each r electron of the systems moves in an effective field resulting from the field of the u-core including the nuclei and the averaged repulsions of the other a electrons. Defining the corresponding individual effective Hamiltonian,Heff,the solving of an individual Schrodinger equation : Hew

=

Ep

yields the individual energy E and orbital Q.

ELECTRONIC

STRUCTURE

337

OF NUCLEIC ACIDS

A search fora molecular orbital of an LCAO form leads to the equations:

1c m , , - ES,,) 0 =

with the definitions:

H,, = Jx:Hx,d~ A%, = J x : x S d r where H is the individual effective Hamiltonian. The individual energies are solutions of the equation

IH,,- fiLYr81

=

0

Formally, these equations are similar to t,he SCF LCAO MO equations. Thus, a good definition of Heifmay yield satisfactory solutions, without the tedious iteration procedure of the SCF method. In practice, two calculation procedures have evoIved from these equations, the Hiickel and the Wheland-Mulliken approximation. Neither of them approximation specifies the analytical form of H , but instead treats some of the matrix parameters. Their main difference resides in the elements H,,as adjustable neglect (Huckel) or nonneglect (Wheland-Mulliken) of overlap. Their approximation, namely the neglect of common feature is the tight-binding all matrix elements involving nonbonded atoms.

C. The Representation of the u-Bonds Calculations carried out for T electrons may be supplemented, for the sake of a general picture of the electronic distribution, by calculations on the underlying o-system. A procedure used forthis purpose in purines and pyrimidines is a refinement of R method formerly proposed by Del Re (29, SO) for representing the u-bonds in saturated compounds. Its essential features are as follows: a. Each bond is treated as a two-electron problem. b. Each electron in the bond is described by a molecular orbital $0

= aXa

fbXb

linear combination of two atomic orbitals. Huckel-type Hamiltoninn yields c. The use of an effective one-electron the secular equation of second order:

Ha, - E H u b - ESOa with

HaL - ZSab Hbb

-E

338

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

d. By neglecting overlap and putting

all the parameters appear in terms of units a and ,f3. (These units are and I~cH,respect’ively). e. As an original feature of the procedure, one defines

QH

where 6, is a characteristic of atom a,c st,ands for all the atoms bound to atom a,and y o ( c ) is a proportionality factor that introduces what Del Re effect of atom c on atom a. calls the inductive f. The set of 6 O , y, and e values were chosen by Del Re by trial and error for reproducing as many properties as possible for a number of saturated molecules, imposing a few logical conditions on the parameters. In particular, the initial set of So values obeyed a linear law in the global atinnic electronegativities x:

The preceding t,echniquewas adapted by Berthod and Pullman (25,27) so as to make allowance for the possible differences in valence state or hybridization ratio of the same atom. Thus, orbital electronegrrtivities must be used as a guide for the choice of So values. This leads in particular to the use of different u-parameter values for saturated and conjugated moIecuIes, a logical step in a procedure which uses linear combinations of valence orbitals.

D. The Extended Huckel Theory The extended Huckel theory (EHT) method is due to Hoffmann and is a pioneering contribution in the field of computations including all valence electrons in conjugated molecules. In Hoffrnann’s method (45, 46)the molecular orbitals orbitals xIL:

pi are

built up as a linear combination of Slater atomic cpz

=

1

CiILXP

IL

the coefficients and the orbital energies being obtained by solving the set of secular equations:

c

Ci (HIL, - ES,,)=

0

Y

where H isa n effective Hamiltonian.

i

=

1, 2 , .

..,

ELECTRONIC STRUCTURE O F NUCLEIC ACIDS

339

The atomic basis set is made of one Is orbital for each hydrogen atom, one 2s and three d p orbitals for each carbon and each heteroatom (with Slater orbital exponents). The diagonal matrix elements H,, are the atomic valence state ionization potentials for each orbital and the nondiagonal matrix elements Hs9 are approximated by the Wolfsberg-Helmholtz formula:

+

Hpv= 0 . 5 f ( ( H , H,,)Sp,

with K = 1.75everywhere. This method is obviously an extension of the Huckel approximation for ?r electrons in the sense that the molecular orbitals are obtained as the eigenvalues of an effective hamiltonian H that is not explicit, the matrix elements of H being treated as empirical input characteristics of the atoms Hiiekel involved. For these reasons the procedure is called the Extended Theoryalthough it differs from the Huckel set of hypotheses in that it includes overlap as well as aZ2 nondiagonal elements in the secular det,erminant.

E. The Iterative Extended Huckel Theory Despite some remarkable achievements, in particular in conformational chemistry, the extended Huckel theory is obviously not entirely satisfactory from a theoretical point of view and it has been the object of a number of criticisms, some of them constructive enough to give rise to improved procedures. The essential modifications proposed, and which seem worth retaining, include : a. The use of the ‘(best single Slater” orbital exponents as determined by Clementi and Raimondi (&’), instead of the usual set of Slater exponents in the basis set. b. The replacement of the Wolfsbert-Helmholtz formula by Cusachs’ 49). expression (48,

c. The generalization of an old Wheland-Mann idea (50)[developed later under the general designation of a-technique (jl)]for making the diagonal matrix elements of the Hamiltonian depend on the net atomic 63). charges and solving the equations by an iterative process (52, These various modifications have recently culminated in a number of “Iterative Extended Huckel Theory” (IEHT) procedures advocated by 54)to yield results in more satisfactory agreement different authors (53, with experiment than the unmodified Hoffmann technique, particularly in the field of properties deriving from the coefficients of the atomic orbitals, like dipole moments, etc. The computations on nucleic acid bases described in this paper as having

340

BERNARD

PULLMAN

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PULLMAN

been carried out within the IEHT, were performed with the modifications a and b indicated above, introduced in the Hoffmann equations. Moreover, the diagonal matrix elements H have been made dependent on the orbital occupancy n and on the net atomic charge q through the relation (6.2)

+

H,, = A ( 4 & The numerical values used for A and B are the smoothed valence-state ionization potentials of reference (6% ).

F. The SCF Procedures for AllValence Electrons (the CND0/2 Method) The extended Huckel and the iterative extended Huckel methods are both based on an effective Hamiltonian, and are thus similar in nature to the Huckel-Wheland r-electron theory. Other methods have been proposed in recent years for treating all the valence electrons in the framework of a Hartree-Fock theory within the set of approximations known in the r-electron theory as the Parker-Parr-Pople method. The description of the technical details of this method is beyond the scope of this review. We may simply indicate that among the different versions of the SCF methods, the one utilized for the calculations described below on the purines and pyrimidines is the so-called CNDO/Z procedure (55-57).

IV. Problems Investigated The principal problems related to the structure and properties of the nucleic acids and their constituents that have been studied from the quantum-mechanical point of view are listed on p. 341,together with the essential references. As an illustration of the scope and the possibilities of the method, we shall discuss now some of these problems. They will be chosen among the representative fields : (a) some fundamental electronic properties of the purine and pyrimidine bases; (b) intermolecular interactions between the bases, defining some of the aspects of the structure of the nucleic acids; (c) some problems in radio- and photobiology; (d) possible electronic aspects of a biological phenomenon such as mutagenesis.

V. Electronic Properties of the Purine and Pyrimidine Bases A. Electron Distribution and Dipole Moments Because of the multiplicity and abundance of the available data, the examination of which would be lengthy, we are obliged to limit our dis-

ELECTRONIC

STRUCTURE

O F NUCLEIC

341

ACIDS

Problems Electronic distribution and chemical reactivily Dipole moments Basicity Ionization potentials and electron affinity Polarography Tautomerism Interatomic distances Electronic transitions Properties of the triplet states Hypochroniicity Optical rotatory dispersion Interbase interactions (van der Wattlb-Lundun forces in H-bonding and stacking, in solution and in crystals) Interactions with conjugated aromatics (aminoacridmes and carcinogenic conjugated hydrocarbons) Solvent effects Involvement in charge transfer complexes Proton tunneling and the structure of the hydrogen bonds Conformational problems Radiosensitivity Photobiology Diamagnetic anisotropy, ring curreiits, proton shifts Spin densities in free radicals Mechanisms ofmutagenesis Semiconductivity Coding illass spectroscopy Degradation by xanthine oxidase Structure of analogs

Principal references (2,58-62) (20,26, 27,3 4 fi.2) ( 2 , 6,7,2% 63) (2,17,2.2, 64,65) (62,66) (1,26, 67-69) (2,70) (22-25, 72-75) (20,22-25, 76-79) (16, 80-87) (88) (10,28, 88-103)

(204-209)

(93, 220-122) (2,113,114) (116-123) (32, 124) (126-229) (230-236) (137-242) (243-249) (67, 69,116-118, 230,160, 252) (13, 6.2, 162-265) (256, 167) (258) (269, 160) (2,12,262-165)

cussion here to the most outstanding features. We shalI first center our Figures 1-5 indicate their values (in attention on the l o t d net charges. electron units) for uracil, cytosine, guanine, and adenine, respectively, in the EHT method ( S I )the , IEHT method (SZ), the CNDO method (SS), and in coinputations resulting from the superposition of separated u-electron calculations (following the Berthod-Pullman refinement of the Del Re’s procedure) with either an SCF (22, 24) or R Huckel r-electron calculation

(fw.

Without entering into a detailed discussion of the results [to be found which contains also the division ofthe total charges into their u and in (34), B components and a comparison of the distribution of the r charges obtained in all-valence electrons calculations witfh the results of the usual previous

-1.408

-0.348

0

+o.oos

-1.395

-1 -418

4-0.916

+0.315

Cytosine

Uracil

- 1.409

+0.319

+0.321

\ /

0

N-0.848

+o.tmJ+o+o~ /

,081

+0.112

Guanine

N+O.Z89

1

-0.930

Adenine

FIQ.1. ExtendedHuckeltheory(EHT).Totalnetcharges.

+o.y

-0.488

+ys2

N-0,194

-0.038

-0.296N

+0.168

4-0.055

A

A

0

-0.519

-0.451

7-0.071 +0.094

+0.270

+0.260

Uracil

Cytosine +o.a28

-0.494

+o.m

\ /

0

N-0.125

-0.055

-0.266

4-0.115

+O.lSl

-!-0.242

Guanine

Adenine

FIG. 2.Iterative extended Hiickel theory (IEHT). Totalnetcharges.

342

+a.iso

-0.358

\

+0.117 / N-0.222 +0.020

-0.240

-0.344

NI

+ o ~ N ) y 9 ~ 0 3

0

0

-0.373

-0.419

I

-0.175

+o.iz5

+0.139

Uracil

Cytosine +0.107 4-0.115

-0.379

\

0

/ N-0.288

-0.172

I

N-0.103 +O. 139

-0.257 +0.120

+0.118

Guanine

Adenine

FIG.3.“Complete neglect of differential overlap(CNDO). Total net charges. f0.229 +0.229 -0.451

y 6 . 324

+0.326 -0.164

1

+0.195

+o.i9s

Cytosine

Uracil 4-0.230

-0.493

0

+0.230

Ax:k “5.361 +o.s5d

-*El7

- 1 J . 5 5 0+0.098 ~ -40.069

+0.369 4-0.070 +O.hSl

+o.igs

Guanine FIQ.4. Total net charges (u

-0.556

-0.066

1 +o.iss

Adenine

+ SCF 343

T)

(SCF

=

self consistent’ field).

+O.O60

344

BERNARD

PULLMAN

AND

ALBERTE

+0.229

-0.353

PULLMAN

+0.229

4-0.055

+0.481

1

1+0.192

*N/.aso

-0.350

I

-0.283

+o.ies

Cytosine

Uracil

+o.aso

+0.230

-0.371

0 -0.540

-0.510

I

1

+0.231

I

+0.196

+0.1m

Adenine

Guanine FIG.5. Total netcharges (U

+ Huckel

T)

*-electron computations], we may observe that although the general aspects of the global electron distribution are similar in all methods (the oxygens carry a net negative charge, the nitrogens, whether pyrrolic, amino, or pyridine-like are also negative, etc. . . .), differences appear naturally in the numerical values from one method to another, Their relative ordering is, however, generally parallel, as is illustrated in Figs. G and 7, which show graphically the evolution of the gross charges along the molecular framework in adenine and cytosine, respectively. Some essential features of the electronic structures of the bases are constant in all computations: the most negative oxygen is always that of cytosine, the carbon atoms of the C5-C6bond of the pyrimidines are of opposite polarity, with C-5negative, whereas all carbons are globally positive (with the exception of C-5of the purines in CNDO). The overall image of the NH2 group also shows a fair constancy. Generally speaking, the net charges obtained by EHT are much larger (whether negative or positive) than those obtnined by the other procedures, a known defect of the EHT. From a more general point of view it is particularly instructive to examine comparatively the values and the directions of the dipole moments predicted by the different procedures. The results are summarized in Table I and Fig. 8.

ELECTRONIC

STRUCTURE

O F NUCLEIC

ACIDS

345

I I

II

1I I

I

346

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

TABLE I DIPOLEMOMENTS (DEBYEUNITS)" ~

~~

U / d

A

U

<: C

3.2 3.6 6.8 7.2

77 39 -36 97

U / i d

2.0 73 35 3.3 7.2 -35 7.1 110

EHTd 6.1 12.2 17.2 17.3

65 41 -16.5 106

IEHTd 4.4 7.6 13.6 12.9

104 36 -33 125

CNDOd 2.9

4.6 7.5 7.6

64 36 31 102

Expl. valueo

3.0 39 -

FIG. 8. Relative magnitudes and directions of the dipole moments (from left t,o right: IEHT, EHT, CNDO, u SCF T).

+

347

ELECTRONIC STRUCTURE O F NUCLEIC ACIDS

distribution (26) with either a Huckel or the Pariser-Parr-Pople (PPP) a-distribution; the IEHT method yields intermediate values. What is very striking is that all methods predict greater dipole moments for guanine and cytosine than for uracil, which in turn should have a slightly greater moment than adenine. The comparison of the theoretical values with the experimental ones, available only for simple derivatives of adenine and uracil, indicates that the CNDO and the u a (Huckel or PPP) calculations are in very satisfactory agreement with experiment. It may therefore reasonably be expected that the values predicted for G and C in these last two procedures are the reliable ones. What is still more striking is the similarity in the directions of the moments predicted for the bases by all methods. Obviously, in spite of the quoted differences in charge distributions, the overall polarity of the molecules is well reproduced by all procedures although somewhat exaggerated by some of them.

+

B. Energies of Molecular Orbitals. Ionization Potentials and Electron AtTinities

Table 11,which gives the calculated energies of all the occupied molecular orbitals forthe five fundament,albases in the CNDO method, illustrates the type of results obtainable by the all-valence electrons procedures. Table I11 contains for the sake of comparison the energies of the a-electronic orbitals obtained in a a-electronic SCF computation. The most striking feature of the CNDO computations, which in fact is common to all the three all vaIence-electrons computations, is the indication of a large intermingling of the u- and a-levels with no appearance of a superficial “a-shell” as is implicitly assumed in a-electron calculations. Although the details may differ, the interspersing of the u-levels among the a-orbitals seems thus to be a constant feature in all-electron calculations. Such a constancy in the outcome of procedures that are quite different in their principles is probably a strong indication in favor of the picture obtained. It may be added that this mixing of u- and a-levels occurs much less in the virtual (empty) orbitals in which the a’-levels are often grouped below the u*-levels. orbitat (HOMO) is a B orbital in the The highestoccupiedmolecutar CNDO calculation. This is the case also in the EHT computation, but not in the IEHT one, in which the highest orbital is a CJ one. It seems probable (Sd) that this last result is an artifact of the IEHT parametrization. If we consider therefore, as seems highly plausible, that the highest occupied ?r molecular orbital is the one that gives, in fact, the measure of the first ionization potential of the molecule, the order of its increasing energy is,

348

PULLMAN

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AND ALBERTE

PULLMAN

TABLE I1

OCCUPIED ELECTRONIC ENEEGYLBVELS(eV) (SIGN REVERSED)IN

TEE

CNDO

METHOD^

~

U 52.87 45.09 43.64 40.22 36.88 32.25 27.41 R 26.38 26.36 25.71 21.39 R 20.07 19.83 19.12 T 18.65 17.72 16.42 13.83 R 13.18 12.90 R 11.88

0

T

C

G

A

53.03 45.87 43.96 41.59 39.61 32.88 31.98 27.38 u 26.90 26.75 24.74 R 22.21 21.35 20.00 R 19.66 19.22 18.58 T 17.56 17.42 15.88 13.74 a 13.10 12.85 a 11.37

52.36 44.01 41.69 38.85 36.61 31.66 27.75 25.74 B 25.53 25.45 20.96 20.01 R 19.91 18.40 17.91 R 17.20 15.56 13.68 R 13.21 11.81 R 10.78

55.49 51.27 48.23 42.73 39.50 38.98 32.08 30.57 28.50 27.82 A 27.60 26.48 u 22.82 22.46 21.61 20.60 ?r 19.66 19.55 17.38 16.75 15.65 T 15.34 15.08 R 14.21 u 13.11 12.77 11.55 R 9.06

55.33 50.39 44.13 38.86 37.34 33.94 30.41 29.94 27.73 R 27.07 25.64 22.62 a 21.15 20 51 19.82 18.07 7r 17.89 17.31 15.76 15.11 a 14.14 13.11 a 12.57 11.46 7r 10.08

The a orbitals are indicated

&s

such. All other orbitals are u.

TABLE I11 OCCUPIED

a

R-ELECTRON ORBITALS

IN THE PPP-SCF” METHOD (ev)(SIGNREVERSED)

U

T

C

G

A

15.18 13.12 12.34 9.90 9.15

15.12 13.07 12.39 10.71 9.84 8.82

14.23 12.23 11.25 9.33 8.16

14.84 13.68 11.96 10.66 10.09 9.46 7.59

14.58 12.78 11.32 9.93 9.43 7.92

Pariser-Pam-Pople Self-Consistent Field.

349

ELECTRONIC STRUCTURE O F NUCLEIC ACIDS

TABLE I V IONIZATION POTENTIALS(eV)

Expt. MasS ZEHTa

Compound

EHTa

Guanine Adenine Cytosine

11.84

10.31

11.95

10.43

12.50

Uracil

12.71

10.39 11.08

CNDOa 9.06

10.08 10.78 11.88

SCFu

~ C -

trometry

7.59 7.92 8.16

9.15

8.91 8.90 9.82

CTCO 7.8 8 8.1

Abbreviations defined in text and in legends to Figs. 14.

in absolute values (Table IV) G < A < C < U in the EHT and IEHT procedures and G < C < A < U in CNDO. Thus in all cases, guanine appears as the best T donor and uracil as the worst one among the four bases, in agreement with previous SCF ?r calculations (22). It is very rewarding that the recently measured values ofthe ionization potentials for the bases are in the order (65) A = C < T < U. The measurements for guanine are unfortunately missing. However, a strong electron-donor ability of this compound seems indicated by its easy charge-transfer complexation with chloranil (166). As to the absolute values of the ionization potentials, they are overestimated in the all-va~IenceCalcuIations and somewhat underestimated in the SCF calculation quoted. (Generally they are overestimated in the SCF calcdations too.) The theoretical data on the energies of the HOMO'S (and of the lowest emptymolecular orbitals, the LEMO's) of the bases were useful in interpreting and predicting polarographic ionization and oxidation 66, 167-170).2 potentials of the bases (62,

C. Electronic Transitions This is, generally speaking, an important chapter in the analysis of quantum-mechanical calculations. In this respect it must, however, be remarked immediately that none of the three all-valence electrons procedures outlined here has initially been devised or carefully adapted for the calculation of transition energies, i.e., that no numerical agreement with experimental results is expected from these methods in this respect in the present stage of their development. In fact their possibilities for the interpretation of the electronic spectra of conjugated heterocyclics such as the purines and pyrimidines is only in the course of investigation. It is also well known that the Huckel x-electron method is of onIy a limited validity 4

Seearticle by Paiebkinthis volumeiEds.1.

350

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

in the field of electronic transitions. On the other hand, the SCF r-electron procedure, in particular with configurational interaction included, has proved its value in this field. In the particular case of the purines and pyrimidines it has contributed significantly to the elucidation of the 73-75). We shall summarize the principal features of the spectra (22-25, contribution briefiy. Experimentally, the near ukraviolet absorption spectra of the purine and pyrimidine bases have been studied by different workers (171-174) down t o 180 mp in aqueous solutions, and more recently a systematic study of the various spectra has been performed with an attempt at classification (176). This has been completed recently by measurements on Mention must some of the bases in the vapor phase by Clark et al.(176). also be made of a recent detailed study of both the absorption and emission ofpurine and ofsome of its derivatives in different solvents (177-179). Clark and Tinoco (175) classify the observed transitions in the nucleic bases into three groups. As both the classification and the theoretical investigation described below have been extended to a number of related bases, such as purine (P), hypoxanthine (HX), xanthine (X), and uric acid (UA), we shall include them in the following discussion. The three groups of bands (Table V) are the following: TABLE V EXPERIMENTAL SPECTRA(mp

- eV)

ofabsorption Regions

Firstb

Compound"

U A P C G HX UA

x

mp

258 (244) 260 (252) 265 277 (290) 275 (293) 278 (281) 286 267

eV

mp

eV 5.4 -

237 256 247 -

mp

203 208 5.1.5 200 5.2 204 4.S 203 5.0 198 -

4 . 8( 5 . 1 ) 230

4 . 8( 4 . 9 ) 4 7 240 4 . 5(4.3) 4 . 5(4.2) 4 . 4 5(4.4) 4.36 4.6

Third

Second

eV

mp

6 . 1 181 5.96 185 6 . 1 188 6 . 1 185 6 . 1 190 6.26 185

eV

6.8 6.7 6.6 6.7 6.5 6.7

-

Abbreviations defined intext. Vaporphasedatainparentheses,

1. I n all these molecules, the first band corresponding to the longest wavelength of absorption is situated between 260and 278mp (4.764.4 eV) with HX, G, and C absorbing toward the longer wavelength limit and A

ELECTRONIC

STRUCTURE

O F NUCLEIC

ACIDS

351

and U (or T) toward the shorter. In the vapor phase, the distinction between HX, G, C, on the one hand and P, A, U, on the other is still more marked since the first band of hypoxanthine, guanine, and cytosine is eV) while the corshifted t o 290,293,and 281 mp, respectively (4.2-4.4 responding absorption in uracil and adenine occurs a t 244 and 252 mp, respectively (5.1 and 4.9eV). From the data available (180), uric acid seems t o belong to the group HX, G, and C. 2.Toward the short wavelength end of the spectrum (the third region), all these molecules exhibit two strong maxima, one around 200-208mp (6-6.2 eV) and another toward 180-188mp (6.8-6.6 eV). 3.Between these extremes is located the second transition, which, however, docs not appear in all molecules: it is present in HX a t about 247 mp (5eV), it is seen as a shoulder in the spectrum of C (237mfi or 5.2eV), of G (256mp or 4.8eV). and of 1 (240mp or 5.2eV), whereas it is not visible in the spectrum of U or A. However, in the case of U, the existence of a weak hidden band around 230mp (5.4 eV) has been suggested On the on the basis of experiments in optical rotatory dispersion (175). other hand, the first band in adenine has been suggested to be a composite one (172)formed by the superposition of two transitions, a suggestion 182)and studies of polarized supported by dichroism measurements (181, fluorescence (183). As Table VI clearly shows, the SCF results, even without conjigurational interaction (CI), correlate in a very satisfactory way with the previous classification. As concerns the transition of longest wavelength, the calculations distinguish obviously two groups among the bases : guanine, cytosine, hypoxanthine, and uric acid absorbing toward longer wavelengths and adenine, uracil, and purine absorbing toward shorter wavelengths. At the other end of the spectrum, all molecules exhibit a calculated band in the eTJ corresponding clearly to the first band of the t$hird region 5.9-6.1 region. Then comes a group of maxima that may reasonably be correlated with the second band in this region. As to the intermediate region, all compounds exhibit a calculated absorption in the range 5.3-5.5 eV except purine, in which the corresponding maximum is shifted t o 6 eV so as to be accidentally degenerated with t,he next transition. Interestingly enough, the calculated intensity of this theoretical band in uracil is exceptionally low, in accordance with the absence of an observable maximum in the intermediate region in this compound. As to the case of adenine, it seems tempting to consider the 5.5eV transition, which is very close to the first (at 5.3eV), as representing the second component of the composite band system. The calculated intensities are reasonable without being particularly good, n feature to be expected from surh x calculation.

TABLE VI C.4LCULATED

Compound"

u A P C G

m UA

x U A P

C G HY 1%

s a

First region

Second region

TRANSITIONS

(eV) Third region

5.1 (0.7) 5.2 (0.4)-5.5 (0.3) 5.5 (0.65) 4.2 (0.1) 4.4 (0.4) 4.4 (0.3) 4.6 (0.3) 5.0 (0.4)

SCF (intensities i nparentheses) 5.5 (0.02) 5.9 (0.2)-6.3 (0.5)-7.2 (0.01) 6.1 (0.5)-6.3 (0.01)-6.4 (0.2)-6.9 (0.2) 6.0 (0.4) 6.0 (0.7)-6.7 (0.03) 5.3 (0.4) 6.0 (0.5)-6.5 (0.05)-6.8 (0.3) 5.4 (0.6) 6.0 (0.1)-6.3 (0.1)-6.6 (0.2)-6.8 (0.1) 5.5 (0.7) 6.1 (O.lb6.3 (0.1)-6.7 (0.4)-6.7 (0.04) 5.5 (0.1) 6.0 (0.4)-6.1 (0.3)-6.2 (0.2)-6.5 (0.4)-7.0 (0.2) 5.9 (0.3) 6.1 (0.3)-6.2 (0.1)-6.4 (0.2)-7.2 (0.01)

4.8 (0.3) 4.8 (0.1)-5.0 (0.1) 4.9 (0.1) 4.1 (0.1) 4 . 3 (0 3) 4.2 (0.2) 4.3 (0.2) 4.7 (0.5)

SCF GI (intensities inparentheses) 5.4 (0.06) 5.8 (0.2)-6.2 (0.6)-7.2 (0.04) 5.8 (0.2F6.3 (0.1)-6.5 (0.5)-6.9 5.5 (0.2) 6.3 (0.3)-6.7 (0.3) 5 . 1 (0.1) 5.8 (0.4)-6.3 (0.3)-6.9 (0.2) 5 . 1 (0.3) 5 . 7 (0.1F5.9 (0.1)-6.2 (0.3)-6.6 5.1 (0.2) 5 . 8 (0.1k5.9 (0.2)-6.6 (0.2)-6.8 5.4 (0.3)-5.5 (0.4) 5.8 (0.1)-6.0 (0.2)-6.2 (0.4)-6.8 5 . 4 (0.1b5.6 (0.3) 6.0 (0.2b6.3 (O.lb6.8 (0.01)

Abbreviations defined in text.

(0.2) (0.03) (0.5) (0.2)

td

M 0

2

P

m U

ELECTRONIC STRUCTURE O F NUCLETC ACIDS

353

The results after configuration mixing (Table VI) confirm entirely the previous assignments with the following characteristics : 1.As concerns the absorption band of longest wavelength, the calculations continue t.0distinguish two groups among the compounds-one group, absorbing farther toward the red, comprising G, C, HX, and UA; and another group, absorbing toward shorter wavelengths, composed of P, A, and U. This distinction is slightly emphasized after configuration mixing where the respective ranges are 4.14.3eV for the first group, and 4.84.9 eV for the second, in very satisfactory agreement with the observed values. The location of this band was, as we have seen, already fairly satisfactory without configuration mixing, with the notable exception, however, of purine, which shows now a very strong lowering of the energy ofthis transition (and a parallel improvement of the results) through the mixing of essentially the first threeexcited configurations. As a rule, the calculated intensities of the first band are closer to the experimental values after configuration mixing in these compounds. In particular, the anomalously high value obtained for uracil before mixing is appreciably lowered and brought much closer to the observed one. 2.Leaving aside for the moment the second calculated transition, we observe in all the compounds a series of maxima ranging from 5.7to 7.2eV (6.0 to 7.2eV before mixing) which clearly correspond to the third region. 3. In the intermediate region there is a calculated transition between 5.1and 5.5eV in all compounds after configuration mixing. Obviously, this transition can be correlated with the second band. Its position is improved with respect to the SCF results and compares very favorably with the experimental wavelength in the compounds in which it can be seen. I n uracil the low value of its calculated intensity is in agreement with the fact that it seems to exist as a hidden transition. On the other hand, in adenine, it appears very close to the first transition (much closer than in any of the other compounds), thus confirming the interpretation of the observed farthest absorption of this molecule as being composite. This phenomenon, already visible in the SCF results, becomes much clearer after configuration mixing. I n contrast, no such phenomenon appears in purine, where the second band is appreciably separated from the first one. Thus, on the whole, the classification and the interrelations of the observed spectra are reproduced in quite a satisfactory way by the calculations. The main features of this agreement are already present in the SCF results. Configurationinteraction does not bring about any drastic modification although it yields a general improvement of the results, particularly as concerns the position of the second transition and the intensities of the bands. Purine is the only compound for which the SCF results are relatively poor and for which configuration mixing has a rather strong improving

354

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PULLMAN

AND

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PULLMAN

effect. This is obviously related to the accidental degeneracy of the 5 + 7 and 4 - 6 configurations and to their strong mixing with the 5 + 6 configuration, yielding a first excited state in which the three are equally weighted. In general the discrepancy between the theoretical and the experimental energies of the electronic transitions, in particular as concerns the first and the second transitions, does not exceed 0.2eV in the calculations described here, in any of the compounds studied. In no way is such a satisfactory agreement general in all SCF calculations of the bases. Unsatisfactory parametrization and other GomputationaI deficiences are responsible for a number of defects in some of the published calculations. A number of other purines as well as other aspects of electronic transitions, such as the directions of the polarizations of the transitions, the energies of the triplet states, the structure of the excited states, in particular the spin densities in the first triplet and the zero-field splitting parameters have also been the subject of extensive research (22-25, 73-79).

VI. Interbase Interactions A. The Forcesinvolved Having illustrated in the preceding section the nature of the results susceptible to be obtained in the study of the electronic structure of the purine and pyrimidine bases, we would like presently to evoke briefly the problem of the interactions between such bases, in particular of those leading to the type of associations that occur in the nucleic acids. These interactions related to interactions are obviously of two types: in-plane hy drogen-bonding, and vertical interactions between stacked bases. This is of course a very fundamental problem directly related to the essential question of the nature of the forces responsible for the stability and ordered structure of the nucleic acids. For a long period and in a number of publications there prevailed a definite duality of conception as to the roles in this respect of hydrogen bonding and stacking. Thus, when the doubly-helical structure of DNA was established, the majority ofmolecular biologists seemed to believe that the stability of this edifice arose essentially from hydrogen bonding. When, lder, different arguments (184), among which a prominent one was the discovery of ordered (“stacked”) single-stranded homopolynucleotides, indicated that such a conception was deficient or at Ieast insufficient, a number of authors proposed that the stability of the nucleic acids must be due mostly to van der Waals-London interactions between vertically superposed, stacked bases or base pairs. Even today the question is still

ELECTRONIC STRUCTURE O F NUCLEIC ACIDS

355

frequently raised whether the stability of the nucleic acids and polynucleotides is due essentially to hydrogen bonding between horizontal pairs of bases or to van der Waals-London forces between the stacked ones. Stated in this way, the question is essentially incorrect because it implicitly assumes that there is a fundamental difference in the nature of the forces operating between the horizontally linked or vertically stacked bases. It implies in particular that the van der Waals-London forces do not play any role in the interaction between the horizontally associated bases. Such a conception is totally erroneous as was shown in 1962 by De Voe and Tinoco (10) on the very example of the interactions between the bases of DNA and as can be demonstrated st,iII more significantly today. The situation is that it is the same type of force, namely the general intermolecular forces, which are in fact responsible for the interactions both between the horizontally and the vertically associated bases. Before demonstrating this situation in more detail, a few words need to be said about the modes of compilation of these forces. The intermolecular forces are usually evaluated and have been so initially for the interactions between the nucleic acid bases (10) in the I( ) the sum dipole” approximation which considers these interactions ( E D as of three principal contributions :

ED = IC,,

+ E,,+ E L

where E,,, are the dipole-dipole, E,,the dipole-induced dipole, and EL the London or dispersion interaction energies. These are defined, respectively, as follows:

where and p~ are the respective dipole moments of molecules A and 13 (with RABthe distance between the points of location of these dipoles), (YA and CYB their polarizabilities, and IAand iB their ionization potentials. It may, however, be observed (28,185) that because of the shortage of the intermolecular distances with respect to the moIecuIar dimensions, this “dipole” approximation may be rather inaccurate in the particular case of the interactions between the purine and pyrimidine bases and that it may be preferable to treat the problem in t2he“monopole” approximation, that is, by considering all the negative and positive charges in the system as

356

BERNARD PULLMAN AND ALBERTE PULLMAN

interacting in a simple coulombic fashion. In this “monopole” approximamay then be considered rn the sum of three main tion the total energy (EM) contributions : =

+

+

EPP Epa

E L

where E,,are the monopole-monopole, E,,the monopole-induced dipole, and EL the dispersion interaction energies. The first two are defined by

since with

where index i designates the atoms of molecule A and j those of molecule B, the p s are the net charges of the corresponding atoms, UA-B represents the dipole moment induced in B by the distribution of charges in A and E A + B the field induced in B by the charges in A at the point of location of the induced dipole ( R i B designates the vector from atom i in A to this point), etc. This transformation from the dipole to the monopole approximation in the calculations of the intermolecular forces has very significant consequences, as can be illustrated in the following example (89). Let us consider a system formed of the molecule of adenine, fixed in space, and the molecule of uracil rotating around it in the same plane (and rotating moreover around an axis perpendicular to its plane and passing through its center) and let us lookfor the minima of the potential energy that may appear as a function of the mutual arrangement of the two bases. When the calculations are carried out in the monopole approximation, well-defined minima appear that correspond always to arrangements equivalent to the different possible “hydrogen bondings” between the bases. This result is due to the predominant contributions to the interaction of the electrostatic energy between the closely located atoms involved in the “hydrogen bonds” and may be considered as a confirmation of the essentially electrostatic nature of such bonds. No such minima are visible when the calculations are carried out in the dipole approximation. Because most molecular

ELECTRONIC

STRUCTURE

OF NUCLEIC

357

ACIDS

associations in biology occur i)etwceii relatively large nioIecules at small distnnccs from each ot,licr, tlic iinportnnce of thc motiopole refinement hardly needs stressing. The technical designatiou of the niouopole approximation as defined above is in fact “the monopole induced-dipole approximation.” Still more recently, further technical refinements have been introduced in the evaluation of the van der Waals-London forces between purines and pyrimidines They involve in or, more generally, between conjugated molecules (203). particular the replacement of the molecular total polarizability by bond polarizabilities and the introduction of a repulsion component at short distances, this last refinement having the important consequence of enabling the theoretical determination of the equilibrium distance. The roles of these improvements are presently being investigated.

B. In-Plane van der Waals-London Interactions and the Configuration of Hydrogen-Bonded Base Pairs In recent years, a large amount of data has been gathered experimentally on hydrogen bonding between purine and pyrimidine bases in crystals and in solution. The results point to an extremely large variety of possibilities and offer therefore an interesting challenge for theoretical interpretation. Among the most outstanding facts are the following: 1. The discovery by Hoogsteen (186, 187) that methylated derivatives of adenine and thymine (both carrying a substituent at their glycosyl nitrogen) cocrystallize as a hydrogen-bonded complex whose configuration is, however, different from the Watson-Crick one observed in the nucleic acids : thymine is linked t o N-7of adenine instead of N-1 (see Table VII the “Hoogsteen configuration”). This type of mixed crystals is quite general for associations between derivatives of adenine and uraci1; in some cases (when derivatives of bromouracil are present) they may beof the “reversed Hoogsteen” type (Table VII) in which the pyrimidine, while still linked to N-7of adenine utilises 0-2 instead of 0-4 for the hydrogen bonding (188, 189-192). On the other hand, derivatives of guanine and cytosine COcrystallize only following theWntson-Crick pairing scheme (188, 193). 2. Cocrystallization of the nucleic acid bases or interaction in nonaqueous solution can be obtained only with base pairs showing complementarity in the Watson-Crick sense (A T, A U, G C), no interaction occurring, e.g., betlween guanine and thymine, adenine and cytosine, cytosine and thymine or guanine and adenine (188, 194, 195). This is a phenomenon that, at first sight, has some magic flavor, because chemically there are no immediately obvious reasons why such associations should not be formed. As illustrated in Fig. 9, the corresponding chemical formulas can

-

-

-

358

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

TABLE V11 INTERACTION ENERGIES IN DIFFERENT POSSIBLE ADENINE URACILPAIRS(KCAL/MOLE)

-

Configuration

*L

’M

-0.22

-0.88

-6.96

-0.17

-0.94

-6.74

Epp

Epcu

-5.86

-5.63

Watson-Crick

ReversedWatson-Crick

Hoogsteen

H

ReversedHoogsteen

H

ELECTRONIC

STRUCTURE

O F NUCLEIC

Guaninethymine

359

ACIDS

Adenine. guanine

14 1~. 9. Non-Wat,son-Crick base pairings.

be drawn easily. It niust, however, be remarked that the exclusive nature of this association in cocrystallisatiori or in solution does not eliminate, of course, the possibility of the estzLblishnient of such noncoinplementary pairings in other more particular circumstances in which they could be imposed by some external factors. In fact, such a possibility has been considered, e.g., by Crick in his “wobble” hypothesis of codon-anticodon interactions (196). 3. When instead of utilizing bases substituted at their glycosyl nitrogens, free bases are employed, no cocrystallization whatsoever is observed (197). 4. Specific types of hydrogen bonding are observed in the crystal structure of the different bases. We shall not discuss this problem here, but shall only mention that it has been dealt with satisfactorily in detail elsewhere (92). A very curious and more recent observation deserves, however, to be considered. It concerns the crystal structure of purine itself. This structure (198)involves long chains of molecules, linked together by single hydrogen bonds, as illustrated in structure I for a dimer. The toN-7 surprising feature of this configuration is that a protoni sattached rather thanto N-9 of thepurine skeleton, whereas in most biological derivatives of purines, including the nucleosides and nucleotides, the substituent is fixed at N-9. 5.Interesting and diversified behavior is observed also in connection with triplets of hydrogen-bonded coplanar bases, which appear sometimes in cocrystalline complexes or in associations of polynucleotides. Thus, for example, a cyclic homo-triplet of hypoxanthines of the form of structure I1 most probably exists in the three-stranded polyinosinic acid (199),and it is of an obvious theoretical interest to know its stability relative to a possible hydrogen-bonded dimer of hypoxnnthines, as in structure 111. But still more striking is certainly the comparative behavior of the two

360

BERNAWD

PULLMAN

AND

ALBERTE

PULLMAN

Purinedimer (1)

Hypoxanthine "trimer"

Hypoxanthine "dimer"

(n)

A.2U;A trimerofadenine and twouracils (Watson-Crickreversed Hoogsteen type)

DAP. 2 U, A trimer of2,B-diaminopurineand two uracils (reversed Watson-CrickHoogsteen type)

(IV)

(V)

+

open hetero-triplets, adenine 2 uracils (A . 2U), postulated to exist in the three-stranded poly A a2 poly U and 2,6-diaminopurine 2 uracils (DAP 2U) observed in the 2:1crystalline complex of 1-methyl-5-iodourac~ with 9-ethyl-2,6-diaminopurine. Both triplets are of the general type

+

u 1

uz

P'

in which the two urmils are "on both sides of the purine," i.e., hydrogen-

ELECTRONIC

STRUCTURE

OF NUCLEIC

361

ACIDS

bonded to the &amino group of the purine and to its N-1 or N-7. However, the detailed configurations ofthe two associations seem to be different in the two cases. I n principle, there are four possible arrangements for each group A 2U or DAP 2U,corresponding to the four possible simultaneous arrangements of the two uracils, which can be denoted as

-

-

1. W atson-Crick-Hoogs teen 2.Watson-Crick-reversed Hoogsteen 3.Reversed Watson-Crick-Hoogsteen 4. Reversed Watson-Crick-reversed Hoogsteen In practice, while the trimer observed in poly A * 2 poly U is of the Watson-Crick-reversed Hoogsteen type IV (200), the trimer observed for DAP 2U is of the reversed Watson-Crick-Hoogsteen type V (201). We shall stop here the listing of the different cases. The question now rises whether this considerable number of observations may be accounted for by a homogeneous interpretation. The answer to this question is affirmative and the homogeneous interpretation is based on the interplay of the van der Waals-London intermolecular forces between the bases, as follows. 1.Table VII represents the interaction energies evaluated for the different possible pairing schemes of the A U dimer (96, 202). (As most of the calculations have been carried out in the monopole-induced dipole approximation, the calculations listed here are generally given, udess otherwise stated, in this approximation.) It is observed that the calculations predict the Hoogsteen arrangement t o be the most stable one, followed rather closely by the reversed Hoogsteen arrangement. Table VIII represents the same results for the different pairing schemes of the G C dimers. In this case it is the Watson-Crick configuration that is the most stable one. (The Hoogsteen type configurations would imply in this case the rare imine form of cytosine.) These results are therefore in agreement with the previously quoted observations in cocrystallization phenomena. It may also be remarked how much stronger the Watson-Crick G C pairing is predicted to be with respect to the Watson-Crick or the Hoogsteen A . T pairing. This strong distinction is conserved in all more sophisticated calculations on these fundamental pairs (203). 2. The exclusivity of cocrystallization or of interaction in solution (in nonaqueous solvents, see below) among the bases complementary in the Watson-Crick sense is accounted for in Table IX (94). Table TX indicates, in the first place, the maximum interaction energies calculated for hydrogen-bonded self-a.ssociations of the different bases (which, as already mentioned, correspond regularly to the associations observed in the cryst,als of these substances) and, in the second place, the

-

-

362

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

TABLE VIII

INTERACTION ENERGIESIN DIFFERENT POSSIBLE

.

GUANINE CYTOSINEPAIRS(KCAL/MOLE) Configuration

EPP

E~(Y

EL

EM

Watson-Crick

Watson- Crick Reversed

Hoogsteen

-3.98

-1.33

-0.44

-5.15

-2.42

-0.91

-0.34

-3.61

H Reversed Hoogsteen

I

H

H

ELECTRONIC

STRUCTURE

363

OF N P C L E I C ACIDS

T-LE IX INTERACTIOX ENERGIES I N HYDROGEN-BONDED BASE PAIRS(KCAL/MOLE) A .A

T.T G*G C.C A .A

-5.8 -5.2

A .T

-14.5 - 13

G*C

-5.8

A .C

-7.8

-7.0

-19.2

C.C

- 13

G.G

-14.5 -5.2

G-T

-7.4

T.T C.c T-T

- 13 -5.2

C*T

-6.5

A*A

-5.8 -14.5

A .G

-7.5

G.G

A *T

>

G-C A. c G.T C*T A .G

> < < < <

A -A T.T G.G

c-c c.c G*G C. C G.G

energies of the corresponding cross-associations. The calculations cover, of course, both the observed and the nonexisting associations. (It is one of the advantages of the theoretician over the experimentalist to be able to study things that do not exist.) It is then observed, in the first place, that the self-associations may obviously be divided into two groups, the relatively strong G . G and C . C self-associations and the relatively weaker A A and T . T self-associations. The cross-associations may also be divided into two groups: on the one hand, we have the two pairs A T and G . C whose interaction energies are greater than the energies of the self-associations of their two constituents (or the mean of these energies, Table X) ; and on the other hand, the four remaining non-Watson-Crick pairs for which the interaction energies are always smaller than the energies of self-association of one of their constituents (or smaller than the mean of the energies of the two constituents, Table X). This situation which is represented symbolically in the lower part of Table IX, suggests by itself an explanation for the exclusivity of the A T and G C pairings, the only ones strong enough to be able to be formed a t the expense of the self-associations of their constituents. It may be interesting to remark that recent investigations on the

-

-

-

-

364

BERNARD

PULLMAN

AND

PULLMAN

ALBERTE

TABLE X INTERACTION ENERGIES IN HYDROQEN-BONDED PAIRS(KCAL/MOLE) Mean value

T .T

-5.5

<

A.T

-13.8

<

G-C

-9.4

>

A .C

-7.8

-9.9

>

G *T

-7.4

-9.1

>

C-T

-6.5

-10.2

>

A .G

-7.5

c.c T *T T *T G-G

-7.0

-19.2

strength of the different types of observed associations in solution have confirmed, as indicated in Table XI, the different detailed predictions of the theory. 3.As concerns the appaxent absence of any interaction between bases unsubstituted at their glycosyl nitrogen, this is accounted for, at least in part, in Table XII. In this case, it is obvious that one has to consider a complementary mode of self- or cross-association of the bases, which would involve the glycosyl nitrogens themselves, as exemplified in structures VI and VII forthe self-association of adenine and the cross-association of adenine with thymine. The modification which this new situation introduces with respect to the previom results, as summed up in Table X, is indicated in Table XII. The essential modification concerns the maximum value of EXPERIMENTAL RESULTSON Result

A .U

-

>A.A

or IT.U

-

G C > G G or C . C G - C > A Tor A . U

THE

TABLE XI RELATIVE STRENGTHOF HYDROGENBONDTNO Solvent

CDClS CDCla CDC1, CDC13 Me2S0

CHC18 G*G>C.C U-U>A*A

+ CHCls

Method

IR IR IR IR NMR

IR

MezSO

NMR

CHCb CHCla

IR IR

Refereu ces

ELECTRONIC

STRUCTURE

O F NUCLEIC

365

ACIDS

TABLE XI1 INTERACTION ENERGIES BETWEEN UNSUBSTITUTED BASES (KCAL/MOLE) A-T

-8.13

-- 7 . 0

-5.2

G.G

c.c

-14.5 - 13

. .4

-8 . 1 3

A

c .c G.G T-T

-19.2 7.8

- 13 -14.5

G.T

-7 . 4

C.T

-- 6 . 5

A.G

-7 . 5

-5.2

T.T

- 13 -5.2

A .A G.G

-14.5

C.C

c.c

-8.13

A-C G-T C.T A.G

< < < <

c.c G-G c .c G .G

the energy of self-association of adenine, which is now greater than the energy of the A . T association. Consequently, in agreement with our preceding rule, unsubstituted A and T should not interact. On the other hand, the new mode of association does not change the previous results relating to the interactions between G and C, and one could therefore expect these free bases to associate. As stated before, they do not appear to do so. The explanation of this disagreement may probably be found in the fact that experimentation with unsubstitued guanine and cytosine cannot be considered as decisive because of the extremely limited sohbility of guanine in the solvents used. 4. The interesting case of the crystal structure of purine, involving the N-7(H) tautomers, may be accounted for along the same lines (98). Thus, quantum-mechanical (EHT) calculations on the relative stabilities of the two tautomeric purines, one with a proton attached to N-7 and the other with a proton attached to N-9, indicate comparable stabilities. On the other hand, these different calculations indicate a difference in one of the electronic characteristics of the tautomers that may possibly account for the presence of the N-7(H)tautomer rather than the N-9(H) one in the crystal. This characteristic is the electronic distribution in the two forms.

366

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PULLMAN

Selfassociation ofadenine (A.A)

(VI)

The two distributions are sufficiently different to lead to a prediction that the dipole moment of the two forms should also be appreciably different. In a refined Huckel approximation, the predicted dipole moment is 4.1D for the N-9(H) form and 6.3 D for the N-7(H)form. I n the SCF approximation, the two dipole moments are respectively 3.6 D and 5.5 D. The moment predicted for the N-9(H) form is in satisfactory agreement with the moments known for related compounds: 4.3 D in 9-methylpurine ( l o ) , 3.8 D in 6,9-dimethylpurine (209). In these circumstances, it appears possible that the van der WaalsLondon forces (their electrostatic component in particular) may also be greater in the crystals involving the N-7(H)tautomer, and possible that this conclusion may hold when the calculations of the interaction energies are carried out in the “monopole” rather than in the “dipole” approximation. In order to check this hypothesis, calculations have been performed on the van der Waals-London interaction energies for dimer I and for the hypothetical dimer VIII, which differs from dimer I only in the shift of the proton from N-7 to N-9 of the bases involved. The results of the calculations, carried out in this case in different approximations, are indicated in Table XIII. It is believed that they are listed in the order of increasing refinement. The results of Table XI11 indicate, as expected, that theintermolecular interaction energies aresignificantly greater indimerI thanindimerV I I IIt; may certainly be extrapolated that the Same situation would prevail in higher polymers of the two kinds.

367

ELECTRONIC STRUCTURE O F NUCLEIC ACIDS

H

Hypothetical purine "dimer"

(VIE)

5 .Finally, we may consider the problem of the hydrogen-bonded coplanar triplets of bases (96). I n the case of the triplet of hypoxanthines, the calculations indicate that the total energy of interaction in the triplet is - 18.69 kcal/mole, evenly distributed (- 6.23 kcal/mole) per hydrogen bond or per hypoxanthine present. As remarked before, it may be particularly significant to compare these values with those predictible for a hypothetical hypoxanthine-hypoxanthine base pair such as I11 linked together by two hydrogen bonds, which represents probably the most stable such pair. This energy would be equal to - 10.83 kcal/mole, which therefore that represents - 5.42 kcal/mole per hypoxanthine ring. Itappears TABLE XI11 INTERACTION ENERGIES IN PURINEDIMERS(KCAL/MOLE)

Cl imponents Compc)t i n (1s

Approximation

Dinier I (observed)

Dipole-induced dipole Monopole-induced dipole Monopole-bond polarieabilities Monopole-bond polarieabilities repulsion Dinier VIII Dipole-induced dipole (hypothet,ical) Monopole-induced dipole Monopole-bond polarieabilities Monopole-bond polarieabilities repulsion

+

+

Electro- Indric- Disper- Repulstatic tive sion sion

Total

- 1.12 - 0.30 -6.84 -0.73

-0.39 -0.39

-

-1.81 -7.96

-6.84

-1.36

-1.32

-

-9.52

-6.84

-1.36

-1.32

+2.42

-7.10

-0.21 -5.68

-0.12 -0.43

-0.37 -0.37

-

-0.69 -6.48

-5.68

-1.03

-1.34

-

-8.05

-5.68

-1.03

-1.34

$4.37

-3.68

368

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ENERGIESOF

PULLMAN

AND

ALBERTE

PULLMAN

TABLE XIV WAALS-LONDONINTERACTIONS IN COMPLEXES ADENINE WITH URACIL (KCAL/MOLE)

VAN DER OF

Configuration

Interaction E,,

E,,

EL

Em

Watson-Crick-Hoogsteen

A*UI -4.64 -0.25 -0.69 -5.58 A.Uz -5.86 -0.22 -0.88 -6.96 Ui.Uz +0.96 -0.02 -0.03 fO.91

Watson-Crick-reversed Hoogsteen

A.Ui -4.64 -0.25 -0.69 -5.58 A * U z -5.63 -0.17 -0.94 -6.74 u1.U~ +0.57 -0.01 -0.03 +0.53

A.2U -9.54

-0.49 -1.60 -11.63

A.2U -9.70 -0.43 Reversed Watson-Crick-Hoogsteen A*Ui -3.98 -0.25 A * U z -5.86 -0.22 Ui*Up +0.60 -0.02 A.2U -9.24 -0.49 Reversed Watson-Crick-reversed A - U i -3.98 -0.25 Hoogsteen A.Uz -5.63 -0.17 u1.U~ 4-0.42 -0.01 A.2U -9.19 -0.43

-1.66 -11.79 -0.71 -4.94 -0.88 -6.96 -0.03 +0.55 -1.62 -11.35 -0.71 -4.94 -0.94 -6.74 -0.03 +0.38 -1.68 -11.30

TABLE XV ENEROIEBOF OF

WAALS-LONDONINTERACTIONS IN COMPLEXES 2,6-DIAMINOPL?INE (DAP) WIT11 URACIL (KCAL/MOW) VAN

Watson-Crick-Hoogs teen

DER

DAPmU1 -6.49 -0.22 -1.16 -7.87 DAP-Uz -5.80 -0.24 -0.48 -6.52 Ui*Ue +0.96 -0.02 -0.03 +0.91

DAP.2U -11.33 -0.48 Watson-Crick-reversed Hoogsteen DAP U1 -6.49 -0.22 DAPJJz -5.55 -0.21 u 1 - U ~ +0.57 -0.01 DAPmPU -11.47 -0.44 &versed Watson-Crick-Hoogsteen DAP. U1 -6.69 -0.21 DAP*U, -5.80 -0.24 Ui-Ue +0.60 -0.02 DAP.2U -11.89 -0.47 Reversed Watson-Crick-reversed DAP U1 -6.69 -0.21 Hoogsteen DAP-Uz -5.55 -0.21 Ui*Ut $0.42 -0.01 DAP.2U -11.82 -0.43

.

-

-1.67 -13.48 - 1.16 -7.87 -0.51 -6.27 -0.03 +0.53 -1.70 -13.61 -1.16 -8.06 -0.48 -6.52 -0.03 +0.55 -1.67 -14.03 - 1.16 -8.06 -0.51 -6.27 -0.03 $0.38 - 1 70 -13.95

ELECTRONIC

STRUCTURE

OF NUCLEIC

ACIDS

369

inthis casetheformaftion ofthetriplet represents a moreadvantageous organization by aboztt 1.dha2perbase. The interesting comparison of the two open triplets, A . 2U and DAP . 2 U is shown in Tables XIV and XV (97). In these tables, the calculations concerning these triplets are dissociated into their constituent elements, representing the partial interactions between each pair of bases. It is interesting to observe that while the interactions between the linked bases correspond to attractions (equal at k s t approximation to the attractions between the corresponding isolated pairs), the interaction betweenthenonlinked terminal basesintroduces a repulsion. Although the value of thiq repulsion term is relatively small, it, seems to have in this particular case important structural consequences. Thus, for example, the most stable configuration for the A . 2 U trimer is predicted to be the Watson-Crick-reversed Hoogsteen one, a result due largely to the effect of the repulsion between the two uracils. Should this repulsion be neglected, the most stable configuration would be the Watson-Crick-Hoogsteen one. It is the somewhat lower repulsion observed in the Watson-Crick-reversed Hoogsteen model that ensures the overall greater stability of this model over the Watson-Crick-Hoogsteen model, On the other hand, the most stable cordiguration for the DAP * 2U trimer is predicted to be the reversed Watson-Crick-Hoogsteen one, followed closely by the reversed WatsonCrick-reversed Hoogsteen one. Both predictions are in agreement with the previously quoted observations. It must nevertheless be observed that some of the experimental results refer t o structures of polynucleotides in solution, the overall stability of which also involves similar van der WaalsLondon interactions between stacked bases (see below) and is influenced by the effect of the solvent. It is nevertheless probabIe that the preference for one of the configurations over the other springs at least in part from the factors analyzed here. Thus, altogether, uniform calculations of the intermolecular forces are obviously able t o account correctly for a large variety of observed facts. This situation may be considered as signifying that these forces play an important if not a dominant role in these facts.

C. Vertical Interactions between Stacked Bases and the Interaction of the Base Pairs in the DNA Helix Similar calculations may and have been performed (93) for the stacking mode of association of the bases, whether free bases (or their nucleosides) in aqueous solution (as studied in particular by Ts’o and his co-workers (142, 210-214) or linked bases as in the various di-, oligo-, and polynucleotides. The forces involved me the same as those considered for the

370

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

horizontal association of the bases, although the calculations indicate that, while it is the electrostatic component of the van der Waals-London forces that predominates in hydrogen bonding, it is the dispersion component that is the most important one in the vertical interactions, a conclusion in agreement with the experimental deductions of Ts’o (214; see also 215). Altogether, the calculations for the stacking type of interactions are more difficult, and much longer to carry out, because they do not correspond to a geometry known in advance and imply therefore a search for the position(s) of maximum interaction. This may be a long and a tedious process (93,202). We shall not discuss the results of such studies here. As an example, we may just indicate the results obtained for the stacking interaction of purines. Figure 10 indicates the most stable configuration predicted for the stacking of two purines in refined calculations, including the repulsion term at short distlances.The equilibrium distance is found to be 3.4A, and the overall arrangement is of the “alternate stack” type (21.4, 216). This configuration corresponds very closely to the one proposed by Ts’o etal.(214, 216)for the stacking of two purine nucleosides in water from nuclear magnetic resonance studies of the concentration dependence of proton shifts. (A different and a less probable arrangement following our own has been suggested in ref. 217.) The value found for the interaction energy corresponding to the configuration represented in Fig. 10 is 4.9 kcalj mole, in satisfactory agreement with the experimental enthalpy of self(The daerent contributions association of purines in water (218,1219,219a). to this value are, in kcal/mole: E,, = -2.1;E,,= -0.6; E L = -4.9; Erepulsion = +2.7.) The variation of the interaction energy as a function of the relative position of the two purines is illustrated partially in Fig. 11. In the calculations corresponding to this figure, the purines are held at a fixed intermolecular distance of 3.4A. For convenience in the calculations, the molecular axes are fixed with respect to the dipole moment (parallel and perpendicularly). Their origin is in the center of the moment. I n Fig. 11, starting from the original position, noted (0,O) and corresponding to the antiparallelism of the dipole moments, one purine is displaced by 1A relative to the ot8heralong the in-plane axis; these translations are noted (1,O) etc. Each displacement is followed by a rotation of one of the purines around the third vertical axis, giving rise to the different curves of Fig. 11. The complete study implies translations in the intermediate directions and along the vertical axis. On the other hand, we may go over directly to the results obtained by this type of calculation for the neighbor base pairs of the DNA helix, in which the two types of interaction, horizontal and vertical, exist simultaneously and in which a fixed geometry may he assumed for such neighbor

FIG.10. The most probable arrangement for the stacking of two purines (interplane distance 3.4 A).

Mutual angle of the dipoles

FIG.11. Evaluation of stacking interactions between twopurines.

372

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

TABLE XVI NEARESPNEIGI~BOR BASE-BASEINTERACTIONS IN THE DNA HELIX IN A v.%CUUM (KCA1./2 MOliES OF BASE)

Vertical interactions Adjacent basepairs

EPP

Eon

EL

Average contribution Total ofthe Total stacking in-plane interaction energy interactionsenergy

+o. 9

-2.0

-10.2

-11.3

-19.2

-30.6

-1.6

-2.5

-4.0

-8.5

-19.2

-27.7

+2.6

-2.0

-8.3

-7.7

-19.2

-26.9

f1.2

-0.8

-10.3

-9.9

-12.2

-22.1

-0.6

-1.7

-4.9

-7.2

--1 2.2

-19.4

-0.1

-1.7

-5.2

-7.0

-12.2

-19.2

f1.8

-1.0

-7.8

-7.0

-12.2

-19.2

+0.5

-0.5

-7.4

-7.4

-5.5

-12.9

+0.4

-0.3

-6.2

-6.1

-5.5

-11.6

f1.5

-0.7

-5.8

-5.0

-5.5

-10.5

pairs. These results are reproduced in Table XVI and represent in fact, in this particular case, a reevaluation in the “monopole” approximation of the earlier results of De Voe and Tinoco (lo), obtained by these authors in the “dipole” approximation. The notations are those utilized by De Voe and Tinoco, the arrows designating the direction of the chain pointing from the 3 carbon on one sugar tothe 5‘ carbon on the adjacent sugar: e.g., 14 represents dThd3‘-P-B’-dAdo (or d-TpA). Table XVI requires some explanation. The first four columns of numbers indicate the calculations concerned specifically with the stacking or vertical interactions. Column four indicates the values of the total stacking energy. Although this energy is particularly high for some couples of neighbor G C pairs and particularly low for some couples of neighbor A . T pairs, it is nevertheless

ELECTRONIC

STRUCTURE

O F NUCLEIC

373

ACIDS

+

not possible to relate its value in a regular way to, say, the G C content of the helix. It is, however, evident that the total interaction energy between near-neighbor base pairs must involve also the in-plane horizontal interaction energies between the hydrogen-bonded bases. When the average contribution of the in-plane interactions [deduced from the values listed in Tables VII and VIII for the Watson-Crick pairing of guanine with cytosine (- 19.2 kcal/mole) and adenine with thymine (- 5.5kcal/mole)] are added to the vertical interactions, one obtains the last column, which then indicates the total interaction energy for each group of adjacent base pairs. It is now evident from these last results that the dgerent combinations of the base pairs may be clearly divided into three groups. The most stable combinations are those formed by the G C pairs only. Next in order of decreasing stabilities come the mixed combinations comprising an A T and a G * C pair. Finally, the less stable combinations are those containing only A T pairs. It must be emphasized that this result, evidently consistent with the well-established greater thermal stability of nucleic acid rich in guanine and cytosine over those rich in adenine and thymine ( 2 2 )enables , to bring a tentative answer to the ill-debated question whether it is the horizontal, in-plane interactions between the hydrogen-bonded bases or the vertical interactions between the stacked bases that are responsible for the overall stability of the double-helical structure of DNA. The calculations indicate that the two types of interaction are apparently of a comparable order of magnitude and contribute therefore nearly equivalently to the overall stability. It may be observed that while the electrostatic E,,term makes the predominant contribution to the in-plane interactions, it is the dispersion EL term that contributes mostly to the stacking type of interaction. Although we have not discussed here in any detail the contribution of the solvent to this stability, and although information about this effect seems much less quantitative than that concerning the van der Wads!221-226) that London interaction, it nevertheless indicates (93,110-112, the contribution of solvation effects are certainly far from negligible and possibly of the same order of magnitude as those of the horizontal or vertical interactions. The overall stability may then be the resultant of three nearly equal contributions.

-

-

-

D. Related Problems of Molecular Associations Involving Purines and Pyrimidines

At least two problems closely related to those investigated above have been studied quantum-mechanically along the same main lines of approach.

374

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

One of them is the probable intercalation of aminoacridines between adjacent base pairs of DNA as advocated in particular by Lerman (226’236). The quantum-mechanical computations, which appear compatible with an intercalation model, have been carried out by two groups: Gersch and Jordan (106, 107) and Gilbert and Claverie (108, 109). The second problem concerns the physical interaction between carcinogenic aromatic hydrocarbons with purines or with the nucleic acids in solution, as exemplified by the solubilizing effect of the bases or the nucleic The stacking nature of the acids upon the hydrocarbons (237-242). interactions with the bases seems to have been confirmed more recently by a number of physicochemical techniques, and an intercalation model for interactions with the nucleic acids is under consideration (1.42, ,%?.4-2.48). Calculations confirm the predominant role of van der Waals-London forces 105). At the same time, they lead, however, to in these interactions (104, the prediction that this loose, physical type of interaction should not show any specificity with respect to the carcinogenic activity of the hydrocarbons. This prediction is verified by experiment. A recently discovered, strong, chemical interaction with the nucleic acids may be of a more direct significance for carcinogenesis (249, 250). In a somewhat related field of research, two types of calculations, one making use of the intermolecular forces approach (124) and a second one investigated theconformation of applying the extended Huckel theory (31), the nucleosides of the purine and pyrimidine bases of the nucleic acids. Both calculations predict that although there should be two minima in the conformation for uridine, potential energy curves, there is a preferred anti cytidine, and adenosine and a preferred syn conformation for guanosine. These predictions appear to be in agreement with the available experimental data (142,651, 252). Finally, it has also been shown that a number of molecular associations involving biological purines (and to a much lesser extent pyrimidines), which have frequently been somewhat loosely attributed to charge transfer complexes, with the purines generally considered as the electron donors, can be accounted for in a much more satisfactory way as loose van der WaalsSuch is the case of the much-investigated London associations (114). purine-isoalloxazine interaction. In fact, an important general conclusion from this particular research is that, in molecular associations between conjugated molecules, the van der Waals-London forces are generally responsible for the major part of the stability of the ground state of the association, even if the complex may definitely be characterized as a charge-transfer one (say by the appearance of the characteristic, new charge transfer band). Altogether, it seems that the possible involvement of purines, or to a lesser degree pyrimidines, in charge transfer complexes has been overestimated.

ELECTRONIC

STRUCTURE

OF NUCLEIC

ACIDS

375

VII. Problems in Radio- and Photobiology A. Spin Densities in Free Radicals Derived from Nucleic Acid Bases Among the problems of radio- and photobiology that recently have undergone a theoretical investigation are the nature and structure of the free radicals derived under the influence of radiation from the nucleic acid bases and the mechanism of photodimerization of thymine. The mechanism of photodimerization of thymine, which appears to be one of the most important reactions in photobiology, is discussed in the next section. In this section, we limit ourselves to the problem of the free radicals produced upon irradiation of purines and pyrimidines. The most significant development in this field is the demonstration that the principal species formed upon irradiation of purine and pyrimidine derivatives in the powdered state or in single crystals are free radicals formed by hydrogen addition to selected positions of these bases. The first such radical to be discovered is the radical IX derived from thymine, which

Radical ofthymine

(MI

appears also to be the principal rntlicnl species observed upon irradiation of the nucleic acids (253-259). Since this discovery, Gordy and co-workers have demonstrated the formation of a similar type of free radicals, in particular through proton or deuteron bombardment, from other bases of the nucleic acids and from also 262,263). The related analogs and polynucleotides (14’7-149,260,261; structure of the principal free radicals obtained in this way from the nucleic acid bases is indicated in Fig. 12. The contributions of theory to the study of the problems connected with the formation and structure of these free radicals have been manifold. In particular, rules have been proposed for the prediction of the relative and for the prediction of the radioresistance of the different bases (126,126) Calculations have also been site of addition of the hydrogen atoms (146). performed on the distribution of the spin densities in the radicals. It is obvious, in fact, that the lone electron is not locatledentirely at one atom as indicated in Fig. 12,but, being of the ?r type, is spread out partially over the whole molecular periphery. The distribution of the spin densities and

376

BERNARD

PULLMAN

AND

0

PULLMAN

H

H

Uracil

ALBERTE

Cytosine 0

Guanine

Adenine

FIG. 12. Free radicals from H-addition to purines and pyrimidines.

COMPARISON

OF

TABLE XVII OBSERVED AND CALCULATED SPIN DENSITIES OF 13-ADDITION RADICALS I N P U R I N E S AND PYRIMIDINESa Spin density

Radical source

H-addition on

Thymine Uracil 5-OH-uracil 5-C1-uracil Cytosine

C-6 c-5 C-6 C-6 c-5

Deoxyadenosine (monohydrate)

c-2

Guanine (hydrochloride dihydrate)

C-8

4

b

Spin density on

c-5 C-6

c-5 c-5 C-6

j 1;

1

Reproduced from Gordy (149). Derived from observed value of 0.75 at 300°K.

Observed a t 300 K 0.70 0.71 0.64 0.80 0.71b 0.15 0.37 0.38 0.08

Calculated 0.72 0.72 0.67 0.77 0.71 0.12 0.38 0.39 0.08

ELECTRONIC

STRUCTURE

OF NUCLEIC

ACIDS

377

in particular the location of the maximum spin densities can be determined, and it has been, by Gordy and his co-workers, through the study of the hyperhe coupling constants in the electron spin resonance spectrum of the radicals. We present here only the comparison of the computed and observed values of the maximum spin densities. The comparison is indicated in Table XVII, reproduced here by permission of Dr. Gordy from a review paper (149). The agreement between theory and experiment is excellent. It may be worthwhile to stress that the indicated spin densities have been computed within the simpIe Huckel method and that in all having been computed before the expericases they represent predictions, mentation was carried out. In some cases the knowledge of the theoretical values was useful in assigning the appropriate structure to the observed radical.

B. The Mechanism of Thymine Photodimerization Among the different problems investigated theoretically in relation with the photochemistry of purines and pyrimidines, we would like to recall here the contribution of the molecular orbital studies to the problem of the mechanism of thymine photodimerization. This reaction, first discovered upon ultraviolet irradiation of frozen 271)irradiation of DNA, is presently thymine (264-269) and invivo(270, recognized as one of the most important in photobiology. It takes place through the C5-C6 bond of thymine and involves the formation of a cyclobutane ring. The reaction also occurs, with more or less facility, with different thymine analogs (272). The first molecular orbital calculations on the problem (131, 13.2) were carried out within the general frame of the Huckel approximation. They aimed a t the determination of the electronic characteristics of the first excited state of the compounds studied and upon a search for a correlation between these characteristics and, say, the rates of photodimerization of a series of pyrimidine bases. Among the characteristics of the excited states, two appear in fact to be particularly outstanding in connection with their possible significance to the problem under investigation. 1.The first characteristic concerns the distribution on the molecular periphery of the “uncoupled” electrons of the first excited state, which in this approximation may be confused with the “spin densities.” This distribution is illustrated in Fig. 13 for the specific examples of thymine, uracil, and cytosine. Its examination in these molecules and in a series of analogs shows the following. ofthepyrimidines, thehighest concentrations of thelone a. In themajority

378

BERNARD PULLMAN AND ALBERTE PULLMAN 0.151

0.162

0.641

co.013

I

0.033 0.611

Thymine

Uracil

o.3f30.350

0.017

0.507

0 0.161

~ 0 ‘ 2 g 2

Cytosine FIG.13. Distribution of the uncoupled electrons in the first excited state (HMO).

electrons i ntheir jirst excited state occurat C-5and (7-6. The only cases for which this does not happen are those of the bases that do not dimerize. In 5-nitrouracil or 2-thiothymine, the maximum concentration of the lone electrons is at the extracyclic substituent, the NOzor SH group, respectively. b. A most, striking parallelism exists between theyield of photodimerization TABLE XVIII CONCENTRATION OF THE UNCOUPLED ELECTRONS AT THE 5-6BONI) O F THE PYRIMIDINES I N THEIRFIRSTEXCITED TRIPLET The yield of photodimerization Great,

Alean

Small None

Compound Thymine Uracil 6-Methyluracil Orotic acid N1,N3-Dimethyluracil Isocytosine 5-Aminouracil 5-Methylcytosine Cytosine 2-Thiothymine 6-Nitrouracil 6-Azathymine

Concentration of the uncoupIed electrons at the C5-C6 bond 1.21 1.25 1.21 1.12 1.16 1.16 1.05 0.88 0.86 0.71 0.64

1.14

ELECTRONIC

STRUCTURE

OF NUCLEIC

379

ACIDS

0

0

o*cy7 /J N

yy

0.818 0 . 3 1

0.811

H

N H

Ground state

First excited state

00.368

0.466

00.356

0.449

FIG. 14.Mobilebond orders.

and thetofal concentration of theuncoupled electrons attheC5-CG bond of the pyrimidines in their first excited state. This parallelism is illustrated in Table XVIII. The only exception to this rule is offered by G-szatJhymine. It is possible that the inability ofthis molecule to undergo photodimerization may be due to its general relative insensitivity to the effect of ultraorto the difficulty of incorporating a nitrogen atom violet irradiation (27’3) into a cyclobutane ring. 2. The second characteristic concerns thestriking decrease of themobile orderof theC5-C6 bond upon excitation, as opposed to the rather very limited modifications of the mobile orders of all the other bonds of the molecules. The phenomenon is illustrated in Fig. 14 for the specific example of thymine, and Table XIX indicates the modification ofthe mobile order of the C5-C6 bond in all the pyrimidines investigated. An approximate overall inverse correlation can be seen again between the rate of photodimerization and the bond order of the C5-C6 bond in the first excited state (or a direct correlation between the rate and the variation of this bond TABLE XIX MOBILEORDER OF THE C5-C6 Boxu ~

~~

Pyrimidine

Ground state

First excited state

Thymine Uracil &Me thyluracil Orotic acid xi1 Nl,N3-Dimethylura Isocytosine 5-Aminouracil 5-Methylcytosine Cytosine 2-Thiothymine 6-Nitrouracil 6-Amthp i n e

0,811 0. 819 0.790 0,809 0.811 0.817 0.806 0.749 0.758 0.804 0.733 0.831

0.287 0.298 0.296 0.323 0.348 0.319 0.300 0.430 0.459 0.474 0.57s 0.295

~

Variation upon excitation 0.524 0.521 0.494 0.486 0.463 0.498 0.506 0.319 0.299 0,330 0.155 0.536

380

BERNARD

PULLMAN

AND

ALBERTE

1.409

1.356

PULLMAN

a; 1.814

0.817

1.149 0.995

HN

N

1.7391

1.079

1.2991

1.231

0 1.394

Electronic charges

0

0.477

0.346

H

H Bond orders

0

0 0.852 0.855

NH2

I

X Y 0.588

0

H

0.860

H

13:::

0

H

Free valences on the C5-C6 bond FIG.15.First excited singlet of uracil, thymine, and cytosine.

order upon excitation). Azathymine is again a striking exception to such a correlation. These results clearly show, within the limit of the method employed, that in these biological pyrimidines the electronic excitation seem to be localized largely in the C5-C6 double bond and that some features of this excitation may apparently be related tothe rate and the mechanism of the photodimerization reaction. There is one important point, however, for which the preceding conof the siderations are insufficient. This concerns the question of thenature first excited state actually involved inthis reaction. Thus, as is well known, the

-

PPO 0

~666’0

215'0

666’0 OSO'O

Y"'"'" 562’0

0

PPS’O

puoq 9 3 - 4 9 a q uo saauapAaa.qg

H

H

16B’O

("yo

- IQS’O 3 EH

I

’HN

7

PBL’O

0

NH 0

sJap.ro puos

H

H

/N

966’0

I

928’0

LZS'O

WE’O

O

ISP’O

166’0

0

I

'HN

68L’I

T8E

382

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

Huckel approximation of the molecular orbital method does not distinguish between the first excited singlet and triplet. Itrs results concern the “first excited state,” without it being possible explicitly to relate it to thc different multiplicities. Although it may probably be considered, and has been so by Mantione and Pullman (1%),that the distribution of the unpaired electrons may give a first approximation to the spin densities distribution in the first excited triplet, it may also represent some feature of the excited singlet. In order to be able to solve this question, more refined approximations of the molecular orbital method, which remove the degeneracy, must be used. Attempts in this direction have been carried out recently, but before describing them, it may perhaps be useful to indicate how this problem appears from the experimental point of view. Because of the inhibiting effect of oxygen and paramagnetic ions, Beukers and Berends (269)favored the hypothesis of an intermediate triplet state. At least three groups of workers studying in detail the kinetics of photodimerization of orotic acid also favor the involvement of the triplet state in the reaction (274-277). On the other hand, Shulman and coworkers, who some time ago seem to have favored a singlet state intermediate (27 8), recently adopted a more delicate viewpoint by estimating that while the triplet state is probably the precursor of the dimer in liquid solution, the dmer formed in frozen aqueous solution most likely originates from an excited singlet state (279). Lamola and Mittal (280) have emphasized the importance of the structure of the base and of that of the solvent by showing, with dienes as specific triplet quenchers, that the photodimerization of thymine in acetonitrile proceeds entirely through the triplet state while the photodimerization of uracil in acetonitrile and in water proceeds in par2through the triplet state, More recently, Lamola and Yamane (281) have shown that a sensitized photodimerization of thymine occurs upon irradiation of water solutions of DNA containing acetophenone (a sensitizer that allows only the thymine triplet to be populated), which obviously shows at least that the thymine triplet state can lead to a dimer. Therefore, a competition between the singlet and triplet in photodimerization of the biological pyrimidines seems possible, and this situation may perhaps account both for the success of the simple Huckel type calculation and-a point to which we are now coming-the difficulties of the more refined calculations to correlate unambiguously the yield of photodimerization with the structural properties of either the excited singlet or triplet. Figures 15 and 16 represent some typical aspects of the results obtained 7’8) for the first excited singlets and by the SCF CI procedure (22-26, triplets of thymine, uracil, and cytosine, the only molecules for which they

EL ECTR O N I C STR UCT UltE OF NUCL E IC ACIDS

383

seem to be available at present. The results obtainable by the other refined methods are quite similar. It may be observed in particular that the calculations and their comparison with the previously published results on the properties of the ground states of these molecules indicate: (a) a high concentration of spin densities on the C5-C6 bond in the triplet state of the pyrimidines, the concentration being, however, much greater for uracil and thymine than for cytosine; (b) a decrease in the bond order of the CbC6 bond upon excitation, again much more pronounced in the case of uracil and thymine than in the case of cytosine; (c) an increase in the free valences of C-5 and C-6 upon excitation, these free valences being, in both excited states, greater in uracil and in thymine than in cytosine. The particular feature of these results is that they allcorrelate with the greater tendency to photodimerization ofuracil and thymine over cytosine, and are therefore insufficient to indicate whether it is one or the other of these excited states that is involved in the reaction. Obviously more extended calculations are needed before a theoretical conclusion may be reached, inasmuch as it can be reached, about the preferential involvement of the excited singlet or triplet in the photodimerization mechanism (136). Very recently these studies have been extended to the evaluation of the energies and the electronic structures of the different possible conformers of the thymine photodimer (282). A numerical error is present in the diagrams indicating the distribution of spin densities in the first triplet of purines in refs. (22,94,78). The corrected values are reproduced in Fig. 17.

VIII.Electronic Factors in Mutagenesis3 One of the consequences of the recent rapid development of our general knowledge of the biochemical role of the nucleic acids is the recognition of the physicochemical nature of biological mutations. It is generally accepted today that the genetic code resides in the sequence of the purine . pyrimidine must therefore consist of a changeinthis base pairs of DNA. A mutation sequence, a change that may occur as a result of a substitution an inversion, a deletion, or an addition of the bases. [For general reviews see (283-289)l. The existence of such a precise definition of mutation makes some aspects of its mechanism quite accessible to a direct quantum-mechanical investigation (160). Of course, although the fundamental nature of the event representing the mutation is common to all cases, the molecular aspects of the phea

See article by Singer and Fraenkel-Conrat, in this volume [Edh.].

384

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

.457

Adenine

Purine

0.640

0 I

Guanine FIG. 17.Spindensities in thefirst excited triplet.

nomenon depend on the circumstances that induce the transformation. From that point of view, mutations may be divided, in the first place, into spontaneous and induced mutations, the spontaneous ones being those that apparently occur without the obvious intervention of any visible external agent. Induced mutations may in turn be subdivided into those produced by radiations and those produced by chemicals. Among the radiationinduced mutations, we may distinguish those due to UV light and those due to ionizing radiations. Among the chemically induced mutations we may distinguish those due to base analogs or antimetabolites and those due to chemicals that bear no structural relationship to the purine or pyrimidine bases, etc. The subject is thus very broad, and in this review we can deal explicitly with only some of its aspects. We shall therefore try to choose those that are particularly illustrative of the way in which the methods of quantum chemistry may help in understanding the mechanism, at the electronic level, of the mutational transformations corresponding to these different possibilities. A particularly important problem from this point of view is the possible role of the tautomerimtion of the bases, whether spontaneous or induced, in producing mutations.

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As indicated by a large number of experimental investigations, based on X-rays, UV- and infrared spectroscopy, the determination of ionization constants, etc., the purine and pyrimidine bases of the nucleic acids may be considered as existing essentially in their lactamand aminoforms (Fig. 18).This situation is in agreement with the quantum-mechanical calculations on the relative stabilities of the different possible tautomeric structures of the bases (1, 26). The specific pairing of adenine with thymine and of guanine wit.h cytosine, characteristic of the Watson-Crick model of DNA, is dependent on the predominance of these usual tautomers.3 The possibility that spontaneous mutations may involve the rare tautomeric forms of the bases was advanced as early as 1953by Watson for uracil and and Crick. Such rare tautomeric forms would be lactim for adenine and cytosine (Fig. 19). The presence of a guanine and imine rare tautomeric form may give rise to a coupling, through hydrogen bonds, of unusual bases (Fig. 20) and may thus lead to a perturbed sequence of base pairs in later generations-that is, to a mutation. Although other mechanisms, such a4 miscoupling of ionized rather than tautomeric bases (151,290-292) or base deletions (293), may also play a role in spontaneous mutations, the intervention of rare tautomenc forms remains plausible. It may also be considered in connection with mutations induced with base analogs or by the effect of radiation. Hence, from the quantum-theoretical point of view, it may be interesting to investigate a number of questions connected with the possibility of tautomerization of the bases. Among these questions are the problems of which of the purine and pyrimidine bases of the nucleic acids has the greatest probability of existing in a rare tautomeric form and thus to be particularly involved in spontaneous mutations, what are the principal characteristics of the rare tautomeric forms of the bases, and finally, what are the consequences of their interference (i.e.; of a miscoupling) on a number ofphysicochemical properties of the nucleic acid, e.g., their stability, relevant to their biological function? The problem of the tendency of the bases to exist in their rare forms, for which very limited experimental data are available, can be dealt with relatively simply at least in a first rough approximation. Thus it can be shown (294,295) that when a given type of tautomeric equilibrium is being studied in a related series of compounds, the essential varyingfactor tendency of the compounds to exkt in a rare responsible for the relative tautomeric form is the variation of the resonance energy accompanying the tautomeric transformation. In the case of the purine and pyrimidine bases, and there are two such transformations to be considered :the lactam-lactim transformations." In the l ~ ~ u m tautomerism, - l u ~ ~ the ~ ~ the amino-imine transformation of the Zactamform (the most stable) to the Zactim form

386

BERNARD PULLMAN AND ALBERTE PULLMAN

Adenine

Thymine

Guanine

Cytosine

FIG.18. The usual forms of the purine and pyrimidine bases.

Adenine

. 5 C %

0A N H Thymine

Guanine

..$ A

N H

Cytosine

FIG.19. Rare tautomeric forms of purines and pyrimidines.

I H

Cytosine {rare imino form). adenine (normalform)

H

Guanine ( r a r e enolform). thymine (usual form)

FIG.20.Examples of misccupling of the bases of the nucleic zcids.

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ACIDS

(less stable) is associated with an increase of resonance energy so that the proportion of the Zactim form will be the greater, the greater this increase. i ~ z n e the transformation of the aminoform I n the a ~ ~ n ~ tautomerism, (the more stable) to the iminoform (less stable) is accompanied by a decrease in resonance energy and will therefore be the grent.er, the smaller this decrease. Explicit, calculations of resonance energies for the different tautomeric forms of the purine and pyrimidine bases of the nucleic acids, lead to the prediction that the bases that, from this point of view, should forms are have the greatest tendency to exist in a rare iminoand lactam TABLE XX RESONANCEENERGIES OF THE TAUTOMERIC FORMSSUSCEPTIBLE TO BE PRESENTIN DNA (IN 0 UNITS = 16 KCAL/MOLE) Compound ~

Giianine Uracil Thymine Adenine Cytosine

Resonance energies of the tautomeric forms ~~~

Lactam: 3.84 1.92 2.05 Amine: 3 . 8 9 2.28

Ah!= ~

Lactim: 4.16 2.14 2.27 Imine: 3 . 6 2 2.15

~~

0.32 0.22 0.22 -0.27 -0.13

AR = Variation of resonance energies accompanying the transformation from l.he stable to the less stable form.

cytosine and guanine, respectively (Table XX) (67). These are therefore the bases that have the greatest probability to be involved in spontaneous mutations insofar, of course, as tautomerization may be considered as a cause of such mutations. The transformation G . C -+ A . T should then be more frequent than the reverse one. It may be interesting to note that the fact that the G . C pairs constitute the unstable part of the genome and that they mutate spontaneously more frequently than the A T pairs has been reported in a number of publications (28s) 284,23296). At this point it may also be interesting to inquire about other effects that the presence of rare tautomers and subsequent miscouplings may introduce in the physicochemical properties of DNA. Without going into details of this problem, which would require a complete description of the electronic characteristics of the rare forms of the bases, we may point out a difference in one of the essential characteristics between the normal and the rare forms of the bases that may have a significant bearing on one of the properties of the nucleic acids we studied earlier, namely, their stability. It is recalled that the dipole moments of guanine and cytosine ( = 7 D) are predicted to be appreciably greater than those of adenine and thymine ( ~ 3 -D). 4 The dipole moments predicted for the rare farms Qf the bases

388

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(which we shall mark with an *) are (297) A* = 7.3D,T' = 5.7D, G' = 3.6D, and C* = 4.9 D. Thus one observes a complete reversal of the relative classification of the bases, a situation suggesting that large changes may be expected in the stabilities of nucleic acids involving such rare forms [the more so as variations predicted in the ionization potentials between the common and the rare tautomers are much less important (297)l. That such TABLK XXI INTERACTION ENERGIES IN THE WATGON-CRICK AND MISCOTTPLEDBASE PAIRS(KCAL/MOLE)

A .T G*C A’ . Ca C’. A G * .T T'.G

-4.61 -15.91 -16.78 -7.85 -6.83 -14.86

-0.27 -2.02 -1.67 -0.49 -0.05 -1.60

-0.77 -1.25 -0.99 -0.90 -1.11 -0.99

THE

-5.65 -19.18 -19.44 -9.24 -7.99 -17.45

~

a

Asterisk designation defined in text.

is really the situation may be seen from the data of Table XXI indicating the van der Waals-London interaction energies calculated for miscoupled base pairs involving the rare tautomers. It may be predicted that miscoupled pairs containing the rare forms of guanine or cytosine should be appreciably less stable than those containing the rare forms of adenine or thymine. The formation of the first ones (which as we have seen previously is more probable than that of the second ones) may therefore introduce local elements of instability into the nucleic acids. Finally, it may be interesting to investigate the situation that might occur in base pairs if both components were in their rare forms. In such a case, the coupling would continue to occur between the Watson-Crick complementary bases, but would then be of the type A’. T', G* C'. Calculations show that, within the same approximations, the van der Waals-London interactions energies corresponding to such two pairs should be - 19.1and - 12.3kcal/mole, respectively. One would observe thus a reversion of the relative order of interaction energies with respect to the usual pairs A T and G . C. This prediction may perhaps be interesting in connection with the concept advocated by Lowdin and his school (116-118) that mutations are the result of a double proton tunneling between the bases. Such tunneling would result in the formation of the unusual pairs A* . T* and G* . C'. When considering the possible role of tautomerization of the bases in rnutagsnesis, one need not restrict oneself to spontaneous mutations. The

-

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O F NUCLEIC ACIDS

same factor may be consiclercd as playing a certain role in induced mutations, whether by physical or chemic. 1 mcans. Thus it is generally acknowledged that ultraviolet and ionizing radiations may be mutagenic either by direct action on DNA or through different indirect effects, for insfance by producing reactive substances able to interact with DNA or base analogs susceptibleto be incorporated into DNA. As one of the essential consequences of, say, the absorption of UV r d i a tions, the purine and pyrimidine bases of DNA are raised into excited states. I n these circumstances, a reasonable hypothesis capable of accounting for the enhancement of mutageiiesis by UV irradiation consists in postulating that the irradiation increases, among other effects, the chances of the events that are already responsible for the occurrence of mutations in the ground states of the molecules, in particular spontaneous mutations. Specifically, attention may thus again be focused on the relative tendencies of the bases to exist in different tautomeric formsin their excited states. The calculations of the resonance energies have therefore been extended to the first excited state of the purine and pyrimidine bases (the involvement of the higher excited states being less probable in chemical phenomena because of the rapidity of internal conversion) and compared with the same quantities determined previously for t.he ground state. The results are TABLE XXII VARIATION OF RESONANCEENERGIES ( A R ) UPON TAUTOMERIZATION (IN j3 UNITS) Tautameric transformation Lactam-lsctim

Amino-imino

Compound

Uracil Thymine Guanine Cytosine Adenine

A R in the groundstate

A R inthefirst

0.22 0.22 0.32 -0.13 -0.27

0.43 0.40 0.36 -0.01 -0.14

excited state

summed up in Table XXII for the essential tautomeric transformations susceptible of involvement in the mispairing of the bases and thus in mutations. It is shown in Table XXII that (a) the gain of resonance energy accompanying the lactam-lactim tautomerization of uracil, thymine, and guanine is greater in the first excited state of these molecules than in their ground state; (b) the loss of resonance energy accompanying the aminoimino tautomerization of cytosine and adenine is smaller in the first excited state of these molecules than in their ground state. Consequently, the ease of the tautomeric transformation into the rare form should be greater for all these compounds in their excited states, and this situation may account,

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PULLMAN

at least in part, for the enhancing effect of UV radiations on the rate of mutagenesis. Cytosine, for which the two tautomeric forms appear to be of nearly identical energy in the excited state should, from this point of view, be the essential site of mutation. Although this factor certainly cannot be considered as the only one or even the most important one in UV-induced mutagenesis, it is interesting to remark that cytosine does seem to be the primary target of such mutations in a number of cases (298, 299). Of course, other modifications occurring in the electronic structures of the bases upon excitation may also be responsible for the increased mutagenesis. Such other factors could be, for instance, the modification of the electronic charge of the nitrogen atom of the bases involved in the glycosyl linkages. For example, an increase of its net positive charge induces an increase in the rate of the enzymatic or acidic hydrolysis of the linkage (300), a situation that may facilitate the incorporation of a wrong base or the breaking of the chain. Calculations carried out for the distribution of electronic charges in the first excited state of the molecules indicate that such an increase should actually accompany the excitation of the pyrimidines. With the usual restrictions concerning the validity of Huckel-type calculations for the study of specific excited molecular states, the results are illustrative of some ways in which UV radiations may induce or increase physicochemical changes leading to mutagenesis. On the other hand, apart from raising the molecules into their excited states, radiations have also some indirect effects. Thus, as is well known, among the jmportant products of the UV irradiation of the bases of the nucleic acids are the hydrohydroxy derivatives of the pyrimidines (Fig. 21), in which the elements of water have been added to the C5-C6 bond of these bases (and the dimer of thymine, of course, in which the C5-C6 bond is also saturated). Although the hydration itself is not a mutagenic transformation, it represents an example of a UV-induced abnormal component. Of course, the tautomeric equilibrium may be displaced in these components, and if the displacement is in the direction of the rare form, the chances of the miscoupling of bases and thus of mutation will be increased. That such may actually sometimes be the case is indicated by the calcultltions carried out for the C5-C6 saturated derivative of cytosine, which show that the loss of resonance energy on passing from the aminoto the

FIG. 21.The hydrohydroxy derivative of cytosine.

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iminoform is reduced in the saturated derivative of the compound. On the contrary, the tautomerization of uracil is predicted to be more difficult in the hydrated form (61). The same type of consideration may, of course, be applied also to similar modifications introduced by chemical means. Thus the study of the action of hydroxylamine on polynucleotides has shown that the alterations of structure due essentially to the saturation of the C5-C6 link of the pyrimidine are capable of introducing miscouplings capable of leading to mutations; e.g., the enzymatic replication of poly C treated with hydroxylamine (or irradiated) shows that C5-C6 saturated cytosine is able to behave 302). Although the mutagenic effect of a C5-C6 saturated like uracil (301, into cytosine may sometimes be attributed to its deamination in situ uracil, it seems in other cases to be linked to the activity of the saturated form itself (303, 304). This is, in particular, the case for the mutagenic effect of hydroxylamine, where the species responsible for the errors in coupling would be the 5,6-dihydro-6-hydroxylamincytosine,st,ructure X NH

H 5,6-Dihydro-6hydroxy laminocytosine

(X)

(305), in which the effect could be due to the increased probability, indicated by previously quoted calculations, of the saturated form to exist in configuration. (See article by Phillips and Brown in Vol. 7 of this the imine series.) The perturbations produced in the coupling by the saturation of the 5-6 double bond of uracil seem to be of a more ambiguous nature (302,303, 306, 307). The previously quoted calculations point to the conclusion that they cannot be due to an increased probability of its existence in the form. In this respect, it is interesting to note the recent conrare lactim to the effect that the mutagenic clusion of Rottman and Cerutti (308) transitions due to irradiation of poly U cannot be considered as a simple result of the saturation of the C5-C6 bond but involve perhaps a spwial direct effect of the hydroxyl group fixed at position C-6. In these investigations on mutagenesis, definite conclusions as to the most probable mechanism are difficult and the preceding considerations are essentially illustrative ofthe possible contributions of quantum-mechanical calculations to their investigation. In no way do they mean that we consider the tautomeric shifts as the most important such mechanism. In

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PULLMAN

some cases, connected in particular with chemical mutagenesis, it seem rather obvious that they are not and that other mechanism are much more likely. Such is, for example, the case of the action of the alkylating agents. These compounds, which besides their mutagenic activity display also carcinogenic and antitumor action, are known to attack the nucleic acids. It has been an outstanding success of the theory to have been able to predict which should be the most significant centers of the bases and of the nucleic acids to undergo these attacks (1, 6, 63). These centers have been shown to be the most basic nitrogens of the purine and pyrimidine rings. Thus it may be predicted that, while in guanine the most basic nitrogen shodd be located in the imidazole ring and be in fact N-7,the situation should be quite different in adenine, where the most basic nitrogen should be located in the pyrimidine ring. I n the nucleic acids, the calculations predicted the attack to occur primarily on N-7of guanine. Experimental results on the action of alkylating agents on nucleic acids, purine ribonucleotides orribonucleosides, and purine bases confirm these general predictions. They indicate in particular that, in the nucleic acids, guanine derivatives and guanine, the alkylating agents attack essentially N-7 of the guanine ring (509-316). The questions may then be raised, what are the possible consequences of these attacks and how do they relate to the mutagenic properties of the alkylating agents? A tentative answer to these questions may be obtained if we look at the electronic structure, in particular the charge distribution, in the guanine alkylated at N-7and compare it with that of guanine itself (Fig. 22). The most important perturbations affect the imidazole ring ofthe guanine skeleton, and among them we may note, in particular, the great increase of the net positive charges of N-9and C-8.This situation suggests immediately the possible occurrence, under the influence of alkylation at N-7of two phenomena: 1.An appreciable increase of the susceptibility of the glycosyl linkage of guanine to enzymatic hydrolysis. In fact, it has been shown, as already 1.481

1.468

?

0.789

I

+.!

0.991

1.803

Guanine

N-7-Alkylguanine FIG. 22.Electronic charges.

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mentioned (SOO), that the rate of such a hydrolysis of purine and pyrimidine ribonucleosides and ribonucleotides is parallel to the value of the net positive charge of the nitrogen atom of these bases engaged in the glycosyl linkage. 2. An appreciable increase of the susceptibility of C-8 to undergo attacks by nucleophilic agents, such as hydroxyl ions, with the concomitant increase of the degradation of the guanine moiet'y through the opening of its imidazole ring. Both phenomena may have as a result the perturbation of the normal base-pair sequences and may contribute therefore to the mutagenic activity of the alkylating agents. In fact, both effects have been suggested, on experimental grounds, as contributing to this activity (284, 316, 317). The preceding considerations refer to the production of point mutations through the specific interaction of an alkylating agent with a selected purine. Some of these agents have, however, a wide variet,y of action; they may produce interstrand cross-linking in DNA (318) or they may attack The mutagenic effects they the phosphate groups of the nucleic acids (319). may produce in these ways would consist in large deletions. In concluding, it may be added that similar investigations have also been carried out for the electronic aspects of the mutagenic activity ofother chemicals such as nitrous acid, formaldehyde, aminoacridines, etc. (160, 320, 321). Quite generally, the correlation with the electronic properties of DNA is satisfactory. On the whole, the physicochemical nature of the basic phenomena involved in mutagenesis seems daily to become more and more evident, and the interpretation of these phenomena at the electronic level appears relatively conclusive. This situation raises new hopes for the similar understanding of the mechanism of carcinogenesis, whether spontaneous, radiation-induced, or provoked by chemical means, the two processes, mutagenesis and carcinogenesis, possibly being interrelated to some extent, although there is no doubt that the problem of carcinogenesis, asunderstood today in molecular and quantum biology, is probably a more complex one than that of mutagenesis. Thus, while carcinogenesis also undoubtedly involves in its fundamental mechanism an alteration of the genetic apparatus of the cell, it is in no way certain yet whether the principal, direct site of action of the carcinogens is the nucleic acids or other cell constituents, in particular some essential proteins (320-322).

IX. Conclusion This paper is by necessittylimited to the discussion ofonly a few selected subjects in the broad field of qunntiini-mechanical calculations of the

394

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PULLMAN

properties of the nucleic acids and their constituents. The subjects considered are, however, different enough to indicate some of the general features of this mode of approach. In the first place, it is obvious that the calculations on these molecules have been carried out, by now, by a large variety of methods. To some extent, insofar as relatively large molecules are concerned, these conjugated heterocycles are probably among the most thoroughly studied ones. Although the intrinsic values of t,he different types of calculation are uneven, we have today as a result of these concentrated efforts a large amount of data about a number of aspects of the electronic structure of these type of compounds. Presently it may even be said definitely: a large data. number of reliable In the second place it m ~ be y useful to stress that, for a number of years, marly of these properties-e.g., dipole moments, ionization potentials, electron affinities, spin densities in free radicals originating from the bases-have been known only theoretically. It is only in very recent years that some of these molecular properties could be determined experimentally. The agreement between theory and experiment, remarkable in most respects, needs t o be refined in some cases. Because of the slowness of the experimental development, the theory still retains largely, in this field, the importance of its predictive character. From a different point of view, one of the most impressive features of the procedure is its universal character, its unlimited applicability. Thus, while the usual experimental methods of chemistry and physics are intended t o study essentially one (sometimes more, but never too many) specific molecular property, the quantum-mechanical studies aim at obtaining, through the operation of solving the wave equation, a multiplicity of results that in principle (i.e., if we were really able to solve rigorously the extensive equations) yield complete information about all the structural properties of the system under investigation. Even if, as is the case inpractice, we can solve only approximately somewhat reduced equations, the amount of information-which is, of course, approximate and partial-still generally covers a wide variety of aspects of the problem studied. In this respect, the extension of the calculations beyond the usual r-electrons approximation toward the simultaneous study of all valence (an even simple all) electrons represents a broadening of the possibilities of the procedure, which may be of particular significance in biochemistry (and pharmacology) in that i tenables thequantitative inclusion of steric and con&-mational efects into what was previously an essentially electronic description. Finally, this recent evolution of the theoretical procedures, together with the rapid development of the computational techniques and of the

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capacities of the computers themselves, open the possibilities for an effective direct investigation of the macromolecules themselves and of their essential properties, which up to now had to be deduced by extrapolation from smaller systems. Such an enlargement is essential forthe transformation of quantum biochemistry into a quantum biology.

REFERENCES 1.B. Pullman and A. Pullman, “Quantum Biochemistry.” Wdey (Interscience), New York (1963). 2. A. Pullman, B. Pullman, and G. Berthier, Compt.Rend.Acad.Sci. 243, 380(1956). 3. B. Pullman and A. Pullman, Biochim. Biophys. Acta36, 343 (1959). Q. A. Pullman and B. Pullman, Bull. SOC.Chim.Francep. 766 (1958). 6.A. Pullman and B. Pullman, Bull. SOC.Chim.Francep. 594 (1959). 6. T. Nakajima and A. Pullman, J. Chim.Phys.66, 793(1958). 7. T.Nakajima and B. Pullman, Bull. SOC. Chim.Francep. 1502(1958). 8.T.N a k a j b and B. Pullman, Bull. SOC.Chim.Francep. 663 (1959). 9. B. Pullman and A. Pullman, Nature189,725(1961). 10.H . De Voe and I. Tinoco, Jr., J. Mol.Biol. 4,500 (1962). 11.A. Pullman, Compt.Rend.Acad.Sci. 266, 5435(1963). 12.J . I. Fernandez-Alonso, in“The Structure and Properties of Biomolecules and Biological Systems” (J. Duchesne, ed.), p. 3.Wiley (Interscience), New York 1964. 13.J. Ladik, in“Electronic Aspects of Biochemistry” (B. Pullman, ed.), p. 203 Academic Press, New York,1964. 14. A. Veillard and B. Pullman, Compt.Rend.Acad. Sci. 263, 2277(1961). 15.A. Veillard and B. Pullman, J .Theoret. Biol. 4,37 (1963). 16.R. X. Nesbet, Biopolymers Symp.1,129(1964). 17. Ch. Nagata, A. Imamura, Y.Tagashira, and M. Kodama, Bull. Chem.SOC.Japan 38, 1638(1965). 18.J. Ladik and K. Appel, Theoret. Chim.Acta4,132(1966). 19.M . Tanaka and S. Nagakura, Theoret. Chim.Acta6, 320 (1966). 20. V. A. Kuprievich, Intern. J. QuantumChem.1,515 (1967). 21. A. Imamura, K.Fujita, and C. Nagata, Bull. Chem.SOC.J a p a n40, 522 (1967). 22. H. Berthod, C. Giessner-Prettre, and A Pullman, Themet. Chim.Acta6, 53 (1966). 23.H. Berthod, C. Giessner-Prettre, and A. Pullman, Compt.Rend.Acad.Sci. 262, 2657(1966). 24.H. Berthod, C. Giessner-Prettre, and A. Pullman, Intern. J .Quantum Chern. 1,123 (1967). 95.A. Denis aridA. Pullman, Theoret. Chim.Acta7,110(1967). 26.H. Berthod and A. Pullman, J .Chim.Phys.62, 942 (1965). 27. H. Berthod and A. Pullman, Biopolymers 2,483(1965). 28.D. F. Bradley, S. Lifson, and B. Honig, in“Electronic Aspects of Biochemistry” (B. Pullman, ed.), p. 77.Academic Press, New York, 1964. 29. G. Del Re, J. Chem.SOC.p.4031(1958). 30. G. Del Re, in “Electronic Aspects of Biochemistry (B. Pullman, ed.), p. 221. Academic Press, New York, 1964. 31.F. Jordan and B. Pullman, Theoret. Chim.Acta9,242 (1968). 3.2.A. Pullman, E. Kochanski, M. Gilbert, and A. Denis, Theoret. Chim.Acta10,231 (1968).

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BERNARD

PULLMAN

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ALBERTE

PULLMAN

33.C. Giessner-Prettre and A. Pullman, Yheoret. Chim.Acta9, 279 (1968).

34.A. Pullman, Ann.N . Y .Acad.Sci.(1968) in press. 36.E. Clementi arid D. R. Davis, 6. Compul. Phys.1,223(1966). 36.E. Clementi, H. Clcmenti, and D. It. Davis, J .Chem.Phys.46,4725(1967). 37.A. Streitwieser, Jr., “Molecular Orbital Theory forOrganic Chemists” Wiley, New York,1961. 38. I(. Higasi, H. Baba, and A. Remhaurn, “Quantum Organic Chemistry.” Interscience, New York, 1965. 39. H. H. Jaff6,in “Comprehensive Biochemistry” (M. Florkmand E. H. Slots, eds.), Vol. 1,p. 34.Elsevier, Amsterdam, 1962. 40.A. Pullman, in “Molecular Biophysics” (B. Pullman and M. Weissbluth, eds.), p.81.Academic Press, New York, 1965. 4i. R. G. Pam, “Quantum Theory of Molecular Electronic Structure.” Benjamin, New York, 1963. 42. L. Salem, “The Molecular Orbital Theory of Conjugated Systems.” Benjamin, New York, 1966. 43.K. Ohno, Advan.QwzntumChem.3,239 (1967). 44.A.Pullman, Ann.N .Y .Acad.Sci.in press. 46. R. Hoffmann, J .Chem.Phys.39, 1397(1963). 46.R. Hoffmann, J.Chem.Phys.40, 2745(1964). 47.E. Clementi and D. L.Raimondi, J .Chem.Phys.38,2686(1963). 48.L.C. Cusachs, Quantum Theory Project, University ofFlorida, Rept. No. 50, p. 36 (1964). 49. L. C. Cusachs, J .Chem.Phys.43, 5157(1965). 60. G. W. Wheland and D. E. Mann, J .Chem.Phys.17, 264 (1949). 61. A. Streitwieser, Jr., J. Am. Chem.Soc. 82,4123(1960). 6.9. L. C. Cusachs and J. W. Reynolds, J .Chem,Phys.43,S160(1965). 63. D. G. Carroll, A.T. Armstrong, and S. P. McGlynn, J .Chem.Phys. 44,1865(1966). 64.R. Rein, N. Fukuda, H. Win, and G. A. Clarke, J .Chem.Phys.46,4743(1966). 66.J. A. Pople and G. A. Segal, J. Chem.Phys.43, S129(1965). 56.J. A. Pople and G. A. Segal, J .Chem.Phys. 44,3289(1966). 57.J.A. Pople and G. A. Segal, J. Chem.Phys.43,S136(1965). 68. R. L. Muller and P.G. Lykos, Tetrahedron Letters 11, 493(1962). 69. B. Pullman, Tetrahedron Letters No. 4,231(1963). 60.B.PuUman, J .Org.Chem.29,508 (1964). 61.N. Dupuy-Mamelle andB. Pullman, J .Chim.Phys.64,708 (1967). 62.B. Pullman, in “Molecular Biophysics” (B. Pullman and M Weissbluth, ds.), p. 117.Academic Press, New York, 1965. 63. H. Berthod, C.Giessner-Prettre, and A.Pullman, Theoret. Chim.Acta8,212(1967). 64.A. Pullman and M. Rossi, Biochim. Biophys. Actu 88,211(1964). 65.Ch. Lifschits, E. D. Bergmann, and B. Pullman, Tetrahedron Letlers 46,4583 (1967). 66.B.Pullman, J .Chem.Phys.43,S 233 (1965). 67. B.PulIman and A. Pullman, Biochim. Biophys. Actu64,403(1962). 68.A. Pullman, Biochim. Biophys. Acta87,365 (1964). 69. 0.V. Shramko, V. I. Danilov, and Yu A. Kruglyak, Biofirika 10, 561 (1965). 70.B. Pullman, ActaCryst. 17, 1074(1964). 71. A. Pullman, Bull. SOC.Chim.Francep. 641(1958). 72.H. G. Mautner, and G. Bergson, ActaC h . Scand. 17,1694(1963). 73.M. Tanaka and 5. Nagakura, Theoret. Chim.Acta6 ,320 (1966).

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ACIDS

397

74.S. Kwiatkowski, ActaPhys.Polon. 29, 573 (1966). 76.J. S. Kwiatkowski, Mol.Phys.13, 229(1967). 76.A. Imamura, H. Fujita, and C. Nagata, Bull. Chem.SOC. Japan40, 21 (1967). 77.J. P. Malrieu, Compt.Rend.Acad.Sci. 264, 662 (1967). 78.A. Pullmsn, i n“The Triplet State” (A. B.Zahlan, ed.), p. 515.Cambridge Univ. Press, London and New York, 1967. 79.A. Pullman and E. Kochanski, Intern. J .Quantum Chem.lS,251 (1968). 80.I.Tinoco, J .Am. Chem.SOC.82,4785(1960). 81. W. Rhodes, J .Am. Chem. SOC.83, 3609(1961). 82. W. Rhodes, J .Chem.Phys.37, 2433(1962). 85.H. De Voe and I. Tinoco, J. Mol.B i d .4, 518 (1962). 84. I.Tinoco, in“Molecular Biophysics” (B. Pullman and M. Weissbluth, eds.), p. 269. Academic Press, New York, 1965. 86. A. D. Mc Lachlan and M. A. Ball, Mol.Phys.8, 581 (1964). 86.G. Felsenfeld and S. Z. Hirschman, J. Mol.Biol. 13, 407 (1965). 87.G. N. Fowler, Mol.Phys.11, 31 (1966). 88.C. A. Bush, and I. Tinoco, Jr., J. Mol.Biol. 23, 601(1967). 89.H. A. Nash, and D. F. Bradley, J .Chem.Phys.46, 1380(1966). 90.H. Jehle, W. C. Parke, and A. Salyers, i n “Electronic Aspects of Biochemistry” (B. Pullman, ed.), p.313.Academic Press, New York, 1964. 91.H. Jehle, W. C. Parke, and A. Salyers, Biophysics 9, 433(1965). 92. B. Pullman, P. Claverie, and J. Caillet, Proc. Natl. Acad.Sci. U .S.66,904(1966). 93.B.Pullman, P. Claverie, and J. Caillet, J .Theoret. Biol. 12, 419(1966). 94.B. Pullman, P. Claverie, and J. Caillet, J. Mol.Biol. 22, 373(1966). 96.B. Pullman, P. Claverie, and J Caillet, Compt.Rend.Acad.Sci. 263, 2006(1966). 96.B. Pullman, P. Claverie, and J. Caillet, Proc. Natl. Acad.Sci.U .S .67,1663(1967). 97.B. Pullman and J.Caillet, Theoret. Chim.Acta8, 223(1967). 98.B. Pullman, H. Berthod, and J. Caillet, Theoret. Chim.Acta10,43 (1968). 99.M. Pollak and R. Rein, J. Theoret. Biol. 11, 490(1966). 100.R. Rein, P. Claverie, and M. Pollak, Intern. J. QuantumChem.2, 129 (1968). 101.R. Rein, N. S. Goel, N. Fukuda, M. Pollak, and P. Claverie, Ann.N . Y.Acad.Sci. in press (1968). 102.F. Hofelich, J .Chem.Phys.47, 2433(1967). 103. P. Claverie, i n “Molecular Associations in Biology” (B. Pullman ed.), p. 115. Academic Press, New York, 1968. 104. B. Pullman, P.Claverie, and J. Caillet, Science 147, 1305(1965). 106. J.Caillet and B. Pullman, i n“Molecular Associations in Biology” (B. Pullman, ed.), p.217.Academic Press, New York, 1968. 106. N. F. Gersch and D. 0. Jordan, J. MoZ. Biol. 13, 138(1965). 107.D. 0. Jordan, i n “Molecular Associations in Biology” (B. Pullman, ed.), p. 221. Academic Press, New York, 1968. 108.M. Gilbert and P. Claverie, in“Molecular Associations in Biology” (B. Pullman, ed.), p. 245.Academic Press, New York, 1968. 109.M. Gilbert and P. Claverie, J. Theoret. Biol. 18, 330 (1968). 110. 0. Sinanoglu and S. Abdulnur, Photochern. Photobiol. 8, 333 (1964). i l l0. . Sinanoglu and S. Abdulnur, Federation Proc.24, 512 (1965). 112.0.Sinanoglu, in “Molecular Associations in Biology” (B. Pullman, ed.), p.427. Academic Press, New York, 1968.

113. A. Pullman and B. Pullman, i n ‘LQuantumTheory of Atoms, Molecules and the Solid States (P. 0. Idwdin, ed.), p. 345,Academic Press, New York, 1966.

398

BERNARD

PULLMAN

AND

ALBERTE

PULLMAN

114.M. J.Mantione, in“Molecular Associations in Biology” (B. Pullman, ed.), p. 411. Academic Press, New York, 1968. 116.P. 0. Uwdin, Rev.Mod.Phys.36, 724(1963). 116.P. 0. Liiwdm, Mutation Res.2, 218 (1965). 117.P. 0. Lowdin, in“Electronic Aspects of Biochemistry” (B. Pullman, ed.), p.167. Academic Press, New York, 1964. 118.P. 0. Lijwdin, Advan.QuantumChem.2, 213(1965). 119.R. Rein and I?. E. Harris, J .Chem.Phys. 41, 3393(1964). 120.R. Rein and F. E. Harris, J. Chem.Phys. 42, 2177(1965). 191.R. Rein and F.E. Harris, J. Chem.Phys.43, 4.415(1965). 122.R. Rein and F. E. Harris, J. Chem.Phys. 46,1797(1966). 123.S. Lunell and G. Sperber, J .Chem.Phys. 46, 2119(1967). 1.94. I. Tinoco Jr., R. C. Davis, and S. R. Jaskunas, in “Molecular Associations in Biology” (B. Pullman, ed.), p. 77.Academic Press, New York, 1968. 125.B. Pullman and A. Pullman, in“Comparative Effects of Radiation” (M. Burton, J.S. Kirby-Smith, and J. L. Magee, eds.), p. 105.Wiley, New York, 1960. 126.B. Pullman, Mem. Acad.Roy.Belg. 33, 174(1961). 127.J.Duchesne and A. Van de Vorst, Bull. Cl.Sci. Acad.Roy.Belg. 61, 778 (1965). 128.A. Van de Vorst, M. Richer, and K. V. Rajalakshmi, Bull. C1. Sci. Acad.Roy.Belg. 62, 276 (1966). 129.M. Lacroix, J.Depireux, and A. Van de Vorst, Proc. Natl. Acad.Sci.U.S .68, 399 (1967). 130.A. Pullman and B. Pullman, Biochim. Biophys. Acta76, 269 (1963). 131.M. J.Mantione and B. Pullman, Biochim. Biophys. Acta91, 387 (1964). 132.B. Pullman and M. J. Mantione, Biochim. Biophys. Acta96, (1965). 133.C. Nagata, A. Imamura, Y. Tagashira, and M. Kopama, J .Theoret. Biol. 9, 357 (1965). 134.A. Imamura, H. Fujita, and C. Nagata, Bull. Chem.SOC.Japan40,21 (1967). 135.V. I. Danilov, Photochem. Photobiol. 6, 233 (1967). 136.B. Pullman, Photochem. Photobiol. 7, 525 (1968). 137.A.Veillard, B.Pullman, and G. Berthier, Compt. Rend.Acad.Sci. 262, 2321(1961). 138.A. Veillard and B. Pullman, Compt.Rend.Acad.Sci. 263, 2277(1961). 139.A. Veillard, J .Chim.Phys. 69, 1056(1962). 140.C. Giessner-Prettre and B. Pullman, Compt.Rend.Acad.Sci. 261, 2521(1965). 141.M. P. Schweizer, S. I. Chan, G. K. Helmkamp, and P. 0. P. Ts’o, J .Am. Chem.SOC. 86,696 (1964). 142.P. 0. P. Ts’o, M. P. Schweizer, and D. P. Hollis, Ann.N .Y .A d .Sci.(1968), in press. 143.J. Baudet, G .Berthier, and B. Pullman, Compt.Rend.Acad.Sci.264, 762(1962). l&. A. Pullman, J .Chim.Phys.61, 1666(1964). I&. B.Pullman and M. J. Mantione, Compt.Rend.Acad.Sci. 260, 5643(1965). 146.B.Pullman and M. J. Mantione, Cmnpt.Rend.Acad.Sci. 261, 5679(1965). 14Y. J.N. Herak and W. Gordy, Proc. Natl. Acad.Sci. U.S.64, 1287(1965). 148. J.N. Herak and W. Gordy, Proc. Natl. Acad.Sci.U.S.66, 1354(1966). 149.W. Gordy, Ann.N .Y .Acad.Sci. in press (1968). 160.A. Pullman, ed., in“Electronic Aspects of Biochemistry,” p.135.Academic Press, New York, 1964. 161.C. Nagata, A. Imamura, H. Salto, and K. Fukui, Gann 64, 109(1963). 162.B.Pullman, Cmnpt.Rend.Acad.Sci. 269, 3101 (1964). 263. T. A.Hoffman and J. Ladik, in“The Structure and Properties of Biomolecules and Biological Systems” (J.Duchesne, ed.), p.84. Interscience, New York, 1964.

ELECTRONIC

STRUCTURE

OF NUCLEIC

ACIDS

399

lii4. J. Ladik and K. Appel, J .Chern. Phys.40, 2470(1964). 155.J.Ladik and G. Bicso, J .Chem.Phys.42, 1655(1965). 156.J. Ladik, ActuPhys.Hung.13, 473(1961). 157.H. A. Nash and D. F. Bradley, Biopolymers 3,261 (1965). 168.S. Diner, G .Del Re, and B. Pullman, Compt.Rend. Acad. SCi. 262, 826 (1966). 159.A. M. Perrault, C. Valdemoro, and B. Pullman, J .Theoret. Biol. 2,180(1961). 160. P. S.Song and T. A. Moore, Intern. J .Quantum Chem.1, 699 (1967). 161.B. Pullman and A. Pullman, Bull. SOC.Chim.France, p. 973 (1958). 162. A. Pullman and B. Pullman, Compt.Rend.Acad.Sci. 246, 611 (1958). 163.H. Berthod and A. Pullman, Compt.Rend.Acad.Sci. 262, 76 (1966). 164.P. Pithova, A. Piskala, J. Pitha, and F.Sorm, Collection Czech. Chem.Commun. 30, 1626(1965). 166.J.Pitha, P.Fiedler, and J. Gut, Collection Czech. Chem.Commun. 31, 1864(1966). 166.P. Machmer and J. Duchesne, Nature206, 618 (1965). 167.D. L.Smith and P. J. Elving, J .Am. Chem.Soe.84,1412(1962). 168.D. L. Smith and P. J.Elving, J .Am. Chem.Soc.84,2741(1962). 169.W. A. Struck and P. J.Elving, Biochemistry 4, 1343(1965). 170.P. J.Elving, Ann.N .Y .Acad.Sci., in press. 171.S. F. Mason, J .Chem.Soe., p.2071(1954). 172.G. H. Beaven, E. R. Holiday, and E. A. Johnson, in “The Nucleic Acids” (E. (Chargaff and J.N. Davidson, eds.), Vol. I, p. 493.Academic Press, New York, 1955. 173.I. Tinoco, Jr. and H. De Voe, J .Mol.Biol. 4,518 (1962). 174.D. Voet, W. R. Gratzer, R. A. Cox, and P. Doty, Biopolymers 1, 193(1963). 175.L. B. Clark and I. Tinoco, Jr., J .Am. Chem.SOC.87,11 (1965). 176.L. B. Clark, G.G. Peschel, and I. Tinoco, Jr., J .Phys.Chem.69,3615(1965). 177.J. Drobnik and L. Augenstein, Photochem. Photobiol. 6, 13 (1966). 178.J. Drobnik and L. Augenstein, Photochem. Photobiol. 6, 83 (1966). 179.V. Kleinwachter, 6.Drobnik, and L. Augenstein, Photochem. PhotoSiol. 6, 579 (1966). 180.F. Bergmann and S. Dickstein, J. A m . Chem.Sac.77, 691(1955). 181.R. I?.Stewart and N. Davidson, J .Chem.Phys.39, 255 (1963). 182.11.F. Stewart and N. Davidson, Biopolymers Symp.1,465 (1964). 185.P. R. Callis, E. J.Rosa, and W. T. Simpson, J. Am. Chem.SOC.86, 2292(1964). 184.A. M. Michelson, T. L. Ulbricht, T. R. Emerson, and R. J. Swan, Nature209, 873 (1966). 185.J. 0.Hirschfelder, i n “Molecular Biophysics” (B. Pullman and M. Weissbluth, eds.), p. 325.Academic Press, New York, 1965. 186.K. Hoogsteen, ActaCryst. 12, 822 (1959). 187.K. Hoogsteen, ActaCryst. 16, 907 (1963). 188.A. E. V. Haschemeyer and H. M. Sobell, ActaCryst. 18, 525 (1965). 189.A. E. V. Haschemeyer and H. M. Sobell, Proc. Natl. Acad.Sci. U .S .60,872(1963). 190.L. Katz, K. Tomita, and A. Rich, J .Mol.Biol. 13, 340 (1965). 191.F. S. Mathews and A. Rich, Nature201, 179(1964). 192.F. S. Mathews and A. Rich, J. Mol.Biol. 8,89 (1964). 193.H. M. Sobell, K. Tomita, and A. Rich, Proc. Natl. Acacl. Sci. U.S.49, 885(1963). 194.R. R. Shoup, H. T. Miles, and E. D. Becker, Biochem. Biophys. Res.Commun. 23, 194(1966). 195.Y. Kyogoku, R. C. Lord, and A. Rich, Sciences 164, 518 (1966). 196.F. H. C. Crick, J .MoZ. Biol. 19, 548 (1966). 197.J.H. Miller and H. M. Sobel, Proc.Natl. Acad.Sci.U.S.66, 1201(1966).

400

BERNARD

PULLMAN

AND ALBE RTE PULLMAN

198. D. G. Watson, R. M. Sweet, and R. Marsh, A d a Cryst. 19, 573(1965). 199. A. Rich, Biochim. Biophys. Actu29, 502 (1958). 900. H. T.Miles, Proc. Natl. Acad.Sci. U .8.61, 1105(1964). ,901. L.L. Labana and H. M. Sobell, Proc.Nail. Acad.Sci. U.S.67,459(1967). ,202.B.Pullman and A. Pullman, Advan.QuantumChem.4, 267 (1968). 203. J. Caillet, Compt.Rend.Acud.Sci. 266,288 (1968). 204. R. M. Hamlin Jr., R. C. Lord, and A. Rich, Science 148,1734(1965). 206.Y.Kyogoku, R C. Lord, and A. Rich, J.Am. Chem.SOC.89,497 (1967). 206. J. H. Miller and H. M. Sobell, J .Mol.Biol. 24, 345(1967). ,207.2. Katr and S. Penman, J. Mol.B i d .16, 220 (1966). 208. J. Pitha, R. Norman Jones, and P. Pithova, Can.J. Chem.44,1045(1966). ,209. E.D. Bergmann, private communication. ,210. P. 0. P.Ts’o, in“The Nucleohistones” (J. Bonner and P. 0. P. Ts’o, eds.), p.149. Holden-Day, San Francisco, 1964. 811. M. P. Schweizer, S. I.Chan, G. K. Helmkamp, and P. 0. P. Ts o, J. Am. Chem.Soc. 86, 696 (1964). 212. M. P. Schweizer, S. I. Chan, and P. 0. P. Ts’o, J .Am. Chem.SOC. 87,5241 (1965). 813. A. D. Broom, M. P. Schweizer, and P. 0. P. Ts’o, J. Am. Chem.SOC.89, 3612 (1967). ,214. P. 0. P. Ts’o, in “Molecular Associations in Biology” (B. Pullman, ed.), p.39. Academic Press, New York, 1968. 216.S. Hanlon, Biochem. Biophys. Res.Commun.23, 861(1966). ,216. A. D. Broom, M. P. Schweirer, and P. 0. P. Ts’o, J. Am. C h m . Soc.89,3612 (1967). 217. G. K. Helmkamp and N. S. Kondo, Biochim. Biophys. Acta146,27 (1967). 818. S. J. Gill, M. Downing, and G. F.Sheats, Biochemistry 6 ,272 (1967). 219. K. E.Van Holde and G. P.Fbssetti, Biochemistry 6, 2189(1967). ,219~.P. R. Stoesser and S. J. Gill, J. Phys. Chem.71,564(1967). $20. J. Marmur and P. Doty, Nature201, 179(1959). ,221. N. R. Kestner and 0. Sinanoglu, J. Chem.Phys. 38, 1730(1963). 2.88.A. Sinanoglu, Ann.N .Y. Acud.Sci. in press. ,223. M. Falk, K. A. Hartman, Jr., and R. C. Lord, J. Am. C h m . SOC.86, 387 (1963). 884.L. Levine, J. A. Gordon, and W. P. Jencks, Biochemistry 2, 168(1963). $26.S.Levin, Proc. Biochem. SOC.93, 16P (1964). 286. L.S. Lerman, J .MoZ. Biol. 3, 18 (1961). 387. L.S. Lerman, Proc. N d .Acad.Sci. U.S.49, 94 (1963). 228. L. 5.Lerman, J .MoZ.Biol. 10, 367 (1964). 229. G. Weil and M. Calvin, Biopolymers 1, 401(1963). 230. R. K. Tubbs, W. E.Ditmars, and Q. Van Winekle, J. Mo1. Biol. 9, 545 (1964). 231. J.Chambron, M. Daune and C. Sadron, Biochim. Biophys. Acta123, 306 (1966). ,239. I. Isenberg, R. B. Leslie,S. L.Baird J., R. Rosenbluth, and R. Bersohn, Proc.Natl. Acad.Sci. U.S.62, 379 (1964). 233. B. J. Gardner and S. F. Mason, Bwpolymers 6, 79 (1967). 23.4. G. V. Gurskii, Biophysia 11, 849 (1966). 236. A. J. Blears and S. S. Dangluk, Biopolymers 6, 535 (1967). 236. D. M. Neville, Jr. and D. R. Davies, J. Mol.Biot. 77, 57 (1966). 237. H. Weil-Malherbe, Biochem. J. 40,351(1946). 338.E. Boyland and B. Green, Bm t. J. Cancer16, 347 (1962). $39. E.Boyland and B. Green, Brit. J. Cancer16, 507(1962). 240. E. Boyland and B. Green, Bwchim.Biophys. Acta87,653(1964).

ELECTRONIC

STRUCTURE

OF NUCLEIC

ACIDS

401

241. B. C. Giovanella, L. E. Mc Kinney, and C. Heidelberger, J .Mol.Biol. 8,20(1964). 242. A. M. Liquori, B. De Lerme, F. Ascoli, C. BotrB, and M. Frascicatti, J .Mol.BWZ.

6,527 (1962). 24.9. B.L. Van Duuren, J .Phys.Chem.68,2544(1964). 2a. B. L. Van Duuren, Nature210, 622 (1966). 246.Ch. Nagata, M. Kodama, Y. Tagashira, and A. Imamura, BiopoZymers 4,409 (1966). 246.M. Kodama, Y. Tagashira, A. Imamura, and Ch. Nagata, J .Biochem. 69, 257 (1966). 247.I. Isenberg and S. L. Baird, Jr., Biopolymers 6, 477 (1967). 248. B. Green and J. A. Mc Carter, J .Mol.B i d .29, 447 (1967). 249. P. Brookes and P. D. Lawley, Nature202, 781 (1964). 260. P. Brookes, CancerRes.26, 1994(1966). 261. A. E. V. Haschemeyer and H. M. Sobell, ActuCryst. 19, 125 (1965). 262.A. E. V. Haschemeyer and A. Rich, J. Mol.Bid. 27, 369 (1967). 263. H. Shields and W. Gordy, Proc. Natl. Acad.Sci.U.S.46, 269 (1959). 264. R. A. Patten and W. Gordy, Nature201, 361 (1964). 266. A. Ehrenberg, L. Eherenberg, and G. Lofroth, Nature200, 376 (1963). 256.R. Salovey, R. G. Shulman, and W. M. Walsh, J .Chem.Phys.39, 839 (1963). 257.J. Eisinger and R. G. Shulman, Proc.Natl. Acad.Sci.U.S.60, 694(1963). 268. W. Gordy, B. Pruden, and W. Sniper, Proe. Natl. Acad.Sci.U.X.63,751 (1965). 269. B. Pruden, W. Sniper, and W. Gordy, Proc.Natl. Acad.Sci.U.S.63,917 (1965). 260.J. N. HBrak and W. Gordy, Proc.Natl. Acud.Sci.U.S.66, 698(1966). 261.Ch. Alexander and W. Gordy, Proc.Natl. Acad.Sci.U.S.68, 1279(1967). 96g. H. C. Heller and T. Cole, Proc.Natl. Acad.Sci. U.S.64, 1486(1965). 26s. D. E. Holmes, L. S. Myers, Jr., and R. B. Ingalb, Nature209, 1017(1966). 264. R. Beukers, J. Ijlstra, and W. Berends, Rec.Trav. Chim.77,729(1958). 266. R. Beuken, J. Ijlstra, and W. Berends, Rec.Trav. Chim.78, 879(1959). 266. R. Beuken, J. Ijlstra, and W. Berends, Rec.Trav. Chim.79, 101(1960). 267. R. Beukers andW. Berends, Biochim. Biophys. Acta41, 550 (1960). 268. W. Berends, J .Chim.Phys.68, 1034(1961). 269.R. Beuken and W. Berends, Biochim. Biophys. Actu49,181 (1961). 270.A. Wacker, H. Dellweg, and D. Weinblum, Nutumoissenschuften 20, 477 (1960). $71. A. Wacker, J .Chim.Phys.68,1041(1961). 272.A. Wacker, This series, 1, 369 (1963). 27s. H. L. Gunther and W. H. Prusoff, Biochim. Biophys. Actu149, 361 (1967). 274. E. Setumpf and D. Shugar, Photochem. Photobiol. 4 ,719(1965). 875.E. Setumpf-Kulikowska, D. Shugar, and J.W. Boag, Photochem. Photobio7. 6 ,41 (1967). 276.A. Haug and P. DOUZOU, 2. Naturforsch. 20b, 509(1965). 277.M. Charlier and C. Helene, Photochem. Photobiol. 6 ,501 (1967). 278. R. 0. Rahn, R. G. Shulman, and J. W. Longworth, J .Chem.Phys. 46, 2955(1966). 279.J. Eisinger and R. G. Shulman, Proc. Natl. Acad.Sci.U.S.68, 895(1967). 280. A. A. Lamola and J. P. Mittal, Science 164, 1560(1966). 281. A. A. Lamola and T. Yamane, Proc. Nut[. Acad.Sci.U.S.68, 443 (1967). 28.9. F. Jordan and B. Pullman, Theoret. Chim.Actu9, 242 (1968). 283. E.Freese, Proc.Intern. Congr.Biochem., MOSCOW, Vol. I, p.204.Pergamon Press, Oxford, 1961. 284.E. Preese, i n “Molecular Genet,irs” (a. H. Taylor, ed.), Part T, p. 207.Academic Press, New York, 1963.

402

BERNARD

PULLMAN

AND ALBERTE

PULLMAN

285.E. Freeze and E. Bauts Freese, Radiatioih Res.SuppZ.6 ,97 (1966). 286.D. R. Krieg, This series, 2, 125(1963).

287. W. Hayes, “The Genetics of Bacteria and Their Viruses.” Blackwell, Oxford, 1964. 288. R. M. Herriott, CancerRes.26,1971(1966). 289. L. E. Orgel, Aduun.Enzymol. 27, 289(1965). 290.P. Brookes and P. D. Lawley, New Scientist No. 342, 543(1963). 291.P. D.Lawley and P. Brookes, Nature192, 1081(1961). 292.P . D . Lawley and P. Brookes, J. Mol.Biol. 4, 216 (1962). 293.S.Brenner, L.Barnett, F. H. C. Crick, and A. Orgel, J. Mol.Biol. 3,121(1963). 294.G .W. Wheland, "Resonance in Organic Chemistry.” Wiley, New York, 1955. 295.B. Pullman and A. Pullman, “Les Theories Electroniques de la Chimie Organique.” Masson, Paris, 1952. 296.J .W . Drake, Proc.Null. A c d .Sci.U.S.66, 738 (1966). 897.B. Pullman and J. Caillet,, Compt.Rend.Acad.Sci. 264, 1900(1967). 298.J. W. Drake, J .MoZ. Biol. 6,268 (1963). 299.B. D. Howard and I. Tessman, J. Mol.B i d .9,372 (1964). 300. A. Pullman and B. Pullman, PTOC. Natl. Acad.Sci. IT.S.46, 1572(1959). 301.J.Ono,R.G. Wilson, and L.Grossman, J. Mol.Bid.11, 600 (1965). 302.J .H. Philipps and D. M. Brown, J. Mol.Bwl.12, 816 (1965). 2703.H . E . Johns, J.C. Le Blanc, and K. B. Freeman, J. MoZ. Biol. 13,849(1965). S04.D . M . Brown and 3.H. Phillips, J. Mol.B i d .11, 663(1965). 306. J. H. Phillips, D. M. Brown, and L. Grossman, J. Mol.Biol. 21, 405(1966). 306.J .Ono, R.G. Wilson, and L. Grossman, J .Mol.Biol. 11, 600(1965). S07. L. Grossman, Proc.Natl. Acad.Sci. U.S.60,657 (1963). 308.F. Rottman and P. Ceruttli, Proc. Natl. Acad.Sci.U. S.65,960(1966). SO9. P.Brookes and P. D. Lawley, Biochem. J. 77, 478 (1960). S10.P. Brookes and P. D. Lawley, Biochem. J. 80,496 (1961). Sll. P. Brookes and P. D. Lawley, J. Chem.SOC.p.3923(1961). 3 2 .P. D. Lawley and P. Brookes, Biochem. J .89, 127(1963). 31s. P. Brookes and P. D. Lawley, J .CeZZuZaT Comp.PhysioZ. 64 (Suppl. l), 111(1964). 314.P. Brookes, in “Chemotherapy of Cancer” (P. A. Plattner, ed.), p.32 Elsevier, Amsterdam, 1964. 316.P . D . Lawley, This series, 6, 89 (1966). 316.J.A. Haines, C. B. Reese, and Lord Todd, A.,J. Chem.SOC.p.5281(1962). 517.L. B. Townsend and R. K. Robins, J. Am. Chem.Soc.86, 242 (1963). 918. P. D. Lawley and P. Brookes, J. MoZ. BioZ. 26, 143(1967). 319.J.T .Lett, G. M. Parkins, and P. Alexander, Arch.Biochem. Biophys. 97,80 (1962). 520. B. Pullman, Biopolyme~s Symp.1, 141(1964). 511. A. Pullman, Biopolymers Symp.1, 47 (1964). S22. B. Pullman, J .CeUdarComp.Physiol. 64 (Suppl. l), 91 (1964).

The Chemical Modification of Nucleic Acids N. I<. KOCHETKOV" AND E. I. BUDOWSKYt *Institute of Organic Chemistry, and tlnstitiite for Chemistry qf Natural Products, of USSR, Academyof Sciences MOSCOW,USSR

I. Introduction . . . . . . . . . . . 11. Possibilities of Chemical Modification in the Study of the Primary and Secondary Structures of Nucleic Acids . . . . . 111. Importance of the Chemical Modification Methods for Studies of Nucleic Acid Function . . . . . . . . . IV. Reactions Used for the Chemical Modification of Nucleic Acids A. Reactions at the Carbohydrate Moiety . . . . . B. Reactions at the Phosphate Group . . . . . . C. Cleavage of the N-Glycosyl Bonds . . . . . . D. Reactions at the Heterocyclic Base Residues . . . . . V. Modification of the Uracil Nucleus with Hydroxylamine . A. Reaction of Uridine with Hydroxylamine . . . . B. The Preparation and Some Properties of Deuridylic RNA's C. The Effect of the Secondary Structure of Polynucleotides upon Hydroxylaminolysis . . . . . . . . . VI. Modification of the Cytosine Nucleus with Hydroxylamine . A. Reaction of the Cytosine Nucleus with Hydroxylamine . B. Application of the Reaction of Hydroxylamine with Cytosine Residues to Functional Studies . . . . . . . References . . . . . . .

.

403

.

404

.

407 405 4053 409 410 411 417 417 421

.

. . . . .

. .

. . .

.

425 427 428 432 433

1. Introduction Modification of the units of polymer chains by reaction with chemical agents is one of the promising methods for studying the structure and function of biopolymers, particularly polynucleotides. The method can be applied at all stages of nucleic acid investigation-isolation of individual polynucleotide species, studies of primary and secondary structure, studies of nucleoprotein complexes, and studies of polynucleotide functional specificity. In the ideal case, the method of chemical modification should yield polymers with chemical alterations of known extent, character, and distribution along the polymer chains. Only in this case is it possible to 403

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interpret unequivocally the results of a chemical modification. However, the realization of these severe requirements is still a problem, and while methods only approximating such an ideal are presently practical, they may still meet with considerable success. Most convenient for the chemical modification would be highly selective single-stage reactions leading to stable modified units of known structure and properties. However, no such reactions have as yet been elaborated. Usually the action of a chemical reagent results in simultaneous, modifications of several types of nucleoside units that may, however, proceed at different rates. The reactions are usually multistage and yield mixtures of products even with monomer models. Still more complicated are the reactions with polynucleotides, which must be regarded as parallelconsecutive ones. For this reason, elaboration of a method of chemical modification should always involve a kinetic study of the reactivity of different nucleosides under the conditions (pH, temperature, concentration, etc.) used subsequently for polymer modification. When applying a chemical modification method to the solution of biochemical or biological problems, one must also consider the possibility of complications due to intra- or intermolecular interactions in polynucleotide complexes that may alter the reactivity of nucleoside monomer units, such as interaction of base residues along a polynucleotide chain or within complementary regions of different chains, interaction with proteins and metal ions. The data on the reactivity of nucleoside units serve as a basis for understanding in chemical terms the specificity of biochemical processes occurring in the cells with polynucleotides and also nucleotide coenzymes. Hence, progress in molecular biology needs further advances not only in analytical and physical chemistry, but also in the organic chemistry of nucleic acids, starting with the reactivity of nucleic acid components.

II. Possibilities of Chemical Modification in the Study of the Primary and Secondary Structures of Nucleic Acids Studies of the primary structure of nucleic acids may be performed principally by three alternative routes : 1. Sequential cleavage and identification of terminal groups. 2.Cleavage of a polymer chain into blocks a t known types of linkage. 3.Direct electron microscopic observation of base residue distribution along polymer chains,

The first route, whether enzymatic (exonucleases) or chemical-enzymatic, presents very strict requirements as to the synchronous action of enzymes, or to the selectivity and quantitative yield of the chemical

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405

reactions applied to remove terminal units. At present there exist neither enzymes nor chemical procedures meeting these requirements to a more or less complete degree, and this first route is used practically either to determine short terminal sequences of polynucleotides or to analyze oligonucleotides built of less than 10 nucleotide residues (1-4). The second route, i.e., block structural analysis, is based on selective cleavage at known types of linkages, subsequent structure determination of the oligonucleotides, and reconstruction of the initial polymer structure (see article by Holley in Volume 8 of this series). As the cleavage is performed only once, the requirements of specificity and quantitative yield are much less rigorous. The specificity of cleavage by guanylyl- and pyrimidinyl-ribonucleases considerably increases when the polynucleotides have a rigid secondary structure (5-6) ; this increase ofspecificity was the major basis of the recent success in determining the sequences of severa1 However, one is limited to a rather small tRNA’s and of 5 S RNA (7-13). number of enzymes. For example, no enzymes are known to be capable of selective cleavage of RNA at uridylic, cytidylic, or “minor” nucleoside units. However, selective cleavage at the units mentioned may be performed by chemical or combined chemical-enzymatic methods. In the first case, it is necessary that the modification weaken the internucleotide bonds adjacent to the modified units. Methods ofthis type with group specificity, like apyrimidination or apurination, have been widely applied in studies of the distribution of nucleotides in DNA (f4-18). Analogous methods having a higher selectivity and enabling chemical cleavage at base residues of only one type may be also elaborated for polyribonucleotides and subsequently applied to studies of RNA primary structure. Another approach provided by the chemical modification method for the selective cleavage of polynucleotide chain into blocks is the alteration of base residue structure in such a way that the corresponding internucleotide bond becomes more resistant to nucleases. This approach is based on an increase in the specificity ofaction of nucleases due to modification of the substrate. For example, the destruction of uracil nuclei by hydroxylamine (19, 20)or their modification by N-cyclohexyl N’-p-(ll-methyl morpho22)results in stabilinium) ethyl carbodiimide (CME-carbodiimide) (21, lization of the adjacent phosphodiester bonds to pancreatic ribonuclease, so that the enzyme splits the modified polynucleotide at cytidylic acid residues only, and thus may be regarded as a highly selective enzyme, “cytidylyl-ribonuclease.” After acrylonitrile action on RNA, pancreatic ribonuclease does not split the polymer chain a t the modified pyrimidine Modification of the guanine nuclei in tRNA with glyoxal moieties (23). results in units stable to T1-ribonuclease, so that the enzyme splits the

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N. K. KOCHETKOV AND E. I. BUDOWSKY

modified chain at only inosine and 2-dimethylamino-6-oxopurineresidues (% 26). Such uses of the chemical modification method make it possible not only to study the primary structure of nucleic acids, but also t o determine the structural features of the nucleoside units determining the selective action of ribonucleases. For obvious reasons, the block method cannot be applied to the sequence determination of high-molecular-weight nucleic acids. The most promising method for determining the structure of polynucleotides (both DNA and RNA) of any length seems to be electron microscopy. The technique is now under development in a number of laboratories (26-30). One of the major problems-making visible in the electron microscope bases of given t y p e c a n be solved by chemical modification methods. To this end, the base must be bound to a contrasting group, capable of extensive electron scattering (one or several heavy metal atoms), i.e., the modifying agent must contain a moiety capable of binding a contrasting reagent. Such alteration does not usually change the specificity of modification. The application of this approach has already been attempted, but not yet with considerable success. However, if one remembers that elucidamolecules of polytion of structure by this method will need only 102-103 nucleotide, it will be realized that further development of the electron microscopic approach is very desirable. Finally, it is noted that chemical modification is at least as sensitive to the secondary structure of polynucleotides as is the action of enzymes, so that fixation of conformation of a polymer chain will increase the specificity of both enzymatic cleavage and chemical modification. For this reason, chemical modification may also be used to study the secondary structure of polynucleotides. The effect of complementary base interaction upon their reactivity is so great that it seems possible to modify units belonging to nonhelical regions selectively and to determine their composition and size, or even to identify “looped” units, if the primary structure of the polynucleotide is established. Studies of this kind are widely performed in many laboratories with a number of modifying agents-acrylonitrile ( S I ) , hydroxylamine (32, SS), CME-carbodiimide (34, 35), perphthalic acid (Sf?), etc. When studying the primary or secondary structure of nucleic acids, one attains usually a maximum (ideally, a quantitative) degree of modification of a given type of unit. Hence, the mechanism of reaction, the kinetics of intermediate stages, and the structures of intermediate products are not very important. Sometimes even a relatively high degree of nonspecific side reaction is permissible.

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111. Importance of the Chemical Modification Methods for Studies of Nucleic Acid Function Use of the chemical modification methods in functional studies of nucleic acids requires an essentially different approach. First of all, a small degree of modification usually leads to a great change in functional activity. For this reason, the reaction must be stopped a t early stages, and to find a correlation of the structural change in the polynucleotide with the change of its functional activity, it is necessary to have reliable data on the mechanism and the kinetics of reaction, on structure and properties not only of final, but also of intermediate reaction products. Second, as only a small fraction of the polymer chain is modified, there arises a new problem, because change of a functional property depends on both the extent of modification and on the distribution ofthe modified units along polymer chain. There is a considerable difference between functional studies of nongenetic nucleic acids (messenger and tRNA, etc.) and those of genetic DNA and RNA. I n the first case, change ofthe functionaI properties reflects the average alteration of a number of molecules; modification is randomly distributed over the chains of all the molecules. The functional properties of each of the modified macromolecules make their contributions to the average value, and each contribution is proportional to its content in the resulting mixture. Hence, with the nongenetic nucleic acids we obtain results that are a statistical average. Both the obtaining of experimental data and their interpretation become more difficult when chemical modification methods are applied to studies of the function of genetic (replicating) nucleic acids, because in this case we observe the properties and fates of single macromolecules differing in both the extent of modi€ication and the distribution ofmodified units along the polymer chain. In this case as well, the results will reflect the relationship between the monomer unit structure and the functional specificity of the polynucleotide. However, in such studies it is necessary to obtain a considerable amount of reliable experimental data and to interpret the results much more carefully than in studies of nongenetic nucleic acids. Obviously, the more precise and reliable are the data on the specificity of modifying agents, on the kinetics and mechanism of the reactions, on the structure and properties of the modified units, the more unequivocal will be the interpretation of the chemical basis of the functional specificity. From this viewpoint, it is easy to realize that successful application of the chemical modification method to functional studies must be based upon detailed knowledge of the organic chemistry of nucleic acids and their

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components. Data obtained in the course of such studies will permit chemical explanations of the biological processes involving nucleic acids.

IV.

Reactions Used forthe Chemical Modification of Nucleic Acids

In this section, we survey briefly the most important chemical reactions of the nucleic acid components used for the chemical modification of nucleic acids. Included in this survey are only the reactions that have been, or axe now being, applied to study the structure and function of the nucleic acids. At the same time, we include also a number of reactions promising to become important for the purpose. We do not discuss here a number of known nucleic acid reactions whose nature prevents them from application to selective chemical modification, nor reactions so briefly studied that the usefulness of the corresponding reagents cannot yet be evaluated. It will be noted that there axe many reactions, mainly with the heterocyclic nuclei, that seem very interesting for future application to nucleic acids modification, but the very poorknowledge of their specihity, mechanism, and kinetics considerably limits their use. Moreover, the lack of such data often leads to incorrect conclusions and invalid speculations, which sometimes result in discrediting the chemical modification method. The reactions used forthe chemical modification of nucleic acids may be divided into four groups of different importance :

(A) Reactions at the carbohydrate moiety (B) Reactions at the phosphate grouping (C) Cleavage of N-glycosyl bonds (D) Reactions at the heterocyclic base residues The reactions of the first group are widely applied during the isolation and terminal sequence analysis of nucleic acids. The second group of reactions is used very rarely and is not ofsufficient importance. The most important at present are the reactions of the third and, especially, of the fourth group, which have found wide application in the study of the structure and function of nucleic acids.

A. Reactions at the Carbohydrate Moiety Modification of the carbohydrate moiety involves principally the free hydroxyls. The reactions may be divided into (a) alkylation and acylation and (b) oxidation. (a) Most studied in detail is the acylation of 2’-hydroxyl groups in polyribonucleotides. Acetylation with acetic anhydride in neutral aqueous

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solution proceeds quantitatively, does not affect any nucleic bases, and results in stabilization of the modified polynucleotide to ribonucleases (37-39). Introduction of an acyl grouping lowers the stability of single- and double-stranded polynucleotide helices (4G42). 2l-O-Acetyl radicals may be removed quantitatively under mild conditions, leaving the polynucleotide molecule practically intact. Alkylation of the 2’-OH group is a much less specific reaction, usually complicating the modification of base residues by alkylating agents. (b) Among the oxidation reactions, the most important is that of periodate oxidation of the terminal 2’,3’-glycol groups of polyribonucleotides. This reaction is highly selective and results in the formation of two carbonyl groups in the /%positionrelative to the phosphate residue, and in

I

M-H !€ 0I. bH

OH

OH R = Polynucleotide chain B = Nucleic acidbase X = Product ofribose interconversion

a).

the a-position relative to the glycosyl center (1, 43, Such alteration of a ribose residue leads to labilization of both the phosphodiester bond of the modified unit in the polynucleotide residue and of the N-glycosyl bond to the base residue. Cleavage of the labilized bonds proceeds under mild conditions; it is catalyzed by amines. The reaction is widely applied to RNA terminal analysis (45-48) and to the isolation and purification of polyribonucleotides. It is to be noted that, under more drastic conditions, periodate reacts with some base residues, especially with pseudouridine (49)*

The free hydroxyls (5 and 3 -OH in polyribo- and in polydeoxyribonucleotides, 2’-OH in polyribonucleotides) may also be oxidized under appropriate conditions. However, the methods used for the purpose, catalytic oxidation with oxygen over platinum (50)and oxidation with dimethyl sulfoxide (51), are not selective and, for this reason, cannot be applied to the modification of polynucleotides.

6. Reactions a t the Phosphate Group The esterification and amidation of terminal phosphate groups in polynucleotides were proposed a few years ago (62,53), but have not yet found application in the structural analysis or functional study of nucleic acids.

410

N. K. KOCHETKOV AND

0

0

1

R-0-P

OH OH

0 II/OH OH

R-0-P,

E. I. BUDOWSKY

CH,OHCDI

PhNH,

cDI

II

R-0 -P-OCH, I OH

0 II

R-0-P-NHPh

I

OH R = Polynucleotide chain

The formation of intrachain phosphate triesters results in facile splitting of internucleotide bonds. The action of alkylating agents that are esters of strong acids, such as dimethyl sulfate, leads practically only to alkylation of nucleic bases under proper conditions (65). On the other hand, alkylation with diazomethane leads to side reactions; the esterification of intrachain phosphate groups and the nonspecific degradation of polynucleotide chains have been reported (56). Alkylation of the primary hydroxyls of internucleotide phosphate residues proceeds under rather drastic conditions and is complicated by simultaneous alkylation of the bases. Several workers have demonstrated that the formation of phosphate triesters leads to facile cleavage of internucleotide bonds, but this reaction cannot be used for a specific degradation of polynucleotide chains. Phosphodiester bonds in DNA are relatively stable, and their cleavage occurs under conditions altering nucleoside residues: in acid, there takes place apurination; in alkali, deamination and degradation of base residues occur. As is well known, the phosphodiester bonds in RNA are readily hydrolyzed owing to the presence of free hydroxyls in the 2‘-position, this results in a mixture of 2’- and 3’-phosphates. However, acidic cleavage of RNA phosphodiester bonds is accompanied by and alkaline cleavage leads to considerable changes some apurination (57), in the base residues (68,69). For this reason, the use ofacidic and alkaline hydrolysis for the determination of RNA nucleotide composition may lead to error. It will be noted, that the rates of phosphodiester bond hydrolysis depend on the structure of the adjacent nucleoside residues and the catalyst used for hydrolysis (60-6s).However, this dependence is not very great, and thus it cannot be used for the selective cleavage ofpolynucleotides.

C. Cleavage of theN-Glycosyl Bonds Selective cleavage of the N-glycosyl bonds of the purine nucleoside residues in DNA in an acidic medium was the first selective reaction proposed for the modification of polynucleotides (64,65). This reaction has

T H E CHEMICAL

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been subsequently studied in more detail and is now often applied to studies of the primary structure of DNA (14, 16). The necessary acidity , 1

ArNH, or Ar,NH

4.

X

.

/ /

DNA

Apurinic acid

Pyrimidine clusters

Pu = Purinebase;Py = Pyrimidine base; X = Product ofdeoxyribose interconversion

(pH 2) is achieved by adding either mineral acids or sulfated cationites. Apurinization and further splitting of the DNA chain can be performed also by other methods: by the action of mercaptans in the presence of zinc by reaction with formic acid chloride followed by alkali treatment (66,67), in the presence of amines, (68, 69) etc. The N-glycosyl bonds in ribonucleoside monomer units are considerably more stable than those in 7 1 ) and , the hydrolysis of RNA phosphodiester deoxyribonucleosides (70, bonds proceeds faster than does the cleavage of purine base residues (57). Selective removal of the heterocyclic bases may be performed also by other methods that are discussed later.

D. Reactions at the Heterocyclic Base Residues When considering the reactions of the heterocyclic base residues, one must remember that, besides the common bases (A,G, U, C, T), DNA and RNA contain a number of the so-called “minor” bases (729, whose reactivity varies very markedly. Moreover, base reactivity depends not only upon structure, tautomeric form, and charge distribution at the moment of reaction, but also on the interaction with other bases, with metal ions, etc. The effect of these factors upon the rate and specificity of modification are not considered below in detail, but we shall briefly outline the most important of the reactions used for base modification and the prospects of their further investigation and use. The present survey is not concerned

412

N. K. KOCHETKOV AND E. I. BUDOWSKY

with the photochemistry of the nucleic acids because the problem is comidered in a number of comprehensive reviews (see Setlow in Volume 8 of this series; also 73-76). We alsodo not discuss the alkylation of nucleic acids, discussed in an earlier volume of this series by Lawley (76; see also 77). The present review does not cover the interaction of nucleic acids with protons and hydroxyl ions, with metal ions and with dyes, nor the photodynamic decomposition of nucleic bases. The carbonyl groupings in nucleic bases are rather inert, whereas the amino groups react in the usual manner. Of the reactions with the amino groups, the most important are those with nitrous acid and with formaldehyde. Reaction with nitrous acid results in deamination of the heterocyclic bases, so that the cytidine, adenosine, and guanosine residues become uridine, inosine, and xanthosine residues, respectively (78-82) (see article by Shapiro in Volume 8 of this series). The rates of deamination of the bases differ; cytidine is deaminated more slowly than are adenosine and guanosine. The rate of reaction essentially depends on pH and on the participation of the base in complementary interaction. Among side reactions, mention should be made of the formation of interstrand crosslinks on treatment of doublestranded DNA with nitrous acid. Deamination of the purine residues results in alteration of the mode of cleavage of the RNA polymer chain with guanylyl-ribonuclease, as the latter does not split the bonds adjacent to xanthosine residues, but does split those adjacent to inosine (83). The deamination reaction is widely used in functional studies, particularly in the genetic ones (84, 86),and in investigations of DNA structure (86). Two major reactions are important during the modification of polynucleotides with formaldehyde (87-92). These are the formation of methyl01 derivatives and the formation of methylene crosslinks between base amino groups (mostly, between purine nuclei) : R-NH,,

CHnO

It

=

R-NH-GHzOH

RHNi __+

R-NH--CH*-NH-R

nucleic bwe containing an amino group

The first reaction is essentially reversible. Hence, the interpretation of the functional and physical-chemical evidence obtained with formaldehyde is complicated. However, some very important data on the structure and function of the nucleic acids have been obtained in investigations of polynucleotides modified by formaldehyde (93-96). Of other carbonyl-containing reagents, the most interesting is probably glyoxal, a highly selective reagent for the modification of guanosine residue; the reaction involves both the amino group a t position 2 and the amide

T H E CHEMICAL

MODIFICATION R

I

O F NUCLEIC

413

ACIDS R

I

0 0 II II H-C--C-H

HN

HO

0

0

nitrogen at position 1 (97-99). The modification proceeds under mild conditions, and guanosine residues in RNA (99) and DNA (100) can be modified quantitatively. The modified unit is stable in a weakly acidic medium, but elimination of glyoxal, regenerating guanosine residues, occurs in an alkaline medium (98,99). Modification of the guanosine residue with glyoxal protects the adjacent phosphodiester bond from cleavage with T1-ribonuclease, (99, l O l ) , but does not protect it from the attack of guaiiylyl-ribonuclease from Actinomyes u ~ r e o v e r t ~(99). c~~Za~~s Analogous modifications of guanosine nuclei can also occur with other dicarbonylic compounds (97, 98), but the mechanism of the reaction and the structure of the modified base residue has been firmly established only with glyoxal and with a less selective reagent, affecting also cytidine residues, ninhydrin (98, 102). Water-soluble CMEcarbodiimide is widely applied to modify nucleic acids. The reagent most probably adds to the amide nitrogen atom losing a proton in alkaline medium-i.e., N-3 in uridine, N-1 in guanosine and inosine, N-1 and N-3in pseudouridine (21, 22, 105-1 08).

OHB

The reaction proceeds quantitatively under mild conditions (room tem5-24hours). The adducts with uridine, guanosine, perature, pH 7.5-8.5, and inosine residues are stable in neutral and weakly acidic solution, but are readily hydrolyzed under alkaline conditions (pH 10) to yield the original nucleosides (21, 103, 105, 107). One of the two groups added by pseudouridine remains intact after mild alkaline treatment (21, 109). The phosphodiester bonds adjacent to nucleoside residues modified by CMEcarbodiimide are not attacked by some nucleases (21, 22,35,110,111). The reagent has been successfully used in studies of the structure and function

414

N. K. KOCHETKOV AND

E. I. BUDOWSKY

of nucleic acids; the basis of this application have been the above-mentioned stability to nuclceses and the marked dependence of the rate of inodifica111). tion upon the participation of bases in complementary pairing (36, Acrylonitrile reacts with the nucleic bases in the same way as does CME-carbodiimide, but under somewhat more drastic conditions. However, cyslnoethylation does not result in introduction of a positively

charged residue, so that it leads to protection only from ribonucleases, but not from phosphodiesterases (23, 112). Peroxide compounds are widely used for the modification of nucleic acids. Hydrogen peroxide in alkaline medium modifies the uracil, cytosine, and guanine nuclei (113). The chemistry of the reaction and the structures of the products are unknown, but a homolytic mechanism may be anticipated, as inhibitors of radical processes inhibit the modification (114). More specific is the act.ion of peracids. Reaction of perphthalic acid with nucleosides and polynucleotides results in the modification ofcytosine

(ql

R

R I

<:lrJ\o I

perphthalic

acid

NH,

NH*

s

B perphthalic acid I

NH,

NH2

and adenine residues (116,116). The ratje differences are not very great; participation of the base in complementary pairing strongly decreases the The products, cytidine N-3-oxide and adenosine rate of the process (117). N-1-oxide, are stable in usual conditions. In alkaline medium, the N-loxide of adenosine rearranges to isoguanosine (118), the N-3-oxide of cytidine is disrupted (119). The reaction of halogens with nucleic acids has a low specificity. In aqueous solution, the pyrimidine bases add the elements of HOBr at the

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415

5,B-double bond affording 6-hydroxy-5-bromo-5,6-dihydropyrimidines, which are subsequently transformed to 5-bromopyrimidines by the elimination of water. The reaction takes place also with purine base residues (120-1.27). In nonaqueous solvents, the action of halogenating agents results in substitution of the hydrogen atoms at C-8 of purines (168, 129) and at C-5of pyrimidines (13G132). In aqueous solvents, the mechanism of the reaction with purine bases and the structure of the final products are unknown. It may be claimed only that, after halogenation in aqueous solution, the guanine and inosine bases lose their aromatic character. The reaction is alniost insensitive to the participation of bases in compIementary pairing. For the above reasons, its application to the investigation of nucleic acids seems rather limited (but see [84]). A very interesting reaction is the peculiar reduction of the double bond of pyrimidines on ultraviolet irradiat>ionof polynucleotide aqueous solutions in the presence of NaBH4.

R = Sugar residue; R' = H or C H ,

The dihydropyrimidine residue formed is subject to hydroIytic cleavage and to further reduction (133-135). Nucleophilic agents-hydrazine, hydroxylamine, and their derivatives -are also widely applied for the selective modification of pyrimidine bases of the nucleic acids (see article by Singer and Fraenkel-Conrat in this volume). Heating nucleic acids with anhydrous hydrazine results in cleavage of the pyrimidine bases, yielding pyrazoIe derivatives. An analogous reaction takes place with methylhydrazine (136-139), but not with phenylhydrazine. The structure of the modified units is unknown, but most probably ribosylurea (or deoxyribosylurea) units are first formed and subsequently transformed to ribosylhydrazine (deoxyribosylhydrazine) units (140). Such alteration of the substituent at the glycosyl center results in labilization of phosphodiester bond. For this reason, extensive depolymerization of the polynucleotide chain takes place during the course of the reaction to give apyrimidinic RNA or DNA. A slow degradation of the adenine residue Hydrazine in the presence of also takes place under this condition (141). oxygen and copper salts reduces pyrimidine nuclei to 5,B-dihydropyrimi-

416

N. K. ICOCHETKOV

R

I H,N-NH,

Ox?

=I

H,N -C -NHR II 0

H ,

0

R"

AND E. I. BUDOWSKY

H,N -N&

-

%N-NHR

/R"

$7 \N/’\OH H

R = Sugarresidue; R’ = H o r CbOH; R" = Hor CH,

dine derivatives (142). Treatment of apyrimidinic DNA with alkali results in complete cleavage at the modified units while polypurine clusters remain intact; the modification is used for studying the primary structure of DNA (16, 18) (see article by Chargaff in Volume 8 of this series).

DNA

Apyrimidinic DNA

Purine clusters

The degradation of a polymer chain may be considerably reduced if one performs apyrimidination by aqueous hydrazine a t a low temperature. The rate of the cleavage of the pyrimidine bases by anhydrous hydraaine and by an alkaline aqueous solution of hydrazine falls in the sequence U > C > T; however, the rate differences are not enough to make the reaction selective. Under mild conditions in neutral or slight acidic solutions, the amino group of a cytosine nucleus is substituted by hydrazine (144-146) or 2,4dinitrophenylhydrazine moieties (147). Acylhydrazines (semicarbazide, Girard's reagent, and so on) (148-1 62) at neutral or acidic pH are very selective reagents, replacing the amino group of cytosine by an acylhydrazine grouping. The reaction proceeds

THE CHEMICAL

MODIFICATION

O F NUCLEIC

R

R I

I

’ y ?

417

ACIDS

I&N-NHR’_

’y? N\

N\ Nfb

R =

HN-NHR’ Sugar residue

R’ = H"%’; C,H,(NO,), ;’’’1C

- /oc148-i501 "N&

'

under mild conditions and can be used for studying the structure and function of nucleic acids. As mentioned above, the absence of data on the mechanism and kinetics of the major and side reactions taking place during the action of modifying agents on nucleic acids and also on the structure and properties of modified units prohibits wide application and common recognition of the chemical modification methods for investigations concerned with the structure and function of nucIeic acids. We have attempted to overcome these weak points by a detailed investigation of one of the most important nucleophilic agents used for the modification of nucleic acids-h ydroxylamine. Below we present the results obtained in the course of this investigation and some aspects of the application of the data obtained to the study of structure and function of nucleic acids.

V. Modification of the Uracil Nucleus with Hydroxylamine In 1961it was demonstrated that hydroxylamine modifies nucleic acid components; it appeared that the cytosine residue reacts at the highest rate at pH 6,and the uracil residue at pH 10; and that the thymine residue is inert over a pH interval from 4 to 11 (153-157).

A. Reaction of Uridine withHydroxylamine The curve of pH vs. rate of reaction between uridine and hydroxylamine shows a maximum in the alkaline region (see Fig. 1);this may be interpreted as evidence for interaction of a neutral molecule and an anion. The same curve for N-3-methyluridine (see Fig. 2 ) ,which cannot form an anion, exhibits no maximum. It may be concluded from these data that the neutral form of uridine and the very nucleophilic NH20- anion are the The reaction of a uracil residue with hydroxylainteracting species (158).

418

N. K . KOCHETKOV

AND

E. I. BUDOWSKY

A8

C

PH

FIG.1. pH dependence of the rate of reaction of 6 M aqueous hydroxylamine with Up(U)and Cp(C).The reaction rate is measured as percentage decrease of extinction in 10minutes during the initial reaction phase (166).

mine is many staged, but at pH 9 and at high concentrations of reagent it may be regarded exactly enough as a two-stage one (159). At the first stage, the uracil nucleus is degraded to ribosylurea (11) and isoxazolidone (111).The former is subsequently transformed into ribosylhydroxylamine (IV). Hence, treatment of a polynucleotide with hydroxylamine could result in the formation of units of both the XI and the IV type. In order to use the reaction for studying structure and function of polynucleotides, it was thus necessary to find conditions affording RNA of not only known extent of modification, but also with a known ratio of the two

PH

FIG.2. pH dependence of the rateconstants of the reaction of N-3-methyluridme 5’-phosphate with hydroxylamine (3.0 M HzNOH, 37’) (168).

419

THE CHEMICAL MODIFICATION O F NUCLEIC ACIDS

R

HONHR

H,N-C-NHR-

types of modified units. Moreover, it was necessary to eliminate, or to keep to a known minimum, side reactions, i.e., modification of residues other than uracil (especially of cytosine residues, see below) and internucleotide bond cleavage. The modifkation of uracil nuclei in polynucleotides has been studied using preparations of highly polymeric RNA devoid of a stable secondary structure. Optimum pH and concentration of hydroxylamhe (for minimal modification ofcytosine nuclei) were chosen after spectral analysis of the reaction with nucleosides. The optimum ratio of the modikation of uracil to that of cytosine is obtained at pH 10. Increase of the concentration of hydroxylamine above 10M results in a considerable increase of the relative rate of cytosine nuclei modification (159) (cf. 160).

I

I

1

I

9

L

0.5

1.0

1.5

2.0

2.5

-Log

I

3.0

I

-

3.5

K

FIG. 3.Arrheuius plot of the rate constants of the reactions ofhydroxylamine (10 M ) with RNA base residues a t pH 10 (169). Conversion of uridine residues into ribosylurea ki

ka

units (I --+ 11) and of ribosylurea units into ribosylhydroxylamino residues (I1--t IV) (leftrhand line). Cytidine residue modifications (K3)(right-hand line).

420

N. K. KOCHETKOV AND E. I. BUDOWSKY

7

H

, mod C

hr

2oa

20

40

60

80

100

120

hr

FIG.4. Calculated amounts of uridine, ribosylurea, ribosylhydroxylamine, and modified cytidine residues in the course of the reaction of hydroxylamine (lOM,pH 10.0) with RNA a t 10"and 20" (the h t three values: percentage of the starting amount of uridine; the fourth value: percentage of the initial amount of cytidine) (169). The reaction with RNA could be studied after elaboration of effective methods to follow its stages separately. In the above scheme, it is seen that the formation of the ribosylurea (11)is followed by accumulation of isoxatolidone (111),so that the concentration of the latter may be taken as a measure of the extent of reaction I -+ 11. The IV formation leads to formation of hydroxylamino residues in place of uracil nuclei. The former may be quantitatively determined after mild acid hydrolysis of a modified polynucleotide. Modification of cytosine nuclei results in accumulation of two moles of bound hydroxylamine per mole of modified cytidine residue. Mathematical treatment of the kinetics of bound hydroxylamine and isoxazolone accumulation during RNA modification enabled determination of the rate constants of the ribosylurea (K1) and ribosylhydroxylamine (Kz) formation, and also the rate constant of cytidine modification (113)over the temperature interval ct40" (Fig. 3).

421

T H E CHEMICAL MODIFICATION O F NUCLETC AC1I)S

011the basis of tlicse rate coustaiits, it is iiowpossible to choose conditions for obtaining a predicted extent of modification and a predicted ratio of modified units (Pig. 4). In Fig. 3,which presents the temperature dependence of the rate constants, it is seen that the highest selectivity is observed at low temperatures. For example, quantitative transformation of uridine units into ribosylhydroxylamino ones takes place with a very small ( 2 4 % ) extent of cytosine modification. I n the presence of methanol and at high ionic strength, the rate of the transformation of ribosylurea units into ribosylhydroxylamino ones is extremely small so that RNA may be obtained with uridine quantitatively transformed into ribosylurea residues.

B. The Preparation and Some Properties of Deuridylic RNA’s The above data served as a basis for the elaboration of methods of preparation of deuridylic ribonucleic acids, i.e., ribonucleic acids devoid of their uracil residues. Treatment of ribosomal RNA with 10 M hydroxylamine in an aqueous solution containing methanol and KCI afforded RNA with ribosylurea units in place of all the uridine residues, the so-called “deuridylic RNA I.” More prolonged treatment, or treatment with 10 M hydroxylamine in the absence of salts afforded the so-called “deuri-

0.2

0

N W

a 1 .o

0.1

0.8 0.6

0.4

C

I

J

0.2

I

I I

80

1

%

z

s

1

120

160

ml

FIG. 5.Chromatographic patterns of poly U (solid line) and poly (ribose phosphate) obtained by poly U hydroxylaminolysis and subsequent treatment ofthe product with pancreatic ribonuclease (dotted line) (pH 7.2,37 ,24 hours, enzyme: substrate 1:20) (161).

422

N. K. KOCHETKOV AND E. I. BUDOWSKY

dylic RNA 11” with ribosylhydroxylamirio residues substituting for all the uridine residues. Deuridylic acids are of considerable interest for both structural and functional studies. Transformation of uridine residues to ribosylurea units does not affect the stability of the neighboring phosphodiester bonds. The degradation of uracil nuclei results in stability of the corresponding internucleotide bonds to pancreatic ribonuclease, as revealed by the stability to the enzyme of poly(ribosy1urea phosphate), obtained by treatment o polyuridylic acid with hydroxylamine (Fig. 5). Terminal analysis of the oligonucleotides obtained by hydrolysis of deuridylic RNA-I with pancreatic ribonuclease revealed that the cleavage takes place only at cytidylate units. Treatment of the oligonucleotide mixture with phosphatase and subsequent alkaline hydrolysis gave cytidine as the only component of the nucleoside fraction, whereas ribosylurea residues were present only as ribosylurea phosphate. Thus the high stability of the phosphodiester bonds adjacent to ribosylurea appeared not less than that between usual nucleotides (Scheme 1). RNA

Deurldylic RNA-I

c-c

oligo nucleotldes

End analysis

SCHEME 1

Such behavior also suggested that the ribosylurea unit exists in the form of N-ribofuranosylua with a stable N-glycosyl bond. Hence, the modification of uracil nuclei with hydroxylamine and subsequent hydrolysis with pancreatic ribonuclease enables selective cleavage of the polynucleotide ciain at cytidylic acid residues. Another interesting aspect of the reaction is also the fact that modification of the cytidine nucleus with hydroxylamine (and also with 0-methylhydroxylamine and even with

T H E CHEMICAL

MODIFICATION

OF NUCLEIC

423

ACID6

0-benzylhydroxylamine) does not affect the susceptibility of the adjacent 165). phosphodiester bond to cleavage by pancreatic ribonuclease (164, On the other hand, substitution of a uracil nucleus by hydroxylamino residue (formation of deuridylic RNA-11) results in labilization of the neighboring phosphodiester linkage (166).Under conditions affording deuridylic RNA-11, the polynucleotide chain is cleaved only at the uridine units converted to ribosylhydroxylamino residues. Practically no nonspecific degradation of the polynucleotide chain takes place. For this reason, deuridylic RNA-I1 may not be used for specific splitting of the polynucleotide chain at cytidine units with pancreatic ribonuclease, but quantitative chemical degradation of deuridylic RNA-I1 at uridine units may be achieved. Removal of the heterocyclic nuclei and subsequent treatment under appropriate conditions is widely applied to cleave apurinic and apyrimidinic DNA. The reaction is based upon p-elimination of the phosphate group in the ribose residue with a free glycosyl center. The same principle of @-eliminationis used for the end-group analysis of RNA (see above). To facilitate the selective splitting of deuridylic RNA, the hydroxylamino residue must be removed to give a free glycasyl at the modified uridine residue. For this purpose, deuridylic RNA-I11 was obtained from deuridylic RNA-I1 by mild acidic hydrolysis of the labile N-glycosyl bond in rybosylIn the resulting ribose phosphate residues, hydroxylamino residues (167). there is a potential carbonyl grouping in the @-positionto the 3'-phosphate; this considerably weakens the phosphodiester bond. During alkaline hydrolysis of deuridylic RNA-111, nucleoside 2'(3'),5'-diphosphates are formed along with nucleoside 2'(3')-monophosphates in an amount equal Hence, @-elimination to the content of uridine in the starting RNA (Fig, 6). of the phosphates of the modified units proceeds at a high rate in alkaline medium (168). The method may be used particularly for evaluating the relative frequencies of the occurrence of pairs . . . UpNp . . , as the N units afford nucleoside 2'(3'),5'-diphosphates. However, the conditions are too drastic to cleave the polyribonucleotide chain selectively only at modified uridine units. As it was known that the /3-elimination of phosphodiester bond proceeds under mild conditions in the presence of amines, their effect has been studied with ribose 3’phosphate as a model compound. It appeared in fact that amines considerably facilitate the @-elimination.The reaction proceeds most readily in the presence of aromatic amines possessing electron-donor substituents. Subsequent studies with polynucleotides revealed that deuridylic RNA-111 is completely cleaved into blocks in the presence of p-anisidine in 4-5 hours at pH 5 and 30".Under these conditions, the original RNA remains practically intact (169).Hence, cleavage of the uracil nuclei by hydroxyla-

.

I

I

HO-P=O

HO-y=O

b

OH

I

I

0

II

(HO),-P-O

0 (D

t V,I

FIQ. 6. Chromatographic pattern of the alkaline hydrolyzate of deuridylic tRNA-111 (0.3M KOH, 37", 24 hours), separated according to Todinsou on DEAE-Sephaclex A-25; broken line: alkaline hydrolyzate of the starting tRNA (168).

THE CHEMICAL

MODIFICATION

OF NUCLEIC

425

ACIDS

mine, removal of the hydroxylamino residue, and treatment with amine leads finally to selective cleavage of the polynucleotide chain at uridine residues. (See Scheme 2.) RNA

Deurldylic RNA-I

Deurldylic RNA-II

Deurldylic RNA-III

P-ellminotion

CHO

A A A

SCHEME 2

C. The Effect of the Secondary Structure of Polynucleotides upon Hydroxylaminolysis

The reactivity of the uracil nucleus considerably decreases in polymers compared with monomers. For example, the rate of uracil hydroxylaminolysis in commercial RNA is about half that in uridine (e.g., at 37",K, = 0.445 and 1.05 h-l, respectively). The rate constant values of uracil modification in the 10"-40" interval fit well a straight line on an Arrhenius plot (Fig. 3),suggesting a noncooperative character of the forces, reducing the reactivity of uracil nuclei in polunucleotides, the effect is probably the consequence of base-stacking interactions (159). Still greater is the effect of the complementation interactions, leading to formation of stable double-stranded structures. The rate of tRNA hydroxylaminolysis at low temperatures considerably decreases after modification of a part of the uridine residues (5'3) (Fig. 7). Obviously, the stable secondary structure prevents the uridine residues involved in complementary pairs from reacting. In favor of the assumption is the fact that the Tm of tRNA does not change after modification of 50% of the uridine residues (at 10"); the decrease of the hyperchromic effect after such treatment is small enough to be explained by a decrease of the molar extinction coefficient of RNA resulting from degradation of the uracil residues rat>herthan by :in at t,anli. 011the double-st,rancledregions.

426

N. K. KOCHETKOV AND E. I. BUDOWSKY

20

40

60

80

100

120

140

hr

FIG.7.Degradation of the uracil residues in tRNA with hydroxylarnine. Curve a : 7 M NHzOH, pH 10, 10";6 :7 M NH20H, pH 10,37"; c : 7 M NH20H, pH 10,10" in the presence of 7 M urea. Broken line: change of hyperchromism in the reaction a t 10" without urea (33).

When the secondary structure of the tRNA is destroyed by the addition of urea, or by increasing the temperature of the reaction mixture, all the uridines become subject to modification (Fig. 7).Direct support to the suggested effect of complementation interactions in tRNA upon the reactivity of its uridine residues has been obtained in the experiments with complexes of poly U and poly A. As seen in Fig. 8, all the uridine residues of poly U in excess of a 1:l complex are readily modified, whereas the remaining ones, forming the poly A poly U complex, react many t h e s more slowly ( I70). Hence, the rate of the modification of uracil nuclei by hydroxylamine is affected by both the stacking and the complementation interactions. The rates of the modification of uridine residues in poly U and in poly U . poly A differ by a factor of 5 to 6;still greater is the difference in heteropolymers, particularly in tRNA, containing more stable G C pairs. The approach may serve for detection and for analysis of the composition, size, and localization of the sites with weakened (or absent) secondary structure, and for the selective modification of RNA a t 'ilooped" sequences, which is of interest for functional studies, or for more selective cleavage of polynucleotide chains a t either loop cytidine or loop uridine residues. Finally, the rate of uracil residue modification is affected by the presence of ferric ions in the reaction mixture. An increase in the rate of reaction occurs when 3 to ti metal ions are prescnt per 100 nucleotide residues, but only during modification of tRNA rather than with uridylic or polyuridylic acid. Copper ion, oxygen of air, and hydrogen peroxide do not

-

THE

CHEMICAL

MODIFICATION

01%'

20

OF NUCLEIC

"

-

40

I

’

427

ACIDS

60

00 hr

FIG.8. Modification of theuracil nuclei of poly U in mixtures of poly U and polyA; ordinate: themolar ratio ofuridme t o adenosine residues (170).

change the rate of RNA uracil modification. The effect of ferric ions is eliminated by addition of Versene to the reaction mixture (171). The outstanding opportunities of hydroxylamine as a modifying agent attracted attention to the reactivity of its derivatives. The most interesting type of reagents of the group appeared to be the 0-alkylhydroxylamines, selectively modifying cytidine (160,164,165,17,2) (this reaction is discussed in detail in the following paragraph). N-Acylhydroxylamines do not, react with the bases in nuclek acids. However, hydroxylamine, formed during hydrolysis of hydroxamie acids, reacts with uridine residues in the usual way. The first stage of the processtransformation of uridine into ribosylurea residue-is considerably facilitated by the presence of hydroxamic acids (17S,174).

VI.

Modification of the Cytosine Nucleus with Hydroxylamine

Modification of the cytosine residues by nucleophilic reagents (hydroxylamine, hydrazine, semicarbazide, etc.) is widely applied to the

428

N. K. KOCHETICOV AND E. I. BUDOWSKY

study of structure and furictioii of tlic Iiucleic acids. The reagent more frequently applied to functiorial studies is hydroxylamirie (175-177). Data on the mechanism arid lcirietics of the reaction of cytosine residues with hydroxylamine and knowledge of the structure and properties of the intermediate and final reaction products are of basic importance for correct and complete understanding of the results of the above studies.

A. Reaction of the Cytosine Nucleus with Hydroxylamine The studies by Brown (163), Shuster (154))and other workers revealed, that two products may be isolated from the reaction mixtures of cytidine with hydroxylamine (see article by Phillips and Brown in Volume 7 of this series). These are N-4-hydroxycytidine (VI) and 6-hydroxyamino-5,6 dihydro-N-4-hydroxycytidine (VIII) ;the latter compound is presumably formed via the intermediate 6-hydroxyamino-5,6-dihydro-N-4-hydroxycytidine (VII). R OY? I

N$

"YN

N\

>r $ R

H

O Y N

, N

R

O Y N N,

NH,

HNOH

NH,

HNOH

(V)

(VI)

(VH)

(MI)

Comparison of the structure of the reaction products of uridine and cytidine with hydroxylamine revealed that both of the two functional groupings of hydroxylamine are involved in the reaction with uridine, whereas only the amino group of hydroxylamine is involved in the reaction with cytidine. It could be thus anticipated, that substitution of the hydrogen atom in the hydroxy group of hydroxylamine makes impossible the reaction with uridine and does not essentially change the reactivity toward cytidine. Experiment confirmed these considerations. It appeared that 0-methyl- and 0-benzylhydroxylamine do not attack the uracil nucleus, The reaction hut readily modify the cytosine nucleus (160, 164, 165,17.2). proceeds according to a mechanism similar to that with hydroxylamine and In view of this fact, we results in products of analogous structure (178). shall discuss the mechanism of the reactions of cytidine with hydroxylamine and 0-alkylhydroxylamine in parallel. The most convenient approach to studies of the mechanism and kinetics of the reaction of the cytosine nucleus with hydroxylamine appeared to be via the spectral method, eliminating the difficulties and artifacts typical of the analytical procedures based on separation and quantitative isolation of the reaction products. To this end, the spectral

THE CHEMICAL

MODIFICATION

OF NUCLEIC

ACIDS

429

characteristics of VI and VIII have been investigated (178). Attempted isolations of compound VII failed because of its lability, but it could be assumed that the compound does not absorb at X > 240mp to any practical It was very extent, like dihydrocytidine or 6-hydro~y-5~6-dihydrocytidine. important for these studies that, of all t.he reaction components, only compound VIII absorbs at 300 mp, so that its concentration may be followed by direct spectrophotometry of the reaction mixture. proposed a scheme that assumed a In 1965,Brown and Phillips (179) sequence of reactions V -+VII+ VIII-+ VI. However, the scheme seemed to contradict tlie modern concept of the cytosine reactivity. Cytosine contains two centers of nucleophilic reactivity, C-4 and the 5,6 double bond. For this reason, it seemed more probable, that the correct schemeis in fact the following sequenceof parallel and consecutive reactions:

Obviously, the relative rates of reactions at each of the stages should depend on both the nature of nucleophilic agent and the relative electrophilic character of the two centers in the cytosine nucleus, determined by the nature and the position of substituents. Spectral studies of the conversion of VI

K-7 F? VIII Ki

revealed that under conditions usually applied to nucleic acid modification, these reactions proceed much slower than that of cytosine residue modification (178) (cf. 186). Hence, the scheme of the reactions of hydroxylamine and O-methylhydroxylamine with cytidine could be described exactly enough asFfollows:

The direct conversion of V into VI was proved in our laboratory by the facts that the compound V I accumulates in the reaction mixture without any lag period and that the rate Constant, calculated on the assumption of direct conversion of V into VI, remiLined constant during the course of reaction (186). Analogous evidence has been obtained simultaneously and independently by Lawley (181). Analysis of the system of differential

430

N. K. KOCHETKOV AND E. I. BUDOWSKY

equations corresponding to the last scheme revealed that the final products ratio must be described by relationship (Eq.1) (186),and thus must depend on the concentration of modifying reagent.

I I the concentrations of the corresponding compounds when CVIand C ~ I are Cv = 0 ; CH~NORis the concentration of reagent in the reaction mixture; K,,etc., are the second-order reaction rate constants (excluding L). The experimental data on the reaction of cytidine with hydroxylamine and with 0-methylhydroxylamine showed, that this relationship is in fact the case (Fig. 9) and thus confirmed the validity of the proposed scheme of

4

L

3A

0.6

=E

-

--

2-

e

>

0 k '.

43"

40"

>

>

V

u

I

1

2

'

3

4

5

t

I

I

L

1

1

0.1 0.2 0.3 0.4 0.5 0.6 0.7

-

-

NH20H

FIG. 9.Dependence of the ratio of the reaction products of cytidine with (A)hydroxylrtmine (pH 6.0) and (B)0-methylhydroxylamine (pH 4.9) on the reciprocal of the concentration ofthe modifying reagent (180).

reaction. After direct determination of CvI/G'vm and RBI the values of K 4 and K4/K6 could be readily calculated from Eq.(1). The pH of the reaction mixture considerably affects the rate of the reaction of cytosine with hydroxylamine. The dependence of the rate of the modification on pH is described by curves with maxima at pH 6 for hydroxylamine and at pH 5 for 0-methylhydroxylamine. The data suggest that the protonated cytosine residue reacts with a neutral hydroxyIamine molecule. Although the above experimental pH optima do not coincide with the calculated values (pH 5.1and 4.5, respectively), the discrepancy finds a reasonable explanation. First, the rates of the conversions V -+ VI and V + VII must depend on pH in an andogous manner, whereas the rates of reaction VII+ V must increase con-

431

THE CHEMICAL MODIFICATION O F NUCLEIC ACIDS

siderably with decreasing pH (cf. 74,187). Second, the optimum pH of reaction VII + VIII must be higher than that of reaction V + VI as the pK of the compound VII is higher than that of cytidine (cf. 185, 184). Hence, the optimum pH of the overall reaction, measured by the decrease of cytidine concentration, must be somewhat higher than that calculated from the known pK values of cytidine and hydroxylamine. The conclusion is in accord with the experimental data. An important prediction following II by increasing pH, from the data is the increase of the ratio C ~ I / C ~ Icaused also confirmed by experiment (Fig. 10).

0

4.5

5.5

6.5

PH FIG. 10. The dependence on pH of the reaction product ratio of cytidme with hydroxylamine (curve A, 30 ,1M NH20H)and with O-methylhydroxylamine (curve B, 37",2.5M NHpOCs) (180).

The rates of the stages ofthe reaction depend also on the ionic strength of the reaction mixture. Change of the ionic strength from 0.1 to 4.5M results in a change of the CVI/CVIHratio for the reaction with O-methylhydroxylamine by a factor of 1.5. A considerable isotopic effect is also the casesubstitution of D20for HzO results in increase of the CVI/CVIIIratio from 0.55 to 1.05. The values and ratio ofthe rates of reactions at C-4 and C-5, C-6, and, respectively, the CVI/CVIII ratio depend essentially on the nature and position of the cytosine substituents. For example, Janion and Shugar demonstrated that the presence of a methyl group a t C-5 or C-6 and also the presence of a hydroxymethyl group at C-5 ofthe cytosine inhibits the reaction V + VII, so that the reaction proceeds as a simple conversion ofV into VI (186, 186).

432

N. K. KOCHETKOV AND E. I. BUDOWSKY

The rate of this reaction is rather low. An analogous scheme may be applied to explain the reactivity of 4-akoxy- and 4-thio-2-ketopyrimidine with hydroxylamine. I n this case, the reaction at C-4 is much faster than that at C-5, C-6,and faster than that at C-4 of cytosine and of 5-or &methylcytosine (186). On the other hand, the substitution of methyl groups for hydrogen atoms of the exocyclic amino group inhibits the reaction at C-4 and increases the CVI/CVIII ratio (187). The effect of the substituents upon the rates of reactions at C-4 and (3-5, C-6 correlates with the electron localization energies a t these centers, calculated by Hiickel’s MO method (188). (See article by Pullman and Pullman in this volume.) The obtained parameters of the reaction of cytosines with hydroxylamine and 0-methylhydroxylamine enables a choice of conditions to obtain a given ratio of modified cytidine residues of the types V I and VIII over a wide range. For example, it appeared possible to convert the cytidines in poly C quantitatively to units of the type VIII orVI (189) (cf. 252). An inverse problem may be also solved-knowing the conditions of reaction and the structure of the starting pyrimidine nucleus, one may predict the extent of modification and the amount and ratio of the various types of modified units. At the same time, it is to be remembered that the action of hydroxylamine upon DNA involves also a number of side reactions; the most important of these are the following: (i) Modification of the adenines to N-6-hydroxyadenine. The reaction consists of the substitution on the amino group of a hydroxylamino (or 0-methylhydroxylamino) residue. The rate of the reaction is about the same as that of the substitution of the amino group in cytidine (190). (5)At low concentrations of hydroxylamine, the latter is decomposed to afford peroxides. The peroxides may modify not only cytidine, but also thymidine and adenosine (191, 192). The formation of peroxides is inhibited by mercapto compounds, pyrophosphate, Versene, etc. (178,193), together with the corresponding side reactions. Allthe side reactions mentioned are slower than those modifying the cytosine nucleus. However, they must be taken into account, or, if possible, eliminated in functional studies.

B. Application of the Reaction of Hydroxylamine with Cytosine Residues to Functional Stud,ies

The above data are of essential importance for the correct interpretation of evidence on functional alterations in nucleic acids caused by the action of hydroxylamine or its derivatives.

THE CHEMICAL MODIFICATION O F NUCLEIC ACIDS

433

Hydroxylamiiie and 0-methylhydroxylamine are widely applied to genetic studies because of their extremely high mutagenicity and the small extent of inactivation of the genetic material. Both the genetic studies (19.4) and the studies of the effect of modification by hydroxylamine and 0-methylhydroxylamine on the template activity of poly C in the RNA polymerase system (195-197) revealed that the action of the reagent results in transitions of the C + U (C -+ T) type along with inactivation. As only two types of stable modified units are formed during the modification of cytosine nuclei, it seemed reasonable to expect that one of them is responsible for the transition, and the other for the inactivation. Most probably, the t#ransitionsare due to units of type VI. The following evidence is in favor of this conclusion. The transitions are observed with T-even phages (198, 199), containing 5-hydroxymetliylcytosine instead of cytosine. The former may only form units of type VI, as demonstrated by Yanion and Shugar (185, 186). The inactivating effect of hydroxylamine upon transforming DNA is higher than that upon DNA of T-even phages by a factor of 104 (dOO), in accord with the fact that units of types VI and VIII may be formed in transforming DNA, whereas only units of type VI are formed in DNA of the T-even phages. Analysis of the data of Phillips et al. (195) leads to the conclusion that the units leading to transitions are formed by a single-stage process, whereas the inactivating ones are formed in a two-stage reaction. Taking into account the instability of units of type VII, only units of types VI and VIII need be considered; of them, units VI are formed in a single-stage reaction, and units VIII in a two-stage reaction. Finally, the increase of the inactivating action with increasing pH (194, 201) is in accord with the decrease of the CvI/CvIII ratio at acidic pH values. Allthe above evidence is in favor of the general mechanism involving the CVIICVIII ratio, which can be deduced from the reaction conditions and the nature of the nucleus subject to modification. The availability of the data on side reactions also makes it possible to attempt a rational treatment of the mechanisms of template synthesis, particularly of the mechanism of chemical mutagenesis caused by hydroxylamine.

REFERENCES 1.C. T.Yu and P. Zamecnik, Bzochirn. Biophys. Acta46, 148 (19ti0). 2. H. Weit and P. Gilham, J .Am. Chem.Soc. 89, 5473 (1967). 3. B. Singer and H. Fraenkel-Conrat, Biochim. Bzophp. Acta72,534 (1963). 4. A. Steinschneider and H. Fraenkel-Conrat, Biochemzstry 6, 2735 (1966). 6.E. Wagner and V. M. Ingram, Biochmistry 6,3019(1966). 6. A. Annqtrnng, IT. Tlntlopinn, V. hr. Ttlgritm, :indE. W:igner, 13roc hrmzslrr/ 6, 3027 (100ti).

434

N . K. KOCHETKOV

AND E. I. BUDOWSKY

7 .G. Brownlee, F. Sanger, and B.Barrel, J. Mol.Biol. 34, 379 (1968). 8.R. W. Holley, J. Apgar, G. A. Everett, J.T. Madison, M. Marquisee, S. H. Merrill, J.R. Penswick, and A. Zamir, Science 147, 1462 (1965). 9.H. G. Zachau, D. Dutting, and H. Feldman, Angew.Chem.78, 392 (1966).

10. A. A.Bayev, T. V. Venkstern, A. D.Mirsabekov, A. I. Krutilina, L. Li, andV. D. Axelrod, Mol.Biol. 1, 7541 (1967). 11.J. Madison, G. Everett, and H. Kung, Science 163, 531 (1966). 12.V. RajBhandary, S. Chang, A. Stuart, R. Faulkner, R. M. Hoskinson, and H. G. Khorana, Proc. Natl. Acad.Sci.U.S. 67, 751 (1967). 18.H. M. Goodman, J. Abelson, A. Landy, S. Brenner, and J. D. Smith, Nature217, 1019 (1968).

14.R.Rudner, H. Shapiro, and E. Chargaff, Biochim. Biophys. Acta129, 85 (1966). 15.H. S. Shapiro and E. Chargaff, Biochemistry 6, 3012 (1966). 16.V. Habermann, Collection Czech. Chem.Commun. 26, 3147 (1961); 28, 510 (1963). 17.G. B. Petersen and J. Reeves, Biochim. Biophys. Acta129,438 (1966). 18.A. Masin and B. Vanjushin, Biokhimiya 33, 377 (1967). 19.N. Kochetkov, E. Budowsky, M. Turchinsky, and V. Demushkin, Dokl.Acad. Nauk SSSR162, 1005 (1963). 20.W. Seifert and W. Zillig, 2.Physiol. Chem.348, 1017 (1967). 21.R.Naylor, W. Nancy, and P. Gilham, J. Am. Chem.SOC. 87,4209 (1965). 22.0. Ivanova, D. Knorre, and E.Malygin, MoZ. B i d .1, 335 (1967). 2.9. R. Chambers, Biochemwtry 4, 219 (1965). 24.P. R.Witfeld and H. Witzel, Biochim. Biophys. Acta72, 338 (1963). 26. N. Kochetkov, E. Budowsky, N. Broude, and L. Klebanova, Biochim. Biophys.

Acta134,492 (1967).

W.M. Beer and E. N. Moudrianakis, PTOC. Natl. Acad.Sci.U . S. 48,409 (1967). 27.M. Beer and C. Zobel, J. MoZ. Biol. 3,717 (1961). 28.B.Ulanov and A. Malyshev, Biophysics (USSR) 12, 325 (1967). 29. B. Ulanov, A. Serebrjany, and R. Kostjanovsky, Dokl. Acad.Nauk SSSR176,474 (1967). SO. P. J.Highton and M. Beer, J. Mol.B i d .7 ,70 (1963). S1. M. Yoshida and T. Ukita, Biochim. Biophys. Acta167, 466 (1968). 32.T. Morosova and R. Salganik, Biokhimiya 29, 17 (1964). SS. N. Kochetkov, E. Budowsky, and

V. Demushkin, Dokl.Akad.Nauk SSSR 168,

102 (1966).

S4. G. Augusti-Tocco and G. L.Brown, Nature206,683 (1965). 36.V. Drewitch, D. Knorre, E. Malygin, and R. Salganik, Mol.Biol. 1, 249 (1967). 36. H. Seidel and F. Cramer, Biochim. Biophys. Acta108, 367 (1965). 37.A. Stuart and H. G. Khorana, J .Biol. Chem.239, 3885 (1964). 38.D. Knorre, N. Pustoshilova, N. Teplova, and G. Shamovsky, Biokhimiya 30, 1218 (1965).

39. D. Knorre, A. Malysheva, N. Pustoshilova, A. Sevastyanov, and G. Shamovsky, Biokhimiya 31, 1181 (1966). 40.D. Knorre and G. Shamovsky, Biochim. Biophys. Acta142, 555 (1967). 41.D.Knorre and G. Shamovsky, Mol.B i d .2,33 (1968). 42.D. Knorre, N. Pustoshilova, and A. Sevastyanov, Biokhimiya 33, 56 (1968). 4s.P. Whitfeld, Biochem. J .68,390 (1954). 44.D. M. Brown, M. Fried, and A. Todd, J. Chem.SOC.p. 2206 (1955). 45.H. Ney and L. Heppel, J. Hiol. Chem.239, 1927 (1964). 46.M. Ogur a n d J. Small, J. Biol. C h m . 236, P. C .ti0 (19fiO).

THE CHEMICAL

MODIFICATION

OF NUCLEIC

ACIDS

435

47.J.X. Khym, Biochemistry 2,344 (1963). 48.D. Livingston, Biochim. Biophys. Acta87, 538 (1964). 49.M. Tomass, Y. Sanno, and It. Chambers, Biochemistry 4, 1710(1965). 60. G. Moss, C. B. Reese, K.Sehofield, R. Shapiro, and A.Todd, J. Chem.Soc.p.1149 (1963). 61. K. Pfitsner and J. Moffat, J .Am. Chem.Soc.87, 5661(1965). 52.U. RajBhandary, R. Young, and H. G. Khorana, J .B i d .Chem.239, 3875(1964). 63. R. Ralph, R. Young, and H. G. Khorana, J. Am. Chem.SOC. 86,2002(1963). 64.J. Cox and 0. Ramsay, Chem.Rev.64, 317 (1964). 55.K.-H. Scheit and A. Holy, Biochim. Biophys. Acta149, 344 (1967). 56.A. Holy and K.-H. Scheit, Biochim. Biophys. Acta138, 230 (1967). 67.R. Markham and J. D. Smith, Biochem. J .49, 401(1951). 68. H. Fritz and B. Rottger, Z.Naturforsch. 188, 124(1963). 69.E. Budowsky and L. Klebanova, Vopr.Med.Khim.13,299 (1967). 60. B. Lane and G .Butler, Biochim. Biophys. Acta33,281 (1959). 61. B. Lane and G .Butler, Can.J .Biochem. PhysioZ. 37, 1329(1959). 6R. K. Dimroth and H. Witzel, Ann.Chem.620, 109(1959). 63.W. Farkas, Biochim. Biophys. Acta166,401 (1968). 64.C.Tamm, M. Hodes, and E. Chargaff, J. Biol. Chem.196,49 (1952). 65. C. T a m and E. Chargaff, J .Biol. Chem.203, 689 (1953). 66.A. Jones and D. Letham, J .Chem.SOC.p. 2573(1956). 67.A. Jones, D. Letham, and M. Stacey, J .Chem.SOC.pp. 2579,2584(1956). 68.K. Burton and G. B. Petersen, Biochem. J .76, 17 (1960). 69.K. Burton, Biochem. J .77, 547 (1960). 70.F.Micheel and A. Heesing, Chem.Ber.94, 1814(1961). 71.H. Venner, Z.Physiol. Chem.339, 14 (1964). 72.T. V. Venkstern, Usp.Biol. X h i m .6,3 (1964). 73.D. Shugar, in“The Nucleic Acids” (E. Chargaff and J. Davidson, eds.), Vol. 111, p.39.Academic Press, New York, 1960. 74.J. McLaren and D. Shugar, “Photochemistry of Proteins and Nucleic Acids,” Macmillan (Pergamon), New York, 1964. 7 5. A. Wacker, This series 1, 369 (1963). 76.P. D. Lawley, This series 6,89 (1966). 77.G.Wheeler, CancerRes.22, 651 (1962). 78.K. Lohman, Biochem. Z.264, 381 (1932). 79.H. Schuster, Z.Naturforsch. 16b, 298 (1960). 80.R.Shapiro, J .Am. Chem.Soc. 86,2948(1964). 81.I.Carbon, Biochim. Biophys. Actu96,550 (1965). 82.T.Kotaka and R. L. Baldwin, J. Mol.Biol. 9, 323 (1964). 83. T.Ushida and F. Egami, J .Bioclrem. (Tokyo) 67, 742(1965). 84.J. Nelson, S.Ristov, and R. Holley, Biochim. Biophys. Acta149, 590(1967). 85.H. G.Wittman, Nnturwissensrhaften 48, 729(1961). 86.T. I. Tikhonenko, E. N. Dobrov, T. Velikodvorskaya, and N. Kisseleva, J .Mol. Biol. 18, 58 (1966). 87.S.Zamenhof, H. Alexander, and G. Leidy, J .Exptl. Med.98, 373(1953). 88.D. E. Hoard, Biochim. Biophys. Acta40,62 (1960). 89.M. Feldman, Bi0chi.m. Biophys. Acta149, 20 (1967). 90.S.Lewin, J .Chem.SOC.p. 792 (1964). 91. S. Lewin, Arch.Biochem. Biophys. 113,584 (1966). 92.S.Lewin and M. Barnes, J .Chem.SOC.(B), p. 478 (1966).

436

N. K. KOCHETKOV

AND

E. I. BUDOWSKY

93.L. Grossman, S. Lewin, and W. Allison, J .Mol.Biol. 3, 47 (1961).

!/4.J. Penniston and P. Doty, Biopolymers 1, 145(1963). 95. E. N. Dobrov, T. Tikhonenko, and S. Klimenko, Biophysics USSR 12, 957 (1967). 96. E. Triphonov, Yu. Lazurkin, and M. Frank-Kamenetsky, MoZ.Biol. 1, 164(1967). 97. M. Staehelin, Biochim. Biophys. Acta31, 448 (1959). 98. R. Shapiro and J.Huchmann, Biochemistry 6,2799(1966). 99. N. Broude, E. Budowsky, and N. Kochetkov, Mol.Biol. 1, 214(1967). 100. N. Broude and E. Budowsky, in preparation. 101. P.Witfeld and H. Witzel, Biochim. Biophys. Acta72, 338 (1963). 102. R. Shapiro and S. C. Agarwal, J .Am. Chem.SOC.90,474(1968). 103. P. T.Gilham, J. Am. Chem.SOC.84, 687 (1962). 104. S. Brostoff and V. Ingram, Science 168,666 (1967). 106. N. W. Ho and P.Gilham, Biochemistry 6, 3632(1967). 106. A. Girshovich, M. Grachev, and L. Obuchova, MoZ. Biol. 2,351(1968). 107. D. Knorre, E. Malygin, G. Mushinskaya, and V. Favorov, Biokhimiya 31, 334 (1966). 108. R. Naylor and P. Gilham, Biochemistry 6,2722(1965). 109. D. Knorre and G. Mushinskaya, Izv. Sibirsk. Otd.Akad.Nauk SSSR7, 132(1967). 110. R. Salganik, V. Dashkevich, aiid G. Dynuhits, Biochim. Biophys. Acta149, 603 (1967). 111. V. Drevitch, R. Salganik, D. Knorre, and E. Malygin, Biochim. Biophy.?. Acta 123, 207 (1966). 118. M. Yoshida and T. Ukita, J. Biochem. (Tokyo) 67,812(1965). 113. H. Priess and W. Zillig, 2. Physiol. Chem.342, 73 (1965). 114. N. Kochetkov, E. Budowsky, V. Shibaev, and G. Eliseeva, in press. 115. F. Cramer, K. Randerath, and E. Schiifer, Biochim. Biophys. Acta72, 150(1963). 116. F. Cramer and H. Seidel, Biochim. Biophys. Acta72, 157 (1963). 117. H. Seidel and F. Cramer, Biochim. Biophys. Acta108, 387 (1965). 118. F. Cramer and G. Schlingloff, Tetrahedron Letters 43, 3201(1964). 119. H. Seidel, Biochim. Biophys. Acta138, 98 (1967). 120. R. Bergquist and A. Deutch, ActaChem.Seand. 8,1880(1954). 12f. R. Caputto, L. F. Leloir, C. E. Cardini, and A. C. Paladini, J. Biol. Chem.184, 333 (1950). 192. T. Suzuki and E. Ito, J. Biochem. 46, 403(1958). 123. C. T.Yu and P. C. Zamecnik, Bioehim. Biophys. Acta76, 209(1963). 124. T. Venkstern, A. Mirsahekov, B. Gorshkova, and A. Bayev, Biokhimiya 28,712 (1963). 125. R. Shapiro and S. C. Agarwal, Biochim. Biophys. Res.Commun.24, 401 (1966). 126. A. Jones, J. Tithemor, and R. Walker, J .Chem.Soe. (C), p.1635(1966). 127. K. Brammer, Biochim. Biophys. Acta72, 217 (1963). 1% R. Holmes and R. Rolins, J. Am. Chem.SOC.86, 1242(1964). 129. F. Ascoli and I?. Kahan, J .Biol. Chem.241, 428 (1966). 130. W. Seer and D. Shugar, ActaBiochim. Polon. 8, 363 (1961). 131. M. Bessman, I. R. Lehman, J. Adler, A.S. Zimmerman, E. S. Simms, and A. Kornberg, Proc. Natl. Acad.Sci. U.S.44, 633 (1958). 132. A. M. Michelson, J.Dondon, and M. Grunberg-Manago, Biochim. Biophys. Acta 66, 529 (1962). 133. P. Cerutti, H. Todd Miles, and J. Frazier, Biochem. Biophys. Res.C o m m m . 22, 5 (1966).

T H E CHEMICAL MODIFICATION OF NUCLEIC ACIDS

437

134.P. Cerutti and N. Milles, J .Mol.Biol. 26, 55 (1967). 236.G. Belle, P.Cerutt,i, and B. Witkop, J .Am. Chem.SOC.88, 3946 (1966). 136. F. Baron and D. Brown, J. Chem.SOC. p. 2855 (1955). 137.S. Takemura, Biochim. Hiophys. Acta29, 447 (1958); Bull. SOC.Chem.Japan32, 920 and 926 (1959). 138. D. Hayes, F. Hayes-Baron, J .Chem.Soc.( C )p. , 1528 (1967). 139. A. Temperli, H. Turler, P. Rusl,, A. Danon, and E. Chargaff, Biochim. Biophys. Actu91, 462 (1964). 240.D. M. Brown, in“Methods in Enzymology,” (L. Grossman and I(. Moldave, eds.), Vol. 12, Part A, p. 31,Academic Press, New York, 1967. 141.B. Ellery and R. Symons, Nature210, 1159 (1966). 142,D. Brown, A. McNaught, and P.Shell, Biochim. Hiophys. Res.Comnzun. 24, 967 (1966). 143.E. Budowsky, J. A. Haines, arid N . Kochetkov, Dokl.A k d . h uukSSSR168, 379 (1964). 144. F. Lingens and H. Schneider-Bernlohr, Biochim. Biophys. Acta123, 611 (1966). 146.F. Lingens and H. Schneider-Bernlohr, Ann.Chem.686, 134 (1965). 146. R. Shapiro and R. Klein, Biochemistry 6, 235 (1966); 6, 3576 (1967). 247.A. B. Patel and H. D. Brown, Nature214, 402 (1967). 148. K. Kikugawa, A. Muto, H. Hayatsu, K. Miura, and T. Ukita, Biochim. Biophys. Acta114, 85 (1966). 149. H. Hayatsu and T. Ukita, Biochim. Biophys. Acta123, 458 (1966). 160. M. Yoshida and T. Ukita, Biochim. Biophys. Acta 123, 214 (1966). 151. L. Gal-Or, J. Mellema, E. N. Moudrianakis, and M. Beer, Biochcmistry 6, 1909 (1967). 166. C. Janion and D. Shugar, ActaBiochim. Polon. 16, 107 (1968). 153.H. Shuster, J .Mol.Biol. 3, 447 (1961). 154. D. Brown and P. Shell, J.Mol.Biot. 3, 709 (1961). 166. E. Freese, E. Bautz-Freese, and E. Bautz, Proc. Natl. Acad.Sci. U.S .47,845 (1961). 156. D. Verwoerd, H. Kohlhage, arid W. Zillig, Nature 192, 1038 (1961). 167. N. Kochetkov, E. Budowsky, and N. Simukova, Biokhimiya 27, 520 (1962). 268. V. Demushkin and E. Budowsky, Unpublished. 159. N. Kochetkov, E. Budowsky, V. Demushkin, M. Turchinsky, N. Simukova, and

E. D. Sverdlov, Biochim. Biophys. Acta142, 35 (1967). 160. D. Brown and P. Shell, J .Chcm.Soc.1966, 268.

161, N. Kochetkov, E. Budowsky, M. Turchinsky, and V. Demushkin, Dokl.Akad. Nauk SSSR 162, 1005 (1963). f62. S. Takemura, J .Biochim. (Tokyo) 46, 1285 (1959). 163. E. Rosenberg and S. Zamenhof, J .Uiol. Chem.236, 2845 (1961). 164.N. Kochetkov, E. Budowsky, and N. Simukova, Dokl.Akad.Nauk SSSR 163, 597 (1963). 265.E. Budowsky, N. Simukova, R. Slubaeva, and N. Kochetkov, Biokhimiya 30, 902 (1965). 166.E. Budowsky, N. Simukova, and L. Gus’kova, Biochim. Biophys. Acta166, 755 (1968). 167. M. Turchinsky, L. Gus’kova, arid E. Budovsky, Mol. BioZ.1, 792 (1967). 168. N. Kochetkov, E. Budowsky, M. Turchinsky, and N. Simukova, Biochem. Biophys. Res.Commzin.19, 49 (1965). 169. E.Budowsky, M. Turchinsky, aid L. Gus’kova, in prepsrstion. 170.N. Kochetkov, E. 3udowsky, and V. Demushkin, Mol.Bid.1, 583 (1967). 171. M. Turchinsky, E. Sverdlov, and E, Budowsky, Mol.Biol. 1, 623 (1967).

438

N. K. KOCHETKOV AND E. I. BUDOWSKY

17.2. N. Kochetkov, E. Budowsky, and R. Shibaeva, Biochim. Biophys. Acta88,493 (1963). 173.N. Kochetkov, E. Budowsky, and V. Domkin, Mot.Bio2. 1, 558 (1967). 174.N. Kochetkov, E. Budowsky, and V. Domkin, Khim.Prorodnych Soedin. p. 230 (1968). 176.J.Phillips and D. M. Brown, This series 7,349 (1968). 176.L.Frolova and L.L. Kiselev, Dokl. Acad.Nauk SSSR167, 1466(1964). lY7.S. Belman, T. Huong, E. Levinc, and W. Troll, Biochem. Biophys. Res.Commun. 14, 463(1964). 178.E.Budowsky, R. Shibaeva, E. Sverdlov, and G. Monastyrskaya, MoZ. Biol. 2,321 (1968). 179,D. Brown and J.Phillips, J. Mol.BioZ.11, 663(1965). 180.E.Budowsky, E.Sverdlov, R. Shibaeva, and G. Monastyrskaya, MoZ. Biol. 2,329 (1968). 181.P. Lawley, J. Mol.Biol. 24,75 (1967). 182.H. Johns, J. LeBlanc, and K. Freeman, J .Mol.Biol. 13, 861;883(1965). 18s.C. Janion and D. Shugar, ActuBiochim. Polon. 7 ,309 (1960). 184.I.Tessman, R. Poddar, and S. Kumar, J .Mol.Biol. 9,352 (1964). 186.C. Janion and D. Shugar, Biochem. Biophys. Res.Commun. 18, 617 (1965). 286. C. Janion and D. Shugar, ActuBiochim. Polon. 14, 293(1967). 187.R. Shibaeva, E.Budowsky, and N. Kochetkov, in press. 188.N. Kochetkov, E. Budowsky, E. Sverdlov, R. Shibaeva, V. Shibaev, and G. Monastyrskaya, Tetrahedrvn Letters 34, 3253(1967). 189.E. Budowsky, E.Sverdlov, and G. Monastyrskaya, in press. 190.E. Budowsky, E. Sverdlov, G. Monastyrskaya, in press. 191.F. Freese, E. Bautz-Freese, and S. Graham, Biochim. Biophys. Actu123, 17 (1966). 192.H.-J. Rhaese, E. Freese, and H. 8. Melzer, Biochim. Biophys. Acta166,491(1968). 19s. E.Freese and E. B. Freese, Biochemistry 4,2419(1965). 194.E. Bautz-Freese and E. Freese, Proc. Nutl.Acad.Sci.U.S.62, 1289(1964). 196.J.Phillips, D. Brown, R. Adman, and L. Grossman, J. MoZ. Biol. 12,816 (1965). 196.R. 0. Wilson and M. J.Caicuts, J .BioE.Chem.241, 1721(1966). 197.J. Phillips, D. Brown, and L. Grossman, J .Mol.Biol. 21, 405 (1966). 198.E. Freese, E. Bautz, and E. Bauta-Freese, Proc.Nutl. Acid.Sci.U.S. 47,845 (1961). 199.V. Chubukov, J .Mierobiol. Epidemiol. Immunobiol. (USSR), p. 129(1965). ,900.E.Freese and H. Strack, Proc. Nutl. Acad.Sci. U . S .48,1796(1962). 201.J. H. Van de Pol and G. Van Arkel, Mutat. Res.2 ,466(1965).

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 italics show the page on which the complete reference is listed.

A Abdulnur, S., 341(110, l l l ) , 373,(110, 111), 397

Abelson, J., 80(60), 113, 405(13), 434 Abrams, R., 160(227), 219 Achey, P. M., 118(64, 72, 73), 120(64, 72, 73), 139, 142054, 160), 151(64, 72, 73, 154), 152, 153(72, 160), 154(72), 213 (64,72, 73, 154, 160), 216, 217 Adam, R., 184(296), 221 Adams, A,, 270,271(143), 299 Adman, R., 6(21), 7(21), 14(21), 27, 145 (182), 181(182), 218, 433(195), 438 Adelberg, E. A., 9(51), 15(51), 24(51), 28 Adkins, B. J., 231(27), 296 Adler, A,, 249,260,297 Adler, J., 5(15), f l , 415(131), 4% Afanaseva, T. P., 96(120), 116 Agarwal, S. C., 413(102), 415(125), 4.90 AjdaEiE, Z., 186(306), 221 Alexander, Ch., 375(261), 401 Alexander, H. E., 18(85), 29, 412(87), 435 Alexander, M., 76(12), 112 Alexander, P., 21(101), 29, 118(1, 78, 80, 105), 120(78, 80, 1051, 128(78, 80), 130(78, 801, 131(1), 133(137, 147), 156 (78, 80), 213(78, 80), 214,2lG,217, 393(319), 402 Alfert, M., 15812221, 211(222), 213(222), 219 Allison, W., 412(93), 436 Alpers, D. H., 79(36), 112 Anderegg, J. W., 156(210), 164(210), $19 Anderlovb, A., 68(111), 73 Anden, M., 80(51), 94(113), 101(113), 113,114, 170(260), 820 Anderson, E. C., 187(308), 221 Anderson, J. S., 98(127), 116

Anderson, W. F., 118(117), 126(117), 116 Anthony, D. D., 105(153), 110(153), 116 Antoni, F., 194(324), 198(324), i?11 Aoki, A., 45(60), 72 Aoyagi, S., 245(83a), 246(83a), 197 Apelgot, S., 118(19), 151(19), 211(19), 212 (19), 213(19), 214 Apgar, J., 278(154), 299, 405(8), 434 Appel, K., 330(18), 341(154), 396, 399 Appleman, A. W. M., 155(200), 164(200), 218

Applequist, J., 240(64), ,997 Arbogast, A., 118(27), 120(27), 213(27), 214

Armstrong, A., 405 (61,433 Armstrong, A. T., 339(53), 396 Arnott, S., 273, ,999 Arya, S. K., 251(96a), 255, 256, 297 Asano, K., 107(159), 116 Ascione, R., 271(143), 299 Ascoli, F., 374(242), 401,415(129), 430 Attardi, G., 76(14), 77(16), 112 Augenstein, L., 350(177, 178,179), 399 August, J. T., 76(6), 84(6), 116 Augusti-Tocco, G., 406(34), 434 Axelrod, V.D.. 405(10), 4.94

B Baba, H., 332(38), 396 Babinet, C., 80(55), 113 Bach, D., 32(5), 45(80), 48(80, 82, 83), 70, 78

Bacq, Z. M., 118(1), 131(1), 214 Bar, H., 45(74), 46(74), 48(74), 64(74), 68(74), 72

Baird, S. L., Jr., 374(232, 2471, 400, 401 Balabuha, V. S., 133(142), 217 Baldwin, R. L., 14(81), 17, 19(81), 21(81),

439

28, 85(77), 114, 259(119, 122), 198, 304(11), 310(11), 326,412(82), 435

440

AUTHOR

INDEX

Belts, R. E., 164(236), $19 Ball, M. A., 341(85), 3997 Bennet, E.L., 160(228), 219 Ballard, D.,76(3), 112 Bertinek, J., 45(62), 7’2 Baltimore, D., 88(93), 114 Berenbom, M.,165,220 Bangerter, B. W , 240(67), 297 Berends, W.,377(264, 265, 266, 267, 268, Banghart, F. W., 162(233), 213 269),382,401 Baptist, J. E., 118(65), 120(65), 151(65), 213(65), 216 Berg, H., 45(73, 741, 46(74), 48(73, 74, 84, 85),57(85), 64(74), 68(74), 72 Barker, G.C., 38(26), 5O(26), 71 Berg, P.,80(50), 85(77, 781,86,90(98), 92 Barnabei, O.,198,222 (1051, 94(50), 99(98, 1301, 100(98), Barner, H.D.,118(31), 120(31), 213(31), lOl(135, 1381, 102(50, 98), 105(98, 214 130), 108(105), 110(98), 113,11/, 116, Barnes, M., 412(92), 436 170(259), 10,304(11,15), 310(11,15), Barnett, L.,286(174), 300, 385(293), 402 326 Baron, F., 415(136), 437 Barone, T.F., 118(90), 173, 174(90), 211 Berg, T. L., 189,205(317), 221 Bergmann, E. D., 341(65), 349(65), 366 (90),226 Barrel, B., 405(7), 434 (209),396,.boo Bergmann, F., 351(If@), 399 Basilio, C.,5(16), 14(16), 2’7 Bergquist, R., 415(120), 436 Baudet, J., 341(143), 398 Bergson, G ., 341(721,696 Bauer, H.,37(14), 71 Baugnet-Mahieu, L.,164(238), 187(238), Bernardi, G.,57(96), 73 Berns, K.,118(117), 126(117), 217 205(238), 213(238), 219 Berry, R.,118(42), 120(42), 155(42), 216 Bauts, E., 417(155), 433(198), 437 Bauts, E. K. F., 116,307, 308, 310(28), Bersohn, R.,374(232), 400 Bertani, G.,118(48), 120(48), 121(48), 135 3% (481,213(48), 2f6 Bauts, F. A.,308,326 Bautz-Freese, E., 1(3), 6(28), 7(34, 351, Berthier, G.,328(2), 341(143), 396,396 23, 24, 26, 271, 338, 8(34), 12(28),13(3), 14(28, 34,46,95. Berthod, H., 330(22, 341(22,23,24,26,27,63,98, 163), 346 961, 15(95), 16(3), 19(35, 95, 96), 21 (22,26), 347(26), 349(22), 350(22, 23, (95), 24(3), 25(95), 27,28, 118(94), 24), 354(22, 23, 24), 365(98), 382(22, 204(94), $is, 383(285), 402,417(155), 23), 383(22, 24), 385(26), 392(63), 432(191, 193), 433(194, 198, 437,4% 396,396,397,399 Bayev, A. A.,405(10), 415(124), 434,43C Bessman, M.J., 5(15), 67, 143(165), 218, Beaven, G. H.,350(172), 399 415(131), 436 BebareviE, A., 118(44), lZO(441, 165, 166 (249), 190(318), 191(318), 193(318, Betel, I.,155,164(201), 818 321), 197(318), 199, 200(334), 201 Beukers, R.,377(264, 265, 266, 267, 269), 382, 401 (3351, 202(335), 203(337), 213(318), Beychok, S., 224(5), 296 a 4 , 220, ,@i, 9.@ Biczo, G.,341(155), 399 Becker, E. D., 357(194), 364(194), 399 Billen, D.,118(33, 66, 66a, 71, 72), 120 Becker, E. F., Jr., 19(93), 21(93), d9 (33, 66, 66a,71, 721, 124(66a), 142, Beckwith, J. R.,111 (174), if 6 151(66, 66a, 71, 72), 153, 154, 173 Bedford, J. S., 130(124), 156(208), 217, (270), 213(33,66,%a, 71,72, 154, 160, 219 269), $14,916,9i7 Beer, M., 406(26, 27, 30), 416(151), 434, Birnstiel, M., 311(39), 326 437 Beers, R. F., 249(88), 254(88), 286(88), Bishop, C.W., 163,219 Blake, A., 280,286(179, 183). 287,299,300 297 Blather, F. R., 103(150), 116 Belle, G.,415(135), 437 Blazsek, V.,68(106), 73 Belman, S., 428(177), 438

AUTHOR

441

INDEX

Blears, A.J., 374(235), 400 Blohina, V. D., 118(4), 21.4 Blout, E. R., 224, 225(8), 281(163), 296, 299

Boag, J.W., 382(275), 401 Boedtker, H., 289(193, 194), 300 BorBni, E.,194(323), 198(323), 221 Bogdanova, S.L., 96(120), 116 BohitEek, J., 45(71), 61(103), 72, 73 Bolle, A.,78(21, 24), 79(46), I06(21), 112, 113

Bollum, F. J., 143(167, 1741, 151(177), 156(210), 163(235), 164(210), 218,819, 244(80), 249(80), 254(80), 257(80), 259(80), 260(80), 261(80), 274(80), 297, 305, 306,526 Bolton, E. T., 310(37), 317(44), 326 Bonner, J.,168,170,22'0 Botrk, C., 374(242), 401 Boublik, M., 57(98),73 Boudnitskaya, E. V., 185, 205(303), 221 Bourgeois, S., 93(108), 110(108), 114 Boxer, G. E., 89(94), 109(94), 114 Boyce, R. P., 118(13, 91, 97, 113), 119 (13, 91), 120(13), 123(13, 91), 125 (113), 151(13), 154,210(13), 213(13),

(174, 175), 300, 385(293), 402,405 (13),4S4 Brent, T. P.,118(79), 120(79), 128(79), 130(79), 151(79), 155(79), 156(79), 157(79), 158,213(79), 216 Bresler, S. E., 14(91), 15(91), 18(91), 19 (91),20,21,25(91), 29 Breyer, B., 37(13, 14), 71 Bkzina, M., 32(7), 36(7), 68(7), 70 Britten, R. J., 302,317,318,319(5), 321, 324

Brody, E., 94(112), 114 Brody, E.N., 96(119), 116 Brookes, P., 20,21(100), 22(106), 29, 374 (249, 2501, 385(290, 291, 292), 392 (309,310, 311, 312, 313, 314), 393 (318), 401, 402 Broom, A. D., 238(49), 296, 369(213), 370 (2161,400 Brostoff, S.,413(104), 4%’ Broude, N.,406(25), 413(99, IOO), 434, 436

Brown, D., 415(136), 416(142), 417(154), 419(160), 427(160), 428(160), 429,433 (195,1971,437, 438 Brown, D. D., 322(52), 326 Brown, D. M., 1(6), 6(18, 19,20,21,25, 214,816 291, 7(6, 21,291, 14(21, 291, 17(20), Boy de la Tour, E., 78(21), 106(211, 113 19(18), 27, 391(302, 304, 3051, 402, Boyland, E., 374(238,239,240), 400 409(44), 415(140), 42811751,434,437, Bradbury, E.M., 286(172), 300 Bradley, D. F., 225(15, 16), 274115, 161, 438 286(173, 1781,295, 300, 305(17), 326, Brown, G. B.,39(32), 71 330(28), 341(28, 89, 157), 355(28), Brown, G. L.,406(34), 434 Brown, H. D.,416(147), 437 356(89), 395, 397, 399 Brahms, J.,57(99), 73, 225, 238(56, 57), Brown, J. C., 79(41), 89(41), 113 241,242(56, 771,243,247(57,86), 248 Brown, R. A.,202 (571,249(87a), 250(106), 252661,253, Brownlee, G.,405(7), 434 254(87a), 255(86, 87a),261, 267(14), Bruner, R.,99(133), 104(151), 105(133, El), 106(133), 116 274,275,279,296, 297, 698 Brunfaut, M.,185(303), u)5(303), 221 Brammer, K. W., 12,28, 41511271,436 Buc, M. H., 183(294), 221 Brdar, B., 174(265), ,920 Buchanan, J.M., 78(22), 112 BrdiEka, R., 32,YO Bremer, H.,79(38), 82.(67), 83(67), 87 Bucher, N.L., 187(313), 221 (86),88(38), 89, 90(86), 92(86, 101, Budilova, E. V., 164(240), 187(240), 205 (240),219 1021, 93(101), 98, 99(133), 104(38, 1511, 105(86, 133,1511,106(38, 1331, Budnikov, G., 37(20), 71 107(86), llO(67, 1661, 113, 114, 116, Budowsky, E. I., 6(24), 7(24, 331,B', 405 (19, 33),406(25), 410(59), 413(99, 116 loo), 414(114), 417(157, lSS), 418 Brenner, S., 78(29), 79(46, 471, 110(29), (158, 159), 419(159), 420(159), 421 112,113, 207(348), 210(349), 222, 286

442

AUTHOR

INDEX

(161), 423(164, 165, 166, 167, 168, Carrier, W. L., 118(12), 119(12), 120(12), 123(12), 151(12), 154, 210(12), 213 169), 424(168), 425(33, 159), 426(33, 1701, W(164, 165, 170, 171, 172, 1731, (121,214 428(164, 165, 172, 178), 429(178), 430 Carroll, D. G., 339(53), 396 Caskey, C.T., 320(48), 326 (1801,431(180,187),432(178,187,188, Cassani, G.,305,306,326 189, 1901, 434,434 436,437,438 Cassim, J. Y., 230,296 Bugarski, M., 186(306),221 Cavalieri, L. F., 40,71 Bukaresti, L.,68(106), 73 Cecil, R., 70,73 Bulant, V.,45(61), 72 Burgess, R.R., 80(591,83(59), 84(59, 651, Cernf, R.,45(72), 72 Cerutti, P., 145(181), 218,391, 402,415 113,116 (133,134, 135),@6,437 Burki, H. J., 118(122), 120(122), 130(122), Chamberlin, M. J., 80(50), 85(77, 781,86, 2f7 87(85), 94(50), 101(138), 102(50,139), Burton, A.,87(89), 114 113, 114,116,259(121), 273(150), 298, Burton, K.,102(142), 116,411(68, 691, 299, 304(11, 12, 13, 14, 151, 310(11, 437 15), 3.96 Bush, C. A., 240, 242(77), 262(59), 275, Chambers, R., 405(23), 409(49), 414(23), 297,341(88), 397 Butler, G.,410(60,61), 436 434,436 Butler, J. A.V., 118(79), 120(79), 128(79), Chambon, P., 76(5), 105(154), llO(154, 165), 112,116,118(47), 120(47), 816 130(79), 133(134), 151(79), 155(79), 156(79), 157(79), 158(79), 213(79), Chambron, J., 374(231), 4UU Champe, S.P., 78(20), 11% 216,817 Chan, S. I., 240(65, 66, 67, 701, 297,341 C (141), 369(211, 212), 398, 400 Cabrera-Juarez, E., 12(77), 15(77), 19 Chandra, P.,9, 11(62), 15(47,62,127), 23, 28,29 (77),28 Caicuts, M. J., 6(22), 7, 14(22), 27,433 Chang, L. O., 162(233), 219 Chang, S., 405(12), 434 (I%), 438 Caillet, J., 341(92, 93, 94, 95, 96, 97, 98, Chantrenne, H.,173(271), 213(271), 220 104, 105), 359(92), 361(94, 96, 203), Chapman, J. D., 139, 217 365(98), 367(96), 369(93,97),370(93), Chargaff, E.,102(141), 116,405(14, 151, 410(64, 65), 411(14, 15), 415(139), 373(!33), 374(105), 388(297), 397, 400, 402

Cairns, J., 142(164), 218 Caldwell, I., 118(80), 120(80), 128(80), 130(80), 156(80), 213(80), 216 Calendi, E., 48(81), 72 Callis, P.R., 351(183),399 Calvin, M.,374(229), 400 Cammarano, P.,199,822 Cannelakis, E.S., 143(168), 218 Cantor, C. R., 244,245,254,277,278,279, 297,2998

Cantor, S. M., 39(28), 71 Caputto, R.,415(121), 436 Carbon, I.,412(81), 436 Cardini, C. E., 415(121), 436 Carlson, J. G., 158(224), 211(224), 213 (224),219

434,435,437

Charlier, M., 382(277), 4 O i Chase, M.,122 Chen, G., 9(51), 15(51), 24(51), 2s Chen, J. H., 9(42), 28 Cheng, Ping-Yao, 288, 289(189), SO0 Chevalley, R.,78(21), 106(21), 112 Chinali, G.,199(333), 922 Chipchase, M.,321 (49), 386 Cho-Chung, Y. S., 204(339), 205(339), 213(339), 222 Chubukov, V. F., 14(114), 29, 433(199), 438 Clark, B. F. C., 98(127),116 Clark, J. B.,131(128), 217 Clark, L. B., 234(46), %96,350(175), 351 (175), 399

AUTHOR

443

INDEX

Clarke, G. A., 339(54), 396 Claverie, P., 341(92, 93, 94, 95, 96, 100, 101, 103, 104, 108, 109), 357(103), 359(92), 361(94,96), 367(96), 369(93), 3701931, 373(93), 374(104), 397 Cleaver, J. E., 155(194), 156(194), 157 (1941, 218 Clementi, E., 331(35, 36), 339, 396’ Clementi, H., 331(36), 396 Cohen, J. A., 174(276), 175(276), 220 Cohen, S. S., 118(27, 31), 120(27, 31), 213 (27, 31), ,914 Cohen, S. N., 95(117), 96(124), 116 Cole, L. J., 133(136), 917 Cole, T., 375(262), 401 Colvill, A. J. E., 83(70), 94(111), 95(111), 96(122), 108(122), 113, 114, 116 Commings, D. E., 210(353, 3541, 222 Contesse, G., 79(43), 113 Cotter, R. I., 279,299 Coulson, C. A., 118(6), 214 Cox, J., 436 Cox, R. A., 133(148), 217,231(29), 232 ( B )233(29), , 296, 350(174), 399 Crab%, P., 225 (19),296 Cramer, F., 406(36), 414(115, 116, 117, 1181, 434, 436 Crane-Robinson, C., 286(172), 500 Crathorn, A. R., 118(79), 120(79), 128 (79), 130(79), 151(79), 155(79), 156 (79), 157(79), 158(79), 213(79), 216 Crawford, E. M., 83(71, 74), 99(74), 100 (741, 102(74), 113 Crawford, L. V., 83(71, 74), 88(91), 99 (741, 100(74), 102(74), 113,114 Creasey, W. A., 155(195), 164(195), 187 (310), 218, 221 Crick, F. H. C., 24,29, 207(346), 222,286 (174), SOU, 253(99), 298, 320(47), 321 (50), 326, 326, 359(196), 385(293), 399, 402

Crothers, D. M., 306, 309, 325 Curtis, H. J., 206(343, 3441, ,922 Cusachs, L. C., 339(52), 340(52), 39G Cushley, R. J., 238(54), 296 Cusin, F., 210(349, 3521, 222

D Daljanski, F., 194(326), f?22 Damjanovich, S, 158(225), 213(225), 2f9

Damle, V., 240(64), 297 Dangluk, S. S.,374(235), 4UO Daniels, M., 12(74), 28 Danilov, V. I., 341(69, 1351,396,398 Danon, A., 415(139), 437 Darnell, J. E., 85(172), 116, 195, 221 Das, H. K., 79(34), 112 Das, N. K., 158(222), 211(222), 213(222), 219

Dashkevich, V., 413(110), 436 D a m e , M., 374(231), 4OO Davidson, J. N., 143(166), 163, 218, 219 Davidson, N., 258(118), 298, 304(8, 101, 306, 309(30), 316, 317(8), 326,351 (181, 1821, 399 Davidson, P. F., 10(54), 28, 133(145), 135 (145), 217 Davie, E. W., 80(61), 90(61), 92(61), 94 (611, 96(61), 110(61), 113 Davies, D. R., 9(42), 28,241(76), 252(99, 101, 1021, 257(117), 259(122), 297, 298, 374(236), 400 Davis, D. R., 331(35, 361,396 Davis, R. C., 243,297, 341(124), 374(124), 398

Davison, P. F., 18(87), 29 Dawid, I,. B., 322(52), 326 Dean, C. J., 118(18, 80, 1051, 119(18), 120(18, 80, 105), 124(18), 128(80), 130(80), 151(18), 154, 156(80), 210 (18), 213(18, 80), 214,216 Delahay, P., 43(52), 45(52), 72 Delbriick, M., 11(69), 28, 154(192), 218 De Lerme, B., 374(242), 401 Dellweg, H., 15(127), 29, 377(2701, 401 Del Re, G., 330, 337, 341(158), 396, 399 De Mars, R. I., 109(163), 116 Dembitzer, H. M., 194(327), 222 Demerec, M., 118(25), 120(25), 213(25), $14

Demushkin, V., 405(19, 331, 417(158), 418 (158, 1591, 419(159), 420(1591, 421 (161), 425(33, 1591, 426(33, 1701, 427 (1701, 434, 437 Denliardt, D. T., 118(101a), 149(101a), 216 Denhardt, G, H., 78(21), 106(21), 11.9 Denia, A., 330(25), 331(321, 341 (25, 32), 350(25), 354(25), 382(25), 3g5 Depireux, J., 341(129), 398

444

AUTHOR

INDEX

DuBois, K. B., 118(38), 120(38), 160(38), Deutch, A., 415(120), 436 916 de Vellis, J., 206(342), 213(342), 2.92 De Voe, H., 240(72), 297, 329(10), 341 DuBois, R. P., 205(341), 213(341), 2Z.8 (10, 831, 346(10), 350(173), 355, 366 Duchesne, J., 341(127), 349(166), 398, 399 Diitting, D., 405(9), 434 (101, 372,396, 397 Dunn, J. J., 116 Devreux, S., 173(271), 213(271), 220 Dupuy-Mamelle, N., 341(611, 391(611, Dewey, D. L., 133(151), 136(151), 217 396 Dewey, K. F., 79(41), 89(41), 113 Dymshits, G., 413(110), 456 Dewey, W. C.,156(209), 157, 219 di Bitonto, G., 198(332), 222 E Dickstein, S., 351(1801,399 Dicckmann, M., 92(105), 99(130), 105 Earle, J. D., 118(110), 124(110), 125(110), 126(110), 127(110), 216 (1301, 108(105), 114,116 Ebal, J. P., 225(13), ,996 Di Marco, A,,48(81), 7.8 Echols, H., 78(25), 119 Dimroth, K., 410(62), 436 Eckstein, H., 118(86), 120(86), 126, 151 Dirksen, M., 78(22), 112 (86), 155(86), 158(86), 165(86), 205 Diner, 8., 341(158), 399 (861, 213(86), 216 Ditmars, W. E., 374(230), 400 DjordjeviE, B., 118(74), 120(74), 127, 151 Edelman, A., 174(276), 177(276), 920 Edgar, R. S., 78(21), 106(21), 111 (741, 156(74), 158, 213(74), 816 Edmonds, M., 181(284), ,920 Djordjevi6, N., 118(106), 124(106), 216 Dobrov, E. N., 57(97), 73, 284(163), 300, Edwards, P. A., 21 (103), 29 Egami, F., 412(83), 436 412(86, 951, 456,436 Eguchi, S., 46(57), 72 Doermann, A. H.,12.2 Ehrenberg, A., 375(255), 401 Doida, Y., 187(307), 221 Doly, J., 76(5), 105(154), 110(165), 112, Ehrenberg, L., 375(255), 401 Eigner, J., 31(1), 70 116 Eisenstadt, J. M., 173(273), 213(273), 220 Domkin, V., 427(173, 174), 438 Eisinger, J., 375(257, 2791, 401 Dondon, J., 415(132), 436 Doty, P., 18(86), 29, 79(41), 89(41), 113, Ekert, B., 118(19, 201,151(19), 211(19), 212(19, 201, 213(19, 20),,914 118(117), 126(117), 184(296), ,917, 221, 224(2, 8),225(8), 229(24), 231(2, 291, Eliseeva, G., 414(114), 4% 232(29), 233(29), 267(133), 280(180), Elkind, M. M., 158, 211(216, 217, 2181, 213(216, 217, 218), 219 281(161), 285(2), 286(170), 289(193), 296, 298,,999, 300, 301, 303, 304(7), Ellery, B., 415(141), 437 316, 317(42), 324,326,350(174), 373 Ellis, F., 156(203), 218 Ellis, M. E., 133(136),617 (220), 399, 400, 412(94), 436 Elson, D., 76(2), 112 Douzou, P., 382(276), 401 Dove, W. F., 78(23), 11.2,304(8), 317(8), Elving, P. J., 38, 39(27, 34, 351, 40(35), 41(27, 45), 42(34, 41, 451, 43(34, 45), 3.26 71, 349(167, 168, 169, 170), 399 Downing, M., 370(218), 400 Drake, J. W., 15(117, 119), 29, 387(296), Emerson, T. R., 233(38, 39, 40, 41), 234 (40, 41, 45), 235, 237, 238, 247(84), 390(289), 40.2 .89G, 697, 354(184), 399 DmkuliE, M., 118(59), 120(59), 138(59), 151(59), 173(59), 174(265), 175(59), Emery, A. J., Jr., 80(63),82(63), 83(63), 113 213(59), 216,220 fl Drewitch, V., 406(35), 412(35, I l l ) , 414 Emmelot, P., 8(37, 38), Emrich, J., 79(40), 89(40), 113 (35,I l l ) , 434,436 Englesberg, E., 77(19), 112 Drobnik, J., 350(177, 178, 179), 399 Dubinin, N. P., 118(93), 120, 204(93), ,916 Ennis, H. L., 93(106), 114

AUTHOR

445

INDEX

Ephrussi-Taylor, H., 12(76), 15(76), 19 (76), 28 Epstein, R. H., 78(21, 22, 241, 80(61), W (611, 92(61), 94(61), 96(61), 106(21, 1571, 110(61), 112, 113,116 Ermolaeva, N. V.,133(142), 217 Errera, M., 118(2,30,32), 120(30,32), 133 (32, 1351, 151(32), 173(32), 174(32), 185(301, 303), 205(301, 305), 213(30, 32), 214,221 Evans, A., 94(113), 101(113), 114, 170 (2601, 220 Evans, H. G., 130(123), 217 Everett, G. A., 278(154, 1 5 3 , 299, 405f8, ll),494 Eyring, H., 231(30, 33), 235(30), 238(30), 296

F Fancher, H., 99(130), 105(130), 115 Falk, M., 373(223), 400 Farkd, J., 234(47), 296 Farkas, W., 410(63), 436 Fasman, G. D., 231,249(87), 254,260(87), 262(124), 296, 297, 2D8 Faulkner, R., 405( 121,434 Fausto, N., 118(83), 120(83), 160(83), 162, 163(83), 164(237), 187(316), 189(316, 316a), lW(83, 316, 316a), 205(237, 316), 213(237), 216, 221 Favorov, V., 413(107), 436 Feldman, H., 405(9), 434 Feldman, M., 412 (89), 436 Feldschreiber, P., 118(18), 119(18), 120 (18), 124(18), 151(18), 154(18), 210 (is), 213(18), 214 Felsenfeld, G., 31(2), 57(101), 70, 73, 240 (63), 245, 249(93), 252(102, 103, 1041, 277(83b), 296, 297, 298, 341(86), 397 Fernandes-Alonso, J. I., 329(12), 341(12), 395

Fernindez-Morin, H., 83(70), 113 Fiala, S., 44(59), 72 Ficq, A , 185(301), 205(301), 221 Filipovich, I. V., 118(4), 214 Filipowicz, B., 44(58), 72 Fisher, H., 109(163), 116 Fitch, F. W., 118(38), 120(38), 160(38), 215

Flamm, W. G., 309, 323(32), 325 Flessel, C. P., 252(100), 298 Fluke, D. J., 150(188), 218 Fontaine, F., 267(134), 298 Forejt, J., 37(17), 71 Forssberg, A,,118(2, 35, 36), 120(35, 361, 160(35, 36), $14 Fowler, G. N., 341(871,397 Fox, C. F., 93(lG9), 99(129), 100(129), iio(iog), 114, 116 Fox, E., 118(49), 120(49), 121(49), 135 (49), 213(49), 216 Fox, J. J., 234(44), 238(54), 296 Fradkin, G. E., 118(100), 121(100), 213 (1001, 216 Fraenkel-Conrat, H., 3(7), 7(32), 9(32, 43, 48, 49), lO(32, 57), Il(32, 57, 631, 12(32, 78), 13(32), 14(32, 431, 15(32, 48, 57), 17(48, 49), 18(83), 19(32), 20 (43), 21(48), 22(32, 48, 49, 1051, 23 (32, 83), 24(48), 26(32), %7, 28, 433 (3, 4), 433 Frampton, E. W., 118(71), 120(71), 142 (159), 151(71), 153(159), 173(268), 178, 179, 198(268), 213(71, 159), 216, 217, 220 Frank-Kamenetsky, M., 412(96), 436 Franklin, R. M., 23(107), 29 Frary, B. D., 45(76), 46(76), 50(76), 60 (76), 61, 68(76), 72 Frascicatti, M., 374(242), 401 Frazier, J., 145(181), 218, 415(133), 436 Fredericq, E., 267(134), 298 Freeman, E. J., 106(156), 116 Freeman, K. B., ll(68, 701,12(68), 15 (68), 28, 391(303), 402, 438 Freese, E., 1(3), 6(28), 7(34, 35), 8(34), 9(46), 12(48), 13(3), 14(28, 32, 46, 94, 95, 96), 15(95), 16(3), 19(35, 94, 95, 961, 21(95), 23, 24(3), 25(95), f l , 28, 29, 118(94), 204(94), 216, 383(283, 284,285), 387(283,284), 393(284), 401, 402, 417(155), 432(191, 192, 1931, 433 (194, 198, 200), 437,438 Freifelder, D., 10(54), 18(87), 28, 29, 118 (51, 52), 120(51, 52), 121(51, 521, 133 (51, 52, 1451, 13561, 52, 129, 1 4 8 , 136(51, 52, 1291, 138, 150(51, 52, 129), 213(51, 52, 1291, 215,217

446

AUTHOR

Freifelder, D. R., 118(52), 120(52), 121 (52), 135(52), 136(52), 150(52), 213 (52), 216 Frenkel, E. P., llP(82), 120(82), 160(82), 816

Fresco, J. R., 224, 225, 252(100), 256, 270 (142), 271(142), 289(193, 194, 195, 198), 290(198), ?296,298, 899, 300, 310 (351, 326 Freund, A. M., 57196),73 FriE, I., 234, 296 FriE, J., 268( 136), 299 Fried, M., 409(44), 434 Fritz, H., 410(58), 436 Frolova, L., 428(176), 438 Fromageot, P., 106(158), 109(162), 116 Fuchs, E , 80(53), 83(69), 90(97), 93 (97), 104(152), llO(97, 152), 113, 114, 116 Fujii, I., 269(139, 140), 270(139), 299 Fujiki, H., 143(175), ,918 Fujioka, M., 156(211), 186(211), 187(312), 219,2?,91 Fujita, H., 330(21), 341(76, 134), 354(76), 3996,397, 398 Fuke, M., 269 (140), 899 Fukuda, N., 339(54), 341(101), 396, 397 Fukui, X., 341(151), 385(151), 398 FulIer, W., 273(149a), 89g Furth, J. J , 76(4, 12),80(51), 94(113), 101(113), 112, 1f3, 114, 170(260), E20 Futai, M., 238(55), 296

G

INDEX

Gellert, M., 241(76), 297, 309(33), 326 Georgiev, G. P., 168, ,980 Gersch, N. F., 341(106), 374, 3.97 Gerson, J., 14(95), 15(95), 19(95), 21(95), 25(95), 29 Gerstenberger, A., 10(56), 28 Gierer, A., 4, 14(12), f l Giessner-Prettre, C., 330(22, 23, 241, 331 (331, 341(22, 23, 24, 33, 63, 1401, 346 (22), 349(22), 350(22, 23, 241, 354(22, 23, 241, 382(22, 23, 241, 383(22, 241, 392(63), 396, 396, 398 Gilbert, M., 331(32), 341(32, 108, 1091, 374,396,397 Gilbert, W., 76(11), 77(17, 181, 111(18), 112 Gilham, P. T.,405(2), 413(21, 103, 105, 1081, 43% 434, 436 Gill, S. J., 370(218, 219a), 400 Gillespie, D., 85, 116, 301(3), 307, 323(3), 324,326 Ginosa, W., 150(184), 218 Giovanella, B. C., 374(241), 401 Girshovich, A., 413(106), 436 Gladstone, L., 76(1), 112 Glaubiger, D., 243, 997 Goel, N. S., 341(101), 397 Gold, M., 101(137), 116 Goldberg, I. H., 109(160), 126 Goldstein, A., 79(34), 112 Goldthwait, D. A., 105(153), 110(153), 116 Gollmick, F. A., 45(73, 741, 46(74), 48 (73, 74, 84), 64(74), 68(74), 72 Gomatos, P. J., 269(137), 299, 310(36), 3% Goodgal, S. H., l4(89), 18(89), 29 Goodman, H. M., 76(10), 112, 405(13),

Gage, 1,. P., 78(28), 111 Gaines, K., 79(38), 88(38), 98(38), 104 (381,106(38), 113 Gal-Or, L.,416(151), 437 Ganesan, A. T., 210(351), 222 434 Gordon, J. A., 373(224), 400 Gardner, A. W., 38(26), 50(26), 71 Gardner, B. J., 286(182), 287(182), 300, Gordon, M. P., lO(58, 59, 60),12,27 Gordy, W., 341(147, 148, 1491, 375(147, 374(233), 400 148, 149, 253, 254, 258, 259, 260,2611, Gartland, W. J., 271(145), 299 376,377,398,401 Gayevska, E., 118(117), 126(117), 917 Geiduschek, E. P., 10(54), 19(92, 931, 21 Gorlenko, Zh. M., 96(lm), 116 (92, 9 3 , 28, 29, 78(24, 28), 84, 85 Gorn, R., 224(8), 296 415(124), 436 (75), 94(111), 95011, 115, 118), 96 Gorshkova, B., (119, 121, 1221, 97, 108(122), 118, 113, Goutarel, R., 288(188), 300 Goutier, R., 164(238), 187(238), 189, 205 11.4, 116 (238, 3171, 213(238), 219, 281 Gelbard, A. S., 185(304), 281

AUTHOR

447

INDEX

Grachev, M., 413(106), 436 Grady, L. J., 139,141,217 Graham, S.,7(35), 19(35), 27,432(191), 438

Granick, S., 286(177), 300 Grasemann, J. M., 118(49), 120149), 121 (491,135(49), 213(49), 215 Gratzer, W. B., 231(29), 232(29), 233(29), 350 279(15&.),280,281(158),296,299, (1741,599 Graziosi, F.,172,220 Green, B., 374(238, 239,240,248),400,4Ol Green, D. M., 14(115), 29 Green, D.W., 249(89), 2.97 Green, G., 288(188), 300 Greenberg, J., 9(49, 50), 15(50, 1241,17 (491,22(49), 28,29 Greer, S., 118(109), 124(109), 216 Grimison, A.,12(74), 28 Gros, F.,76(11), 77(16), 78(27), 79(43, 48), 92(104), 93(108), 96(123), 110 (108), 112, 113,114,116 Grosjean, M.,224(4), 225(4), 230(4), 296 Grossman, L., 6(21,29), 7(21, 29), lI(61, 73), 12(73), 14(21, 291, 15(61, 71,72, 73), H,28, 145(182), 171(262), 181 (182), 218, 220, 231(31), 249(87), 254 (311,260(87), 296,297,391(301, 305, 306,307),402,412(93), 433(195, 1971, 436,438

Grunberg-Manago, M., 9(41), 14(41), 19 (411,28,89(95), 114, 415(132), 436 Giinther, H. L., 379(273), 401 Guild, W. R.,118(8), 214 Gulyas, S.,118(75), 120(75), 128(75), 130 (75), 156(75), 213(75), 215 Gumport, R. I., 99(129), 100(129), 115 Gurskii, G . V.,374(234), 400 Guschlbauer, W., 63(104), 64(104, 1051, 73,249(92), 29Y Gus kova., I,., 423(166,167,169), 437 Gut, J., 43(50), Y2,341(165), 399 Guttes, S.,158(223), 211(223), 213(223), 219

H Habermann, V.,45(72), Y2, 405(16), 416

(IF), 434 Harhagen, J. M., 267(135), 298 Hadopian, H., 405 (6),433

Hadeib, Lj., 118(44), 120(44), 816 Hagen, U.,118(88, %a), 133(150), 144 (881, 145(88), 148(88), 167(88, 88a), 169(88, 88a), 172,213(88, %a), 216, 2lY

Hsger, G., 216 Haines, J. A.,393(316), 402,43Y Haines, R. B.,118(6), a14 Hall, B. D.,90(99), 114, 116, 289(193), 300,301(21,324 Hall, C. E., 83(73), 100(73), 113 Hall, E. J., 130(124), 156(208), 217,219 Hall, J. B., 102(143), 115 Hamer, D., 39(30), 40,42(30), 68(30), Yl Hamilton, 11.D., 273(147), 299 Hamlin, R. M., Jr., 36412041,400 Han, A,,158(219, 2201,211(219,220), 213 (219,220), 219 Hanawalt, P. C.,118(11), 119(11), 120 (Il), 123(11), 151(11), 153,174(275), 210(11), 213,214,220 Hanlon, S., 370(215), 400 Hanson, K.P., 155(199), 164(199), 218 Harbem, E., 184,185(300), 221 Hariharan, P. V.,11 (701,28 Harm, W., 118(98, 991, 121(98, 991,213 (98,991,816 Harrington, H., 118(37, 84,85), 120(37), 144,145(84, 851, 146,147,160(37), 167 (84,85), 169(84,851, 213(84, 8 5 ) , 215, 216

Harris, F. E,, 341(119, 120,121,1221,3.98 Hartman, K.A,,Jr., 373(223), 400 Hartman, Ph. E., 143(163), 207(163), 218 Hartmann, G., 110(167), 111(170), 116 Haschemeyer, A. E. V.,238(52), 296,357 (188,189),374(251, 2521,399, 4 O l Haselkorn, R., 18(86), 29, 257(115), 289 (193,194), 2.98, 300 Hashizumc, H., 253, 269(139), 270(139), 280,698,299

Hastings, R.,11(65),25 Hatch, F. T., 118(87), 213(87), 616 Haug, A.,382(276), 401 Hayashi, M., 87(88), 94(110), 114,175, 260

Hayashi, M. N., 94(110), 114 Hnyatsu, H., 416(148, 149), 43Y Haves, D., 415(138), 43Y HRyes, F.N., 82(68), 113, lXlf286), 221

448 Hayes, W., 24(109), 29, 383(287), 40B Hayes-Baron, F., 415(138), MY Haynes, R. H., 118(15, 65, 92, 1161, 119 (15, 92), 120(15, 651, 123(15, 92), 151 (15, 65), 154(15),210(15), 213(15, 65), 214, 216, 216, 917 Heath, J. C., 39, 40(29), 41(29), 42(29), ri Heesing, A., 411(70), 436 Heidelberger, Ch., 184(300), 185(300), 221, 374(241), 401 Helene, C., 382(277), 4Ol Heller, H. C.,375(262),401 Helmkamp, G. K., 240(65), 254(109, 110), 289(109, 110, 197, 200), 290(109>,291 (109, 197, 2001, 297, 298, 300, 341 (lal), 369(211), 398, 400 Henley, D., 271 (1431,299 Hennig, S. B., 143(176),218 Henry, J., 80(52), 113 Heppel, L., 305(17), 325, 409(45), 434 Herak, J. N.,341(147, 148),374(147, 1481, 375(260), 398, 401 Herriott, R. M., 12(77), 14(88), 15(77), 18(88), 19(77), 28, 29, 383(288), 402 Hevesy, G., 118(120), 159, 160, 217, 219 Hewitt, R., 118(66, 66a, 721, 120(66, 66a, 72), 124(66a), 151(66, 66a,721, 153 (72), 154(72), 213(66, 66a, 72), 216 Hewlins, M. J. E., 6(19, 251, 7, 27 Heyrovsk?, J., 32, 36(9), 37(16, 17), 38 (2'2, 241, 39(9), 71 Hiatt, H., 76(11), 118 Hidvbgi, E. J., 194(323, 324), 198(324), 981 Hietbrink, B. E., 205(341), 213(341), 222 Higasi, K., 332(381, 396 Highton, P. J., 406(300),434 Higuchi, S., 269(138), 271(146), 272(146), 273,999 Hillova, J., 118(107), 124(107), 916 Hilz, H., 118(86), 120(86), 125(86), 151 (861, 155(86), 158(86), 165(86), 205 (861, 213(86), 216 Hirschfelder, J. O., 355(185), 6999 Hirschman, S. Z., 245, 277(83b), 297, 341 (86), 597 Hishizowa, T., 186(305), 197(305), 991 Hnilirn, I,., 286(171), 300

AUTHOR

INDEX

Ho, N. W., 413(105), 436 Ho, P., 76(4), 118 Hoard, D. E., 412(88), 136 Hodes, M., 410(64), 436 Hofelich, F., 341 (1021,997 Hoffer, M., 234(43, 44), 296 Hoffman, T. A., 341(153), 598 Hoffmann, R., 338,396 Hofschneider, P. H., 83(69), 119 Holand, J. J., 182(289), 921 Holcomb, D. N.,249(91), 252(95), 297 Holiday, E. R., 350(172), 399 Holland, J., 194(323, 324), 198(323, 3241, 221

Holiey, R. W., 278(154), 299, 405(8), 412 (841, 415(84, 4% 435 Hollis, D. P.,238(49), 896, 341(142), 369 (1421, 374(142), 598 Holmes, D. E., 375(263), 401 Holmes, R., 415(128), 436 Holoway, B. W., 133(145), 136(14!i), 217 Holy, A., 410(55, 56), 436 Holewarth, G. M., 229, 230(25), 281(161), 996, 999 Honig, B., 330(28), 3411281, 355(28), 396 Honikel, K. O., 110(167), 116 Hoogsteen, K., 357, 699 Hooper, C. W., 273(148, 148a), 999 Horn, E. E., 14(88), 18(88), 99 Hory, K., 143(175), 218 Hoskinson, R. M.,405(12), 434 Hotz, G., 118(102, 102a), 121(102), 123, 149(102a),916 Howard, A., 118(39), 120(39), 156(202), 160(39), 216,918 Howard, B, D., 15(118), 29, 390(299), 409 Howard-Flanders, P., 118(13, 91, 971, 119 113, 911, 120(13), 123(13, 91),151(13), 154, 910(13), 213(13), g14, 216 Howsden, F. L., 118(110, l l l ) , 124(110, lll), 125(110, 1111, 126(110), 127 (110), 216 Hoyer, B. H., 317,325 Hradecna, Z., 95(116), 103(116), 115 Huang, P. C., 76(14), 119 Huang, R. C.,168, $20 Huchmann, J., 413(98), 436 Hudnik-Plevnik, T., 118(106), 124(106), 133(138), 142(158), 151(161), 174

AUTHOR

449

INDEX

(138), 175(138, 278, 279), 176, 177, Janik, B., 38, 39(27, 35), 40(35, 42, 43), 41(27, 42), 42(43, 46,47,481,43(46), 180(279), 183,195(330), 196(330), 197, 66(46), 71 198(278, 330), 213(278), 216,217,220, Janion, C., 6(27), f l , 416(152), 429(186), 222 430(186), 431(183, 185,1861,432(152, Hudson, C. S., 234(42), 296 1861,433,437,438 Humlovh, A.,43(51), 45(51), 72 JankoviE, V.,165(249), 166(249), 220 Humphrey, R.M.,156(209), 157,219 Jaskunas, S. R., 254(108a), 277(108a), 278 Huong, T., 428(1771,@8 (108a,156), 279(108a), 298,299, 341 Hurwitz, J.,76(6, 121,79(39), 80(51, 58), (1241,374( 1241,398 81(58), 82,83(58), 84(6), 88(39, 931, 89,92(39), 94(113), 95(117), 96(124), Jehle, H., 341(90, 91), 397 97(39), 98, 101(113,137), I04(39), 105 Jencks, W. P.,373(224), 400 38,71 (39,155), 109(155), 112,113,114, 116, Jenkins, I. L., Jennings, J. P., 233(35), 296 116,170(260), 220 Jensen, R.,258(118), 298,309(30), 325 Huston, D. C., 138(152), 217 Hutchinson, P., I18(8), 214,2730491, Jensen, L.H.,238(53), 296 Jirgensons, B., 286(171), 300 299 Johns, H.E., ll(68, 69, 70), 12(68), 15 (68),28, 391(303), 402, 438 I Johnson, E. A.,35011721,399 Ibuki, F., 45(60), 72 Jones, A.,411(66, 671,415(126), 496,4% Ijlstra, J., 377(264, 265, 2661,401 Jones, D. S., 102(140), 116 Imahori, K.,253,269(139), 270(139), 280, Jones, 0. W., 90(98), 92(105), 99(98, 1341, 298,299 100(98), 102(98), 105(98), 106(134, Imamoto, F., 78(44, 45),113 156), 108, 110(98), 114, 116 Imamura, A.,330(17, 21), 341(17, 76, 133, Jones, R. Norman, 364(208), 400 134,151), 354(76), 374(245, 2461,385 Jordan, D.O., 341(IM), 374,997 (1511,596,597, 598,401 Jordan, F.,331(31), 341(31), 374(31), 383 Ingalh, R. B.,375(263), 401 (282),396,401 Ingram, V. M., 405(5, 6),413(103), 433, Jorgenson, G., ll8(66), 151(66), 213(66), 456

Inman,

R.B., 143(172, 173), 918, 258(119,

1201, 298

Inoue, Y., 245(83a), 246,297 Inouye, M.,79(40), 89(40), 115 Isaars, L.N., 78(25), 112 Isenberg, I.,374(232,247), 400, 401 Ishihama, A.,80(57), 113,170(261), 210 Ishimoto, M.,15(127), 29 Ishiwa, H., 14(125), 29 Isupova, L.S.,133(142), 217 Ito, E., 415(122), 436 Ivanova, O., 405(22), 413(22), 434 Twai, I., 231(34), 237(34), 238(34), 296

216 Josse, J., 31(1), 70

Jovih, D.,158(219), 211(219), 213(219), 219 Jovicki, G., 165(249), 166(249), 220 Joyner, A.,78(25), 112

K

Kabat, S., 76(14), 112 KaFanski, K.,118(44), 120(44), 616 Iiafiani, K. A.,194,212 Kaga, M., 187(312), 121 Kagi, J., 231(32), 249(32), 262(32), 268 (32),29G J Kahan, F., 415(129), 436 Jacob, F., 77(15, 161,112, 210(352), 22.2 Kaiser, A. D., 306(22), 310(22), 3% Kakefuda, T., 210(254), 2% Jacobson, B., 157(214), 819 KalLb, D., 45(64, 65, 66,67, 68,69,70), Jaffk, H.H., 332(39), 396 72 James, T.W., 289(191, 192), 300

450

AUTHOR

INDEX

Kalinin, V. L., 14(91), 15(91), 18(91), 19 Kikugawa, K., 416(148), 437 Kim, J. H., 185(304), 221 (91), 20(91), 21(91), 25(91), 29 Kim, S. XI., 185(304), $21 Kallenbach, N.R., 306, 309(34), 326 Kimball, R.F., 118(119), 126(119), 217 Kalvoda, R.,37(20, 21), 38(22, 23), 71 Kameyama, T.,80(57), 113, 167(254), Kimura, M., 319, 320(46), 326 170(261), 173(273),205(254),213(254, Kin-Ichiro Miura, 183(295), 221 Kirk, J. M., l09(161), 116 273), 220 Kiselev, L.L., 428(176), @8 Kamiya, T., 82(66), loo(%), 113 Kanazir, D., 118(30,32, 106, 115), 120(30, Kiseleva, N.P., 284(167), 300 32), 124(106), 126(115), 1331321, 151 Kishimoto, S., 157(212), 18612121, 187 (309),219,221 (32), 173(32), 174(32), 199(334), 200 (334), 201(335), 202(335), 203(337), Kita, M., 238(58),996 Kjeldgaard, N. O.,76(8), 78(8), 79(49), 213(30, 321,$14,216,217,222 112,113 Kanmir, D. T., 165(249), 166(249), 190 (318), 191(318), 195(330), 196(330), Klamerth, O., 173(267), 175, 178, 213 (2671,,220 197(318), 198(330), 213(318), 220,221, Klebanova, L., 406(25), 410(59), 434,436 228 KIee, W. A,,235,296 Kanner, L. C., 94(lll), 95(111), 124 Klein, G., 118(35, 361, 120(35, 36), 160 Kano-Sueoka, T.,271(145), M9 (35,36), 214 Kaplan, H.S., 118(17, 54, 55, 56, 63, 68, 110, 111, 112), 119(17), ZO(17, 54, 55, Klein, R., 416(146), 437 56, 63, 681, 123(17), 124(110, 111, Kleinschmidt, A. K., 87(89), 114 112), 125(17, 110, 111,1121,126(110, Kleinwachter, V.,350(179), 399 112), 127, 133(17), 137, 138, 140, 151 Klimenko, S.,412(95), 436 (17,54, 55, 56, 63, 681,207W81, 210 Kline, B,,239(62), ,997 (17), 213(17, 54, 55, 56, 63, 68),214, Klouwen, H.M., 155(200), 164(200), 218 Klyne, W.,233(35), 296 215,216,222 Knijnenburg, C.M., 88(92), 114 Kasinski, H. E., 44(59),72 Knorre, D.,405(22, 35),409(38,39,40, 41, Katz, L.,357(190), 399 42), 413(22, 35, 107, 109, lll), 414 Katz, Z.,364(207), 400 (35,Ill), 434,436 Kaudewitz, F.,14(120), 29 Kaufman, B. N., 118(34), 120(34), 160 Kniisel, F., 110(167,168), 116 Kochanski, E.,332(32), 341(32, 791, 354 (341,214 397 (791,396, Kawase, S.,269(140), 299 Kochetkov, N. K., 6(24), 7(33), R,405 Kay, C.M., 262(126), 298 (19, 33), 406(25), 413(99), 414(114), Keck, K.,118(88), 144(88), 145(88), 148 417(157), 418(159), 419(159), 420 (881, 167(88), 169(88), 213(88), 226 (159), 421(161), 423(164, 165, 1681, Keir, H. M., 143(166), 218 424(168), 425(33, 159), 426(33, 1701, Kellenberger, E.,78(21), 106(21), 119 427(1&4, 165, 170, 172, 173, 1741, 428 Kelly, G.W., 194(327), 222 (164, 165, 172), 431(187), 432(187, Kelly, L. S., 118(41), 120(41), 160(41, la), -434, &6,437,4% 228), 216,919 Kelner, A.,118(29), 120(29), 213(29), 214 Kodama, M., 330(17), 341(17), 374(245, 246), 330,401 Kestner, N.R., 373(221), 400 Kodoya, M.,99(128), 116 Khesin, R.B., 96(120), 116 Khorana. H.G., 102(140), 126,305(16), Koga, M.,156(211), 186(211), 219 6(17), 6(17), R,417(156), 307(24), 526,405(12), 409(37, 52, 53), Kohlhage, H., 418(156), 437 434,@6 Kohne, D. E., 302, 317, 318, 319(5), 321, Khuong-Huu, Q., 288(188), 300 Khym, J. X.,409(47), 436 324

AUTHOR

451

INDEX

Kollin, V., 79(37), 113 Kondo, N. S., 370(217), 400 Konrad, M. W., 79(38), 87(86), 88(38), 89, 90(86), 92(86), 98(38), 99(133), 104(38), 105(86, 133), 106(38, 133), 107(86), 110(166), 113, 114, 116,116 Konstantinova, V. V., 133(140), 617 Kopama, M., 341(133), 398 Koranda, J. J , 118(871, 213(87), 216 Kornberg, A., 5(15), 16(80), 27, 28, 95 (114), 114,143(165, 169, 170, 171, 172, 173), 218, 259(120), 298,309(31), 325, 415(131), 436 Kornberg, R. D., 99(130), 105(130), 115 Kornhauser, A., 15(127), 29 Kos,E., 174(265), 220 Koscheenko, N. N., 118(4), 21.4 Kostjanovsky, R., 406(29), 434 Kotaka, R., 14(81), 17, 19(81), 21(81), 28

Kotaka, T., 412(82), 435 KrajinFaniE, B., 118(106), 124(106), 216 Krakow, J. S., 93(107), il4 Kraut, J., 238(50,53), 296 Kreuckel, B., 160(228), 219 Krieg, D. R., 1(1), 13(1), 14(115), 16(1), 27,29, 383(286), 402 Kriek, A. K., N37, 38),27 Kritskii, G. A., 118(43), 120(43), 133 (1391, 165, 225,217 KrSger, H., 118(88, %a), 144(88, %a), 145(88), 148(88), 167(88, 88a), 169 (88, 88a), 172(88a), 213(88, 88a), 816 Kroes, H. H., 174(277), 220 Kruglyak, Yu A., 341(69), 396 KrupiFka, J., 43(50), 45(62), 7 2 Krutilina, A. I., 405(10), 434 Kubinski, H., 102(144, 145, 146), 103(145, 146), 115 Kubitshek, H. E., 15(122, 123), 29 KuFan, Z., 54(93), 73, 118(59, 60, 61, 701, 120(59, 60, 61, 701, 138(59, 60,61, 70), 151(59, 60, 61, 70), 173(59, So), 175 (59, 60, 611, 182(291), 213(59, 60, 61, 70), 115, 221 Icumar, S., 14(125, 126), 29,431(184), 438 Kung, H., 278(155), 299, 405(11), 434 Kuprievich, V. A., 330(20), 341(20), 596 Kurland, C. G., 78(30), 110(30), l l d Kfita, J., 36(9), 39(9), 7 1

Kuzin,A. M., 118(3, 45, 121), 120(45, 1211, 128(121), 130(121), 133(144), 135(3), 138, 155(197), 1&4(197), $14, 217,218 Kwiatkowski, J. S., 341(74, 751, 350(74, 75), 354(74, 75), 397 Kyogoku, Y., 269(138), 299, 357(195), 364(195, 2051, 399, 400

1 Labana, L. L., 361(201), 400 Labaw, L. W., 118(28), 120(28), 150(189), 213(28), 214, 218 Lacroix, M., 341(129), 598 Ladik, J., 329(13), 330(18), 341(13, 153, 154, 155, 156), 396,398,399 Laipis, R., 88(90), l i d Laird, C., 118(49), 120(49), 121(49), 135 (49), 213(49), 816 Lajtha, L. G., 118(42, 46), 120(42, 46), 155(42), 156(203), 216,118 Lakshminarayanan, A. V.,238(51), 296 Lamb, B., 39(31), 42(31), 71 Lamborg, M. R., 231, 249, 262(32), 268 (32), 296,298 Lamola, A. A,,382, 4ffl Landy, A., 405(13), 434 Lane, B., 410(60, 61), 436 I,ane, D., 118(117), 126(117), 817, 317 (41),325 Langridge, R., 86(83), 103(149), 114, 115, 259(123), 269(137), 272(151), 273 (149a, 150), 2998,299 Lapthisophon, T., 118(72), 120(72), 151 (72), 153(72), 154(72), 213(72), 216 Larkiewicz, Z., 118(108), 124(108), 216 Laser, H., 118(14), 119(14), 120(14), 124 (14), 151(14), 210(14), 213(14), 814 I,atarjet, R., 118(19, 25, 261, 120(25, 261, 151(19), 211(19), 212(19), 213(19, 25, 261,214 Lavik, P., 118(37), 120(37), 160(37), 215 Lawley, P. D., l(41, 61231, 8, 13(4), 16 (4), 20(4), 21(4, 100, 1031, 22(106), 27,29,374(249), 385(290, 291, 292), 392(309, 310, 311, 312, 313, 314, 3151, 393(318), 4ff1, 402, 412(76), 455,429, 438 I,awrence, M., 86, il4 Lazar, J., 240(68), 297

452

AUTHOR

INDEX

Limperos, G., 133(143), 217 Lazurkin, Yu.,412(96), 436 Lin, C. Y., 231, 238, 296 Lazurkin, Y. S., 286(178a), 288, 300 Lindahl, T., 270(142), 271(142, 1431, 299 Lea, D. E., 118(5, 61, 214 Leach, W. M., 158(224), 211(224), 213 Lindblow, C., 231(31), 254(31), 262(124), 296, 298 (224), 219 Le Blanc, J. C., 11(68), 12(68), 15(68), Lingens, F., 9(47, 52), 15(47, 52), 23(47), 28, 416(144, 1451, 4.37 28,391 (3031,402,438 Lipsett, M. N., 241(76), 297,289(196), Lebowits, J.,88(90), 114 SOO, 305, 326 Lederberg, J., 210(351), 2.92 Liquori, A. M., 374(242), 401 Ledoux, L., 165(245), 213(245), 219 Litaka, Y ., 269 (138), 273(150a), 299 Lee, T., 157(214), 219 Legrand, M., 224(4), Z25(4), 230(4), 296 Litman, R. M., 12(76), 14(90), 15(76), 18 (901, 19(76), 23(90), 28, 29 Lehman, I. R., 5(15), 27, 143(165, 1701, Little, J. W., 309(33), 3.96 218, 415(131), 436 Livingston, D., 409(48), 436 Leidy, G., 18(85), 89,4120371, 436 Lodeman, E., 15(127), 89 Leive, L., 79(37), 118 Lowdin, P. O., 341(115, 116, 117, Leloir, L. F., 415(121), 4.36’ 388(116, 117, 118), 398 Leng, M., 240(63), .W7 Logan, R., 185, 205(301), 3.91 Lengyel, P., 5(16), 14(16), 97 Lofroth, G., 375(255), 4Ol Lemieux, R. V., 234(43), 296 Lerman, L. S., 10(53), 28, 109(164), 116, Lohman, K., 412(78), 436 286(176), 300,374(226, 227,228), 400 Looney, W. B., 162(!233), $19 Loveless, A., 9(45), 14(455),28 Lesk, A. M., 224(8), 225(8), 896 Lowney, L. I., 79(34), 112 Leslie, R. B., 374(232), 400 Lowy, B. A., 40,71 Letham, D., 411(66, 671, ,436 Lloyd, D. A., 243(79), 297 Lethbridge, J. H., 21(103), 29 Lett, J. T., 21(101), 29, 118(18, 78, 80, London, E. S., 289(190), 300 105), 119(18), 120(18, 78, 80, 105), Longworth, J. W., 382(2'78), 401 124(18), 128(78, 801,130(78, 801, 133 Lord, R. C., 357(195), 364095, 204,205), 373(223), 399,400 (147), 151(18), 154(18), 156(78, 801, 210(18), 213(18, 78, 801,214, 2f6,817, Lord Todd, 393(316), 402 Lowry, T. M., 224(1), 225(1), 296 393(319), 402 Lubin, M., 83(72), 93(106), 113, 114 Levedahl, B. H., 289(191, 192), SO0 Luck, G.,268(136), ,999 Levene, P. A., 289(190), 300 Ludlum, D. B., 9(39, 40), 14(39, 40), 19 Levin, S., 373(225), /to0 (99),20(99), R,g8, 89 Levine, E., 428(177), 438 Lukhovd, E., 54(91, 921, 55(92), 72 Levine, L., 373(224), 400 Lunell, S., 341(123), 398 Levinthal, C., 109(163), 116 Lunt, M. R., 102(142), 116 Lewin, S., 412(90, 91, 92, 93), 436,436 Luria, S. E., 78(22), 109(163), 112,116 Leyko, W., 44(56, 58), 72 Luthy, N. G., 39(31), 42(31), 71 Lezius, A. G.,143(176), 218 Luzzati, V., 57(100), 73,252(98), 298 Li, L., 405(10), 434 Li, T. K., %1(32), 235, 249(32), 262(32), Lykos, P. G., 341(58), 396’ 268(32), 296 M Lichstein, H. C., 173(270), 213(270), 220 Lieberman, I., 156(211), 157(212), 186 Maal@e,O., 76(8), 78(8,30), 110(30), 112 (212), 187(309,312,314,315), 219, 2.91 McArdle, H. H., 143(166), 818 McCaffery, A. J., 286(181), 287(181), 500 Lielausis, A., 78(21), 106(21), 112 McCallum, M., 323(32,53), 326 Lifschits, Ch., 341(65), 349(65), 396 Lifsan, S., 330(28), 341(28), 355(28), 3% McCarter, J, A., 374(248), 401

AUTHOR

453

INDEX

McCarthy, B. J., 194(322), 201(338), 221, 222, 307, 308, 310(37), 311, 317(44),

S26 McConaughy, B. L., 307,308,326 McDonald, C. C., 240(68), 297 McFlya, A. B., 156(210), 164(210), 219 McGinn, F. A., 39(32), 71 McGlynn, S. P., 339(53), 3% McGrath, R. A., 118(16, 691, 119(16), 120 (16, 69), 123(16), 134(16), 137, 138, 139, 151(16, 69), 154, 158(221, 2241, 210(16), 221(221, 2241, 213(16, 691, 213(221, 224), 214, 216, 217, 219 MeGuire, J., 15(117), 29 Maehmer, P., 349(166), 399 McKinney, L. E., 374(241), 401 McLachlm, A. D., 341(85), 597 McLaren, A., 302(4), 309(32), 317, 322, 324, 325, S26 McLaren, J., 412(74), 431(74), 435 McMullen, D. W., 278, 299 McNaught, A., 416(142), 437 McPhie, P., 279(156a), 280, 282(158), 299 McQuillen, K., 79(33), 112 Madison, J. T., 278(154), 299, 405(811),

434 Maestre, M. F., 283, 284, 285(164), 999 Magasanik, B., 167(253), 205(253), 213 (253), 220, 267(131), 298 Mahler, H. R., 181(285), 220, 239, 288, 297, so0 Maidlovh, E., 45(72), 72 Main, R. K., 144(179), 145(179), 163 (179), 218 ‘Maisin, J. R., 165(245), 213(245), 219 Maitra, U., 79(39), 80(58), 81(58), 82, 83 (58), 88(39, 931, 89, 92(39), 96(124), 97(39), 98, 104(39), 105(39, 155), 109 (155), 113, 114, 116, 116 Mak, S., 156(205), 213(205), 219 Malamy, M., 76(12), 112 Malcolm, D., 323(54), 926 Maling, B., 304(13), S26 Malrieu, J. P., 341(77), 354(77), S97 Malygin, E., 405(22, 35), 413(22, 35, 107, 1111, 414135, I l l ) , 434, 436 Malyshev, A., 406(28), 434 Malysheva, A., 409(39), 434 ManEiC, D., 118(44), 120(44), 216

Manclel, M., 9(51), 15(51), 24(51), 28 Mnndel, P., 76(5), 110(165), 112, 116, 118 (471, 120(47), 615 Mandell, J. B., 9(50), 15(50), 28 Mangiorotti, G., 79(35), 112 Mann, D. E., 339, S96 Maooukk, O., 42(49), 72 Mantione, M. J.. 3411114, 131, 132, 145, 1461, 374(114), 375(146), 377(131, 1321, 382, 398 Mantsavinos, R., 143(168),218 Marcker, K. A., 98(127), 116 Marinova, Z., 145(183),918 Markham, R., 410(57), 411(57), 436 Marmur, J., 118(117), 126(117), 217, 267 (133), 272(151), 298, 299, 301, 303, 304(7a, 7b), 316, 317(41), 324, 326, 373(220), 400 Marquisse, M., 278(154), 299, 405181, 434 Marsh, R., 359(198), 400 Marshall, R. E., 320(48), S26 Martinez, A. M., 82(68), 113 Marvin, D. A., 273(148, 148a), 299 Masin, A., 405(18), 416(18), 4 4 Mason, S. F., 286(181, 182), 287(181, 182), 300, 350(171), 351(171), 374 (2331, ~99,400 Masson, F., 57(100), 73, 252(98), ,998 Massoulie, J., 63(104), 64(104), 73, 249 (92), 297 Mathews, F. S., 357(191, 1921, Y99 Mathis, A,, 57(100), 73, 252(98), 298 Mataudaira, H., 186, 197,211 Matsushita, S., 45(60), 72 Matthews, L., 118(117), 126(117),217 Mauger, A. B., 288(187), 300 Maunzot, J. C., 238(56), 241(56), 242(56), 243(56), 249(87a), 252(56), 254(87a), 255(87a), $96 Mautner, H. G., 341(72), 396 Maxwell, Ch. R., 202,292 Mayneord, W. V., 206(345), %1 Mehrotra, B. D., 239(62), 297 Meites, L., 36(10), 71 Meljnikova, H. A., 194(328), 222 Mellema, J., 416(151), 437 Melvin, I. S., 240(69), 297 Melzer, H. S., 432(192), 438 Mennigmann, H. D., 118(114), 126(114),

216

454 Menael, C., 143(176),818 Merrill, S. H., 278(154), 899, 405(8), 494 Meselson, M., 267(132), 298 Metlas, R., 201, 202,288 Meta, E., 143(176), 818 Meyers, D. K., 155(193), 156(193), 157 (1931, 158(193), 162, 165, 186, 918 Michaelis, L., 286(177), 300 Micheel, F., 411(70), 436 Michelson, A. M., 4(14), 9(41, 491, 14 (411, 17(49), 19(41), 22(49), %7', 28, 63(104), M(104), 73, 238(56, 67), 241 (56), 242(56), 243(56), 247(57), 248 (571, 249(87a, 90, 921, 252661, 254 (87a,108, 255(87a, 108), %6(108), 286(90), 296, 897, 898, 305, S26, 354 (184), 399, 415(132), 436 Micka, K., 38(24), 71 Miles, D. W., 231, 996 Miles, H. T., 31(2), 70, 145(181), 818, 249 (93),897,357(195), 364(194), 399,361 (200),400, 415(133), 436 Miletii., B., 118(59, 60, 611, 120(59, 60, 61), 138, 151(59, 60, 611, 158(219), 173(59, 601, 175(59, 60, 611, 182(290), 211(219), 213(59, 60, 611, 213(2191, 816, 219, 881 Miller, I. R., 32(4, 61, 45(80), 48(80), 48 (82,83), 66(86),70,78 MiIIer, J.H., 359(197, 2061, 399, 400 Milles, N., 415(134), 437 Millette, R. L., 80(53), 90(97), 92(100), 93(97), 110(97), 113,114 Mihan, G., 86(83), 114, 273, 899 Milosavljevih, A., 186(306), 221 Milogevib, M., 199(334), 200(334), 928 Mirsabekov, A. D., 405(10), 415(124), 434, @6 Mitsui, H.,%(I%), 116 Mitsui, Y.,269(138), 899 Mittal, J. P., 382, 401 Miura, K., 269(138, 139, 140), 270(139), 299, 416(148), 437 Miauno, S., 111(171), 116 Moffat, S., 409(51), 436 Moffitt, W., 227, 228(22), ,996 Moldave, K , 92(103), 114 Mommaerts, W. H. F. M., 57(99), 73, 225(12, 13, 14), 262(14), 267(14), 274, 279, 296

AUTHOR

INDEX

Monastirskaya, G., 6(24), "(241, 27, 428 (178), 429(178), 430( 180), 431(1&0), 432(178, 188, 189, 1901, 438 Monny, C., 4(14), 27, 252, 898, 305, 325 Monod, J., 77(15), 112 Moohr, J., 95(118), 116 Moore, J. L., 118(87), 213(87), 216 Moore, T. A., 341(1601,399 Mori, J. F., 185, 205(302), 821 Morita, T., 185, 205(302), 8'21 Morosova, T., 406(32), 434 Morris, D. W., 79(49), 113 Moscowita, A., 225(18), 228(18, 22), 229

(It?), 996 Moseley, B. E. B., 118114, 104), 119(14), 120(14), 124(14), 151(14), 210(14), 213(14), $14, 216 Mosher, W. H.,133(143), 817 Mosley, V. M., 118(28), 120(28), 150 (189), 213(28), 814,918 Moss, G., 409(50), 9 5 Moudrianakis, E. N., 406(26), 416(151),

434, .4sr

Mudd, 8. H., 235, $96 Mueller, G. C., 182(288),221 Miiller-Hill, B., 77(17, lS), 111(18), 11% Muench, K., nl, 899 Muller, R. L., 341(58), 396 Mundry, K. W., 4, 14(12), 15(111), 23,

R Mushinskaya G., 413007, log), 436 Muto, A., 416(148), 437 Myers, L. S., Jr., 375(263), 401

N Nadkami, G. B., 173(272),213(272), 220 Nagakura, S., 330(19), 341(73), 350(73), 354(73), 396, S996 Nagata, Ch., 330(17, 21), 341(17, 76, 133. 134, 151), 354(76, 245, 246), 385(151), 396, 397,sgs,40t Naito, T., 21 (1041, 89 Nakada, D., 167(253), 205(253), 213 (2531, 820 Nakajima, T., 329(6, 7, S ) ,341(6, 71, 392 (61,396 Nakamoto, T., 84, 85(75), 113 Nakamura, Ch., 186(305), 197(305), 221 Nakanishi, K., 245(83a), 246(83a), .897

AUTHOR

INDEX

455

Okada, S., 118(122), 120(122), 130(122), NaBata, Y., 105(155), 109(155), 116 144(180), 145(180), 163(180), 187 Nancy, W., 405(21), 413(21), ,434 (3071, 217,218,221 Nandi, U. S., 258(118), 298,309(30), 326 Naono, S., 77(16), 78(27), 79(43, 481, 96 Okada, Y., 79(40), 89(40), 113 Okun, L., 118(49), 120(49), 121(491, 135 (123), 112,113,116 (491, 213(49), 216 Nash, H. A., 341(89, 1571, 356(891, 394, Oliver, R., 118(42), 120(42), 155(42), 216 397,399 Olivera, B. M., 258(118), 298, 309(30), Naylor, R., 405(21), 443(21, 108),434,436 326 Neidhardt, F. C., 76(7), 112 Olner, R., 156(203), 218 Neifan, A. A., 194(328, 329), 22.2 Olson, A. C.,240(69), #7 Nelson, J., 4121841, 415(84), 436 Oltmanns, O., 9(52), 15(52), 28 Nesbet, R. K., 330(16), 3411161, 596 Negkovih, B., 128, 156, 157(207), 186 Ono, J., 11, 12, 15(73), 28,171, 220,391 (301, 306), 402 (306), 219,221 Neville, D. M., 286(178), 300, 374(236), Opara-Kubjnska, Z., 102(144), 103(145), 116, 118(10), 119(10), 120(10), 124 400 (lo), 125(10), 151(101, 210(10), 213 Newton, J., 79(40), 89(40), 113 (lo), 214 Ney, H., 409(45), 434 Ord, M. G., 118(40), 120(40), 155(40), Nikolik, S., 186,221 160(40), 163, 165(247, 247a), 166 Nirenberg, M., 320 (481, 526 (247a), 216, 220 Nishimura, S., 102(140), 116,183(292), Ordy, J. M., 206(343), 222 221 Nishimura, T., 231(34), 237, 238(34, 551, Orgel, A,,286(174, 1751, 300,385(293), 296 409 Orgel, L. E., l(21, 13(2), 16W, 27, 118 Nitta, K., 111(171), 116 (%I), 212(24), 212(24), 214,383(289), Niyogi, S. K., 307, 308, 310(27), 326 40.2 Norman, A., 150(184), 218 Novak, D., 118(61), 120(61), 138(61), 151 Oriel, P. J., 269(141), 285, 286(169), 299, 300 (61), 175(61), 213(61), 216 Novelli, G. D., 1670541, 173(273), 205 Ormerod, M. G., 118(78), 120(78), 128 (781, 130(78), 156(78), 213(78), 216 (254), 213(254,273), 260 Osawa, S., 99(128), 116 Novogrodsky, A., 88(93), 114 Noyes, W. D., 118(42), 120(42), 155(42), Oshinsky, C. K., 309(33), 326 Otaka, E., 99(128), 116 216 Nuesch, J., llO(167, 168, 1691, 111(169), Oth, A,,267(134), 29.8 116 P Nutwell, D., 202, 222 Paduch, V., 1 1 8 ( 8 6 ) , 120(86), 125(86), 151 Nygaard, A. P., 90(99), 114,301(2), 324 (86), 155(86), 158(86), 165(86), 205 Nygaard, 0. F., 118(8l>, 120(81), 158 (86), 213(86), 216 (223), l60(81), 161, 162, 163(230), Paigen, K., 85(79), 114,118(34), 120(34), 211(223), 213(223), 216,219 160(34), 914 Painter, R. B., 118(76, 771, 120(76, 77), 0 128(76, 771, 130(76, 771, 155(76), 156 (76, 77), 157(76), 158, 184(76), 211 Obuchova, L., 413(106), 436 (215), 213(76, 77, 215), 616 Ochoa, S., 5(16), 14(16), 2’7 Paladini, A. C., 415(121), 436 Ogur,M., 409(46), 434 PnleFek, E., 38(18, 19), 39(33), 40(19, 33, Ohno, K., 332(43), 396 39, 42, 43), 41(33, 39, 42, 441, 42(33, Ohtsuka, E., 305(16), 326 39, 43, 46, 471, 43(33, 44, 46), 45(33, Oikawa, K., 262(126), 998

456 75, 76, 77, 78, 79), 46(18, 19, 75, 76, 77, 78), 48(75, 771, 49(75), MF(18, 75, 761, 51(19), 52(19, 75, 77), 53(19, 44, 87, 88, 89, 901,54(44, 79, 901,55(79), 56(18, 75, 78, 90, 94, 951, 57(75, 77, 781, 58(102), 59(75, 951, 60(18, 19, 76, 79), 61(103), 63(19), 64(19), 66 (19, 33, 461, 67(19), 68(19, 76, 87, 88, 901, 71,72,7 3 Palm, P., 104(152), 110(152), 116 PmteliE, M., 186(306), 881 Panusz, H., 44(56), 72 Pardee, A. B., 167(252, 2551, 205(252, 255), 213(252, 255), 8.20 Pardue, M. L., 162(233), 819 PGizkov6, H., 45(61), 72 Parke, W. C.,341(90,91), 397 Parkins, G. M., 21(101), 29,118(78), 120 (78), 128(78), 130(78), 156(78), 213 (781, 216,393(319), 402 Parr, R. G., 332(41), 396 Patel, A. B., 416(147), 437 Patrick, M. H.,118(65), 120(65), 151(65), 213(65), 216 Patten, R. A., 375(254), 401 Patterson, D. L., 304(12), 3% Peacocke, A. R., 133(149), 817,280,286 (179, 1831, 287, 299,300, 317 (431, 326 Pesevski, I., 182(290, 291), 881 Pelc, S. R., 156(202), 818 Penistan, 0. P., 39(28), 71 Penman, S., 364(207), 400 Penniston, J., 412(94), 436 Penswick, J. R.,278(154), 899,405, 434 Perevertajlo, G. A.,57(97), 73 Permogorov, V. I., 286(178a), 288, 300 Perrault, A. M., 341(159), 399 Perry, R. P., 182(287), 281 Perumov, D. A., 14(91), 15(91), 18(91), 19(91), 20(91), 21(91), 25(91), 89 Peschel, G. G., 350(176), 399 Peter, H. H., 240(67), 897 Peters, E., 165,280 Petemen, D. F., 118(38), 120(38), 160(38), 187(308), 216,221 Petersen, E., 118(88a), 144(88a), 167(88a), 169(88a), 172(88a), 213(88a), 216 Petersen, G. B., 102(142), 115,405(17), 411 (681,434,436

AUTHOR

INDEX

Petrovib, D., 158(219, 220), 211(219, BO), 213(219, 220), 919 PetroviE, J., 192(321), 193(321), 281 PetroviE, 5. P., 165(249), 166(249), 190 (3181, 191, 192, 193(320, 321), 197, 213(318), 880,221 Petters, V. B., 194(327), 288 Pettijohn, D., 82(66), 100(66), 113, 118 ( l l ) , 119(11), 120(11), 123(11), 151 ( l l ) , 153, 210(11), 213(11), 214 Pfitzner, K., 409(51), 436 Phil, A., 153, 158(225), 213(225), 218, 219 Phillips, D. M. P., 286(172), 300 Phillips, J.,428(175), 429, 433(195, 197), 438

Phillips, J. H., 1(6), 6(20, 21, 29), 7(6, 291, 14(21,29), 27,391(302,304,305), 402 Phillips, W. D., 240(68), 897 Piekala, A., 341(164), 399 Pitha, J., 341(164, l a ) , 364(208), 399,400 Pithova, P., 341(164), 364(208), 399,400 Pitot, H. C., 204(339), 205(339, 3401, 213 (339,3401,322 Pivec, L., 57(98), 73 Pleticha, R., 45(63),72 Pochon, F., 17, 88,256,298 Podder, R. K., 14(126), 89,431(184), 438 Pohl, S. H., 4(11), 18(11), 27 Poland, D., 247, 297 Pollak, M., 341(99,100,101), 397 Pollard, E. C.,118(8, 62, 64,73, 89, 90), 120(62, 64), 138(152), 139, 141, 142 (1541, 150(188), 151(62, 64,1541, 152, 173(269), 174(90), 175(62), 210(355), 211(90), 213(62, 64, 73, 154, 2691, 814,215,816,817,8.0, 284, 899 Pons, S., 199(333), 822 Pople, J. A., 340(55, 56, 57), 396 Postel, E. H., 14(89), 18(89), 29 Potter, R. L., 118(81,82), 120(81,82), 160 (21, 82), 161, 162, 163(230), 216 Potter, V. R., 156(210), 163(M5), 164 (2101, 219 Pouwels, P. H., 88(92), l l 4 Prestidgp, L. S., 167(252, 255), 205(252, 255), 213(253, 255), 820 Preston, B. N., 133(149), 817 Preuss, A., 83(69), I13

AUTHOR

457

INDEX

Priess, H., 82(64), 83(64), 113,414(113), 436 Printz, M. P.,32, 57(3), 70, 101(136), 116 Pruden, B., 375(258, 2591,401 P ~ S O R ,w. H., 379(273),401 Pullman, A.,328(l, 231,329(1, 4,5,6,9, ll), 330(22, 23, 24, 251, 331(32, 33, 341, 332(1, 40,44), 338,341(1, 6,22, 23, 24,25, 26,27, 32, 33, 34, 63,64, 67,68,71, 78, 79, 113,125,130, 144, 150, 161, 162, 163), 346(22, 26), 347 (26,341,3491221,350(22, 23,24,25), 354(22, 23, 24,25,78, 791,361(202), 370(202), 375(125), 382(22, 23,24,25, 78), 383(22, 24,78, 150), 385(1, 2,6, 2951,387(67), 390(300), 392(1, 6,631, 393(150, 300,321), 396,396,397,398, 399, 400, 402 Pullman, B., 328(1, 23), 3290,4,5,7,8, 9), 331(31), 332(1), 341(1, 31,59,60, 61,62, 66,66, 67,70, 92,93,94,95, 96,97,98,104,105,113,125,126,130, 131, 132, 136, 137, 138, 140,143,145, 152, 158, 159, 161,1621, 349(62, 65, 661,359(92), 361(94,96,202),365(98), 367(96), 369(93,97), 370(93,202),373 (93), 374(31, 104,105), 375(125, 126, 146), 377(131,132), 382,383(136,282), 385(1, 295), 387(67), 388(297), 390 (300), 391(61), 392(1), 393(300, 320, 322), 396, 396,397,39S,399, 400,401,

Rapaport, S. A.,11(69), 28, 244(80), 249 (801, 254(80), 257(80), 259(80), 260 (SO),261(80), 274(80), 297 Rash, E., 187(311), 291 Rasmussen, R. E., 118(77), 120(77), 128 (77), 130(77), 156(77), 158,211(215), 213(77,215), 216 Ratliff, R. L., 80(56), 82(68), 83(56), 113, 181(286), 221 Rebhum, L.,187(311), 221 Reese, C.B., 393(316), 402,409(50), 436 Reeves, J.,405(17), 434 Reggiani, M.,48181),72 Reich, E.,109(160), 116 Rein, R.,339(54), 341(99, 119, 120,121, 122), 396, 397,398 Reiter, H., 118(103), 123(103), $26 Rembaum, A.,332(38), 396 Resnik, R.A,,286(180), 287,28S,300 Reuschl, H., 118(102), 121(102), 216 Revel, M., 92(104), il4 Reybeyrotte, N.,ll8(19), 151(19), 211 (191,212(19), 213(19), 214 Reynolds, J. W., 339(52), 340(52), 396 Rhaese, H.J.,14(95), 15(95), 19(95), 21 (95),25(95), 29, 432(192), 437 Rhodes, W., 341(81, 821,397 Rich, A,, 76(10), 85(81), 11.3,114,249 (89), 252(99, 101,102,103), 257(114, 116,117), 297, 298,357(190, 191,192, 193, 195), 359(199), 364(204, 205), 374(252), 399,400,401 Richards, E. G., 249,252(100), 297,698 409 Pustoshilova, N., 409(38, 39,421,434 Richardson, C .C., 950141,11.6,143(173, 1731,218 R Richardson, J.P., 80(54, 621,81(54), 82 (54, 62), 83(74), 87(62, 871, 91(54, Rxckus, J. A,,194(328), 29% 131), 92(62), 96(62), 99(62, 74,1311, Radloff, R., 88(90), 114 lOO(74, 1311, lOl(62, 131), 103(62), Rahn, R. O.,382(278), 401 104(62, 1311, 105(87, 1311, 109(87, Raimondi, D .I,., 339,396 131), llO(62, 1311,l f 3 116 , Rajalakshni, K.V., 341(128), 398 Richer, M.,341(128),398 RajBhandary, U.,409(52), 4% Riley, M., 167(252), 205(252), 213(252), RajBhandary, V., 405(12), 434 620,304(13), 335 Ralph, R., 409(53), 436 Ristov, S.,412(84), 415(84), 436 Ramachandran, G.N . ,238(51), 696 Rltchie, D. A.,15(116), 29 Ramsny, O., 436 Ramuz. M.. 76(5), 105(1!54), 110(1 54, Roberts, R . B., 76(13), 112 I&), 112,116' Robertson, F. W., 321(40),326’ Robev, S., 145(183), 218 Randerath, K., 414(515), 436 '

458

AUTHOR

Robins, R. K., 231(33), 29G, 393(317), 402 Rorsch, A., 174(276, 2771, 177(256), 220 Rolfe, R., 267(132), 298 Rolins, R., 415(128), 436 Romancev, E. F., 118(4), $14 Rosa, E. J., 351(183), 399 Rosenbaum, M., 202 Rosenberg, E., 437 Rosenbluth, J. R., 374(232), 4UO Rosenfeld, F. M., 133(133), 217 Rosenfeld, V. L., 229(23), 295 Rossetti, G.P., 370(219), 400 Rossi, M., 341(64), 396 Roth, S. S., 164(239), 187(239), 205(239), 213(239), 219 Rottman, F., 391,402 Rottger, B., 410(58), 455 Rouviere, J., 77(16), 79(48), 112, 115 Rownd, R., 303(7b), 304(7b), 325 Rudner, R., 405(14), 411(14), 454 Ruger, W., 307, 310(28), 325 Rust, P., 415(139), f l Ruet, A,,160(158), 116 Rumano, B., 198(332), 222 Rushizky, G. W., 167(252), 205(252), 213 (252), 2ZU, 245 (83),297 Ruttkay-Nedeckjr, G., 68(107, 108, 109, 110, 1111, 73 Ryter, A., 210(352), 222

S Sadron, C., 225(12), 295, 374(231), 400 Sakaki, T., 269(140), 299 Salas, M., 79(42), 89(42), 113 Salem, L., 332(42), 396 Salganik, R., 406(32, 351, 413(35, 110, 1111, 414(35, 1111, 434, 436 Salovcy, R., 375(256), 4 O l Salser, W., 78(24), 112 Salto, H., 341(151), 385(151), 598 Salyers, A,,341(90, 911,397 Samejima, T., 225(10), 231(9, 28), 232 238(58), 240(28), 251 (28), 233(28), (971, 253(94), 257(97), 258(97), 260 (97), 261(10), 262(10, 971, 264(97), 267(97), 268(97), 269(139), 270(139), 289(28), 290(97), 295, 296, 298, 299 Samorayski, T., 206(343), 29.2

INDEX

Sander, C.,254(109), 289(109, ZOO), 290 (1091, 291(109, 200), 298, 300 Sanger, F., 405(7), 434 Sanner, T., 153, 158(225), 213(225), 218, 219

Sanno, Y., 409(49), 436 Sarabhai, A. S., 79(46), 113 Sarkar, P. K., 231(27), 232(28), 233(28), 240(28), 241(75), 249, 250(94), 251 (94, 961, 252(94), 254, 255(94, 96), 256(96), 257(96), 263(128), 265(128, 129), 267(111), 268(111), 277(94, 961, 279(94, 961, 280, 281(129), 289(28), 290(94, I l l ) , 296, 297, 298 Sarnat, M. T., 94(111), 95(111, 115), 96 (122), 108(122), 114,116 Sasisekharan, V.,238(51), 206 Sato, T., 269(13!3), 299 Sauerbier, W., 150(187), 818 SaviE, D., 118(106), 124(106), 180, 210, 220 Sawada, F., 238(58), $96 Scaife, J. F., 184(298, 299), 8.91 Scarpinato, B., 48(81), 78 Schaechtler, M., 79(33), 11.2 Schafer, E., 414(115), 436 Scheit, K.-H., 410(55, 561,436 Schell, P., 6(18), 19(18), 87 Schepmm, A. M., 174(277), $20 Scheraga, H. A., 245(83), 247(85), 262 ( I n ) , 278,279, $97, 898 Scherrer, K., 85(172), 116, 195, ,222 Schildkraut, C. L.,95(114), 114, 143(171, 172), 218, 259(120), 298, 301, 303(7b), 324, S%5 Schlessinger, D., 79(35), 111 Schlingloff, G., 414(118), 436 Schmidt, H., 37,71 Schmier, I., 281(163), 299 Schneider-Bernlohr, H., 416(144, 145), 437 Schofield, K., 409(50), 435 Scholes, G., 118(21), 133(130, 131, 132), 135(21), 212(21), 213(21), 914, 217 Schramm, G., 4, ?87 Srhuster, H., 4(9), 7(31), 12(13, 751, 14 (13, 751, 18, 22, 23(13), 25(13), 97, 412(78). 455 Schweizer, M. P., 238, 240(65, 66), $96,

AUTHOR

459

INDEX

297,341(141, 142), 369(142, 211,212, Shigeura, J. T., 89(94), 109(84), 114 Shimkin, M.B.,21(104), 29 213), 370(216), 374(142), SDS, 400

Scope, Y.M., 233(35), 296 Scott, J. F., 183(294), 221 Seaman, E., 262(124), 298 Searashi, T.,118(103), 123(103), 216 Seeds, W. E., 273(148), 299 Segal, G.A.,340(55, 56,571,396 Seidel, R.,406(36), 414(116, 117, 119), 434, 436 Seifert, W.,405(20), 434 Sein, K.T., 203,222 Sekiguchi, T., 186(305), 197(305), 221 Selzer, R., 15(127), 29 Semal, M., 164(238), 187(238), 205(2381, 213(238), 219 Sentenae, A.,106(158), 109(162), 116 Serebrjany, A.,406(29), 434 Sereni, F., 198(332), 822 Setlow, J. K., 11, 12(64), 15(64), 19(98), 28,99, 151,218 Setlow, R. B., 118(8, 12,96), 119(12), 120 (12),123(12), 143,151(12, 177, 1901, 154,167(251), 170,174(274, 2751,205 (251), 210(12), 213(12, 2.51, 274), 214,216, $18, 290 Sevastyanov, A.,409(39, 42), 4.94 Shaffer, C .R., 139,217 Shamovsky, G.,409(38, 39, 40, 411,434 Shapiro, H.S.,405(14, 15), 411(14, 15), 434

Shapiro, R., 4110,ll), 18(11). 27,409(50), 412(80), 413(98, 102), 415(125), 416 (146),436,436, 437 Shastry, K. S., lO(58, 59,60), 28 Sheats, G.F., 370(218),400 Sheldrick, P.,102(146, 147), 103(146, 1471, 116 Shell, P.,6(25), 27,416(142), 417(154), 419(160), 427(160), 4281154,160), 437 Shemyakin, M.F.,96(120>,116 Sheppard, D., 77(19), 112 Sherrer, K.W., 194(322), 221 Shibaev, V. N., 6(24), 7(24), 27, 414 (114), 432(188), 436,438 Shibaeva, R. P., 6(24), 7(24, 331,27,423 (165), 427(165, 172), 428(165, 172, 1781, 429(178), 430(180), 431(180, 187), 432(178, 187, 1881, 487,438 Shields, H., 375(253), 401

Shimizu, B.,231(34), 237(34), 238(34, 551, 286

Shin, D. H., 92(103), 114 Shooter, K. V., 21(103), 29 Shoup, R. R., 357(194), 364(194), 399 Shmurak, S.X., 286(178a), 300 Shramko, 0. V.,341(691,996 Shugar, D., 6(27), 11(66), 19(97), 27,28, 29, 382(274, 275), 401,412(73, 741, 415(130), 416(152), 429(186), 430 (186), 431(74, 183, 185, 186), 432 (152,1861,433,436,436,437,438 Shulman, R. G., 375(256, 257, 279), 401 Shuster, H., 417(153), 428,437 Siebke, J. C., 102(142), 116 Siekevitz, P., 802 SimiE, M. M., 195,196,198(330), 222 Simmons, N.S., 224,281(163), 296, 299 Simms, E. S., 5(15), 27, 143(165, 170), 218, 415(131), 436 Simon, M. I.,10(55), 11(61), 15(61), 28 Simpkins, H., 249,252,297,298 Simpson, E., 118(91), 119(91), 123(91), 216

Simpson, W. T., 351(183), 899 Simukova, N., 417(157), 418(159), 419 (159), 420(159), 4’23(164, 165, 166, 168), 424(168), 425(159), 427(164, 165), 428(164, 165), 437 Sinanoglu, O.,341(110, 111, 1121, 373 (110, 111, 112, 221,2221,397, 400 Singer, B., 7(32), 9(32,48,491,lO(32, 57), ll(32, 57,63), 12(32), 13(32), 14(32), 15(32, 48,571,17(48, 49), 19(32), 21 (48), 22(32, 48,49, 105), 23(32), 24 (48), 26(32), Y7,28, 405(3), 433 Sinsheimer, R.I,., ll(65, 67), 28,84(76), 86,87(89), 102(143), 114, 116, 118 (lola), 149(101a), 21G Sippel, A.,111(170), 116 Skalka, A.,78(27), 112 Skoda, J.,45(62), 78 Skov, K.,155(193), 156(193), 157(193), 158(193), 162,165,1%), 818 Slayter, H. S., 83(73,74), 99(74), lOO(73, 74), 113 Sly, W. S.,78(25), 11.2 Small, G.D., 12,Q

460

AUTHOR

INDEX

Steinberg, C. M., 78(21), 106(21), 11% Small, J., 409(46), 434 Smekal, J., 234(47), 296 Steiner, R. F., 24(3(88), 254(88), 286(88), Smellie, R. M. S., 143(166), 181(283), 818, 297 220 Steinschncider, A., 405(4), 43s Smith, D. A., 80(56), S2(68), 83(56), 113, Stent, G. S., 78(29), 79(38), 82(67), 83 181(286), 921 (671, 85(80),88(38), 92, 98(38), 104 (38), llO(29, 67), 112,113,U 4 , 150 Smith, D. E., 37(15), 71 (185), 118, 284(168), 300 Smith, D. L., 39(34), 40, 41(45), 42(34, 41, 45), 43(34, 45), 71,349(167, 1681, Sternberger, N., 80(63), 82(63), 83(63), 99(132), 100(132), 106(132), 110(132), $99 113,116 Smith, J. D., 405(13), 410(57), 411(57), Stevens, A., SO(52, 63), 82(63), 83, 99 434, 435 (132), 100(132), 106(132), 110(132), Smith, K. C., 118(55, 561, 120(55, 56), 151 113, 116 (55, 56), 213(55, 56), 216 Smoot, A. O., 118(83), 120(83), 160(83), Stewart, R. F., 3511181, 182), 399 162(83), 163(83), 189(83), 190(83), Stocken, L. A., 118(40), 120(40), 142, 151 (161), 155(40, 195), 160(40), 163, 164 216 (195), 165(247, 247a), 166(247a), 168, Srnrt, J., 45(62), 7 2 215, 217, 218, 820 Sniper, W., 375(258, 2591, 401 Stoesser, P. R., 370(219a), 400 Snyder, L., 96(122), 97, 108(122), 116 So, A. G., 80, 90, 92, 94(61), 96(61), 110 Storrer, J., 130(125), 131(125), 217 Strack, H. B., 14(94), 19(94), 23, ,99, 433 (61), 113 Sobell, H. M., 357(188, 189, 1931, 359 (200), 438 (1971, 361(201, 2061, 374(251), 399, Strauss, B. S., 14(121), 29, 118(103), 123 (103), 816 400,401 Streisinger, G., 79(40), 89(40), 113 Sober, H. A., 245(83), 297 Streitwieser, A,,Jr., 332(37), 339(51), 396 Song, P. S., 341(1601,399 Stretton, A. 0. W.. 79(46, 471, 115 Sorm, F., 45(62), 78,341(164), 399 Struchkov, V. A., 133(144), l65(246), 217, SormovB,Z., 57(98), 73 ,919 Spahr, P. F., 76(11), 89(96), 112,114 Struck, W. A., 349(169), 399 Sparrow, A. H., 130(123), 133(133), 217 Stuart, A,,405(12), 409(37), 434 Spencer, J. H., 102(141), 116 Studier, F. W., 289(200), 291(200), 300 Spencer, M., 273(149), 299 Stuy, J. H., 118(57, 58), 12M.57, 581, 138, Sperber, G., 341(123), 398 151(57, 58), 175(57, 581, 213(57, 58), Speyer, J. F., 5(16), 14(16), ,97 216 Spiegelman, S., 76(9), 85, 94(110), 112, 116, 116, 175(281), lSO(2811, 220, 301 Subirana, J. A., 317(42), 395 Sueoka, N., 267(133), 271, 2.98, 299 (31, 307, 323(3), 324,326 Sudararajan, T. A,,79(41), 89(41), 113 Sponar, J., 57(98), 73 Sussmuth, R., 9(47), 15(47), 23(47), 28 Sreton, A. 0. W., 207(348), ,922 Sugino, Y., 118(82), 120(82), 160(82), 816 Stacey, K. A., 133(137), 144, ,917 Suit, J. C., 118(66a), 124(66a), 151(66a), Stacey, M., 411(67), 485 213(66a), 216 Staehelin, M., 18(84), ,98, 110(168), 116, Summers, W. C., 103(148), 116,118(101), 413(97), 436 216 Stahl, F. W., 118(49), 120(49), 121(49), 818 Suskind, S. R., 143(183), 207(163), 135(49), 213(49), ,916 Stanley, W. M., Jr., 79(42), 89(42), 113 Susman, M., 78(21), 106(21), 118 Stannem, C. P., 118(75), 120(75), 128(75), Sutherland, G. B. B. M., 293, 299 Sutton, H., 158(216), 211(216), 2131216), 130(75), 156(75), 213(75), 816 219 Stead, N. W., 99(134), 106(134), 116

AUTHOR

INDEX

461

Taylor, E. W., 157(213), 219 Susuki, H., 99(128), 116,175, 990 Taylor, J. H., 143(162), 818 Suzuki, T., 415(122), 436 Sverdlov, E. D., 6(24), 7(24), Z7, 418 Taylor, K., 95(116), 103(116), 116 (159), 419( 159), 420(159), 425(159), Temperli, A., 415(139), 437 427(171), 428(178), 429(178), 430 Teplova, N., 409(38),434 (1801, 431(180), 432(178, 188, 189, Terasima, T., 157(206), 157, 819 Terzaghi, E., 79(40), 89(40), 113 I N ) , m, 438 Tessman, E. S.,150(185), 818 Swafficld, M. N., 187(314, 315), 2.91 Swan, R. J., 233(38, 39, 40, 411, 234(40, Tcssman, I., 14(113, 125, 1261, 15(118), 19 (128), 21(102), 23, 89, 150(185, 1861, 411, 235(39, 41), 237(41), 238(39), 247 218,390(299), 402,431(184), 438 (84), 254(108), 255(108), 256(108), Thach, R., 79(41), 89(41), 113 296, 297, 298, 354(184), 399 Theriot, L., 118(91), 119(91), 123(91), 216 Swartz, M. N., 16(80), 98, 309(31), 326 Swartzendruber, C. S., 118(69), 120(69), Thikry, J., 230, 296 Thiry, L.T., 9, 88 139(69), 151(69), 213(69), 216 Thomas, C. A., Jr., 80(60), 103(150), 113, Sweet, R. M., 359(198), 400 116,302(6), 306, 307, 308, 310(27), Swenson, P. A., 164(251), 174(274), 183 324,325 (292), 205(251), 213(251, 2741, 220, Thomas, R., 304(9), 386 221 Thompson, L. R., 204(338), 289 Swift, M., 187(311), 221 Thrower, K. J., 317(43), 3.96 Symons, R., 415(141), 437 Tikhonenko, T. I., 57(97), 73, 284, $00, Ssabo, L. D., 194(324), 198(324), ad1 412(86, 95), 436,436 Szer, W., 415(130), 436 Szybalski, W., 95(116), 102(144, 145, 146, Till, J. E., 118(75), 120(75), 128(75), 130 147), 103(116, 145, 146, 147, 148), 116, (751, 156(75, 204, 2051, 21305, 204, 118(10, 67, 74, 101, 108, 114), 119(10), 205), 216, 219 120(10, 67, 74), 124(10, log), 125(10), Timofeeva, M., 194(328,329), 922 126(114), 127, 133(146), 134(129), 151 Timofieff-Ressowsky, N. W., 118(7f, 214 (10, 67,74), 156(74), 158,210(10), 213 Tinoco, T., Jr., 225, 234(46), 238(59), 239 (10, 67, 74), 214,816,916 (W),240(60, 61, 71), 241, 242(73), Sztumpf-Kulikowska, E., 382(274, 2751, 243(79), 244, 245, 249(91), 252(95), 401 254(108a), 262(59), 274(15, 16, 17, 73), 275(152), !276(153), 277(108a), T 278(108a, 156, 1631, 279(108a), 183, 284, 285(164), 296,996,297,298,$99, Taber, H., 14(95), 15(95), 19(95), 21(95), 329(10), 341(10, 80, 83, 84, 88, 1%), 25(95), 29 346(10), 350(173, 175, 176), 351(175), Tagashira, Y., 330(17), 341(17, 1331, 374 355, 366(10), 372, 374(124), 396, $97, (245,246), 396,598,401 898,899 Takagi, Y., 99(128), 116,143(175), ,918 TissiBres, A,,80(61), 90(61), 92(61), 93 Takemura, S., 415(137), 437 (1081, 94(61), 96(61), 110(61, log), Tarnaoki, T., 182(288), 221 118,114 Tamm, C., 410(64,65), 436 Tittensor, J., 415(126), 4-36 Tamm, I., 310(36), 326 Tanaka, M., 330(19), 341(73), 350(73), Tobey, R. A., 187(308), 221 Tocchini-Valentini, G. P., 94(111), 95 354(73), 396,396 (111, i m , 114 Tanooka, H., 118(50), 120(50), 121(50), A., 409(44, 501, 434,436 Todd, 135(50), 213(50), 816 Tokarskaya, V. I., 118(45, 531, 120(45), Tashiro, Y., 809 166, 816,,920 Tatarinova, S.,14(114), 99

462 Tolmach, L, J., 156(206), 157, ,919 Tomasi, V., 198(332), 222 Tomasz, M., 409(49), 436 Tomin, R., 186(306), 221 Tomita, K., 357(190, 193), 399 Tomkins, G. M., 79(36), 112 Tomlin, P., 118(54, 55, 561, 120(54, 55, 56), 151(54, 55, 561, 213(54, 55, 56), 216 Toropova, G. P., 133(141), 155(198), 164 (198, 2421, 217,218 Townsend, L. B., 393(317), 402 Trager, L., 15(127), 89 TrajkoviE, D., 201(335), 202(335), 22,9 Trautner, K. W., 16,898 Trautner, T. A., 309(31), 325 Travers, A,, 98,115 Travers, A. A., 116 TrgovEeviF, Z., 118(70), 120(70), 138(70), 151(70), 213 (70),216 Trincher, K. S., 155(197), 164(197), 2198 Triphonov, E., 412(96), 436 Troll, W., 428(177), 438 Trujillo, T. T., 80(56), 83(56), 113,181 (286), 221 Ts’o, P. 0. P., 238(49), 240(65,66,69, 701, 244, 249, 254(109, 1101, 257, 259, 260, 26l(SO), 274, 289(109, 110, 197, 199, 200), 290(109), 291(109, 197, 200), 296,297,,998, $00, 341(141, 1421, 369, 370(216), 374(142), 398,400 Tsuboi, M., 269(138), 271(146), 272(146), 273( 150a), 299 Tsugita, A., 12(78), 88, 79(40), 89(40), flJ Tubbs, R. K., 374(230), 400 Tiirck, G., 10(56), 15(127), 298,%9 Turchinsky, M., 405(19), 418(159), 419 (159), 420(159), 421(161), 423(167, 168, 169), 424(168), 425(159), 427 (171), 434,437 Turler, H., 415(139), 437

AUTHOR

INDEX

Ulbricht, T. L. V., 233(35, 36, 37, 38, 39, 40, 41), 234(41, 45), 235(39, 41), 237 (41), 238(39), 247(84), 254, 255, 256, 296,297 , 298,354(184), 399 Umezawa, H., 111(171), 116 Urks, M., 45(61), 72 Urnes, P. J., 224(2), 231(2), 285(2), 296 U r n , D. W., 231(30), 235(30), 238(30), 296 Usida, T., 412(83), 456

V Valdemoro, C., 341(159), 399 Valentini, L., 48(81), 78 Vallee, B. L.,231(32), 249(32), 262(32), 268(32), 296 Van Arkel, G. A., 7(36), 14(36), F7, 433 (2011, 438 Van Bekum, D. W., 155(196), 164(196, 2411, $18,219 van Bruggen, E. F. J., 83(70), 113 Van de Pol, J. H., 7(36), 14(36), 27,433 (201), 438 Van de Vorst, A., 341(128, 1291, 3998 Van Duuren, B. L., 374(243, 2441, &Of Van Holde, K. E., 238(57), 247(57), 248 (57), 296, 370(219), 400 Vanjushin, B., 405(18), 416(18), 434 Van Lancker, J. L., 118(83), 120(83), 160 (831, 162(83), 163(83, 2311, 164(237), 187(316), 189(83, 316, 316a), 190(83, 316, 316a,), 205(237, 316), 207, 209, 213(237), 216,819,221,622 van Rotterdam, J., 88(92), 114 Van Vunakis, H., 10(55), 11(61), 15(61), 28

Van Winekle, Q., 374(230), 4OO VBrt&esz, V.,194(323, 3241, 198(323), 221 Velikodvorskaya, G. A., 284(167), 300 Velikodvorskaya, T., 412(86), 4.96 Veillard, A., 330, 341(137, 138, 1391, 396, 398 U Velluz, L., 224(4), 225(4), 230, 295 Uchiyama, T., 164(237), 187, 189(316, Venkataraman, P. R.,l81(285), 220 316a), 190(316, 316a), 205(237, 3161, Venkstern, T. V., 405(10), 411(72), 415 (124), 434,435,436 213(237), 219,221 Ukita, T., 406(31), 414(112), 416(148, Venner, H., 268(136), 299,411(71), 436 Verwoerd, D. W., 6(17), 7(17), $7,417 149,NO), 434, @6,437 (1561, 418(156), 43 7 Ulanov, B., 406(28,29), 434

AUTHOR

INDEX

Vetter, V., 40(36, 37), 43(36, 53), 44(36, 37, 54, 55), 45(77), 46(77), 48(77), 52(77), 57(77), 71,72 Vielmetter, W., 4, 12(13), 14(13, 1121, 18, 23(13), 25(13), 27, 29 Vinograd, J., 88(90), 114 Voet, D., 231(29), 232(29), 233, 296, 350 (174), 399 Vogler, C., 118(62), 120(62), 151(62), 175 (62), 213(62), 216 Voiculetz, N., 165(245), 213(245), 219 Volkin, E., 118(33), 120(33), 213(33), 214 von Hippel, P. H., 32, 57(3, 1011, 70,73, 101(136), 115 von Stackelberg, M., 37, 71 Vos, O., 155(196), 164(196), 218 Vournakis, J. N., 245, 247(85), 262(127), 278, 279, 297,298

W Wacker, A., 9(47), 10(56), 11(62), 15(47, 62, 127), 23(47), 28,29,118(22), 120, 212(22), 213(22), 214, 377(270, 271, 2 7 3 , 401,412(75), 436 Wagle, M. M., 173(272), 213(272), 220 Wagner, E., 405(5, 6), 433 Wahba, A. J., 5(16), 9(39), 14(16, 39), 27,79(42), 89(42), 113 Wainson, A. A., 118(121), 120(121), 128 (121), 130(121), 217 Wakes, J. R., 181(283), 220 Waldron, D. M., 39(30), 40(30), 42(30), 68(30), 72 Walker, P. M. B., 302(4), 309(32), 317, 319(45), 321(45), 322, 323(32, 53), 324,325,326 Walker, R., 415(126), 436 Wallace, H., 311(39), 325 Walsh, W. M., 375(256), 401 Walter, G., 90(97), 93(97), 104, 110(97, 152), 11.4, 115 Walwick, E. R., 144(179), 145(179), 163 (1791, 218 Wang, J. C., 258(118), 298, 306, 309(30), 325 Wang, S. Y., 267(135), 298 Waring, M. J., 88(91), 114

Warner, R. C., 9(39), 14(39), 27 Warshaw, M. M., 238, 239(60), 240(60, 61), 241, 261(59), 296, 297 Waskell, L. A.,10(60), 28 Watanabe, K. A., 238(54), 296 Watson, D. G., 359(198),@0 Watson, J. D., 24, 29, 76(11), 112,207 (347), 222,252(99), 29s Watson, R., 88(90),114 Webb, B. R., 15(123), 29 Weckcr, E., 23(107), 29 Wchrli, W., 110(168), 116 Weil, G., 374(229), 400 Weil-Malherbe, H., 374(237), 400 Weill, J. H., 225(13), 2996 Weinblum, D., 377(270), 401 Weiss, J. J., 45(63), 72,118(23), 120, 133 (130, 131, 132), 135(23), 167(256), 168 (256), 170(258), 173, 204(23), 212(23), 213(23), 214,817,220 Weiss, S. B., 76(l), 84(75), 85(75), 93 (log), 95(118), 99(129), 100(129), 110 (log), 112,113,114,116 Weit, H., 405(2), 433 Weitzman, P. D. J., 70, 7 3 Weller, P. K., 210(355), 22!? Wells, B., 263(128), 265(129), 298 Wells, R. D., 305, 326 Wempen, I., 234(44), 296 Wetmur, J. G., 304(10), 316, 317, 326 Wheeler, C. M., 144(180), 145(180), 163 (180), 167(256), 168(256), 170(25.8), 173, 218, 220 Wheeler, G., 412(77), 43656 Wheland, G. W., 339, 385(294), 396, 402 Whitfeld, P., 409124, 43), 413(101), 434, 43s Whitfield, J. W., 164, 219 Whitmore, G. F., 118(75), 120(75), 128 (75), 130(75), 213(75), 815 Wiberg, J. S., 78(22), 111 Widholm, J., 309(30), 315 Wieder, C. M., 14(112), 89 Wierschowski, K. L., 11(66), 19(97), 88, 29

Wilder, J., 269(141), 299 Wilhelm, R. C., 4(9), 9(40), 14(40), 20,

n,u, 28

Wilholm, J., 258(118), 298

464

AUTHOR

Wilkins, M. H. F., 273(147, 148, 149, 149~4,299 Wilkinson, A. E., 131(126, 1271, 6f7 Williams, D. L., 82(68), 113,181(286), 661 Williams, R. W., 118(16, 691, 119(16), 120(16,69), 123(16), 134(16), 137,138, 139(69), 151(16,69), 154,210(16), 213 (16, 69), 614,916 Williams-Ashman, H. G., 76(3), 11.8 Wills, E. D., 131(126, 1271, 9f7 Wilson, D. L., 96(119), 116 Wilson, H. R., 273(147, 148, 148a), 699 Wilson, R. G.,6(22), 7, 11(73), 12(73), 14(22), 15(73), 2'7, 68, 171(262), 660, 391(301, 3M), 402, 433(196), 438 Win, H., 339(54), 396 Winkler, U., 1(5), 13(5), 16(5), dY Witkin, E. M., 118(95, 118), 120, 126 (118), 204(95), 216,217 Witkop, B., 415(135), 437 Wittman, H. G., 12(75), 14(75), 28, 412 (89,436

INDEX

233, 240(28), 241(75), 249, 250(94), 251(94, 96, 971, 252(941, 253(94), 254, 255(94, 96), 256(96), 257(97), 258 (97), 260(97), 261(10), 262(10, 971, 263(128), 264(97), 265(128, 1291, 267 (97, 110, 268(91, 110, 277(94, 961, 279(94, 961, 280(160), 281 (129, 162), 285(20), 289(28), 290(94, 97, 1111, 696, 296,297,998 Yanofsky, C., 79(44, 451,115 Yanofsky, S.A., 76(9), 116 Yegian, C., 110(166), 116 Yoshida, M., 406(31), 414(112), 416(150), 4349 @61 437 Youdale, T., 164, 219 Young, R., 409(52,53), 435 Yu, C. T., 405(1), 409(1), 415(123), 433, 436

Yuan, D., 92(102),114 Yung, N. C., 234(44), 996

Z

Zachau, H. G., 183(293), 221,405(9), 434 Witz, J., 57(100), 73, 252(98), Z98 Witzel, H., 4O6(24), 410(62), 413( 101), Za.jec, Lj.,118(59, 601, 120159, 60), 138 (59, 81, 151(59, 601,17369, 60),175 @4, 436,@6 916 (59, 601, 21369, a), Wolf, M. K., 286(173), 300 Wood, W. B., 101(135), 115,170(259), Zamecnik, P. C., 231(32), 249(32), 262 (32, 125), 268(32), 296, 298, 405(1), 220 409(1), 415(123), 433,436 Woodard, L., 187(311), $21 Woodhouse, D. L., 39(30), 40(30), 42 Zamenhof, S., 18(85), 29, 412(87), 436, 437 (30),68(30), 71 Zamir, A., 278(154), 699, 405(8), 434 Woodson, B., 172(264), 173(264), 260 Zampieri, A., 15(124), g9 Woody, R. W., 225(15, 16), 274(15, 161, Zavarine, R., 118(63), 120(63), 126, 151 296 (63), 213(63), 616 Wu, R., 306(22), 310(22), 326 Zemm, W., 206(343), 266 Wyckoff, R. R., 118(28), 120(28), 150 Zeszotek, E., 105(153), 110(153), 116 (189, 914 Ziffer, H., 288(187), SO0 Zillig, W., 6(17), 7(17), 87, 80(53), 82 Y (641,S(64, 69), 90(97), 93(97), 104 (152), llO(97, 1521, 113,114,405(20), Yamane, T., 382,401 414(113), 417(156), 418(156), 434, 436, Yamaoka, K., 286(180, 184), 287, 288 @Y (187), 300 Zirnm, B. H., 309(34), 325 Yamamoto, N., 21(104), 69 Zimmer, C., 268,299 Yamamoto, O., 184, 221 Zimmer, K. G., 118(7), $14 Yamazaki, H., 111(171), 116 Yang, J.T., 225(10, 20), 227(20, 21), 228 Zimmer, K. Z., 118(9),81.4 (% 229(24), I), 230, 231(27), 232(28), Zimmerman, A. S., 415(131), 436’

AUTHOR

IXDEX

Zimmerman, B. X., 19(93), 21(93), 29 Zimmerman, S. B., 5(15), 27, 309, 3215 Zimmermann, F., 118(88), 144, 145(88), 169(88),213(88), 9f6 148, 167(88),

465 Zobel, C.,406(27), 434 Zubay, G.,85(82), 11.4,286(170), 300 Zuman, P.,32(7), 36(7, ll), 42(49), 68 (71,70,71,72

Subject Index A

replication, 142-143 biochemistry, 143 Adenine, polarography of, 39-40 radiation effects, Alkylating agents, chemistry of action, in witro, 143-149 8-9 in vivo, 149-166 ribonucleic acid polymerase binding to, B 99-101 Bacteriophage, deoxyribonucleic acid, role in ribonucleic acid synthesis, 84-88 lethal changes produced by radiaas target for lethal radiation effects in tion, 121-123 living systems, 120-131 Brominating agents, chemistry of action, 12-16 transcription, 1&167 radiation effects, genetic effects in v i m , 171-204 C priming activity, 167-171 Circular dichroism, see Optical Rotatory Dispersion Deoxyribonucleoproteins, optical rotatory Cytosine, dispersion, 281-286 modification hydroxylamine, 427- Dinucleoside phosphaw, optical rotatow 433 dispersion and circular dichroism, 23&244 polarography of, 40-41 Drude equation, optical rotatory disperD sion and, 229 Deoxyribonucleie acid, families in higher organisms, 318-322 E optical rotatory dispersion, Electronic structure, base tilting and, 271-277 interbase interactions, comparison to ribonucleic acid, 262forces involved, 354-357 269 related problems, 373-374 single- veixus double-stranded, 269van der Waals-London interactions, 271 357-369 polarography, 45-16 vertical interactions, 369373 estimation of denaturation, 60-61 methods of calculation, 332-333 native and denatured, 46-54 extended Hiickel theory, 338-339 premelting temperatures and, 55-60 Huckel approximation, 336337 single-stranded breaks and, 54-55 iterative extended Hiickel theory, radiation-induced damage, 339-340 biological consequences, 204-207 representation of o-bonds, 337-338 breakdown, 138-142 self-consistent field method, 333-336 physical and chemical nature, 132self-consistent field procedure for all138 valence electrons, 340 working hypothesis of, 207-213 problems in radioand photobiology, repetition rate, spin densities in free radicals, 375deoxyribonucleic acid-deoxyribonu377 cleic acid interaction, 322 thymine photodimerization, 377-383 deoxyribonucleic acid-ribonucleic problems investigated, 340 acid interaction, 322-324 purine and pyrimidine bases, 34&354 466

467

SUBJECT INDEX

F

cleavage of N-glycosyl bonds, 410-411 complexes, optical rotatory dispersion, Free radicals, nucleic acid bases, spin 280-289 densities in, 375477 components, polarography of, 38-45 G electronic structure, interbase interactions, 354-374 Guanine, polarography of, 41-42 methods of calculation, 332-340 H mutagenesis and, 383-393 problems investigated, 340 Hybrids, purine and pyrimidine bases, 340-354 rates of reassociation, radio- and photobiology and, 375deoxyribonucleic families and, 318383 322 types of calculation, 329-332 factors affecting,316318 heterocyclic bases, reaction at, 411-417 stability, hybrid stability and, 306-308 artificial polymers and, 304-306 phosphate grouping, chemical modifidiscrimination and sequence homolcation of, 409-410 ogy and, 310-316 protein-encased, reactivity and mutanoncomplementary regions and, 308bility of, 21-24 310 visible rotatory dispersion, 289-291 nucleic acids, 306-308 Nucleosides, Hybridization, specificity of, 302-303 optical rotatory dispersion and circular Hydroxylamine, di ch roism, chemistry of action, 5-8 purine and pyrimidine derivatives, cytosine and, 427-432 231-233 functional studies, 432-433 a- versus /?-linkages of sugars, 233uracil nucleus and, 417-427 238 M Nucleotidea, optical rotatory dispersion and circular Mammalian cells, death, damage to dedichroism, oxyribonucleic acid and, 127-131 purine and pyrimidine derivatives, Mercury electrode, nucleic acid com231-233 ponents and, 4-4 a- versus /?-linkage of sugar, 233-238 Microorganisms, inactivation, radiationinduced damage to deoxyribonucleic acid and, 123-127 0 Mutagens, chemistry of action, 4-16 Mutagenesis, electronic factors in, 383- Oligonucleotides, optical rotatory dispersion and circular dichroiam, 238-249 393 Optical activity, origin of,225-227 N Optical rotatory dispersion, calibration of instruments, 229-230 Nitrosoguanidine, chemistry of action, correlation with circular dichroism, 9-10 228-229 Nitrous acid, chemistry of action, 4-5 difference ORD, 230-231 Nucleic acids, Drude equation, 229 carbohydrate moiety, chemical modifiexpressions for, 227-228 cation of, 408-409 nucleic acids, 269-280 chemical modification, nucleic acid complexes and, 280-289 functional studies, 406408 nucleosides and nucleotidea, 231-238 reactions used, 408-417 structural studies, 404406 oligonucleotides, 238-249

SUBJECT INDEX

origin of optical activity, 225-227 synthetic polynucleotides, 249

secondary structure, hydroxylaminolysis and, 425-422 synthctic, optical rotatory dispersion P and circular dichroism of, 249-262 Photobiology, electronic structure and, Polyribonucleotides, polarography of, 61376-383 68 Photochemistry, mutagens and, 10-12 Polyuridylate, optical rotatory dispersion Polarography, of, 249-254 analytical applications, 44-45 Purine (s) , claesical, 3S36 derivatives, optical rotatory dispersion deoxyribonucleic acids, 45-61 and circular dichroism of, 231-233 modern techniques, 36-38 electronic properties, nucleic acid components, 38-45 distribution and dipole moments, polyribonucleo tides, 34m47 natural, 68 molecular orbitals, 347349 synthetic, 61transitions, 34S354 principles of,32-38 polaromaphy of, 4 2 4 3 Polyadenylate optical rotatory dispersion Purine nucleotides, preferential chain of, 24!3-254 initiation and, 97-98 Polyadenylate -2polyinosinate, optical ro- Pyrimidines, tatory dispersion of, 254-257 derivatives, optical rotatory dispersion Polycytidylate, optical rotatory disperand circular dichroism of,231-233 sion of, 264257 clectronic properties, Polydeoxyadenylate, optical rotatory disdistribution and dipole moments, persion of,257-261 340-347 Polydeoxyadenylate poly deoxy thymidylmolecular orbitals, 347-349 ate, optical rotatory dispersion of, transitions, 349-354 257-261 polarography of, 4 2 4 3 Polydeoxy (adenylate-thymidylate) , opR tical rotatory dispersion of, 257-261 Polydeoxycytidylate, optical rotatory Radiation, biological consequences of, 204-207 dispersion of,257-261 deoxyribonucleic acid as target for Polydeoxythymidylate, optical rotatory lethal effects, 120-131 dispersion of, 257-261 deoxyribonucleic acid replication and, Polyguanylate, optical rotatory dispcr142-166 sion of, 254-257 physical and chemical nature of damPolyguanylate - polycytidylate, optical rotatory dispersion of, 254-257 age, 132-138 ribonucleic acid synthesis and, 166-204 Polyinosinate, optical rotatory dispersion Rndiobiology, electronic structure and, of,254-257 Polyinosinate-polycytidylate,optical ro375-383 Ribonucleic acid, tatory dispersion of, 254-257 deuridylic, preparation and properties, Polymers, artificial, hybrid stability and, 421425 304-306 interaction with deoxyribonucleic acid, Polynucleotides, 322-324 doubledranded, reactivity and mutamechanism of chain initiation, 99-107 bility of,17-21 optical rotatory dispersion, hybrid, optical rotatory dispersion of, base tilting and, 271-277 261-262 calculation of, 277-280 reactivity and mutability of,16-17

-

SUBJECT INDEX

comparison to deoxyribonucleic acid, 262-269 strandedness and, 269-271 regulation of synthesis, general concepts, 76-79 selective transcription, asymmetric, 94-96 further restrictions on, 96-97 preferential chain initiation with purine nucleotides, 97-98 transcription mechanism, de mvo synthesis and direction of growth, 88-89 growth rate of chain, 92-93 release of molecules, 89-92 role of deoxyribonucleic acid, 84-88 Ribonucleic acid polymerase, attachment site, nature of, 101-103 binding to deoxyribonucleic acid, 99101 deoxyribonucleic acid priming, radiation effects, 167-171 inhibition of, initiation, 109-11 1 polymerization, 109 initiation reaction of, 104-106 initiation sites, 1061W preparations,

469 physical properties, 81-84 purity, 80-81 termination signals, 107-109 Ribosomes, optical rotatory dispersion, 281-286

T Thymine, photodimerization, mechanism of, 377-383 Transcription, restrictions on, 96-97 selective, 94-98 Trinucleoside diphosphates, optical rotatory dispersion and circular dichroism, 244-249

U Uracil, modification with hydroxylamine, 417-427 Uridine, hydroxylamine and, 417-421

V Van der Waals-London interactions, electronic structure and, 357-369 Viruses, optical rotatory dispersion, 281-286 Visible rotatory dispersion, 289-291

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