Progress in Nucleic Acid Research and Molecular Biology, Volume 28

PROGRESS IN

Nucleic Acid Research and Molecular Biology Volume 28

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PROGRESS IN

Nucleic Acid Research and Molecular Biology edited by

WALDO E. COHN Biology Division Oak Ridge National Laboratory Oak Ridge, Tennessee

Volume 28 7983

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83 84 85 86

9 8 1 6 5 4 3 2 1

Contents CONTRIBUTORS ........................................................

ix

ABBREVIATIONS AND SYMBOLS. ...........................................

xi

SOMEARTICLES PLANNED FOR FUTURE VOLUMES ............................

xv

The Structure of Ribosomal RNA and Its Organization Relative to Ribosomal Protein Richard Brimacombe, Peter Maly, and Christian Zwieb I. 11. 111. IV. V.

Ribosomal RNA Sequences. .............. .......... Secondary Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . RNA-Protein Interactions Three-Dimensional Packing of E. coli rRNA. . . . ................. Outlook . . References

2

5 37

Structure, Biosynthesis, and Function of Queuosine in Transfer RNA Susumu Nishimura

............ I. History of Discovery of Queuosine. 11. Distribution of Queuosine in Anima s and Isolation o Hexose-Containing Queuosine Derivatives. ......................... 111. Biosynthesis of Queuosine in tRNA IV. Properties of tRNA-Guanine Transglycosylase . . . . . . . . . . . . . . . . . . . . . . . ............ V. Biosynthesis of the Queuine Skeleton VI. The Presence of Queuine-Lacking tRNA in Tumor Cells. . . VII. Change of the Ratio of Queuine to Guanine in tRNA during Differentiation. .......................... VIII. IX. Possible Approaches to the Use of Queuine Analogs for Cancer Diagnosis and Cancer Chemotherapy . ................... References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

50 55 60

67 70 71

Queuine: An Addendum Ram P. Singhal I. tRNA-Guanine Transglycosylase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Q in Neoplastic Cells.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Function of Q in Q-tRNAs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References ..................................................... V

75 78 79 80

Vi

CONTENTS

The Fidelity of Translation Abraham K . Abraham I . Agents Affecting Translation Fidelity in Vioo . . . . . . . . . . . . . . . . . . . . . . . .

I1. Message Decoding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

111. Accuracy of in Vitro Translation Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . IV Relationship between Speed and Accuracy., ........................ V . Codon Misreading by Isoacceptor tRNAs ........................... VI . Summary and Future Perspectives ................................. References .....................................................

.

82 85

88 92 94 96 97

Structure and Functions of Ribosomal Protein S1 Alap-Raman Subramanian I. Isolation. Physical Properties. and Shape . . . . . . . . . . . . . . . . . . . I1. Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Discernible Structural and Functional Domains . . . . . . . . . . . . IV . The S1 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ............... V. Location of S1 within the Ribosome . . . . . . . . . . . . . ............... VI . Interactions of S1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII . Functions ofS1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII . Mode ofAction ofS1 ............................................ References ....................... .........

109 120 121 134 137

The Yeast Cell Cycle: Coordination of Growth and Division Rates Steven G . Elliott and Calvin S. McLaughlin I . Methods for Cell-Cycle Analysis .................................. I1. Patterns ofGrowth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

111. Control of Cell Division . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

146 149 167 169

Prokaryotic and Eukaryotic 5 S RNAs: Primary Sequences and Proposed Secondary Structures Ram P . Singhal and Joni K . Shaw I . Structure of Prokaryotic 5 S RNAs ................................. I1. Structure of Eukaryotic 5 S RNAs ..................................

111. Conformation of Prokaryotic 5 S RNAs ............................. IV . Summary ...................................................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix: Sequences of 5 S RNAs ................................. Addenum: References to 32 Additional Sequences . . . . . . . . . . . . . . . . . . .

178 185 191 196 197 198 251

vii

CONTENTS

Structure of Transfer RNAs: Listing of 150 Additional Sequences

.

Ram P Singhal. Edda F . Roberts. and Vikram N .Vakharia I . Organization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1. Abbreviations and Definitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. References and Footnotes for Sequence List ........................ IV . Sequences of126tRNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Addenum: Additional tRNA Sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

211 215 215 219 227 227

INDEX. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

253

CONTENTS OF PREVIOUS VOLUMES ........................................

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Contributors Numbers in parentheses indicate the pages on which the authors' contributions begin.

ABRAHAM K. ABRAHAM (81), Department of Biochemistry, University of Bergen, Arstadveien 19, 5000 Bergen, Norway RICHARDBRIMACOMBE (l), Max-Planck-Institut fiir Molekulure Genetik, Abteilung Wittmann, Berlin-Dahlem, Federal Republic of Germany STEVENG. ELLIOTT(143), Department of Biological Chemistry, California College of Medicine, University of CalijQrnia, Irvine, California 9271 7 CALVINS. MCLAUGHLIN (143), Department of Biological Chemistry, California College of Medicine, University of California, lrvine, Calijornia 9271 7 PETERMALY(l),Max-Planck-Institut f u r Molekulare Genetik: Abteilung Wittmann, Berlin-Dahlem, Federal Republic of Germany SUSUMUNISHIMURA (49), Biology Division, National Cancer Center Research Institute, Tsuktji 5-1-1, Chuo-ku, Tokyo 104, Japan EDDAF . ROBERTS(211),Department of Chemistry, Wichita State University, Wichita, Kansas 67208 JONI K. SHAW(177), Department of Chemistry, Wichita State University, Wichita, Kansas 67208 RAM P. SINGHAL~ (75, 177, 211), Department of Chemistry, Wichita State University, Wichita, Kansas 67208 ALAP-RAMAN SUBRAMANIAN (101), Max-Planck-Znstitutfiir Molekulare Cenetik, Abteilung Wittmann, Berlin-Dahlem, Federal Republic of Germany VIKRAM N. VAKHARIA (211), Department of Chemistry, Wichita State University, Wichita, Kansas 67208 CHRISTIAN Z W I E B(l), ~ Max-Planck-lnstitut f u r Molekulare Genetik, Ahteilung Wittmann, Berlin-Dahlem, Federal Republic of Germany Present address: Laboratory of Cellular and Molecular Biology, Gerontology Research Center, National Institute on Aging, Baltimore, Maryland 21224. Present address: Department of Physiological Chemistry, Brown University, Providence, Rhode Island.

ix

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Abbreviations and Symbols All contributors to this Series are asked to use the terminology (abbreviations and symbols) recommended by the IUPAC-IUB Commission on Biochemical Nomenclature (CBN) and approved by IUPAC and IUB, and the Editor endeavors to assure conformity. These Recommendations have been published in many journals ( 1 , 2 )and compendia (3)in four languages and are available in reprint form from the Office of Biochemical Nomenclature (OBN), as stated in each publication, and are therefore considered to be generally known. Those used in nucleic acid work, originally set out in section 5 of the first Recommendations (1)and subsequently revised and expanded (2,3),are given in condensed form (I-V) below for the convenience of the reader. Authors may use them without definition, when necessary.

I. Bases, Nucleosides, Mononucleotides 1. Bases (in tables, figures, equations, or chromatograms) are symbolized by Ade, Gua, Hyp, Xan, Cyt, Thy, Oro, Ura; Pur = any purine, Pvr = any pyrimidine, Base = any base. The prefixes S-, H2,F-, Br, Me, etc., may be used for modifications of these. 2. Ribonucleosides (in tables, figures, equations, or chromatograms) are symbolized, in the same order, by Ado, Guo, Ino, Xao, Cyd, Thd, Ord, Urd (Prd), 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 three-letter symbols, as in Ara-Cyt (for aCyd) or dRib-Ade (for dAdo). 3. Mono-, di-, and triphasphates of nucleosides (5‘)are designated by NMP, NDP, NTP. The N (for “nucleoside”) may be replaced by any one of the nucleoside symbols given in 11-1 below. 2’-, 3’-, and 5’- are used as prefixes when necessary. The prefix d signifies “deoxy.” [Alternatively, nucleotides may be expressed by attaching P to the symbols in (2) above. Thus: P-Ado = AMP; Ado-P = 3’-AMP] cNMP = cyclic 3’:s’-NMP; BbcAMP = dibutyryl CAMP,etc.

II. Oligonucleotides and Polynucleotides 1. Ribonucleoside Residues

(a) Common: A, G, I, X, C, T, 0, U , P, R, Y, N (in the order of 1-2 above). (b) Base-modified: sI or M for thioinosine = 6-mercaptopurine ribonucleoside; sU or S for thiouridine; brU or B for 5-bromouridine; hU or D for 5,6-dihydrouridine; i for isopentenyl; f for formyl. Other modifications are similarly indicated by appropriate lower-case prefixes (in contrast to 1-1 above) (2, 3). (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 suffix m (e.g., -Am- for 2’-O-methyladenosine, but -mA- for 6-methyladenosine). (d) Locants and multipliers, when necessary, are indicated by superscripts and subscripts, respectively, e.g., -m$A- = 6-dimethyladenosine; 4 U - or -4S- = 4-thiouridine; -ac4Cm- = 2’-O-methyl-4-acetylcytidine. (e) When space is limited, as in two-dimensional arrays or in aligning homologous sequences, the prefixes may be placed ouer the capital letter, the suffixes ooer the phosphodie,ster symbol. 2. Phosphoric Residues [left side

=

5’. right side = 3’ (or 2’)]

(a) Terminal: p; e.g., pppN. . , is a polynucleotide with a 5’-triphosphate at one end; Ap is adenosine 3’-phosphate; C > p is cytidine 2’:3’-cyclic phosphate (1, 2, 3); p < A is adenosine 3‘:5’-cyclic phosphate. xi

xii

ABBREVIATIONS A N D SYMBOLS

(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 triplets should be written without some punctuation separating the residues.) 3. Polarity, or Direction of Chain

The symbol for the phosphodiester group (whether hyphen or comma or parentheses, 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 at right) may be shown by numerals or by right-to-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 is viewed right-side-up. 4. Synthetic Polymers

The complete name or the appropriate group of symbols (see 11-1 above) of the repeating unit, enclosed in parentheses if complex or a symbol, is either (a) preceded by “poly,” or (b) followed by a subscript “n” or appropriate number. No space follows “poly” (2, 5). 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(3 adenylate, 2 cytidylate) = poIy(&C2) or (A3,(&, an irregular copolymer of A and C in 3 :2 proportions; poly(deoxyadeny1ate-deoxythymidylate) = poly[d(A-T)] or poly(dA-dT) or (dA-dT). or d(A-T),, an alternating copolymer 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 subscript “n” may be replaced by numerals indicating actual size, e.g., A;dT,,.,,.

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.: or A, . U, poIy(A) . poly(U) poly(A) . 2 poly(U) or A, . 2U, or (dA-dC), . (dG-dT),. poly(dA-dC) . poly(dG-dT)

2. Nonassociated chains are separated by the plus sign, e.g.: or

2[poly(A) . poly(U)I + PoIY(A) 2[An . U,] + A,, . 2Um + A,,.

. 2 poly(U) + poly(A)

3. Unspecified or unknown association is expressed by a comma (again meaning “unknown”) between the completely specified chains. Note: In all cases, each chain is completely specified in one or the other of the two systems described in 11-4 above.

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

ribonucleic acid or ribonucleate deoxyribonucleic acid or deoxyribonucleate messenger RNA; ribosomal RNA; nuclear RNA heterogeneous nuclear RNA “DNA-like’’ RNA; complementary RNA

xiii

ABBREVIATIONS A N D SYMBOLS

mtDNA tRNA

mitochondria1 DNA transfer (or acceptor or amino-acid-accepting) RNA, replaces sRNA, which is not to be used for any purpose aminoacyl-t RNA “charged” tRNA (1.12.~tRNA’s carrying ammoacyl residues); may be abbreviated to AA-tRNA alanine tRNA or tRNA normally capable of accepting alanine, to form tRNAAIa, etc. alanyl-tRNA, etc. alanyl-tRNA or The same, with alanyl residue covalently attached. alanyl-tRNAAla [Note: fMet = formylmethionyl; hence tRNAme’, identical with tRNAP‘] Isoacceptors are indicated by appropriate subscripts, i.e., tRNAfia. tRNA.fia, etc. V. Miscellaneous Abbreviations

PI, pp, inorganic orthophosphate, pyrophosphate RNase, DNase ribonuclease, deoxyribonuclease melting temperature (“C) t m (not T , ) 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 editor, they increase the ease of reading. Enzymes In naming enzymes, the 1978 recommendations of the IUB Commission on Biochemical Nomenclature ( 4 ) are followed as far as possible. At first mention, each enzyme is described either by its systematic name or by the equation for the reaction catalyzed or by the recommended trivial name, followed by its EC number in parentheses. Thereafter, a trivial name may be used. 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 . JBC 241, 527 (1966); Bchetn 5, 1445 (1966); BJ 101, l(l966); ABB 115, 1 (1966), 129, 1 (1969);and elsewhere. t 2. EJB 15, 203 (1970);J B C 245, 5171 (1970);J M B 55, 299 (1971); and e1sewhere.t 3. “Handbook of Biochemistry” (G. Fasman, ed.), 3rd ed. Chemical Rubber Co., Cleveland, Ohio, 1970, 1975, Nucleic Acids, Vols. I and 11, pp. 3-59. 4 . “Enzyme Nomenclature” [Recommendations (1978) of the Nomenclature Committee of the IUB]. Academic Press, New York, 1979. 5. “Nomenclature of Synthetic Polypeptides,” JBC 247, 323 (1972); Biopolymers 11, 321 (1972);and elsewhere. t

Abbreviations of Journal Titles Journals Annu. Rev. Biochem. Arch. Biochem. Biophys. Biochem. Biophys. Res. Commun.

Abbreviations used ARB ABB BBRC

*Contractions for names of journals follow. tReprints of all CBN Recommendations are available from the Office of Biochemical Nomenclature (W. E. Cohn, Director), Biology Division, Oak Ridge National Laboratory, Box Y, Oak Ridge, Tennessee 37830, USA.

xiv Biochemistry Biochem. J. Biochim. Biophys. Acta Cold Spring Harbor Symp. Quant. Biol. Eur. J . Biochem. Fed. Proc. Hoppe-Seyler’s 2. physiol. Chem. J. Amer. Chem. Soc. J. Bacterial. J. Biol. Chem. J. Chem. SOC. J. Mol. Biol. Nature, New Biology Nucleic Acid Research Proc. Nat. Acad. Sci. U.S. Proc. Soc. Exp. Biol. Med. Progr. Nucl. Acid Res. Mol. Biol.

ABBREVIATIONS AND SYMBOLS

Bchem BJ BBA CSHSQB

EJB FP ZpChem JACS J. Bact. JBC JCS JMB Nature NB NARes PNAS PSEBM This Series

Some Articles Planned for Future Volumes Hypermodified Nucleosides of tRNA

A. ADAMIAK RNA Processing in a Unicellular Microorganism

D. APIRION Nearest Neighbor Effects in the Structure and Function of Nucleic Acids

P. N. BORER The Elongation Factor EF-TU and Its Two Encoding Genes

L. BOSCH Small Nuclear RNAs and RNA Processing

H. BUSCH Participation of Aminoacyl-tRNA Synthetases and tRNAs in Regulatory Processes

G. NASS Neuropolyproteins from Brain

D. RICHTER Viral Inhibition of Host Protein Synthesis

A. SHATKIN AND CARCINOGENESIS VOLUME29: GENETICMECHANISMS

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The Structure of Ribosomal RNA and Its Organization Relative to Ribosomal Protein RICHARDBRIMACOMBE, PETERMALY,AND CHRISTIAN ZWIEB* Max-Planck-Institut f u r Molekulare Cenetik, Abteilung Wittmann, Berlin-Dahlem, Federal Republic of Germany I. Ribosomal RNA Sequences.. ................................. 11. Secpndary Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Derivation of Structures of Escherichia coli rRNA.. . . . . . . . . . . . B. Structures Proposed for rRNA from Other Sources.. . . . . . . . . . . . C. Evidence for Alternative Conformations (“Switches”). . . . . . . . . . 111. RNA-Protein Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Protein Binding Sites on rRNA. . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. RNA-Protein Cross-Linking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Three-Dimensional Packing of E . coli rRNA. .................... A. Intra-RNA Cross-Linking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Electron Microscopy of rRNA within Ribosomal Subunits . . . . . . V. Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 5

8 12 21 24 24 26

37 37 39 41 43

The structure of the Escherichia coli ribosome was last reviewed by our laboratory in this series in 1976 (1).At that time, the emphasis was on the structure and function of the individual ribosomal proteins, and, although a considerable amount of information was already available concerning the ribosomal RNA, progress in this area was hampered by the lack of complete base sequence data for the ribosomal RNA molecules. Since then, as a direct result of the explosion in DNA sequencing technology ( 2 , 3 ) ,the situation has changed dramatically. Complete nucleotide sequences have now been established, corresponding not only to the ribosomal RNA molecules from E . coli, but also to those from a number of different organisms, covering a large portion of the evolutionary spectrum. * Present address: Department of Physiological Chemistry, Brown University, Providence, Rhode Island. 1 Progress in Nucleic Acid Research and Molecular Biology, Vol. 28

Copyright 8 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-540028-4

2

RICHARD BRIMACOMBE

et al.

We begin by reviewing briefly the current status of these sequences. Next, we describe how the sequence information has been used to derive convincing secondary structure models for the RNA from both subunits of the E . coli ribosome, and we compare the various models that have been proposed. We show how extrapolation of these data to ribosomal RNA molecules of widely differing size classes leads to the clear conclusion that the secondary structures, as well as significant regions in the primary sequences, have been conserved to a large extent throughout evolution. Section IV deals with the threedimensional organization of the ribosomal RNA and its arrangement with respect to the ribosomal proteins, concentrating once again on the E . coli ribosome. In particular, we include a review of the application of cross-linking techniques (bath RNA to protein and intra-RNA) to this problem. In general, rather than presenting an exhaustive survey of the literature, we have selected topics or examples to illustrate those problems or points of interest that we consider to be most relevant to the central objective in this field of research, namely, the elucidation of the three-dimensional structure of the ribosomal RNA in situ in the ribosome. I. Ribosomal RNA Sequences The organization and transcription of ribosomal RNA genes is a complex and fascinating subject, itself worthy of review. However, we confine ourselves here to a discussion of the mature ribosomal RNA (rRNA) species, as they occur in the completed ribosomal particles. The “standard” bacterial ribosome, as typified by that of E . coli, contains in its small subunit a single RNA molecule (16 S), which is about 1540 nucleotides in length ( 4 , 5 ) . The large subunit contains a 23 S RNA (ca. 2900 nucleotides) (6) and a 5 S RNA (120 nucleotides) (7). In other organisms, the size of these rRNA molecules varies considerably. The smallest so far reported are those from trypanosome mitochondria, which are only 640 and 1230 nucleotides in length, respectively, from the small and large subunits (8). Next come the mitochondria1 ribosomes from mammals, with RNA molecules of 12 S and 16 S (ca. 950 and 1550 nucleotides) (e.g., 9, lo), and these very small ribosomes contain no 5 S RNA. Other types of mitochondria (e.g., 1 1 ) , and also chloroplasts (e.g., 12, 13), have rRNA molecules corresponding in size to those of the bacterial ribosomes. The chloroplasts and mitochondria from higher plants both contain 5 S rRNA, and in addition the large subunit of higher plant chloroplast ribosomes contains a 4.5 S RNA species (14).

STRUCTURE OF RIBOSOMAL RNA

3

The largest ribosomes are those from the cytoplasm of eukaryotes, with RNA molecules of 18 S (ca. 1800 nucleotides) (e.g., 15, 16) and 26-28 S (up to 4000 nucleotides) (e.g., 17) in the small and large subunits, respectively. The large subunit contains a 5.8 S RNA molecule (e.g., 18) as well as 5 S RNA, and a 2 S RNA species is also observed in the large ribosomal subunit from Drosophila (19). Further, the large subunit rRNA genes for many eukaryotic ribosomes contain introns (e.g., 20, 21), and in Drosophila the final “28 S” transcription product appears in two distinct halves (reviewed in 22). As shown in Section II,B the small rRNA molecules (4.5 S , 5.8 S, and 2 S) all have clear counterparts within the 23 S rRNA from E . coli, and they can therefore better be regarded as products of posttranscriptional processing of the rRNA rather than as “extra” rRNA species. Many complete or partial sequences are now known for all these classes of rRNA, mostly obtained by determination of the corresponding rDNA sequences. A compilation of known 5 S and 5.8 S sequences has been made (23, 24),’ and this list continues to grow at a rate of about one sequence every two or three weeks. [One of the more interesting new sequences here is the “5 S” rRNA from Halococcus morrhuae, which contains a 108-base insertion (%).I The sequence of 2 S rRNA from Drosophila melanogaster is known (26),and also the sequences of 4.5 S rRNA from wheat (27),tobacco (28),and maize (29). More important for the purpose of this article are the sequences of the major rRNA molecules, and those sequences currently complete or nearly complete are listed in Table I (4-6,9-13,15-17,30-40). It can be seen from Table I that almost every common size class of large rRNA mentioned above is represented by sequences from two or more species. In addition, many partial sequences of the major rRNA molecules have been determined. In the case of the small subunit, sequences are available from a total of about 20 species for the 50-200 nucleotides at the 3’ terminus, and compilations of these have been published (41,42).In the case of the large-subunit rRNA, some partial sequences are available for the regions flanking introns in the rDNA of Chlamydomonas reinhardii chloroplast (43) and yeast mitochondrial ribosomes (20, 44), as well as of Drosophila virilis (45),D . melanogaster (46,47),Tetrahymena pigmentosa (21),and Physarum poZycephalum (48,49)cytoplasmic ribosomes. The positions in E . coli 23 S rRNA that correspond to the locations of these inserted sequences have been collated (36).Similar short sequences are known from the 5’ ends of the large-subunit rRNA of Aspergillus nidulans mitochonSee also Singhal and Shaw in this volume. [Ed.]

4

RICHARD BRIMACOMBE

et al.

TABLE I SEQUENCES OF RIBOSOMAL DNA (OR rRNA) MOLECULES" Organism

Small subunit rRNA Large subunit rRNA

Human mitochondrion Mouse mitochondrion Rat mitochondrion

12 s (9) 12 s (10)

Saccharomyces cerevisiae mitochondrion Aspergillus nidulans mitochondrion Paramecium primaurelia mitochondrion

15 S (11) 15 S (31)

Escherichia coli Proteus vulgaris Bacillus brevis Bacillus stearothernophilus Zea mays chloroplast Euglena gracilis chloroplast Saccharomyces cerevisiae cytoplasm Saccharomyces carlsbergensis cytoplasm Xenopus laevis cytoplasm

-

16 S (4, 5) 16 S (34) (16 S) (35)

16 S (12) 16 S (37) 18 S (15) 18 S (16)

16 S (9) 16 S (10) 16 S (30)

20 S (32) 23 S (6, 33)

-

23 S (36) 23 S (13)

-

26 S (17) 26 S (38) (28 S) (39,40)

0 Known sequences are listed, showing the size (S value in Svedberg units) of the corresponding rRNA molecule; parentheses denote that the sequence concerned is not yet fully complete; a dash indicates that the sequence is not available. All sequences were determined from ribosomal DNA, with the exception of E. coli 16 S rRNA (5) and 23 S rRNA (33), and P . vulgaris 16 S rRNA (34);these determinations were made directly from RNA. Numbers in parentheses indicate reference numbers.

dria (50)and Euglena gracilis chloroplasts (51, 52) and from the 3' ends of Aeromonas punctata and Proteus vulgaris (53).Last, a large amount of sequence data is contained in oligonucleotide catalogs made earlier from the rRNA of many species [summarized by Fox et al. (54)l. It has long been known that the sequence of 5 S rRNA is highly conserved (see, e.g., 55),and a corresponding pattern of conservation is seen among the major rRNA molecules. For example, P . vulgaris 16 S rRNA shows 93% homology to the 16 S rRNA from E . coli (34), whereas the homology between the 23 S rRNA molecules from B . stearothermophilus and E . coli is about 75% (36).Escherichia coli rRNA and Z. mays chloroplast rRNA also show about 75% sequence homology, in both subunits (12, 13). Xenopus 18 S and yeast 18 S rRNA are again about 75% homologous to one another (16).The two Saccharomyces 26 S rRNA sequences [ S . cerevisiae (17)and S . carlsbergensis ( I S ) ] are virtually identical, differing only in about 20 positions, and this order of difference is also observed between different

STRUCTURE OF RIBOSOMAL

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5

ribosomal cistrons of the same organism; E . coli 16 S rRNA, for instance, contains 16 single-base cistron heterogeneities (56). When sequences from the different size classes of rRNA (Table I) are compared, it becomes apparent that a number of regions have been conserved in both the large- and small-subunit rRNA molecules of all species, despite the large differences in lengths of the polynucleotide chains. This type of homology becomes even more striking when the secondary structures of the various molecules are considered; it is discussed in more detail in Section I1,B. Since most of the sequence data (Table I) was determined from rDNA, information concerning modified nucleotides in the corresponding rRNA molecules is incomplete in many cases. Escherichia coli 16 S rRNA contains 9 methylated bases (56),and the 23 S rRNA contains 10 methylated bases and 3 pseudouridine residues (33).In P . vulgaris 16 S rRNA, 6 methylated bases are in homologous positions to their E . coli counterparts (34), and the two N6-dimethyladenine residues near the 3’ terminus of the 16 S rRNA seem to be a universal feature of all small-subunit RNA molecules (reviewed in 42). Some methylated bases have been localized in rodent mitochondria1 rRNA, and these also appear to be highly conserved (57).The pattern of nucleotide modification in eukaryotic cytoplasmic ribosomes is rather more complex. Xenopus 18 S rRNA contains 40 methylated bases, of which the majority are 2’-O-methyl groups (16).There is one hypermodified base as well as several base-methylated residues, and in addition there are 40 pseudouridine residues (58),which for the most part have not as yet been localized precisely. The corresponding yeast 18 S rRNA molecule has the same number of base methylations, but fewer 2‘-O-methyl groups (59). These have not yet been localized, whereas in yeast 26 S rRNA, 30 out of the 43 methyl groups have been placed (38).There are also some data on the corresponding sites of methylation in Xenopus 28 S rRNA (60). To conclude this section, it is clear that an enormous amount of sequence information has already been collected, and this provides the raw material for the next stage in the elucidation of the threedimensional organization of ribosomal RNA, namely the construction of models for the secondary structure.

II. Secondary Structures There are, broadly speaking, three different approaches by which a nucleotide sequence can be folded into a double-helical secondary structure: the theoretical, the experimental, and the comparative.

6

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et al.

The theoretical approach relies on the use of computer programs to select the thermodynamically most favorable structure for the sequence, using thermodynamic parameters derived from the melting properties of model oligonucleotides or RNA fragments (e.g., 61-64). A large number of computer algorithms generated for this purpose have been reported (e.g., 65-69), and the newer versions (e.g., 68) have been considerably improved, in that they can select structures for an entire long sequence rather than for just short sections. However, the approach suffers from the obvious disadvantage that the computer program can be only as good as the thermodynamic data put into it, and these data are by no means comprehensive. The effects of imperfections such as “bulges” and “loops” in the helices cannot be computed very accurately, and-more important-the thermodynamic effects of tertiary structure or interactions with protein can hardly be assessed at all. As a result, this approach can lead to erroneous structure predictions, and it has, for example, been shown (70) that the computer may predict quite different structures for two phylogenetically closely related sequences. Nevertheless, when combined with experimental or comparative data, the computer approach is a powerful tool for screening potential secondary structures. The experimental approach to secondary structure determination has made use of a number of different methods; these include chemical modification (e.g., 71-76), analysis of enzyme cutting points (e.g., 56, 77), oligonucleotide binding (e.g., 78, 79), isolation of base-paired RNA fragments (80,81), and intra-RNA cross-linking (e.g., 82, 83). In the case of chemical modification, the RNA is treated with base-specific reagents to test the accessibility (or single-strandedness) of individual residues. The reagents that have been used include kethoxal ( 71)and glyoxal ( 72) (G-specific);monoperphthalic acid ( 73),m-chloroperoxybenzoic acid (72), and diethyl pyrocarbonate (74) (A-specific); and methoxyamine (75) and bisulfite (72)(C-specific). Dimethyl sulfate (74)(C- and G-specific) and soluble carbodiimides (76)(U- and G-specific) have also been applied. The analysis of enzyme cutting points is, in principle, a very similar method, in which the RNA is subjected to a mild digestion by a single-strand-specific nuclease (such as nuclease S1, or ribonucleases A and Tl), and the resulting fragments are analyzed to pinpoint those residues at which the polynucleotide chain has been cut (56).The converse approach, using the double-strand-specific nuclease from cobra venom, has also been applied with success (77).Oligonucleotide binding is a probe for singlestranded regions, in which the putatively exposed sequences can be tested for their ability to bind a short complementary oligonucleotide

STRUCTURE OF RIBOSOMAL

RNA

7

(78,79).Data from all these methods become progressively more difficult to interpret as the sequence under consideration becomes longer, a disadvantage that is not shared by the “base-paired-fragment” approach (80, 81). Here, the RNA is subjected to a mild digestion with nuclease, and the fragments are isolated with base-pairing intact by gel electrophoresis under nondenaturing conditions. This is followed by a second electrophoretic dimension in denaturing conditions, and analysis of the products gives direct information as to which regions of the sequence are base-paired with one another. The last method mentioned above, namely intra-RNA cross-linking (82, 83), also gives direct data on neighborhoods between various parts of the RNA chain; this is discussed in more detail in Section IV,A. The third general approach to secondary structure determination, namely the comparative approach, is without doubt the most powerful and is in essence very simple (84).If similar sequences from different organisms are compared, then a base change in one strand of a putative helical region must be compensated by a complementary base change in the other strand. Thus an A * U pair in one species could become a G C pair at the corresponding positions in the second species, and so on. If the base changes do not compensate each other in this way, then the implication is that the proposed element of secondary structure is incorrect. The approach can also be inverted since, once a secondary structure has been established, similar structures can be sought in more distantly related species by screening for stretches of identical base sequence rather than for differences between the two species concerned. It is clear from the preceding section that a Iarge number of suitable rRNA sequences are available for the application of this approach, and its great advantage is that it generates a “positive feedback” situation in which each new sequence studied not only gives support to the concept that secondary structure in rRNA has been highly conserved among molecules of the same size class, but also serves to confirm and extend the original secondary structure model. In practice, the models derived for the secondary structure of the E . coli rRNA molecules all make use of combinations of several of the methods just outlined, and these models are described in the next section. It should be noted that in general the percentage of residues proposed to be involved in base-pairing is rather less than that predicted by physicochemical studies [e.g., hyperchromicity measurements (85, 8611. This is quite reasonable, since the models on the whole describe minimum secondary structures and take no account, for instance, of tertiary interactions or complex stacking of helical

8

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elements, both of which would contribute to the hypochromicity of the overall structure. A. Derivation of Structures of Escherichiu coli rRNA 1. 5 S RNA Among the large number of secondary-structure models proposed over the years for 5 S rRNA (see, e.g., 55), that of Fox and Woese (84) soon became established as the most reasonable “minimum” structure for prokaryotic 5 S rRNA, the model being based on comparative evidence (see above)’. This model for E . coli 5 S rRNA is illustrated schematically in Fig. 1. More recently, extensions to the base-pairing scheme have been proposed, and two examples of these are also included in Fig. 1. The model of Studnicka et al. (87) was derived by a “filtering” method that combines the theoretical and comparative approaches; minimum energy structures were accepted only if they could be drawn for all the prokaryotic 5 S sequences under consideration. This model also predicts a tertiary interaction between residues 24 and 39. The model of Pieler and Erdmann (88),on the other hand, is based on a combination of comparative evidence and S l-nuclease cleavage data from E . coli 5 S rRNA; it predicts a “tertiary” basepairing between residues 41-44 and 74-77, as well as additional base-pairs in the stem region of the molecule. A similar tertiary base40

,110

1

1 12

40

FIG.1. Three models for the secondary structure of Escherichia coli 5 S rRNA. The sequence is numbered from the 5’ end, and the bars denote base-pairing. (A) The model of Fox and Woese (84), (B) that of Studnicka et al. (87), and (C) that of Pieler and Erdmann (88), indicating the additional tertiary base-pairing proposed between residues 41-44 and 74-77.

STRUCTURE OF RIBOSOMAL

RNA

9

pairing has also been proposed by Hancock and Wagner (89) between residues 37-40 and 73-76, on the basis of an intra-RNA cross-link generated between E . coli residues 41 and 72. It should be noted that short tertiary base-pairings of this nature can be accommodated in the structure, without generating “knots” in the RNA chain; clearly, however, a longer base-pairing of this type (extending over more than half a helix turn) does generate a potential “knot” situation (cf. Sections II,C and IV,A). A model for two eukaryotic 5 S rRNA species, proposed by Troutt et al. (70),is essentially very similar to that of Studnicka et al. (87),and the latest model from Woese and his group (90)offers a general structure for all 5 S rRNA molecules, both prokaryotic and eukaryotic. In this model, additional base-pairing is proposed on the basis of a comparison with 5 S rRNA from the archaebacterium Sulfolobus acidocaldarius, in which the whole region between bases 69 and 111 forms a continuously base-paired hairpin loop. In order to generate such a helix in the case of E . coli 5 S rRNA, a number of mismatched basepairs must be postulated, including three A * G pairs. While it has been shown that A G pairs are common features at the ends of helices in tRNA (91),there is as yet no evidence that they can exist within a continuing helix, and as the authors themselves point out (go), such a helix would certainly be very irregular and readily denatured. The important inference to be drawn here is that it may be possible to replace several Watson-Crick base-pairs by mismatched or nonpaired bases, without altering the overall three-dimensional appearance of the molecule.

2. 16 S AND 23 S rRNA Several research groups have been independently involved in the construction of secondary-structure models for the major rRNA molecules, each using different sets of experimental data derived from E . coli rRNA, and each using sequence comparisons to refine the models. Noller et al. in California, in collaboration with Woese et al. in Illinois (72, 35, 36) have exploited primarily the analysis of sites of chemical modification by kethoxal, glyoxal,bisulfite, and m-chloroperoxybenzoic acid in order to obtain their experimental data. The sequences used for comparative analysis were those of B . breuis 16 S rRNA (35)and B . stearothermophilus 23 S rRNA (36) (see Table I), as well as the extensive oligonucleotide catalogs of Woese et al. (54).The groups in Strasbourg (33,34,92) have made use of single- and double-strand-specific nucleases to generate specific cuts in the RNA, and have used the sequences of P . uulgaris 16 S rRNA (34) and 2. mays chloroplast 16 S

10

RICHARD BFUMACOMBE

et al.

and 23 S rRNA (12,13) for comparison. Our approach has been based and intra-RNA crosson the analysis of base-paired fragments (80,81) links (83) (see also Section IV,A). These methods proved to be very valuable in establishing the “long-range interactions” in the rRNA molecules (Le,,base-pairings between segments of RNA that are remote in the primary sequences), particularly in the case of 23 S rRNA (93), and the models derived from these data (81, 93) were subsequently refined in collaboration with Kossel et al. in Freiburg (94-96) by comparative analysis using the Z. mays chloroplast 16 S and 23 S rRNA sequences, The degree of homology between these sequences and those of E . coli (ca. 75%; see Section I) makes them ideal for comparative analysis, since the two sets of sequences are thus similar enough to be compared directly with one another in detail, and at the same time the degree of divergence is sufficient to allow a large number of “compensating base changes” to be found in double-helical regions of the structures. A comparison of the structure derived for E . coli 23 S rRNA with the corresponding structure for 2. mays chloroplast rRNA shows, for example, a total of over 450 such compensating base changes (95). The latest versions of these models for 16 S rRNA from the three research groups can be found in references 35, 92, 96, and the first complete versions for 23 S rRNA in references 33, 36, 95. It is very difficult, however, to compare these structures with one another, since they have been derived independently and are presented in different formats (particularly the 23 S rRNA structures). For this reason, in Figs. 2 and 3 we have transposed the “California” and Strasbourg” models into our format, in order to allow easy comparison of all the versions for each sequence.2 In those cases where two alternative conformations have been suggested for particular regions of the RNA (35,92),we have chosen the alternative that gives the best agreement among all three models. In the case of the 16 S rRNA (Fig. 2), it can be seen that the molecule is organized into “domains” clearly defined by the longrange interactions already mentioned. These long-range interactions are common to all three models, at least as alternatives, although it should be noted that one of them (between bases 564-570 and 880-886 in the central region) is based only on an interaction obFor the sake of simplicity, the models in Figs. 2 and 3 are referred to as “Berlin,” “California,” and “Strasbourg” models, respectively. In fact, as mentioned above, the “California” model for both 16 S and 23 S rRNA was a collaborative venture between the Californiaand Illinois groups, and the Freiburg group was involved in the construction of both the “Berlin” and “Strasbourg” models for 23 S rRNA.

”

BERLIN

‘I

b

P

m-

“

C A L I F 0 R N I A“ b

P

M

“STRASBOURG“

Ly

w

FIG.2. Comparison of secondary structure models for Escherichia coli 16 S rRNA. The sequence is numbered from the 5‘ end (every 50 bases, with a stroke at every tenth base), and is divided into three domains (a to c) as in reference 96, the bars denoting base-pairing. The “California” (35)and “Strasbourg” (92)models have been rearranged to the “Berlin” (96)format, so that identical structural elements from each model have identical orientations, to facilitate comparison.

12

RICHARD BRIMACOMBE

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served between two rather long fragments of E. coti rRNA encompassing the “binding site” for ribosomal protein S4 (97). This particular interaction is obviously crucial in maintaining the topography of the whole central part of the molecule, and, in contrast to the other longrange interactions, is so far not very well supported by comparative evidence. Figure 2 also shows that the differences between the models are largely differences in the detailed base-pairing arrangements within individual helical elements, but there are a few more serious discrepancies. On the whole, however, the degree of agreement is very satisfactory, in view of the fact that the models are derived from entirely independent sets of data. It should be noted that the interaction between bases 17-20 and 915-918 in the “California” model (domains a and b, Fig. 2) is not incompatible with the hairpin loop (bases 9-25, domain a) in the “Berlin” model; rather, it represents a “tertiary” interaction situation comparable to that proposed for 5 S RNA (see Fig. 1C and 88). The 23 S rRNA (Fig. 3) is also arranged in clear domains, and the whole molecule appears to be a giant loop, closed by an interaction between its extreme 5‘ and 3’ ends (53).Again, there is good agreement between the models with regard to the principal long-range interactions, with the notable exception that bases 578-584 are paired with 1255-1261 in the “California” model (36),as opposed to bases 805-811 in the other two models (33, 95). There are some further significant discrepancies in various parts of the models, in particular the regions between bases 450-510, 1340-1380, 1415-1585, 1900-1975, 2500-2585, and 2765-2785. However, as with the 16 S rRNA, the majority of the remaining differences are again local details of the base-pairing arrangements within well-defined domains; although it will obviously take some time before all these differences are ironed out, it is nonetheless clear that in both 16 S and 23 S rRNA the broad basis of the secondary structures can be regarded as firmly established.

B. Structures Proposed for rRNA from Other Sources The structures just described for E. coli 16 S and 23 S rRNA are, by virtue of the fact that they are based to a large extent on sequence comparisons, simultaneously general models for ribosomal RNA molecules of this size class. The slightly smaller 15 S rRNA sequences from S. cerevisiae mitochondria ( 1 1 ) and A. nidulans mitochondria (31)fit precisely into the same structure as the 16 S molecules (98,31), and this raises the question as to how far the sequence comparison approach can be extended to other ribosomal RNA molecules, outside

STRUCTURE OF RIBOSOMAL

RNA

13

the 16 S and 23 S size class. For this purpose, both the smaller mitochondrial rRNA and the larger eukaryotic cytoplasmic rRNA species (Table I) can be used for comparative analysis. In both cases the overall degree of primary sequence homology with respect to E . coli is at first sight not very high, but nevertheless there are significant regions of conserved sequence, and secondary structures can be built up around such conserved regions, using the E . coli secondary structure models as a basis. The results of these comparisons are very striking indeed. The small mitochondrial 12 S and 16 S rRNA species, although they are only about half the size of the corresponding E . coli molecules, can be arranged in secondary structures for which many elements are precisely equivalent to their 16 S and 23 S counterparts (e.g., 95,96). The reduction in size is achieved by simple amputation of secondary structural loops or by erosion of whole domains. If the E . coli structures are compared with the larger eukaryotic molecules, a precisely analogous situation is observed (e.g., 96), except that here there are extra sequences instead of missing ones. Furthermore, if the sequences within these secondary structure models are compared, it becomes immediately apparent that the degree of sequence homology between the various species is in fact a good deal higher than was at first supposed. Many short stretches of four- to five-base homology, which would not be regarded as significant in a simple comparison of the primary sequences, appear at identical positions in the various secondary structures, and seem to constitute a “core” of conservation running right through the molecules (95,96).The longer stretches of homology tend to be in single-stranded regions (e.g., 72), but there are also significant sequence homologies in double-helical regions. Interestingly, in the case ofA. nidulans 15 S rRNA there appears even to be more conservation of sequence in the double-stranded regions than in the single-stranded areas (31).It should also be noted in this context that the very highly conserved areas of sequence (e.g., bases 1390-1408 and 1492-1502 in the E . coli 16 S rRNA; see reference 42 for review) cannot be placed in secondary structures by the sequence comparison approach, simply because no compensating base changes are to be found in such regions, and this may have led to distortion of the relative importance of sequence Conservation in single-stranded as opposed to double-stranded regions. Some examples of structures deduced for rRNA molecules from different size classes are illustrated in Figs. 4 and 5. Figure 4 shows the models that we have proposed (96) for mitochondrial 12 S rRNA and eukaryotic cytoplasmic 18 S rRNA, compared with the E . coli 16 S

“BERLIN”

’.. ”

.-

CALIFORNIA ”

a

*

w

‘.

”

STRAS B OUR G

‘I

b

FIG.3. Comparison of‘secondary structure models for Escherichia coli 23 S rRNA. The structure is divided into six domains (a to f) as in reference 95, and the “Berlin” (95), “California” (36),and “Strasbourg” (33) models are compared as in Fig. 2.

STRUCTURE OF RIBOSOMAL

15

RNA

"BERLIN" f

"CALIFORNIA" f

"

ST RA S B 0 UR G " f

e

FIG.3 (continued).

IHUMAN MITOCHONDRION 125 rRNA

E.COLI 165 rRNA

5. CEREVISIA E I 0 5 r RNA

b

LIP

FIG.4. Secondary structure comparisons between human mitochondria1 12 S rRNA, Escherichia coli 16 S rRNA and Saccharomyces cereoisiae 18 S rRNA. The diagrams are sketches from the structures proposed in reference 96, arranged as in Fig. 2; corresponding secondary structural elements in the three molecules lie in the same orientation.

16

HUMAN MITOCHONDRION 165 r R N A

a

b

d

f

U E.COLI 23s r R N A

FIG.5. Secondary structure comparison between Escherichia coli 23 S rRNA and human mitochondria1 16 S rRNA. The diagrams are sketches of the structures proposed in reference 95, arranged as in Fig. 3 (cf. Fig. 4). 17

18

RICHARD BRIMACOMBE

et al.

structure; in Fig. 5, there is a corresponding comparison of mitochondrial 16 S rRNA with E . coli 23 S rRNA (95). These models were derived using the mitochondrial 12 S and 16 S sequences from mouse (10) and man (9) and the 18 S sequences from S. cerevisiae (15) and X. laevis (16). It can be seen from Fig. 4 that the extra sequences in the 18 S rRNA occur largely in the region from bases 640 to 850 (equivalent to 590 to 650 in the E . coli sequence), and that precisely this region is entirely missing in the mitochondrial 12 S rRNA. Other regions of the 18 S structure (e.g., domain c) are virtually superimposable on the corresponding E . coli structure, whereas several parts of the mitochondrial 12 S structure in domains a and c have been seriously eroded. Interestingly, in the human (but not the mouse) mitochondrial rRNA, there are a number of instances where a missing hairpin loop is replaced by a stretch of four consecutive C residues (96). Similarly, it can be seen in Fig. 5 that in domains b, d, and e of the large subunit rRNA there is strong conservation of the secondary structure, with a certain number of loops being cleanly amputated in the mitochondrial rRNA. In domains a, c, and f, on the other hand, the mitochondrial domains are considerably reduced in size. [The original publications (95, 96) should be consulted for descriptions of the primary sequence homologies in the various structures.] In a model proposed for yeast 26 S rRNA (38),an analogous situation to that of the 18 S rRNA is observed, with the “extra” sequences relative to 23 S rRNA occurring in large clusters rather than being uniformly distributed throughout the molecule. Further similar structural derivations have been made by a number of authors, proposing models for yeast 18 S rRNA (99, IOO), general structures for the small subunit rRNA (98, lo]), a model for 12 S rRNA (IOZ), and comparisons of bacterial 23 S and mitochondrial 16 S rRNA (33).While there is in general a very high degree of agreement concerning the most strongly conserved features of the structures, there is, not surprisingly, some divergence in areas where the conservation patterns are not so clear. In some cases this arises from ambiguity in the possibIe alignments of the primary sequences under comparison, and one interesting example of this is shown in Fig. 6, where two apparently significant stretches of primary sequence homology have “leap-frogged” each other in the sequence of mitochondrial 16 S rRNA (cf. 33,95),as compared to that of E . coEi 23 S rRNA. As a result, different secondary structures were derived for this part of the mitochondrial molecule (33,95),depending on which stretch of homology

STRUCTURE OF RIBOSOMAL

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FIG.6. “Leap-frogging” sequence homologies. Homologous regions of Escherichia coli 23 S rRNA and mouse mitochondrial 16 S rRNA are compared. The sequences are numbered from the 5’ ends, and the thin dashed lines indicate the homologous residues. The homology in region C occurs at similar positions (around residue 200) in the two sequences, but is less strong than the homologies in regions A and B, which occur 3’ relative to region C in the E. coli sequence, but 5’ relative to region C in the mitochondrial sequence. Homology regions A and B were used in deriving the secondary structure model for the mitochondrial rRNA in reference 95, whereas regions A and C were used for the model of reference 33; not surprisingly, the two proposed mitochondria] models are very different in this part of the molecule.

20

RICHARD BRIMACOMBE

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was taken into consideration (see legend to Fig. 6). Another situation of this type is discussed in Section I1,C. A further consequence of the secondary structure comparisons is that the small rRNA species (5.8S, 4.5 S, and 2 S rRNA; see Section I) can now be accounted for with reasonable certainty. It has already been observed (103) that 5.8 S rRNA shows a significant degree of sequence homology with the 5‘-end region of E . coli 23 S rRNA, and 2 S rRNA in Drosophila corresponds to the 3‘ region of other 5.8 S rRNA species (26).If the stem region of the secondary structure proposed for 5.8 S rRNA (104,105) is opened up (cf. 106), then the molecule becomes very similar to the 5’ region of the E . coli 23 S rRNA structure (95,36,107).This is illustrated in Fig. 7, and a comparison of Figs. 7B and 7C also suggests how the 5.8 S rRNA, which is known to be strongly hydrogen-bonded to the large subunit rRNA (e.g., 108, 109), is attached to the latter (cf. 106,110).However, it should be noted that in the secondary structure proposed for yeast 26 S and 5.8 S rRNA (38) there is no hydrogen-bonding between the extreme 5’ end of 5.8 S rRNA and the extreme 3’ end of the 26 S rRNA [cf. the interaction between the extreme 5’- and 3’ ends of E . coli 23 S rRNA (53)in Fig.

-

Y-

,_---. .

FIG.7. The equivalence of eukaryotic 5.8 S rRNA to the 5‘ terminus of Escherichia coli 23 S rRNA. (A) The “burp gun” model for trout 5.8 S rRNA (105).(B) The same model with the stem region opened up, oriented for comparison with C. (C) The 5’ region of E. coli 23 S rRNA (cf. Fig. 3, domain a). The dashed lines alongside the structure indicate sequence homologies between the 5.8 S and 23 S molecules (103,95).

STRUCTURE OF RIBOSOMAL

RNA

21

31. There is therefore a disagreement with the results of Kelly and Cox ( I l l ) ,who observed an interaction between 5.8 S rRNA and fragments isolated from the 3’ end of Neurospora crassa 28 S rRNA ( 1 1 1 ) . This point remains to be resolved. In a similar manner the 4.5 S rRNA from plant chloroplasts corresponds very clearly to the 3’ terminus of E . coli 23 S rRNA. This was proposed on the basis of sequence homology (113),and the secondary structure of the 4.5 S molecule [which in contrast to 5.8 S rRNA does not appear to be strongly hydrogen-bonded to the chloroplast 23 S rRNA (114)l corresponds precisely to the 3‘-terminal 100 bases of the E. coli 23 S structure (Fig. 3) (53,29). A slightly different secondary structure for 4.5 S rRNA has been proposed (107),but this structure is, in our opinion, both energetically and phylogenetically less favorable.

C. Evidence for Alternative Conformations (“Switches”)

The studies just described demonstrate that, despite differences in size, base composition, and base sequence, all ribosomal RNA molecules so far investigated fall into the same overall pattern of structure. This raises the question of why the high level of conservation has been necessary, and here it is difficult to avoid the conclusion that the ribosomal RNA must be initimately involved in the protein biosynthetic function. This in turn leads to the question of whether the secondary structures are more or less rigid, or whether gross changes in conformation take place in ribosome assembly or during the ribosomal cycle (cf. 84, 115, 116). There is some strong, but unfortunately still not conclusive, evidence that the latter possibility is the correct one; this evidence comes from the analysis of base-paired fragments (80,81,95), where a few RNA sequences from both E. coli 16 S and 23 S rRNA are paired with different partners in the various experiments. In particular, a fragment of 5 S rRNA was identified in association with the 23 S sequence region 1760-1770, which is base-paired to the 1980-1990 region in the secondary structure model of Fig. 3 (domain d) (95). In the 16 S rRNA, the sequence 1050-1070, which is normally associated with the sequence 1190-1210 (domain c, Fig. 2), was also found in association with the sequence 385-400 (domain a, Fig. 2) (81).In such “switch” situations, the sequence comparison approach can again be invoked, to test whether the results are experimental artifacts, or whether they represent genuine multiple conformations of the RNA. In the 5 S-23 S rRNA interaction, a similar but not entirely satisfactory structure can be drawn for the corresponding sequences in 2. mays chloroplasts (95),and also for the equivalent regions (117) in

22

RICHARD BRIMACOMBE

et al.

yeast 26 S rRNA (38) and 5 S rRNA (118). In the case of the 16 S “switch,” the base-pairing concerned is not only conserved from E . coli to 2. mays chloroplast rRNA (96), but a precisely similar switch structure can also be drawn in both S . cerevisiae and X . laevis 18 S rRNA (94,96),i.e., between sequences 1250-1280 (domain c in yeast 18 S RNA, Fig. 4) and sequences 390-410 (domain a). I n every case, this structure is a strong helix composed of 13-15 base-pairs. Similar, although less strong, complementarities can also be drawn (117) for the corresponding regions from A. nidulans (31)and S. cerevisiae (11) mitochondrial 15 S rRNA; all these structures are illustrated in Fig. 8. Additional evidence for an interaction of some kind between these two regions of the E . coli 16 S rRNA comes from a ribonucleoprotein fragment isolated by Spitnik-Elson et al. (119); the RNA from this fragment contained sequences from the regions concerned (among others) and migrated as a single complex after removal of the protein. On the other hand, there is not a trace of a corresponding switch interaction in the 12 S mitochondrial rRNA. This is not too serious, since the small mitochondrial rRNA molecules must inevitably have lost some of the properties of their larger counterparts, and this switch could be an example, since the mitochondrial rRNA is considerably eroded in the region of the 3’-proximal component of the switch (cf. Fig. 4). It should also be noted that the switch structure generates a potential “knot” in the RNA (cf. Section II,A), and it has been suggested (81)that this could be compensated by appropriate twisting of other parts of the structure, leading to a chain reaction of cooperative conformational changes in the RNA. Evidence against the existence of the switch is the fact that the secondary structure of the 18 S rRNA in the region between bases 200 and 500 (encompassing the switch sequence, Fig. 4) is currently in dispute (96,98-100). The controversy arises because two distinct sets of primary sequence homology between yeast and E . coli can be discerned in this region. One set of homology leads to the structure shown for yeast 18 S rRNA in Fig. 4 (domain a) (96),whereas the other set leads to a different structure (98-loo), in which the switch sequence is no longer in an equivalent position to the corresponding E . coli switch sequence. The former structure shows the greatest similarity to the E . coli secondary structure in the region between bases 400 and 500, whereas the latter model shows better primary sequence homology, but less secondary structural similarity. We have already mentioned (Fig. 6) a similar case where two different sets of apparently significant sequence homology led to the deduction of different secondary structures. It may well be that in such cases both sets of

23

RNA

STRUCTURE OF RIBOSOMAL

S.CEREVISIAE

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FIG.8. Possible “switch” structures in small-subunit rRNA molecules. The two structures drawn for Escherichia coli 16 S rRNA were observed experimentally (81),the “normal” structure (cf. Fig. 2) being shown 011 the left and the “switch” structure on the right. The corresponding “switch” structures shown for the other five sequences indicate the possible alternative base-pairings from the equivalent or nearly equivalent regions of the secondary structures in domains a and c (Figs. 2 and 4; and cf. refs. 96,11, 31).See text for further explanation.

homology have significance in the three-dimensional structure of the RNA, but one is also left with the uncomfortable possibility that one or both sets of homology are (despite the extremely low probability) statistical artifacts. Clearly this type of discrepancy needs to be re-

24

RICHARD BRIMACOMBE

et al.

solved before the all-important question can be settled of whether or not major conformational switches do indeed take place during the ribosomal cycle.

I11. RNA-Protein Interactions

The secondary structures described in the preceding sections are in effect “two-dimensional” maps of the RNA, which, in the ribosome itself, undergo further folding to fit into the compact ribosomal particles, together with the ribosomal proteins. The interaction between RNA and protein and the three-dimensional packing of the RNA are the subjects of this section and Section IV, respectively. Unfortunately, in relation to the foregoing sections, the state of our knowledge in both these areas is still extremely fragmentary. A. Protein Binding Sites on rRNA The “classical” approach to the study of RNA-protein interactions in the ribosome has been to make use of the fact that a number of ribosomal proteins can bind singly and specifically to their cognate rRNA molecules. Such complexes can be submitted to mild nuclease digestion, and the RNA fragments remaining bound to the protein can be analyzed. Alternatively, whole ribosomal subunits can similarly be digested and separated into RNP fragments whose protein and RNA content can then be determined. The data obtained from such experiments have been reviewed often and in detail (e.g., 35,120,121).To summarize very briefly, “binding sites” on E. coli 16 S rRNA have been localized at least partially for proteins S4, S7, S8, S15, S17, and S20 (see review reference 121, and references 97 and 122 for original literature). Protein S1 has been implicated in binding to the 3’ terminus of the 16 S rRNA (123,124),and the association of specific regions of the RNA with various groups of proteins has also been reported (120). In addition, some protein attachment sites on 16 S rRNA have been located by electron microscopy (125).In the case of 23 S rRNA, binding sites have been localized for proteins L l , L20, L23, L24 (121) and (more recently) for L11 (126),and other single proteins or groups of proteins have been isolated in association with large subfragments of the 23 S rRNA molecule (120).It should be noted here that in some cases the binding sites described in the reviews (120,121)are based on older partial sequence data for the 23 S rRNA, and precise placings within the complete 23 S sequence ( 6 )have only recently been made (33).In the case of protein L20, the exact location of the binding site is

STRUCTURE OF RIBOSOMAL

RNA

25

still unclear. The more detailed results from all these experiments are included in the summary diagrams (see Figs. 9 and 10). On 5 S rRNA, binding sites for proteins L5, L18, and L25 have been determined (121, 127-129), but there is still disagreement among the various results. To some extent this reflects the fact that the intact 5 S molecule is already smaller than most of the RNA segments isolated as binding sites for proteins on the 16 S and 23 S rRNA; as a result, attempts to localize binding sites within the 5 S rRNA are a priori focused at a more detailed level. An interesting finding emerging from these studies is that all 5 S rRNA molecules contain a bulged base at the same position (A-66 in E . coli; cf. Fig. 1); this residue is implicated in the binding of protein L18 (130).On the basis of similar features in other RNA-protein complexes, it is proposed (130) that such bulged nucleotides may represent primary recognition sites for protein-nucleic acid interactions. Some RNA-protein complexes have been described with 5 S and 5.8 S rRNA from eukaryotes (e.g., 131,132),and it has also been reported that E . coli proteins L18 and L25 can bind specifically to 5.8 S RNA (133).This led to the suggestion that eukaryotic 5.8 S rRNA is the equivalent of prokaryotic 5 S RNA, a proposition in contradiction to the role of 5.8 S RNA indicated by the comparative sequencing studies (Section 11,B). Heterologous complexes can be formed between rRNA from a variety of sources with E. coli ribosomal proteins, including proteins S4, S7, S8, S15, S17, and S20 (e.g., 134,135) and, most dramatically, protein L1(136,137),which binds specifically even to the eukaryotic 26 S rRNA from Dictyostelium discoideum (136).Such observations provide powerful support for the concept presented in the preceding section that secondary structure in the rRNA (and presumably also the three-dimensional structure) has been strongly conserved throughout evolution. The binding sites of E . coli proteins S8 and S15 on several heterologous 16 S rRNA molecules are indeed strikingly conserved (138),and in the case of protein L1, the binding site on D . discoideum 26 S rRNA was shown to correspond precisely to the equivalent binding site isolated from the cognate E . coli 23 S rRNA (136).A similar binding site could be constructed in the 28 S rRNA from X . Zaeuis

(136). However, the application of the binding site approach has limitations. In the first place, the size of the binding site varies considerably from one protein to another. The smallest site, that of L11, is 61 nucleotides long (126),whereas the binding site of protein S4 corresponds to almost one-third of the 16 S molecule (97).The binding sites corre-

26

RICHARD BRIMACOMBE

et al.

spond very closely to secondary structural domains in the rRNA (see Figs. 9 and lo), and sometimes [e.g., in the cases of S4 and L24 (reviewed in 121)] they contain noncontiguous sequences of RNA that migrate as a single complex held together by long-range interactions (see Section I1,A) even when the protein has been removed. Further, although quite a large number of proteins bind singly and specifically to the 16 S or 23 S rRNA [with a strong dependence on the method used for the isolation of both RNA and protein moieties (reviewed in 120 and 139)],it has not yet been possible to localize binding sites for more than a few of these proteins. In many cases, as already mentioned, the proteins (e.g., S7) have been isolated only in association with very large subfragments of the RNA (120). This suggests that binding sites may have been found only in those cases where the protein concerned is indeed associated with a single secondary structural domain of the RNA; in other cases (e.g., if a protein were to be supported between two such domains) then the complex would very likely disintegrate upon mild nuclease digestion, and no binding site would be detected. Regardless as to whether this surmise is correct, it is clear that additional experimental approaches must be applied in order to obtain more detailed information concerning the organization of protein and RNA within the ribosome, and for this reason a number of research groups have turned their attention to RNA-protein crosslinking techniques.

B. RNA-Protein Cross-Linking The cross-linking approach is very simple in concept, but in practice it poses a number of problems, and we therefore discuss the crosslinking of ribosomal proteins to their cognate rRNA molecules within the ribosome at some length. First of all, it is important to remember that cross-linking is a purely topographical probe; that is to say, it offers the possibility of determining neighborhoods or contacts between the various components of a system, but these neighborhoods may or may not reflect strong physical associations between the components concerned. Thus a cross-linking experiment gives a different type of information, compared to the binding site approach described above. We begin by listing the various methods and reagents used to induce RNA-protein cross-links in the E . coli ribosome and go on to discuss the analysis of the cross-linked products. Since the ultimate objective here is the construction of a detailed topographical map of the ribosome at the amino acid-nucleotide level, we pay particular attention to those cases where a precise analysis of the cross-link sites has been possible.

STRUCTURE OF RIBOSOMAL

RNA

27

1. CROSS-LINKING METHODSAND REAGENTS The most direct type of cross-linking is where an amino acid or nucleotide is activated (either chemically or photochemically) and is then able to react directly with a neighboring residue within the ribosome, generating a covalent link. For such a reaction to occur, the two residues concerned must obviously be very close to, if not actually in contact with, one another. The simplest method of this type is irradiation with ultraviolet light (see also Section IV), which generates a number of RNA-protein cross-links in the ribosome (140-144).Crosslinks can be induced in a similar manner in the presence of photoactivatable dyes such as methylene blue (145).Chemical activation leading to cross-linking can be effected by the use of N-ethyl-N’-(dimethylaminopropy1)carbodiimide (146), which presumably functions by activating carboxyl groups of aspartic or glutamic acid, with a subsequent nucleophilic attack by the amino group of a nucleotide residue. A further rather specialized example is the use of periodate to oxidize the 3’-terminal cis-hydroxyl groups of the RNA (147,148); the dialdehyde thus formed can react with protein amino groups to form Schiff bases, which can then be stabilized by borohydride reduction (cf., however, reference 149,and see below). The more usual type of cross-linking involves the use of bifunctional reagents, which generate a bridge between two nucleotide or amino acid residues. Here the distance between the cross-linked residues is obviously dependent on the distance between the two functional groups of the cross-linking reagent, but, since these reagents are seldom rigid molecules, it should be remembered that this distance will be less than or equal to the fully extended length of the reagent. Such reagents are of two kinds, symmetrical or heterobifunctional. A symmetrical reagent must contain functional groups able to react with both nucleotides and amino acids, and this type of reagent is a priori therefore rather nonspecific. Formaldehyde has been used to generate RNA-protein cross-links in the ribosome (150), but with only limited success, since the cross-linking reaction is spontaneously reversible, and therefore a detailed analysis of the cross-linked products could not be made. The chemical reactions involved here are not clear, but presumably lead to methylene bridge formation. More useful are the “nitrogen mustards” (see also Section IV), in particular bis(2-chloroethylfamine (151)and its N-methyl derivative (252,153), which react on the one side with the N-7 atom of guanine residues in the RNA, and on the other side with various amino-acid side chains, thus leading to formation of stable RNA-protein cross-links. Very similar in

28

RICHARD BRIMACOMBE

et al.

reactivity is the reagent diepoxybutane ( 154, 155), which generates cross-linked products containing a cis-diol group that can, if desired, be cleaved subsequently by periodate oxidation. Another example is trichloro-2,4,6-triazine (sym-triazine trichloride) (156),a rather rigid molecule, in which the chlorine atoms can b e substituted one at a time at different temperatures, resulting in the formation of RNA-protein cross-links. The use of a symmetrical reagent obviously leads also to the formation of intra-RNA and protein-protein cross-links, and the inherent lack of specificity can cause problems such as denaturation or aggregation (see below). Considerable attention has therefore been given to the development of heterobifunctional reagents, in which each step in the cross-linking reaction can be independently controlled. One of the first reagents of this type to be applied to the ribosome was 4-nitrophenyl 3-(2-bromo-3-oxobutane-l-sulfonyl)propionate, which contains at one end a bromoketone group (to react with a nucleotidic amino group at p H 6), and at the other end an activated ester (to react with a protein amino group at pH 8) (157). However, this reagent yielded only very low levels of RNA-protein cross-linking (151), partly because of solubility problems, but also very probably because the reagent was “too specific”; that is to say, the type of nucleotideamino acid neighborhood that could be cross-linked by the reagent may have been rather rare in the ribosome. This leads to the concept of a reagent in which one reactive group causes a specific reaction, whereas the second group is designed to react as unspecifically as possible, in order to have a high chance of forming a cross-link with any functional group in its vicinity. Photoactivatable aromatic azides have proved to be very useful for this latter purpose and have been applied in a number of systems. I n the case of the ribosome, methyl azidophenylacetimidate (158,159)and ethyl 4azidobenzamidoacetimidate (160) (both of which carry an imido ester function for specific reaction with lysine residues) have been used successfully, the photoactivatable azide group showing a preference for reaction with the nucleic acid moiety. Azido derivatives of halogenated pyridine (161)belong to this same class of reagents, but probably react with protein in a less specific manner. In another compound of similar type, (4-azidopheny1)glyoxal (162), the glyoxal group is able to react specifically either with unpaired guanine residues in the RNA or with arginine residues in the protein. With all these compounds, the specific reaction is allowed to take place first, excess reagent is then removed, and the nonspecific azide reaction, which proceeds via nitrene formation, is then induced by mild ultraviolet irradiation, The

STRUCTURE OF RIBOSOMAL,

RNA

29

reagent methyl 4-(6-formyl-3-azidophenoxy)butyrimidate (163) is also worthy of mention in this context, although it has up to now been used only as a protein-protein cross-linking agent. Reagents containing sulfhydryl groups can also be used in RNAprotein cross-linking reactions. N-Acetyl-N'-( p-glyoxyloylbenzoy1)cystamine ( 1 6 4 ) reacts with RNA and can be used to generate either intra-RNA cross-links (by coupling adjacent SH groups), or, in combination with a second photoactivatable derivative, RNA-protein crosslinks. More recently, the well-known protein-protein cross-linking reagent 2-iminothiolane (165, 166) (or methyl 4-mercaptobutyrimidate) has been used as an RNA-protein cross-linker (167)by allowing the imidoester function to react first with protein and then giving a mild ultraviolet irradiation treatment, which causes cross-linking to the RNA, presumably by reaction of the sulfhydryl function with excited pyrimidine residues. As already mentioned, the advantage of heterobifunctional reagents is that each step of the reaction can be independently controlled. Such compounds also tend to give a plateau level of reaction as the concentration of reagent is increased, but the yields of the photochemical reactions are usually very low, particularly in the case of the aromatic azides; the nitrene intermediates are so reactive that many of the activated species are lost by reaction with solvent, and the resulting maximum yield of RNA-protein cross-linked product usually corresponds to about 5% of the total ribosomal protein (e.g., 159). In contrast, the symmetrical bifunctional reagents such as diepoxybutane (154) or nitrogen mustard (mechlorethamine) (151) show an accelerating reaction with increasing reagent concentration, which implies that at higher reagent concentrations the reaction is accompanied by a randomization of the ribosomal conformation; this in turn allows more cross-links to form. This point was underscored by experiments with bisulfite, a potentially very useful cross-linking reagent that can substitute the 4-amino group of cytosine with the Eamino group of a lysine residue (168).This reagent showed no RNAprotein cross-linking at all in ribosomal subunits in the presence of magnesium, but when magnesium was removed a large yield of RNAprotein cross-links was obtained (151). In this context it should also be noted that many cross-linking agents are not water-soluble and must be dissolved in organic media, which may have an adverse effect on the ribosomal conformation, even at low concentrations. Ultraviolet irradiation causes destruction of the ribosomal conformation at high doses (142),and, as a general principle with all the cross-linking systems described above, it is clear

30

RICHARD BRIMACOMBE

et al.

that the most specific cross-links are formed under minimal conditions. Loss of ribosomal activity (e.g., in polyphenylalanine synthesizing ability) is usually rather rapid during the cross-linking reactions (e.g., 1/59),but this can be due either to the relatively large number of monovalently reacted cross-linker molecules that become attached to the ribosomal subunits or to a “freezing” of an important part of the ribosome as a direct result of a cross-link, and it does not necessarily indicate a destruction of the ribosomal conformation. Finally, the question of cleavable versus noncleavable reagents must be considered here. Of the reagents described above, only diepoxybutane (154) can be cleaved after the reaction, by virtue of the cis-diol group generated by the reaction. Another cleavable crosslinking system (169) was applied successfully to a synthetic complex between protein L24 and 23 S rRNA, but the system showed virtually no RNA-protein cross-linking in ribosomal subunits (170). In any event, it must be remembered that, after cleavage of the cross-linker, a cross-linked site becomes in general indistinguishable from a site where the reagent has formed a monovalent adduct. In other words, if the ultimate objective of the cross-linking experiment is the analysis of the sites of cross-linking (see below), then the use of a cleavable cross-linking agent may not offer any advantage.

2. ANALYSESOF

CROSS-LINKED PRODUCTS

The first stage in any analysis of the products of an RNA-protein cross-linking reaction with the ribosome is the identification of the proteins involved, and, insofar as they are known, the cross-linked proteins are listed in Table I1 for all of the reagents decribed in the foregoing section. In the earlier experiments the identities of the proteins cross-linked to rRNA were inferred in a negative manner by their disappearance from the pattern of total ribosomal proteins on polyacrylamide gels (140, 141, 147). Since most of the cross-linking reactions are accompanied by a number of side reactions, the disappearance of a protein is obviously a very unsatisfactory criterion, which can in any case be observed only if the level of reaction is very high; as noted in the preceding section, this is not usually the case. Identification with the heIp of antibodies to the individual proteins has been used (148,142), but in most of the recent analyses (e.g., 149151, 159-162), the cross-linked proteins have been identified by removing non-cross-linked protein on dodecyl sulfate sucrose gradients, isolating the RNA plus cross-linked protein, digesting this with nucleases, and analyzing the resulting protein-oligonucleotide complexes on two-dimensional polyacrylamide gels. The existence of a covalent

TABLE I1 RNA-PROTEIN CROSS-LINKING AGENTSO Bridge length

Principal proteins identified as cross-linked to RNAb

Reagent or method of cross-linking

(A)

Ultraviolet irradiation Methylene blue activation N-Ethyl-”-(dimethy laminopropy 1)carbodiimide Periodate oxidation Formaldehyde

<1 <1 <1

s7c s3, s4, s5, s 7 S4, S5, S7, S9/11, S13, S15-18

<1 2

30 S subunit proteins

Bis(2-chloroethy1)amine N-Methylbis(2-chloroethy1)amine

5 5

Diepoxybutane sym-Triazine trichloride (trichloro-2,4,6hiazine) 3-(2-Bromo-3-oxobutane-l-sulfonyl)propionate Methyl 4-azidophenylacetimidate Ethyl 4-azidobenzamidoacetimidate

4 3

s1, S21d S3, S4, S5, S9/11, S13, S21 (S7, S12, S18) S3, S4, S5, S9/11, S13 S2, S3, S4, S5, S6, S7, S9/11, S13 Many s3, s 4

8 6 8

s3, s4, s5, s7, s9

4-Azido-3,5-dichloro-2,6-difluoropyridine 4-Azidopheny lgly oxal N-Acetyl-N’-(4-g1yoxyloylbenzoyl)cystamine 2-Iminothiolane

3 5 >10 5

50 S subunit proteins L2, L4 L2, L3 -

L13, L27, L32 (L2, L6, L16) L1, L2, L3, L5 L1, L2, L3, L5, L20

Reference 142,143 145 146 148 150 151 153

Many L2

154, 155 156

S3, S4, S9/11, S13

L2

157

s3, s4, s5, s 7

L1, L2, L4, L25/29

-

(S12, S13/14, S16, S17, S18) s4, s7, s9 s2, s3, s4, s5, s7, (S12)

160

161 162

-

S3, S4, S5, S7, S8, S9/11, S10

159

164 L2, L4, L6, L21, L23, L27, L29

167, 171

a The cross-linking reagents or methods are listed in the order in which they appear in the text. The bridge length gives the approximate maximum distance between the active functional groups of the cross-linker. b Only positively identified proteins are listed. In many cases other proteins either were not resolved or may have been present as aggregates (see text). Proteins in parentheses are minor products; dashes indicate that no data are available. c Other authors (140, 144) found many more proteins cross-linked (see text). Other authors (149)found many more proteins cross-linked (see text).

32

RICHARD BRIMACOMBE

et al.

cross-link can be demonstrated by using 32P-labeled RNA, in which case the isolated complexes carry a measurable 32P label. In this way, quite low levels of cross-linking of individual proteins can be observed, but the analyses are not unequivocal. In the first place, the cross-linked proteins have altered mobilities in the gels, and this can lead to ambiguities in identification, especially of the smaller proteins. Even if a cleavable cross-linker is used (e.g., 154), the presence of monovalently reacted or cross-linked reagent molecules on the protein can alter its mobility. Further, in all cases the gels of cross-linked proteins show relatively large amounts of protein aggregates; it cannot be excluded that some proteins are particularly prone to aggregation after treatment with the cross-linking reagent, and therefore do not appear on the gels. For this reason the lists of proteins given in Table I1 should be regarded as minimum descriptions of the cross-linked products, and estimates of the yields of individual cross-linked proteins on gels (e.g., by radioactivity measurements) may be quite misleading. A further problem is that the low levels of cross-linking generally observed raise the uncomfortable possibility that only a small, possibly inactive, subpopulation of the ribosomal particles are involved in a particular cross-link, but this uncertainty is common to any study based on chemical modification of the ribosome. Three points from Table I1 deserve special mention. First, in the case of ultraviolet-induced cross-linking, only proteins S7, L2, and L4 become cross-linked at low doses of irradiation (142,143).Turchinsky et al. (144)have described the cross-linking of some additional 30 S proteins, and Gorelic (140, 141) has reported that virtually all the proteins can be cross-linked. At least in the latter instance, the crosslinking was carried out under such vigorous conditions that the ribosomal conformation was certainly destroyed (142).Second, in the case of the periodate oxidation method, Czernilofsky et al. (148) found cross-linking only to proteins S1 and S21, whereas we found a large number of proteins to be involved (149).There are several possible explanations (149)for this discrepancy, which has yet to be resolved. Third, no proteins have so far been reported cross-linked to 5 S rRNA. This is almost certainly a simple consequence of the methods used for the isolation of the RNA-protein cross-linked complexes, which are in general designed only to separate proteins cross-linked to the large 16 S or 23 S rRNA molecules. It should also be noted in this context that some RNA-protein cross-links across the subunit interface (i.e., 30 S proteins cross-linked to 23 S rRNA, or 50 S proteins cross-linked to 16 S rRNA) have been reported using diepoxybutane (154, 1 5 9 ,

STRUCTURE OF RIBOSOMAL

RNA

33

whereas in our experiments with methyl azidophenylacetimidate (159) no such cross-links could be detected in significant amounts. These latter experiments (159) were conducted in such a way as to exclude any possible confusion resulting from low-level cross-contaminations between the 16 S and 23 S rRNA during the analyses of the cross-linked products. Table I1 shows that most if not all of the ribosomal proteins can be cross-linked to their cognate rRNA molecules by one method or another, which reflects an obviously very large number of RNA-protein neighborhoods in the ribosome. It is clear that useful information can be extracted from these data only if the precise sites involved in the cross-links can be identified on both protein and RNA, but so far this has been achieved in only a few instances. In the periodate oxidation cross-linking method (148), the crosslink should a priori involve the 3' terminus of the rRNA, but this has not been positively established (148).The first example of a positive identification of an RNA-protein cross-link site was that of the ultraviolet-induced cross-link between protein S7 and 16 S RNA within the E . coli 30 S subunit (172,173).Here, the site on protein S7 was established by isolating a cross-linked 32P-Iabeled S7-oligonucleotide complex, digesting this with trypsin, and then searching for peptides containing the 32P label. This led to the identification of a short labeled peptide containing five amino acids, in which one amino acid (Met-114) was absent from the sequence ( 1 74) and was presumed to be the site of the cross-link (172).The corresponding site on the RNA was established ( 1 73) by first isolating partially digested 32P-labeled cross-linked S7-RNA fragments containing 30-40 nucleotides, and determining the positions of these fragments in the 16 S rRNA sequence ( 4 ) .This showed unequivocally that the sequence region concerned spanned residues 1215-1255, and that one octanucleotide (CUACAAUG, 1234-1241) was absent. Second, an S7-oligonucleotide cross-linked complex was isolated from a total ribonuclease T1 digest of the S7-RNA complex, and further digestion of this complex with ribonuclease A released C, U, AC, and G. Finally, a third digestion with ribonuclease T2 released two A-residues from the protein-oligonucleotide product after ribonuclease A digestion, which clearly implicates U-1240 as the site of the cross-link. A subsequent report (175), in which only the second of these three digestions was conducted, also led to the same release of C, U, AC, and G from the S7-oligonucleotide complex, but the authors claimed that the crosslink site was within a different oligonucleotide (ACCUCG, positions 1261- 1266).

a

5 20

FIG.9. The secondary structure of Escherichia coli 16 S rRNA (cf. Fig. 2), showing sites of interaction with protein and sites of intra-RNA cross-linking.Protein binding sites are indicated by the regions boxed in with dashed lines, and in the cases of proteins S4 and S8, S15, the smallest of the various published binding sites (97,122)are shown. RNA-protein cross-link sites are indicated by the arrows (e.g., protein S7 at residue 1240),and intra-RNA cross-link sites by the arrowed loops (e.g., at positions 600-650). The site of cross-linking to tRNA is also shown. Sites of psoralen-induced cross-links in the RNA have been localized between approximate positions 930 and 1540, 510 and 1540, 450 and 1540,O and 1540,950 and 1400, and 550 and 870. See text for references.

35

f

L6

5SRNA. L5. L18. L25

FIG.10. The secondary structure of Escherichia coli 23 S rRNA (cf. Fig. 3), showing protein binding sites, RNA-protein cross-link sites, and intra-RNA cross-link sites, as in Fig. 9.

36

RICHARD BRIMACOMBE et

al.

The important point in the analysis of the S7-cross-link site just described is that the presence of the cross-linked product could be positively demonstrated, right u p until the last stage of the analyses. In a number of early experiments, such positive demonstrations were lacking. For example, in our own experiments with formaldehyde (ISO), the spontaneously reversible nature of the cross-linking reaction made it impossible to establish the presence of cross-linked protein on small RNA fragments, and the existence of cross-links on such fragments could only be inferred. Similarly, in attempts (176)to identify sites of cross-linking within synthetic complexes of S20 or S4 with 16 S RNA, the absence of certain RNA fragment bands from digests of the cross-linked complexes was taken as evidence that these bands contained the cross-linked sites, and peptides missing from a corresponding tryptic digest of an S7-16 S RNA complex (177) were assumed to be involved in the cross-linking. In all these cases, the negative nature of the analysis leaves considerable doubt as to the validity of the conclusions. More recently, the site of ultraviolet-induced cross-linking between protein L4 and 23 S rRNA within the E . coli 50 S subunit was determined (178) [Tyr-35 in protein L4 (179)was linked to U-615 in the 23 S RNA (6)l.In this case the analysis of the cross-linking site on the RNA was made using a two-dimensional gel electrophoresis system (178)that allows a number of cross-link sites to be analyzed simultaneously. The system has since been applied to 50 S subunits crosslinked with 2-iminothiolane, and cross-link sites on the 23 S RNA could be positively established for proteins L4, L6, L21, L23, L27, and L29 (167).In the 30 S subunit, a cross-link site for protein S8 was identified (171).On the other hand, analysis of the corresponding sites of cross-linking on the proteins still poses problems, very likely as a result of the monovalently reacted cross-linker molecules present on each protein in addition to the reagent molecule actually involved in the cross-link (171). Finally, the cross-linking of protein S1 to the 16 S RNA in 30 S subunits, using ethyl 4-azidobenzamidoacetimidate, has been described (180). In this experiment, the protein was first treated with reagent, then added to the ribosome and irradiated to generate the cross-link. Protein S1 was therefore the only protein to be involved in any cross-linking, and this is an elegant example of how the independent control of the functional groups in a heterobifunctional reagent can be exploited. It should be noted, however, that the authors were unable to demonstrate positively the presence of the cross-linked protein on the RNA fragment (residues 861-889) they isolated. This

STRUCTURE OF RIBOSOMAL

RNA

37

leaves open the possibility that the protein was in fact cross-linked to a different RNA fragment, which in turn was bound to residues 861889 by hydrogen-bonding. The results of all the cross-linking experiments described above are included in the summary diagrams (Figs. 9 and 10).

IV. Three-Dimensional Packing of €. coli rRNA A. Intra- RNA Cross- Linking Many of the problems discussed in the preceding section are also relevant to the question of intra-RNA cross-linking, and several of the cross-linking agents (Table 11) also lead to formation of intra-RNA cross-links. Compounds such as sulfur mustard (181),nitrogen mustard (152),or diepoxybutane (182)have long been known to generate cross-links in nucleic acids, and all have been shown to cause intraRNA cross-linking in E . coli ribosomal subunits (183,184,185,respectively), as has also simple ultraviolet irradiation (83,95).Another symmetrical bifunctional reagent that has been useful is 1,4-phenyldiglyoxal(1,4-phenylenebisglyoxal)(82,89),and photoactivatable derivatives of psoralens [which intercalate into double-helical regions of the nucleic acid (e.g., 186)]have been successfully applied (187,188). As is the case with the RNA-protein studies, analysis of the sites of cross-linking is obviously essential here, but until recently this has proved to be difficult. Wagner and Garrett (82)were able to isolate a short cross-linked oligonucleotide from the stem region of 5 S rRNA after reaction with 1,4-phenylenebisglyoxal and ribonuclease T1 digestion. By exploiting the G-specificity and ready reversibility of this cross-linking reagent, they could pinpoint the cross-link to residues G-2 and G-112 in the 5 S rRNA sequence (cf. Fig. 1).Similarly, Rabin and Crothers (189) identified a psoralen cross-link in the stem region of 5 S rRNA after partial digestion, making use of the photoreversibility of the psoralen reaction. Monovalently reacted cross-linker molecules cause considerable difficulties in this type of analysis, and recently the problem was overcome for 1,4-phenyldiglyoxal by coupling such monoaddition products to a solid support (89).With this sytem, a new cross-link was identified between residues G-41 and G-72 in the E . coli 5 S RNA (89).The significance of this cross-link for the threedimensional structure of 5 S molecule is described in Section II,A (cf. Fig. l),and these examples serve to illustrate the general principle in intra-RNA cross-linking studies, uiz., the cross-link may be “within”

38

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the secondary structure (as for the G-2 to G-112 cross-link), or it may give an indication of a neighborhood or tertiary structural interaction “outside” the secondary structure (as for residues G-41 and G-72). The analytical methods used in these intra-RNA cross-linking experiments with 5 S rRNA have not so far been applicable to 16 S or 23 S rRNA, since the nuclease digestion products of the larger molecules are too complex. Here, some intra-RNA cross-links have been localized by other methods, involving either two-dimensional gel electrophoretic techniques (83) or electron microscopy (187,188).The first of these methods relies on the fact that RNA fragments from partial digests of rRNA are strongly retarded in denaturing gel systems if they contain an intra-RNA cross-link. The cross-linked fragments can therefore be readily distinguished from the large numbers of noncross-linked fragments on a suitable two-dimensional gel system and can be isolated and subjected to oligonucleotide analysis. Provided that the cross-link lies in a “characteristic” region of the sequence with respect to such an oligonucleotide analysis, the cross-link site can be located, and this has been achieved for a number of ultravioletor diepoxybutane-induced cross-links in both 16 S (83,185)and 23 S rRNA (95).However, so far all the cross-links found by this method are “within” the secondary structure (see above), which almost certainly reflects the selective nature of the partial digestion conditions used. The locations of these cross-links are included in the summary diagrams (Figs. 9 and 10). In contrast, all the intra-RNA cross-links localized by electron microscopy are “outside” the secondary structure. Here Cantor and his colleagues (187,188)used psoralen derivatives to generate cross-links in 16 S rRNA or 30 S subunits, and the cross-link sites were localized to a first approximation by measuring the dimensions of the resulting loop structures that appeared in electron micrographs of the crosslinked RNA. Some of the loops observed are very large, and were originally interpreted as evidence that there are “knots” in the 16 S rRNA structure (190) (cf. Sections II,A and C), since psoralen is an intercalating molecule and long double-helices could be constructed around the cross-link sites. More recently, however, these authors (191, 192) suggested that their cross-links reflect tertiary structural interactions [cf. the tertiary interaction described above for 5 S rRNA (88, 89) (see Fig. l)].In the latest series of experiments, Wollenzien and Cantor used a retarding denaturing gel system to separate the various different cross-linked species of 16 S RNA according to the size of the cross-linked loop (191).In addition, the problem of determining the polarity of the cross-linked RNA molecules in the electron

STRUCTURE OF RIBOSOMAL

RNA

39

microscope has been solved by hybridization and covalent cross-linkage of specific DNA fragments to one end of the rRNA molecule (192). The positions of these psoralen cross-links are indicated in the legend to the summary diagram (Fig. 9) and will obviously be of considerable importance in helping to “fold” the 16 S rRNA secondary structure into three dimensions. A final example of intra-RNA cross-linking is the localization of a cross-link between 16 S rRNA and tRNA. This involves the hypermodified uridine at the 5‘-anticodon position of tRNAVa’or tRNASer, which can be cross-linked very specifically by mild ultraviolet irradiation to 16 S RNA when the tRNA is bound at the ribosomal P-site (193). The cross-link site on 16 S RNA was pinpointed by a series of partial digestions (193-195); it is residue C-1400 in the 16 S rRNA sequence, and a precisely analogous cross-link has since been found in yeast 18 S rRNA (196). The cross-linked nucleotide occurs in a sequence region that is very highly conserved in all small subunit rRNA molecules so far sequenced (see reference 42). The position of this crosslink is also indicated in Fig. 9.

B. Electron Microscopy of rRNA within Ribosomal Subunits

There is now a reasonabIe consensus of agreement among the various models proposed for the E . coli ribosomal subunits on the basis of electron microscopy, and a large amount of data has been collected concerning the distribution of the individual ribosomal proteins on these models (reviewed in 197,198).However, the ribosomal subunits maintain their gross morphological features as seen under the electron microscope even when a large proportion of the proteins has been removed (199). This suggests that it is the rRNA that is largely responsible for determining the overall architecture of the subunits, and some specific regions of the rRNA molecules have been localized by immune electron microscopy. The 3’ ends of isolated rRNA molecules can be oxidized with periodate and allowed to react with a suitable hapten. The modified RNA is subsequently reconstituted into ribosomal subunits and treated with hapten-specific antibody. Using this approach, several groups have made electron microscopic localizations of the 3‘ ends of E . coli 5 S (200,201),16 S (202,203),and 23 S rRNA (201,204);the positions of the 3’ ends in the 30 S and 50 S subunits are illustrated in Fig. 11. A haptenization method has been developed for the 5’-terminal phosphate group of an RNA molecule, and this has been used to locate the 5’ end of 16 S rRNA, by a similar immunological procedure

40

RICHARD BRIMACOMBE

et al.

BASES 925-1395 I PROTllNS 57, 59 510, 511.51?1

BASES 2090-2200

165 RNA Il’-fNOl

165 RNA IS-INOI

N6-OIMETHYL ADENOSINE

30s SUBUNIT 50s SUBUNIT FIG.11. Location of regions of 16 S and 23 S rRNA in Escherichia coli 30 S and 50 S subunits, derived from electron microscopic studies. The sketches crudely summarize the data from several models and several laboratories (see text for details and references).

(205). The 5‘ end of 23 S rRNA has not yet been localized, but it should be noted that, if the secondary structure model (Fig. 3) is correct, then the 5’ and 3’ termini of 23 S rRNA should be virtually coincident, as is also the case with 5 S rRNA (Fig. 1). The use of antibodies against modified bases in the rRNA to pinpoint specific RNA regions by immune electron microscopy has been successful so far only in the case of N6-dimethyladenosine (206,207). Two such residues occur adjacent to one another at a position 23 nucleotides from the 3’ end of the 16 S rRNA ( 4 , 5 ) ,and their location in the 30 S subunit is also indicated in Fig. 11. It is noteworthy that antibodies to a haptenated tRNA molecule crosslinked to the ribosomal P-site (cf. Section IV,A) also bind predominantly to the same region of the 30 S subunit (208), and, taken together with the established involvement of the 3‘ terminus of 16 S rRNA in messenger RNA recognition (209, 210),this region of the 30 S subunit is now rather clearly implicated as being the messenger decoding site. The location of some regions of the 16 S and 23 S rRNA molecules can be inferred from the data on RNA-protein interactions (see Section 111,A). Thus, nucleotides 2090-2200 in 23 S rRNA, which constitute the binding site for protein L1 (136,137),should lie in the vicinity of the antibody attachment site for protein L1, which occupies a very characteristic position on the 50 S subunit (211)(Fig. 11). Similarly, it is highly probable that nucleotides 925-1395 in the 16 S rRNA constitute the “head” of the 30 S subunit, by virtue of the association of this section of the RNA with proteins S7, S9, S10, S14, and S19 (120), all of which have antibody binding sites in the “head” region

STRUCTURE OF RIBOSOMAL

41

RNA

(197,198). Other regions, such as the 16 S rRNA sequences binding to protein 54 or to proteins S6, S8, S15, and S18, can also be tentatively placed (see, e.g., 35). The location of the 5 S rRNA relative to 23 S rRNA within the 50 S subunit is not yet clear; a possible “switch” involving sequences in 5 S and 23 S rRNA has already been discussed (Section II,C), and a ribonucleoprotein fragment containing proteins L5, L18, and L25 together with 5 S rRNA and a short section of 23 S rRNA has also been described (212) (see Fig. 10). These both offer possible contact sites for 5 S rRNA (cf. Fig. ll),but, on the other hand, a strong base-paired interaction previously proposed between 5 S rRNA and residues 143154 of the 23 S rRNA (213) has since been discounted on the basis of comparative evidence (36). Finally, it should be noted in this context that there is a considerable body of evidence concerning the accessibility (i-e.,surface topography) of specific regions of the rRNA within the subunits or at the subunit interface. The evidence comes from modification with kethoxal as well as from nuclease digestion studies, both described in Section I1,A. The identities of oligonucleotides that occur at the subunit interface have been inferred from their differential reactivity toward kethoxal in 70 S ribosomes as opposed to isolated subunits (214, 215). In addition, the identities of oligonucleotides released from the 30 S subunit by digestion with nuclease have been analyzed (216), and digestion with ribonuclease H has been used to probe the sites of binding to the RNA of complementary oligodeoxynucleotides (217, 218). This type of information cannot as yet be interpreted in terms of the precise three-dimensional folding of the rRNA, but will obviously have to be taken into account in any detailed models of the ribosomal subunits in the future.

V. Outlook We show in this article how the sequence information now availa-

ble for ribosomal RNA from many organisms has been used to help construct secondary structure models for these molecules. It is to be expected that these models will become progressively refined during the next few years, as further sequences are determined and more experimental data are collected, but, as we have stressed (Section II,A), the principal features of the secondary structures for both 16 S and 23 S rRNA are already well established. As a result, the emphasis in working out the three-dimensional topography of the ribosomal RNA in situ in the subunits must now focus on the questions of the

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tertiary folding of the RNA, its interaction with ribosomal protein, and the organization of the interface between the subunits. Here, again as we have stressed (Section 111), our knowledge is still very fragmentary. It is not yet clear how far the comparative sequencing approach, so useful in establishing the secondary structures, will be of help in these areas, and the main burden will be the collection of suitably direct experimental information. Cross-linking techniques, despite their drawbacks and the difficulties involved in their application, still offer the best hope for delivering the type of detailed information that is needed, and for this reason we have devoted a considerable amount of space to a description of the current status of these techniques (Sections I11 and IV). As a preliminary picture of the folding of the rRNA takes shape, it will gradually become possible to correlate the structure with data from other sources, in particular from electron microscopy. In this field, progress is being made in the three-dimensional reconstruction and averaging of electron micrographs of ribosomal subunits, both in ribosome microcrystals (219) and as individual subunits (220, 221), and the shape of the ribosomal RNA within the subunits is beginning to emerge. Coupled with improved data from immune electron microscopy (cf. Section IV,B), and with other data on the three-dimensional arrangement of the proteins [e.g., protein-protein cross-linking data (222), or neutron scattering studies of the protein distribution (223)], this should lead to an increasingly coherent picture of the topographical organization of RNA and protein in the subunits. However, here it must be added that establishing the principles that govern the actual physical interaction between proteins and RNA remains one of the most difficult problems to approach in this field. As far as function of the ribosomal RNA is concerned, one emerging technique is particularly worthy of mention, namely the site-directed mutagenesis of rRNA. This involves the construction via rDNA of specific ribosomal RNA mutants, and it offers the possibility of examining the importance of selected areas of the RNA in a manner that has hitherto been unthinkable. Some preliminary results have already been reported (224), and this new technology could well become a vital link in correlating the ribosomal RNA function with the type of structural studies that we have outlined in this article.

ACKNOWLEDGMENTS The authors are very grateful to Dr. H. G. Wittmann for his critical reading of the manuscript, and to many colleagues who have sent us their results prior to publication.

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121. R. A. Zimmermann, in “Ribososmes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 135. Univ. Park Press, Baltimore, Maryland, 1980. 122. R. Miiller, R. A. Garrett, and H. F. Noller,JBC 254,3873 (1979). 123. A. E. Dahlberg and J. E. Dahlberg, PNAS 72,2940 (1975). 124. R. C. Yuan, J. A. Steitz, P. B. Moore, and D. M. Crothers, NARes 7,2399 (1979). 125. M. D. Cole, M. Beer, T. Koller, W. A. Strycharz, and M. Nomura, PNAS 75, 270 (1978). 126. F. J. Schmidt, J. Thompson, K. Lee, J. Dijk, and E. Cundliffe,JBC 256, 12301 (1981). 127. J. Zimmermann and V. A. Erdmann, M o l . Gen. Genet. 160,247 (1978). 128. S. Douthwaite, R. A. Garrett, R. Wagner, and J. Feunteun, NARes 6,2453 (1979). 129. M. Speek and A. Lind, NARes 10, 947 (1982). 130. D. A. Peattie, S. Douthwaite, R. A. Garrett, and H. F. Noller, PNAS 78,7331 (1981). 131. N. Ulbrich, K. Todokoro, E. J. Ackermann, and I. G. Woo1,JBC 255,7712 (1980). 132. K. Todokoro, N. Ulbrich, Y. L. Chan, and I. G. Wool, JBC 256, 7207 (1981). 133. P. Wrede and V. A. Erdmann, PNAS 74,2706 (1977). 134. M. Geisser and G. A. Mackie, EJB 70, 159 (1976). 135. D. L. Thurlow and R. A. Zimmermann, PNAS 75,2859 (1978). 136. R. L. Gourse, D. L. Thurlow, S. A. Gerbi, and R. A. Zimmermann, PNAS 78,2722 (1981). 137. C. Branlant, A. Krol, A. Machatt, and J. P. Ebel, NARes 9, 293 (1981). 138. D. Thurlow and R. A. Zimmermann, in preparation. 139. K. H. Nierhaus, Cum. Top. Microbiol. Immunol. 97, 81 (1982). 140. L. Gorelic, Bchem 14,4627 (1975). 141. L. Gorelic, BBA 390, 209 (1975). 142. K. Moller and R. Brimacombe, MoZ. Gen. Genet. 141,343 (1975). 143. 0. G. Baca and J. W. Bodley, BBRC 70, 1091 (1976). 144. M. F. Turchinsky, N. E. Bronde, K. S. Kussova, G. G. Abduraschidova, E. V. Muchamedganova, J. N. Schatsky, T. F. Bystrova, and E. J. Budowsky, EjB 90,83 (1978). 145. D. E. Zook and S . R. Fahnestock, BBA 517,400 (1978). 146. C. Chiarrutini and A. Expert-Bezancon, FEBS Lett. 119, 145 (1980). 147. R. A. Kenner, BBRC 51,932 (1973). 148. A. P. Czemilofsky, C. G. Kurland, and G. Stoffler, FEBS Lett. 58,281 (1975). 149. J. Rinke and R. Brimacombe, MoZ. Biol. Rep. 4, 153 (1978). 150. K. Moller, J. Rinke, A. Ross, G. Buddle, and R. Brimacombe, EJB 76, 175 (1977). 151. E. Ulmer, M. Meinke, A. Ross, G. Fink, and R. Brimacombe, M o l . Gen. Genet. 160, 183 (1978). 152. P. E. Geiduschek, PNAS 47,950 (1961). 153. P. Maly, unpublished results. 154. H. G. Baumert, S. E. Skold, and C. G. Kurland, EJB 89,353 (1978). 155. S. E. Skold, Biochimie 63, 53 (1981). 156. C. Oste and R. Brimacombe, Mol. Gen. Genet. 168, 81 (1979). 157. G. Fink and R. Brimacombe, Biochem. SOC. Trans. 3,1014 (1975). 158. G. Fink, H. Fasold, W. Rommel, and R. Brimacombe, Anal. Biochem. 108, 394 (1980). 159. J. Rinke, M. Meinke, R. Brimacombe, G. Fink, W. Rommel, and H. Fasold,JMB 137, 301 (1980). 160. R. Millon, M. Olomucki, J. Y. LeGall, B. Golinska, J. P. Ebel, and B. Ehresmann, EJB 110,485 (1980).

STRUCTURE OF RIBOSOMAL

RNA

47

161. R. Millon, J. P. Ebel, F. le Goffic, and B. Ehresmann, BBRC 101, 784 (1981). 162. S. M. Politz, H. F. Noller, and P. D. McWhirter, Bchem 20, 372 (1981). 163. J. A. Maassen, E. N. Schop, and W. Moller, Bchem 20, 1020 (1981). 164. A. Expert-Bezanqon and D. Hayes, EJB 103, 365 (1980). 165. R. R. Traut, A. Bollen, T. T. Sun, J. W. B. Hershey, J. Sundberg, and L. R. Pierce, Bchem 12,3266 (1973). 166. A. Sommer and R. R. Traut,JMB 106,995 (1976). 167. I. Wower, J. Wower, M. Meinke, and R. Brimacombe, NARes 9, 4285 (1981). 168. I. Boni and E. I. Budowsky,]. Biochem. (Tokyo) 73,821 (1973). 169. C. Oste, R. Parfait, A. Bollen, and R. R. Crichton, M d Gen. Genet. 152,253 (1977). 170. C. Oste and R. Brimacombe, unpublished results. 171. J. Wower, I. Wower, and R. Brimacombe, unpublished results. 172. K. Moller, C. Zwieb, and R. Brimacombe, JMB 126, 489 (1978). 173. C. Zwieb and R. Brimacombe, NARes 6, 1775 (1979). 174. J. Reinbolt, D. Tritsch, and B. Wittmann-Liebold, FEBS Lett. 91, 297 (1978). 175. B. Ehresmann, C. Backendorf, C. Ehresmann, R. Millon, and J. P. Ebel, EJB 104, 255 (1980). 176. B. Ehresmann, C. Backendorf, C. Ehresmann, and J. P. Ebel, FEBS Lett. 78,261 (1977). 177. B. Ehresmann, J. Reinbolt, C. Backendorf, D. Tritsch, and J. P. Ebel, FEBS Lett. 67, 316 (1976). 178. P. Maly, J. Rinke, E. Ulmer, C. Zwieb, and R. Brimacombe, Bchem 19, 4179 (1980). 179. M. Kimura and B. Wittmann-Liebold, FEBS Lett. 121, 317 (1980). 180. B. Golinska, R. Millon, C. Backendorf, M. Olomucki, J. P. Ebel, and B. Ehresmann, EJB 115,479 (1981). 181. P. Brookes and P. D. Lawley, BJ 77, 478 (1960). 182. P. D. Lawley and P. Brookes,JMB 25, 143 (1967). 183. R. M. Malbon and J. H. Parish, BBA 246,542 (1971). 184. C. Zwieb, A. Ross, J. Rinke, M. Meinke, and R. Brimacombe, NARes 5, 2705 (1978). 185. C. Zwieb, unpublished results. 186. L. Musajo and G. Rodighiero, Photochem. Photobiol. 11,27 (1970). 187. P. Wollenzien, J. E. Hearst, P. Thammana, and C. R. Cantor,JMB 135,255 (1979). 188. P. Thammana, C. R. Cantor, P. Wollenzien, and J. E. HearstJMB 135,271 (1979). 189. D. Rabin and D. M. Crothers, NARes 7,689 (1979). 190. C. R. Cantor, in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 23, Univ. Park Press, Baltimore, Maryland, 1980. 191. P. L. Wollenzien and C. R. Cantor,JMB 159, 151 (1982). 192. P. L. Wollenzien and C. R. Cantor, PNAS 79, 3940 (1982). 193. R. A. Zimmermann, S. M. Gates, I. Schwartz, and J. Ofengand, Bchem 18, 4333 (1979). 194. B. H. Taylor, J. B. Prince, J. Ofengand, and R. A. Zimmermann, Bchem 20, 7581 (1981). 195. J. B. Prince, B. H. Taylor, D. L. Thurlow, J. Ofengand, and R. A. Zimmermann, PNAS 79,5450 (1982). 196. J. Ofengand, P. Gornicki, K. Chakraburtty, and K. Nurse, PNAS 79,2817 (1982). 197. G. StofRer, R. Bald, B. Kastner, R. Liihrmann, M. Stoffler-Meilicke, and G. Tischendorf, in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Normura, eds.), p. 171. Univ. Park Press, Baltimore, Maryland, 1980.

48

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et al.

198. J. A. Lake, in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 207. Univ. Park Press, Baltimore, Maryland, 1980. 199. V. D. Vasiliev and V. E. Koteliansky, FEES Lett. 76, 125 (1977). 200. I. N. Shatsky, A. G. Evstafieva, T. F. Bystrova, A. A. Bogdanov, and V. D. Vasiliev, FEBS Lett. 121,97 (1980). 201. M. Stoffler-Meilicke, G. Stoffler, 0. W. Odom, A. Zinn, G. Gamer, and B. Hardesty, PNAS 78,5538 (1981). 202. I. N. Shatsky, L. V. Mochalova, M. S. Kojouharova, A. A. Bogdanov, and V. D. Vasiliev,JMB 133, 501 (1979). 203. R. Luhrmann, M. IStoffler-Meilicke, and G. Stoffler, Mol. Gen. Genet. 182, 369 (1981). I 204. I. N. Shatsky, A. G!.Evstafieva, T. F. Bystrova, A. A. Bogdanov, and V. D. Vasiliev, FEBS Lett. 122, 251 (1980). 205. L. V. Mochalova, I. N. Shatsky, A. A. Bogdanov, andV. D. VasiIiev,/MB 159,637 (1982). 206. S. M. PolitL and D. G. Glitz, PNAS 74, 1468 (1977). 207. P. Thammana and C. R. Cantor, NARes 5,805 (1978). 208. M . Keren-Zur, M. Boublik, and J. Ofengand, PNAS 76, 1054 (1979). 209. J. Shine and L. Dalgarno, PNAS 71, 1342 (1974). 210. J. A. Steitz and K. Jakes, PNAS 72,4734 (1975). 211. E. R. Dabbs, R. Ehrlich, R. Hasenbank, B. H. Shroeter, M. Stoffler-Meilicke, and G. Stoffler, JME 149,553 (1981). 212. C. Branlant, A. Krol, J. Sriwidada, and R. Brimacombe, EJB 70,483 (1976). 213. W. Herr and H. F. Noller, FEBS Lett. 53,248 (1975). 214. W. Herr and H. F. Noller,JME 130,421 (1979). 215. W. Herr, N. M. Chapman, and H. F. Noller, J M B 130,433 (1979). 216. N. L. Teterina, A. M. Kopylov, and A. A. Bogdanov, FEBS Lett. 116,265 (1980). 21 7. A. S. Mankin, E. A. Skripkin, N. V. Chichkova, A. M. Kopylov, and A. A. Bogdanov, FEBS Lett. 131, 253 (1981). 218. C. Backendorf, C. J. C. Ravensbergen, J. Van der Plas, J. H. Van Boom, G. Veeneman, and J. Van Duin, NARes 9, 1425 (1981). 219. A. E. Yonath, J. Mussig, B. Tesche, S. Lorenz, V. A. Erdmann, and H. G . Wittmann, Biochem. Znt. 1,428 (1980). 220. V. Knauer, R. Hegerl, and W. Hoppe,JMB, in press. 221. H. Oettl, R. Hegerl, and W. Hoppe,JME, in press. 222. R. R. Traut, J. M. Lambert, G. Boileau, and J. W. Kenny, in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 89. Univ. Park Press, Baltimore, Maryland, 1980. 223. P. B. Moore in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura eds.), p. 11 1. Univ. Park Press, Baltimore, Maryland, 1980. 224. R. L. Course, M. J. R. Stark, and A. E. Dahlberg,JMB 159,397 (1982).

Structure, Biosynthesis, and Function of Queuosine in Transfer RNA SUSUMUNISHIMURA Biology Diuision, National Cancer Center Research Institute, Tsukiji 5-1-1, Chuo-ku, Tokyo,Japan

I. History of Discovery of Queuosine . 11. Distribution of Queuosine in Animals and Plants and Isolation of Hexose-Containing Queuosine Derivati 111. Biosynthesis of Queuosine in tRNA. . . . A. In uiuo Experiments. . . . . . . . . . . . . . B. tRNA-Guanine Transglycosylase . . . . . . . . . . . . . . . . . . . . . . . . . . C. Biosynthesis of Queuosine in the tRNAs of Animal Cells, . . . . . IV. Properties of tRNA-Guanine Transglycosylase . . . . . . . . . . . . . . . . . . V. Biosynthesis of the Queuine Skeleton. . . . . VI. The Presence of Queuine-Lacking tRNA in VII. Change of the Ratio of Queuine to Guanine in tRNA during Cell Differentiation . . . . . . . . . . . . . . . . . . . . . . . . ......... VIII. Biological Function of Queuosine in tRNA. IX. Possible Approaches to the Use of Queuine Analogs for Canc Diagnosis and Cancer Chemotherapy . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

50 55 56 56 57 59 60 62 63 66 67

70 71

The structural complexity of many modified nucleosides, their location at specific sites in tRNA molecule, and their rather ubiquitous presence in a wide range of organisms encourage the belief that they play an important role in tRNA function. However, the elucidation of the functions of these nucleosides in tRNA is difficult, because it seems that each one has only a marginal effect on tRNA function, and biochemical systems in which such marginal effects can be assayed are not easily available. No lethal mutation for a deficiency of a modified nucleoside in tRNA has thus far been found. This is also true for queuosine, even though it is the most complicated nucleoside so far discovered and is located in the anticodon. However, the marginal effect should not be overlooked, because it may be important for the control if higher orders of regulatory processes of cell function. Among many “modified” nucleosides present in tRNA, queu49 Progress in Nucleic Acid Research

and Molecular Biology, Vol. 28

Copyright 8 1983 by Academic Press, lnc. All rights of reproduction in any form reserved. ISBN 0-12-540028-4

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SUSUMU NISHIMURA

osinela is quite unique in its structure and biosynthesis. In addition, its involvement in cell function, cell differentiation, and tumorigenesis has become an important subject, as shown by many recent publications. The first two-thirds of this article is concerned with the chemistry and biochemistry of queuosine, i.e., with isolation, structural characterization, location, and biosynthesis, and is thus quite straightforward. The last third deals with the function of queuosine. Since no final conclusion on the function of queuosine can be drawn, I summarize all the results so far obtained, even though contradictory in some cases, and mention some ongoing experiments, hoping that ideas for future investigations and/or approaches may eventuate. Finally, this is not a completely comprehensive review and is not intended to cover all the literature. Instead, I have tried to describe the history and development of the queuosine problem, mostly along with current work in my laboratory.

1. History of Discovery of Queuosine Queuosine was discovered in the first position of the anticodon of Escherichia coli tRNATyrin 1967, independently by three groups [RajBhandaryet al. (I),Goodman et al. ( 2 , 3 ) ,and Doctor et al. (4j1, during the course of nucleotide sequence analysis of this tRNA. At that time, queuosine, as yet unnamed, was abbreviated as Q, G*, or R by the respective workers,lb and was thought to be a simple derivative of a. The systematic names for queuosine are as follows: IUPAC: 2-Amino-5-{[((1S,4S,5R)-4,5-dihydroxy-2-cyclopenten-l-y~)amino]methy~}1,7-dihydro-7-~-~-ribofuranosyl-4H-pyrrolo[ 2,3-d]pyrimidin-4-one; Chemical Abstracts: [ 1S-( la,4@,5P)]-2-Amino-5-{[(4,5-dihydroxy-2-cyclopenten-ly~)amino]methyl}-l,7-dihydro-7-~-~-ribofuranosyl-4~-pyrrolo[2,3-d]pyrimidine-4-one. The IUPAC name may be simplified by the use of 7-deazaguanosine as a root, thus eliminating both the 2-amino prefix and the whole of the second line, to yield 7-{5-[ (( 1S,4S,5R)-4,5-Dihydroxy-2-cyclopenten-l-y~)amino]methy~}-7-deazaguanosine. b. Soon after the discovery of the new base and nucleoside, and before the structures were known, they were called Q (or base Q) and Q (or nucleoside Q), respectively. Since it was unsatisfactory to have only a one-letter symbol for a nucleoside (or base), the proposal was made that trivial names based on the sound of Q be generated. Hence, the names queuine for the base (which retains the -ine ending of adenine and quanine) and queuosine for the nucleoside (retaining the -osine ending of adenosine and guanosine) were proposed and are now generally accepted (e.g., Chemical Abstracts Name Index, 1977 Cumulative). The three-letter abbreviations Que and Quo (similar to Ado and Guo) for base and nucleoside, respectively, have been proposed, but are in little use. The single-letter symbol Q is retained for the nucleoside, queuosine.

QUEUOSINE IN TRANSFER

RNA

51

guanosine. However, it actually took 8 years to determine its chemical structure. The presence of queuosine in mammalian tRNA was also suggested in 1969 by Rogg and Staehelin (S), although its precise characteristics were not reported. Later, we found that queuosine was present in E . coli tRNAHiS,tRNAA'", and tRNAAspp, again in the first position of the anticodon (6).2Thus it was shown that there is a strict relation between the presence of queuosine and codon recognition of E . coli tRNA: all E . coli tRNAs that recognize U and C in the third position and A in the second position of the codon contain queuosine. In fact, no other E . coli tRNA so far sequenced contains queuosine (10,

11). For the structural characterization of queuosine, large-scale isolations of queuosine from E . coli tRNA were necessary. Several hundreds of grams of unfractionated E . coli tRNA were hydrolyzed by pancreatic RNase, and the QpUp derived from all Q-containing tRNAs was isolated by DEAE-Sephadex and A-25 column chromatography. The QpUp thus isolated was hydrolyzed to give Q p by RNase T 2 digestion, and Q p was separated by Dowex 1 column chromatography (12).Although the amount of queuosine thus obtained was relatively large (-20 mg), the structural eludication turned out to b e quite difficult, because no one thought that the heterocyclic nucleus itself is not guanine but rather 7-deazaguanine (i.e., pyrrolo[2,3-d]pyrimidin-4one). The elucidation of the structure of queuosine was made possible by collaboration with J. A. McCloskey (mass spectrometry) and N. J. Oppenheimer [nuclear magnetic resonance (NMR)] (12).The most important information was obtained by mass spectral analysis of several derivatives of queuosine. [The details of the structural characterization of queuosine have been reviewed (13).1(See Figs. 1-4.) Queuosine is the only nucleoside thus far isolated from ribonucleic acid in which the purine skeleton has a 7-deaza structure. Only 7-deazaadenosine derivatives such as the antibiotics toyocamycin, tubercidin, and sangivamycin, isolated from culture media of Streptornyces, were known at that time.3 A year after the elucidation of the structure of queuosine, the stereochemistry of the cyclopentenediol side chain was established by chemical synthesis of the two stereoisomers and their 'H NMR spectral analysis (17).Finally, Goto et al. determined the absolute configuThe complete nucleotide sequences of E . coli tRNAHss,tRNAA'",and tRNAASP were deduced later (7-9). 3 See Suhadolnik's review of these and other antibiotic nucleosides ( 1 4 ) .Tanaka and his co-workershave reported the isolation of the antibiotic cadeguomycin (7-deaza7-carboxyguanosine)from Streptornyces (15, 16).

R

NH

""YY H O OH

D

FIG.1. Structure of queuosine (A), queuine (B), mannose-containing queosine (C), and galactose-containing queuosine (D).

Wavelength

FIG. 2. Ultraviolet absorption spectra of queuosine (a), hexose-containing 0.1 N HCl; -, HzO; , 0.1 N queuosine (b), and queuine (c). NaOH. 52

_______

I00

1

i

53

A 1 I SN31NI 3 A Ilti13M

l9

25 103

I0j

42 I

1656

911 I

m/e

FIG.3. Mass spectrum of trimethylsilylated queuosine. Data were taken from Kassai et al. (12).

54

SUSUMU NISHIMURA

100

75

>

50

!z

ln

$ 25 z

w

2

FIG.4. Mass spectrum of trimethylsilylated queuine.

ration of queuosine by total chemical synthesis of the two diastereomers of queuosine having the P-D-ribosyl group and comparison of their physicochemical properties with natural queuosine (18,19). I n 1979, the three-dimensional structure of queuosine was determined by X-ray crystallography of queuosine 5’-monophosphate (pQ) (20). (Fig. 5). The coriformation of pQ in aqueous solution has been analyzed by NMR (S. Yokoyama, Z. Yamaizumi, S. Nishimura, and T. Miyazawa, unpublished results). The NMR analysis (spin-coupling constant and nuclear Overhauser effect) showed that the anti form is predominant for pQ.

FIG.5. Three-dimensional structure of queuosine 5’-mononucleotide in stereo view.

QUEUOSINE IN TRANSFER

RNA

55

II. Distribution of Queuosine in Animals and Plants and Isolation of Hexose-Containing Queuosine Derivatives

In 1973, White et al. (21) reported that queuosine and its derivatives are also present in those Drosophila tRNAs that correspond to the same amino acids as the E . coli Q-containing tRNAs. This was before the structure of queuosine from E . coli was known. The development of a simple method for the isolation of queuosine and its identification by mass spectrometry made it possible to determine the distribution of queuosine in tRNAs from organisms other than E . coli (22). Unfractionated tRNAs from various sources were hydrolyzed completely by RNase T2 to yield queuosine 3’-phosphate (Qp), which was separated from most other nucleotides by Dowex-1 column chromatography, since Q p possesses one additional positive charge from the secondary amino group. The nucleotide was finally purified by two-dimensional thin-layer chromatography (23).Mass spectral analysis of trimethylsilylated queuosine was very useful for the identification of queuosine (12, 22). Queuosine has been found in such diverse organisms as rat, rabbit, hagfish, Lingula unquis, and wheat germ (22). It is present in the tRNAs of vertebrates, chordates, brachiopods, echinoderms, and plants, in addition to bacteria. However, yeast tRNA contains no queuosine (22),nor can yeast synthesize it (22a)or incorporate exogenous queuine, unlike other eukaryotes, into their G-tRNAs (22a). Another interesting finding is the isolation of a queuosine derivative from animal tRNAs (22).A queuosine derivative designated originally as Q* behaved differently from queuosine in two-dimensional thin-layer chromatography. The structural characterization of Q* was performed by mass and NMR analysis, using large quantities of Q* isolated from 5 g of rabbit liver tRNA (24). Q* turned out to be a mixture of two components (manQ and galQ), having D-mannose or Dgalactose with a P-glycoside attachment to the oxygen atom on carbon4 of the cyclopentenediol. Recently it was found that manQ and galQ can be separated by liquid chromatography on pBondapak C18 with methanol-formic acid as eluent (K. Nihei, J. A. McCloskey, and S. Nishimura, unpublished results). The question of whether such hexose-containing queuosines are present in any particular tRNA species could easily be answered by application of lectin-Sepharose column chromatography (25).Among Q-containing tRNA species from rabbit liver, rat liver, and rat ascites hepatoma, only tRNAAsPwas retained by concanavalin-A-Sepharose

56

SUSUMU NISHIMURA

(Con A-Sepharose) affinity column chromatography, while tRNATYr from these sources was purified by the use of another lectin, Ricinus communis, bound to Sepharose. The results clearly demonstrate that tRNAAspcontains manQ whereas tRNATyrcontains galQ. The sugars attached in the O4 position of the cyclopentenediol in queuosine are therefore not randomly distributed among the four Q-containing tRNAs, but appear to be present only in specific tRNAs, Con ASepharose column chromatography also proved to be useful for the easy isolation of pure tRNAAsP(25-27). Using this principle, a largescale (0.34 pmol) isolation of tRNAAspisoacceptors 1 and 2 from liver tRNAs (0.3 g) in 88%purity in one step has been described (27). The two, both containing Q, can be resolved by subsequent chromatography on other columns (27). In addition, Garcia and Singhal showed that tRNATyrcan be separated better by the use of a column of lectin immobilized on agarose using a spacer arm with a length of 10 A (28). Q-containing tRNAs can also be purified effectively by acetylated DBAE-cellulose column chromatography, because the cis-diol group of the cyclopentenediol moiety of queuosine interacts specifically with the acetylated DBAE-cellulose (29). This procedure has been used for the isolation of mammalian Q-containing tRNAs (25, 30), as well as presursors of bacterial Q-containing tRNA (31). Group preparations of Q-tRNAs from non-Q-tRNAs have been achieved on CMBcellulose and reversed-phase boronate columns. The latter is especially useful in the preparation of high-specific-activity Q-tRNAs (31a).

111. Biosynthesis of Queuosine in tRNA A. In Vivo Experiments Before the structure of queuosine was known, its precursor was thought to be quanosine. The reason for this conclusion was that tRNATyr,synthesized in large quantity in E . coli cells by infection of 48OSu- phage carrying the E . coli tRNATyr-' gene, contained guanosine in place of queuosine in the first position of the anticodon (3). We showed that the carbon atom at position 2 of guanine is incorporated into queuosine, whereas carbon-8 is not, by feeding [Z-14C1and [8-14C]guanine separately to a guanine-requiring mutant of Salmonella (32). This indicated that carbon-8 in the precursor guanine molecule is excluded together with nitrogen-7 during the course of synthesis of queuosine from guanine. Thus the mechanism of conver-

QUEUOSINE IN TRANSFER

RNA

57

sion of G to Q seems similar to that of the biosynthesis of toyocamycin, 7-cyano-7-deazaadenosine,in Streptomyces rimosus (33, 34) or the biosynthesis of pteridines (35-37). At the time of this discovery, it was generally thought that all modified nucleotides in nucleic acids are derived from parent nucleotides already in the polynucleotide chain. However, it was difficult to imagine how the drastic modification involved in the conversion of G to Q could take place without breaking the N-C glycosyl linkage in the polynucleotide. Therefore, we proposed that conversion of G to Q proceeds either at the nucleotide, nucleoside, or base level, followed by replacement of the guanine moiety in tRNA by the “Q-base” (queuine), leaving the backbone of the tRNA intact (32).

B. tRNA-Guanine Transglyc~sylase~ I n 1962, Marks et al. reported that free guanine is exclusively incorporated into 4 S RNA, but not into ribosomal RNA or mRNA, by rabbit reticulocytes (38).The labeling of4 S RNA by [14C]guanine was thought to be due to the addition of a [ 14C]adenosine moiety to the terminal residue of a tRNA by the tRNA adenylyltransferase (EC 2.7.7.25) reaction after enzymic conversion of the [ l4C]guanine to [14C]ATP. Hankins and Farkas followed up this observation and in 1970 made the following interesting findings (39).

1. [ 14C]Guanine incorporated into tRNA was recovered as [ 14C]GMP after alkaline hydrolysis of the tRNA.

2. When the labeled tRNA was hydrolyzed by RNase T1, most of the radioactivity was recovered in a single long oligonucleotide. This indicated that the incorporation of guanine takes place in a very specific manner, and that it is not a terminal addition to either the 3’or the 5’ end of the tRNA. Farkas et al. later showed that the incorporation of guanine occurs only in a particular tRNA species, probably a minor isoaccepting tRNAHis,by comparison of the pattern of guanine incorporation with amino-acid-acceptor activity of tRNA fractionated by reversed-phased column chromatography (40).In addition, they showed that cell-free extracts of rabbit reticulocytes can catalyze the incorporation of guanine into unfractionated yeast tRNA as well as into homologous tRNA (41).They ascribed this activity to a “guanylating” enzyme. See the summary and amplification in Section I and Tables I and I1 in “Queuine: An Addendum” by R. P. Singhal in this volume. [ed.]

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We tested the possibility that this “guanylating” enzyme might bring about the biosynthesis of queuosine in tRNA by catalyzing the incorporation of queuine by a transglycosylase reaction and incorporating guanine into tRNA by the reverse reaction (12).We tested a variety of purified E . coli tRNA species in a cell-free rabbit reticulocyte system (42). The results clearly demonstrated that only Q-containing tRNAs (tRNATyr,tRNAHis,tRNAAsn, and tRNAAsP)are substrates for guanine incorporation. In addition, sequence analyses of [ 14C]guanine-labeledtRNAHisshowed that the guanine incorporated replaces only the queuine originally located in the tRNA. At that time, it was still possible to argue that the enzyme catalyzes only release of queuine, and that incorporation of queuosine into tRNA proceeds by a different mechanism. There was a discrepancy as to which tRNA species was the substrate for guanine incorporation between homologous and heterologous sytems. Farkas et al. showed that the minor isoacceptors tRNAHiS and tRNAAsnof rabbit reticulocytes that incorporated queuine were the “undermodified” species, having guanosine in the anticodon, and that the major tRNAHisand tRNAAsnspecies that already contained queuosine were not utilized (43,44). For the study of the biosynthesis of queuosine in tRNA, the E . coli system was found to be very useful. A guanine “insertion” enzyme (later called tRNA-guanine transglycosylase) was found in E . coli cells and purified to a homogeneous state (45). In addition, we found that methyl-deficient tRNA and tRNAs from E . coli mutants selected for queuosine deficiency contain analogs of queuosine, i.e., 7-aminomethyl-7-deazaguanosine and 7-cyano-7-deazaguanosine7 which were possibly precursors of queuosine (46, 47). For tests on whether queuine or the bases corresponding to these queuosine analogs are actually incorporated into tRNA by tRNA-guanine transglycosylase, queuine and 7-aminomethyl-7-deazaguanine were chemically synthesized [they could not be obtained from naturally occurring nucleosides because of the resistance of the N-C glycosyl bond to acid treatment (48, 49)]. 7-Cyano-7-deazaguanine was prepared enzymically from tRNA containing 7-cyano-7-deazaguanosine, using E . coli tRNA-guanine transglycosylase (later it too was synthesized chemically) (49). 7-Aminomethyl-7-deazaguanine and 7-cyano-7-deazaguanine were effectively incorporated into isoacceptors of E . coli tRNATYr and tRNAAsn,which contained normal guanine instead of queuine (G-tRNATYrand G-tRNAAsn) (Fig. 6). The bases incorporated were found in the first position of the anticodon. On the other hand, queuine was not incorporated into G-tRNA (49).

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RNA Nlli

FIG.6. Structures of queuosine precursors found in Escherichia coli mutant tRNA and methyl-deficient tRNA: 7-aminomethyl-7-deazaguanosine (A) and 7-cyano-7-deazaguanosine (B).

Since free 7-aminomethyl-7-deazaguanine was found in E . coli cells, it seems to be the actual substrate incorporated into tRNA in uivo (49). Its conversion to queuine must proceed at the polynucleotide level. These results demonstrated that formation of queuosine in tRNA proceeds by a unique posttranscriptional modification mechanism; this is the first example of the insertion of a modified base into the polynucleotide chain without breakage of a phosphodiester bond, catalyzed by tRNA-guanine transglycosylase. The involvement of tRNA-guanine transglycosylase in the biosynthesis of queuosine in tRNA was confirmed by the isolation of an E . coli mutant that lacks this enzyme (50).The tRNA from this mutant is undermodified with respect to queuosine, having guanosine in place of queuosine in the first position of the anticodon.

C. Biosynthesis of Queuosine in the tRNAs of Animal Cells tRNA-guanine transglycosylases have been isolated from rat liver (51), rabbit erythrocytes (52), Ehrlich ascites cells (53),and Drosophila (54), as well as from rabbit reticulocytes (41). Some of these enzymes have been extensively purified. The purified rat liver tRNAguanine transglycosylase catalyzes an exchange of queuine as well as its precursors (7-aminomethyl-7-deazaguanine and 7-cyano-7-deazaguanine) and guanine for the residue originally located in the first position of the anticodon of “undermodified” tRNA (G-tRNA) (51). This result in animal cells is in contrast to the earlier observation in E . coli that queuine is not a substrate for the E . coli tRNA-guanine transglyc~sylase.~ Since the K , value for queuine of the rat liver enzyme is much lower than the values for the bases of queuosine precursors or of guanine (9.2 x M), it was concluded that the actual substrate for tRNA-guanine transglycosylase in queuosine biosynthe-

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sis in vivo in mammalian cells may be queuine, but not queuine precursors. The conclusion that queuine itself may be the substrate for tRNAguanine transglycosylase in mammalian systems was substantiated by recent work of Katze and Farkas. Katze isolated a “Q factor” from animal serum and amniotic fluid (55).Addition of this factor to the medium caused a shift from G-tRNA to Q-tRNA in L-M cells in tissue culture. The in vitro reaction using pure rabbit erythrocyte tRNAguanine transglycosylase and rabbit reticulocyte G-tRNAHiSwith pure Q factor produced tRNA chromatographically indistinguishable from Q-tRNAHis(56). “Q factor” was later shown to be queuine by massspectral analysis (57). When germfree mice are maintained on a chemically defined diet known to contain all essential constituents of the rodent diet but no queuosine or queuine, Q in Q-specific tRNAs gradually disappears with a concomitant increase of G-tRNAs (Q-lacking tRNAs) (58, 59). The decrease of manQ- and galQ-containing tRNAs (tRNAAspand tRNATy*,respectively) was less than that of the Q-containing tRNAs (tRNAHi”and tRNAAsn).After keeping mice for 1 year in a germfree condition on a Q-free diet, all four Q-specific tRNAs become undermodified. Addition of small amounts of queuine, or of Q-tRNAs to the diet induced a shift of G-tRNAs to Q-tRNAs in those mice in several days. These findings strongly suggest that ( a )mice cannot synthesize queuine and must get it exogenously, like vitamins; and ( b ) queuine exogenously added is effectively utilized for the formation of Q-containing tRNA in vivo. Significant amounts of free queuine have been found in common plant and animal food products, such as coconut milk, wheat germ, tomato, bovine milk, yogurt, goat milk, and human milk (60). The biosynthesis of queuosine in tRNA apparently cannot proceed in the primary transcript (61).When cloned yeast tRNATyrgenes were microinjected into Xenopus laevis oocyte nuclei, the tRNA genes were transcribed, and unspliced tRNA precursor was accumulated in the nuclei. The biosynthesis of queuosine as well as of isopentenyladenosine in the anticodon proceeded only after removal of the intervening sequence, although the precursor tRNA already contained five other modifications.

IV. Properties of tRNA-Guanine Transglycosylase As discussed previously, tRNA-guanine transglycosylases have been isolated from a wide variety of sources, such as E . coli, mammalian cells, and Drosophila (45, 52-54). The enzyme has also been

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isolated from wheat germ and extensively purified (W. Farkas, personal communication). The molecular weights of tRNA-guanine transglycosylases differ markedly with the sources of the enzymes. The enzyme from E . coli is a single polypeptide and has a molecular weight of 46,000, whereas that from rabbit erythrocytes, M , = 104,000, consists of two subunits of M , 60,000 and 43,000. The M , of the enzyme from rat liver has been calculated to be 80,000. The K , for guanine is 5.3 x M ( E . coli), 1.5 x lo-' M (erythrocytes) and 8.3 X M (rat liver). The pH optimum is 7.0-7.3, and no energy source such as ATP is required for the reaction. The enzyme from E . coli requires magnesium ion, whereas the rat liver enzyme is not stimulated by magnesium ion. As discussed previously, the bases that can be incorporated into tRNA differ depending upon the source of the enzyme and of the tRNA used as acceptor. The E . coli enzyme catalyzes incorporation of queuine precursors as well as guanine into G-tRNAs, whereas mammalian enzymes (rat liver and erythrocytes) incorporate queuine as well as its queuine precursors into mammalian G-tRNAs. The enzyme from wheat germ can utilize queuine, but not the precursor, 7-aminomethyl-7-deazaguanine (W. Farkas, personal communication). Mammalian enzymes (rat liver, Ehrlich ascites cells, rabbit reticulocytes) can exchange guanine for the queuine in mature E. coli Q-tRNAs of nonmammalian origin only. On the other hand, the E . coli enzyme cannot catalyze an exchange of guanine with any Q-tRNAs. In any case, the bases that can participate in tRNA-guanine transglycosylase exchanges are restricted to guanine or queuine analogs. For example, in the case of the E . coli enzyme, most other purine analogs are not utilized, indicating that the base specificity is very high. An oxygen atom at position 6, an amino group at position 2, and a lone pair of electrons on the nitrogen atom at position 7 in the purine skeleton seem to be necessary for efficient incorporation. Among the various modified bases tested, only 7-methylguanine inhibits the incorporation of guanine into G-tRNA, although 7-methylguanine itself is not incorporated into G-tRNA. The rat liver enzyme does not utilize 7-deazaguanine and 7-methyl-7-deazaguanine. Jacobson et al. (62) showed that various pteridine analogs inhibit incorporation of queuine into G-tRNA in a mammalian tissue culture system, and that pterin inhibits the erythrocyte tRNA-guanine transglycosylase competitively with guanine (Kr = 9 x M). However, the pterin is not incorporated into tRNA (62). It should be noted that the synthetic guanine analogs 8-azaguanine and 6-thioguanine, which act as anticancer agents, are efficiently incorporated into GtRNA by the rat liver enzyme (51).

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A nucleoside doxyribosyltransferase (EC 2.4.2.6) that catalyzes an exchange of bases at the deoxyribonucleoside level occurs in various bacteria (63).The reaction mechanism of tRNA-guanine transglycosylase is similar. Both enzymes catalyze exchange of base by cleavage of a N-C glycoside bond without requiring an energy source. However, tRNA-guanine transglycosylase has a more strict base specificity and, unlike nucleoside deoxyribosyltransferase, catalyzes the reaction in a specific location of the polynucleotide chain. No other tRNAs, or RNAs other than tRNA, or DNA are acceptors for E . coli tRNAguanine transglycosylase (N. Okada and S. Nishimura, unpublished results). Recently, a different type of enzyme that catalyzes incorporation of bases into polynucleotide chains has been isolated. The activity, found in human fibroblasts, catalyzes incorporation of purine bases at apurinic sites in double-stranded DNA without phosphodiester bond cleavage (64, 65). An enzymic activity that inserts adenine and guanine from the corresponding deoxyribonucleoside triphosphates into appropriate apurinic sites of depurinated double-stranded DNA has been also isolated from E . coli (66). These enzymes may effect steps in “repair” mechanisms.

V. Biosynthesis of the Queuine Skeleton

The mechanism of queuine biosynthesis is not fully understood at present. As discussed previously, during the conversion of guanine to queuine, the carbon at 8 position is lost together with the N-7 of the purine skeleton (32). The biosynthesis of queuine may proceed in a fashion similar to the biosynthesis of toyocamycin, pteridine, folate, and riboflavin. The reason why mammalian cells cannot synthesize queuine might be related to the similarity of the enzymic pathway of the synthesis of queuine to that of the vitamins. When methionine is withheld from growing E . coli methionine-requiring mutants, Q-containing tRNAs become undermodified and produce G-tRNAs together with tRNA containing queuine precursors (67,68).However, no radioactivity of [2J4C]methionine or [rnethyL3H or -14C]methionine is incorporated into queuosine in tRNA when E . coli cells are grown in the presence of these radioactive methionines. Thus, how methonine is involved in the biosynthesis of queuine is not clear. One hypothesis is that the ribose moiety of S-adenosylmethionine synthesized from methionine is used for the synthesis of the cyclopentenediol moiety of queuosine.

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RNA

The enzyme responsible for the attachment of mannose to queuosine in tRNA was isolated from rat liver (69).The novel mannosyltransferase catalyzing this reaction using GDP-mannose as donor was purified from a particulate-free soluble enzyme fraction and found to be strictly specific for E . coli tRNAAsP.No other E . coli Qcontaining tRNAs were utilized as acceptor for mannosylation. These results, together with the anomeric configuration of mannose in manQ, indicate that no lipid intermediate is involved in the biosynthesis of manQ.

VI. The Presence of Queuine-Lacking tRNA in Tumor Cells

Discoveries of new isoacceptor tRNAs or increases of particular isoacceptor tRNAs often came about when tRNAs isolated from tumor cells, particular tissues, cells in certain stage of differentiation, or tissue culture cells grown under certain conditions were fractionated by various column chromatographic procedures (70-74). Since 1965, many workers were attracted by these observations because such isoacceptors might have some regulatory function in those particular cells, as proposed by Borek for tumor cells (75).It was recognized that not all amino acids have an abnormal cognate tRNA. Among abnormal isoacceptors found in tumor cells, those for phenylalanine, tyrosine, histidine, asparagine, aspartic acid, and lysine were the most frequent (73, 74). It is now established that the tRNAs for these amino acids in normal tissues contain hypermodified nucleosides, such as wyeine (“Y-base”) (tRNAPh“)( 76,77), queuosine (tRNATy’,tRNAHiS,tRNAAsn, and tRNAAsP),and N-(9-~-~-ribofuranosyl-2-methylthiopurin-6-yl)carbamoyllthreonine (tRNALYs)(78). The presence of such hypermodified nucleosides can influence tRNA behavior in column chromatography. Since queuosine contains one additional positive charge, a tRNA containing guanosine in place of queuosine (G-tRNA) is expected to be eluted later than normal Q-containing tRNA (Q-tRNA) in reversedphase or benzoylated DEAE-cellulose column chromatography. Therefore it is very likely that those isoacceptor species of Q-containing tRNAs that elute later than “normal” tRNAs are formed by a failure in queuosine biosynthesis in the posttranscriptional process. Although the final proof of this hypothesis was obtained later by determination of the nucleotide sequences of tumor-specific tRNAs, as discussed below, several other indirect experiments indicate that it is the case.

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HO

,4

0

I3

FIG. 7. Derivatization of queuine moiety of tRNA by periodate (A) or cyanogen bromide (B).

1. Queuosine reacts with periodate (6,12) and with cyanogen bromide (79,80)to yield queuosine derivatives lacking the extra positive charge (Fig. 7). In fact, elution of the Q-tRNA of Drosophila on reversed-phased column chromatography is specifically altered by treatment with periodate or cyanogen bromide (81). The behavior of mammalian tRNAs with or without treatment by cyanogen bromide on acetylated DBAE-cellulose or Aminex A-28 columns indicates that the Q-specific tRNAs from Walker 256 rat mammary tumor almost totally lack queuosine and hexose-containing queuosine, while normal rat liver Q-specific tRNAs contain queuosine (82). Similarly, the Q-specific tRNAs in tissue-cultures of mouse fibroblasts appear to be hypomodified with respect to queuosine (83, 84). 2. Upon addition of free queuine to growing L-M cells, two isoacceptors of G-tRNAAsPappear in the positions where normal Q-tRNAAsP isoacceptors normally elute (55, 56). In fact, this shift of the elution position of G-tRNA in the L-M cell tissue culture system was later used as a sensitive assay of queuine content (56, 60). Although isoacceptor 2 (the major isoacceptor) of rat liver tRNAAsPdiffers from isoacceptor 2 of rabbit liver in several nucleotide positions (84a),each contains Q in the same p o ~ i t i o n . ~ The complete nucleotide sequences of the major tRNAAspspecies from rat liver and a minor tRNAAsPpresent specifically in rat ascites hepatoma were determined by combined use of several postlabeling procedures (85).5The nucleotide sequence of rat ascites hepatoma tRNAAspis the same as that of rat liver tRNAAsp,except that the manQ of rat liver tRNAAspis replaced by normal G in rat ascites hepatoma tRNAAsP.It is interesting to note that the other modified nucleosides 5

See sequences listed by Singhal et ul. ( 1 1 ) and by Sprinzl and Gauss (10).

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were the same in the two tRNAAspspecies. The same conclusion arises from the determination of the nucleotide sequences of Walker 256 mammary carcinosarcoma tRNAAs"and normal rat liver tRNAAsn(86). Thus, it was concluded that the lack of Q in the tumor tRNAs is due to hypomodification with respect to queuosine in the posttranscriptional process; transcription of different tRNA genes is not involved, Also, queuosine is present in the first position of the anticodon of rat liver mitochondria1 tRNAs, but not in rat hepatoma mitochondria1 tRNAs (K. Randerath, personal communication). Lack of queuosine in tRNA can easily be detected by observing the incorporation of radioactive guanine into unfractionated tRNA by E . coli tRNA-guanine transglycosylase, since the E. coli enzyme catalyzes an exchange of guanine into undermodified tRNAs that contain guanine in place of queuine, but not into Q-containing tRNA (87). With this analytical method, it was demonstrated that tRNAs isolated from various tumors, including the slowly growing Morris hepatoma and SV40-transformed cells, incorporate (exchange) considerable amounts of guanine (hence contain more G-tRNAs), whereas tRNAs isolated from all normal tissues, except regenerating rat liver, incorporate scarcely any. A slight incorporation of guanine was also observed with tRNA from undifferentiated cells, such as rat fetal liver or rabbit reticulocytes (87). Sheep reticulocytes and fetal liver contain significant amounts of G-tRNA whereas adult liver does not (88).In very young rats and in leukemia cells significant amounts of G-tRNA are present (89). Hepatoma tRNA from cancer patients is also a good acceptor for guanine (S. Nishimura, unpublished results). The amounts of G-tRNA in murine erythroleukemic cells decrease markedly as the cells differentiate into mature erythroid cells, without altering the initial growth rate (90). The decrease may be detected at an early stage of erythroid differentiation. Decrease in the amount of G-tRNA in the cells is effectively blocked by 12-O-tetradecanoylphorbol 13-acetate (TPA), which inhibits differentiation of these cells. Similar results using erythroleukemic cells have been obtained by other workers (91). These results clearly demonstrate that the presence of G-tRNA in tumor cells is not due merely to their fast growth rate. Thus, the presence of G-tRNA is probably specific to tumor cells and undifferentiated cells. Since regenerating rat liver tRNA contains a considerable amount of G-tRNA, it is evident that the presence of G-tRNA is not directly linked to tumorigenesis (87). Some properties of malignant cells may transiently be expressed during liver regeneration as in

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the case of the change of the isozyme pattern of pyruvate kinase, a typical indicator of hepatoma cells. The presence of G-tRNA is probably linked to the specific metabolic nature of tumor cells. The reason why tumor cells contain G-tRNA is not known. Transglycosylase with the same specificity was obtained in approximately equal amounts from tumor cells, such as rat ascites hepatoma cells and Ehrlich ascites cells, as from rat liver (51,53).Queuine given to tumor-bearing mice by intraperitoneal injection causes the tumor tRNA to become fully modified with respect to queuine (92). However, it was necessary to keep a high concentration of queuine in the mice in order to maintain Q-tRNA in transplanted tumor cells (N. Shindo-Kkada, M. Iigo, A. Hoshi, and S. Nishimura, unpublished results). A large portion of the queuine was stored in the red blood cells of normal mice when queuine was administered, while the content of queuine in the red blood cells from tumor-bearing mice was drastically decreased (N. Shindo-Okada, M. Iigo, A. Hoshi, and S. Nishimura, unpublished results). It is possible that the supply of queuine in tumor cells is limited, so that the insertion of queuine into tRNA does not proceed fully. Another possibility is that a factor present in tumor cells inhibits the tRNA-guanine transglycosylase reaction, so that the limited amount of queuine in the mice is not enough to convert G-tRNA to Q-tRNA in tumor cells. [Another possibility is presented in Section I1 of “Queuine: An Addendum” by Singhal in this volume. Ed.] A mutant of a fish, Xiphosphorus, that lacks the differentiation gene (Diffl develops malignant melanoma. In the malignant cells, GtRNAAsPand G-tRNAAsnare predominant, whereas benign melanoma develops in a control fish carrying the Dilffgene and containing mostly Q-tRNAASP and Q-tRNATyr(A. Anders, G. Dess, S. Nishimura, and H. Kersten, unpublished results). The results suggest that the lack of queuosine is correlated with the absence of the Diff gene. VII. Change of the Ratio of Queuine to Guanine in tRNA during Cell Differentiation

In 1973, before the structures of queuine and queuosine were known, White et al. (21)made the first observation that queuosine and its derivative (later shown to be mannose- or galactose-containing queuosine) are present in Drosophila tRNA, and that they are located in the same tRNAs, i.e., those for tyrosine, histidine, asparagine, and aspartic acid, as in E . coli tRNA. Their interesting finding is that queuosine biosynthesis in a family of Q-containing tRNA is not com-

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plete in Drosophila. The ratio of Q-tRNA to G-tRNA varies during the life cycle of this organism. The tRNAs from the last larval stage contain G-tRNA almost exclusively, whereas the tRNAs from adults contain predominantly queuosine. The amount of Q-tRNA progressively decreases in larvae during development. During the pupal stage, QtRNA begins to be restored and this process continues on into adulthood. Thus, it was suggested (21)that queuosine in tRNA plays some regulatory role( s) in cell differentiation. Dietary conditions can modify the ratio of G-tRNATYrto Q-tRNATyr in Drosophila melanogaster (93).[This was presumably caused b y the varying content of queuine in the diets.] On a diet of brewers' yeast, the ability to form brown eye pigment, which is deficient in the vermilion mutant, is partially restored. The amount of brown eye pigment is inversely proportional to the concentration of G-tRNATYrin the flies on this diet. These observations support the hypothesis that G-tRNATYr may be closely associated with and regulate tryptophan oxygenase, although the previous observation that the inhibition of tryptophan oxygenase activity by direct interaction of the enzyme with G-tRNATy' must be cautiously reconsidered. (94, 95). In Musca domesticu, only Q-tRNAs appear in both larvae and adults, while in Lucilia sericata the larvae contain a significant portion of G-tRNAs. In Tenebrio molitor, G-tRNATyrappears to be significant only in pupae (96). These results suggest that Q-tRNA does not play a common regulatory function in the development of these insects. A change of the ratio of Q-tRNA to G-tRNA was also noticed with aging in Drosophila (97, 98). As the adults age, the ratio of QtRNA to G-tRNA in tRNATyr,tRNAHis,and tRNAAspincreases, while the chromatographic profiles of other tRNAs, such as tRNAA'", tRNALe",tRNAMet,and tRNASer,do not change. During differentiation of the cellular slime mold Dictyostelium discoideum, quantitative differences appear in the profiles for only three tRNAs: tRNATyr, tRNAAs",and tRNAAsp(99).It is very likely that this is due to a change in the extent of biosynthesis of queuosine. According to the chromatographic profiles, it seems that vegetative cells contain more G-tRNAs, while developing cells contain more Q-tRNAs.

VIII. Biological Function of Queuosine in tRNA As described in Section VII, there is a strong correlation between the appearance of Q-lacking tRNA (G-tRNA) and the stage of the cells in differentiation. A number of approaches have been made to determine the biological function of queuosine in tRNA, but this matter is

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not well understood at present. The following summarizes the results so far obtained with respect to functions of queuosine. Since Q is located in the "wobble" position of tRNAs, it was at first thought that it influences codon-anticodon recognition. I n 1972, we showed that Q in the anticodon of E . coli tRNAs recognizes U in preference to C in the third position of codons (6).This was demonstrated by comparisons of the codon recognition properties of Q-containing tRNATyr,tRNAHis,tRNAAsn,and tRNAAspwith that of tRNAPhe, which contains guanosine in the first position of the anticodon, since G-containing counterparts of the tRNAs were not available at that time. However, it has been shown that even G-containing tRNATyr binds more efficiently to ribosomes with U-A-U than with U-A-C (50). Presumably, the preferential recognition of U in the third position of the codons by E . coli tRNATyr,tRNAHis,tRNAAsn,and tRNAAspis not due to the presence of queuosine, but to another structural feature(s) of those tRNAs. However, Singhal et al. found that mammalian QtRNAAsPprefers G-A-.Cover G-A-U codon more significantly than does G-tRNAAsp( 100).When the three-dimensional structure of queuosine 5'-monophosphate was incorporated into tRNATy' on the basis of the coordinates of yeast tRNAPhe, the bulky 7-substituent group of queuosine takes the planar trans conformation and is fully extended outward, so that it seems not to interfere with codon-anticodon interaction (20). However, Grosjean et al. reported that the presence of queuosine in the wobble position enhances the anticodon-anticodon interaction of tRNA (101). A direct approach to elucidate the function of queuosine would seem to be to isolate a mutant that lacks queuosine in its tRNAs. An E . coEi mutant that does not have the tRNA-guanine transglycosylase was recently isolated (50),and there is no queuosine in its tRNA. However, no clear biological defect was observed in the mutant. Surprisingly, it grew slightly faster than the control iogenic strain under favorable growth conditions, and its rate of protein synthesis, measured by pulse-labeling in uiuo, was 40% faster than the control strain. The only phenotypic defect observed in the mutant thus far is a marked reduction of viability when the cells are kept under conditions unsuitable for growth, indicating that the presence of queuosine in tRNA is important in E . coli for survival in an unfavorable environment. Studies on the function of Q in eukaryote tRNA are more difficult to pursue, because of lack of suitable experimental systems. The Qlacking tRNAHisisolated from rabbit reticulocytes has the same ability to read the codons CAU and CAC as does its Q-containing counterpart in globin biosynthesis in a rabbit reticulocyte cell-free system (102,

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103). Also the Q-lacking tRNATyr isolated from Drosophila can read the amber “stop” codon UAG, resulting in the formation of a 160,000 M,protein directed by TMV RNA in a Xenopus oocyte system, while Q-containing tRNATyrdoes not act as a suppressor (104). Thus, in this particular system, Q in tRNA prevents miscoding of the G in UAG. However, it should be mentioned that the E . coli mutant that lacks tRNA-guanine transglycosylase has no such amber suppressor activity (50). There is no essential difference in the rate or the extent of protein synthesis between Q-containing and Q-lacking tRNA isolated from Drosophila when protein synthesis is carried out in a cell-free, tRNAdependent, mRNA-dependent system (98).On the other hand, addition of queuine to the tissue culture medium of SV40 transformed cells results in marked increase of synthesis of a 75,000 M , protein (Y.Kuchino and S. Nishimura, unpublished results). An interesting observation is that addition of queuine affects the cellular differentiation of mammalian fused cells (isolated nuclei of mouse melanoma cells and cytoplasm of rat myoblastic cells) in a manner that reverses the action of tumor promoter (T. Sekiguchi and S. Nishimura, unpublished results). Induction of ornithine decarboxylase in mouse skin by application of the tumor promoter 12-O-tetradecanoylphorbol 13-acetate is inhibited by intraperitoneal injection of queuine into mice before the phorbol treatment (H. Fujiki and S. Nishimura, unpublished results). It is of interest to examine whether continuous administration of queuine might inhibit the induction of tumors in such experimental animals. These observations indicate that queuine may direct cells into a differentiated stage. However, differentiation of murine erythroleukemic cells into mature erythroid cells is not induced by the addition of queuine to the tissue culture medium (M. Terada and S. Nishimura, unpublished results). Injection of queuine isolated from bovine amniotic fluid into mice bearing Ehrlich ascites tumors can restore queuine to tumor tRNA and consequently may inhibit tumor growth (92). Large amounts of synthetic optically active queuine injected intraperitoneally into mice after transplantation of L-1210 or S-180 tumor cells gave no inhibition of tumor growth, although the tumor tRNAs were completely shifted to Q-containing species by this treatment (N. Shindo-Okada, M . Iigo, A. Hoshi, and S. Nishimura, unpublished results). In view of these conflicting results, it is not possible to draw the conclusion that queuine administration to tumor-bearing animals inhibits tumor growth. No appreciable difference in aminoacylation between G-tRNA and Q-tRNA is observed in the case of E . coli tRNATyr (50) and rabbit

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reticulocyte tRNAHiS(102). However in the case of Dictyostelium discoideum, G-tRNAAsn,which specifically appeared during developmental transition of vegetative amebas by nutrient starvation, was completely deacylated in duo, while Q-tRNAAsnwas aminoacylated (105). Singhal and Kopper ( 100) found the V,,, of aminoacylation of pure rat liver Q-tRNAAspisoacceptors 1 and 2 to be 16-fold higher than that of the G-tRNAAsp(unfractionated but free of Q-tRNAAsp). However, the K,s were the same. In unfractionated preparations from L-M cells, Q-tRNAAsphad a 30%higher V,,, and 55% lower K , than the G-tRNAAspfor the rat liver aminoacyl tRNA synthetase. [See Section 111 of the following Addendum by Singhal. Ed.]. These results indicate that queuosine in tRNA certainly influences aminoacylation of tRNA in some cases. The bulky side chain of queuosine with its unique structure, charge, and reactive group constitute a suitable site for specific interaction with protein or other cell components. Attachment of mannose or galactose to the queuosine molecule should further enhance its specific interaction. Thus Q, manQ, or galQ in tRNA may be an appropriate interacting site, if tRNA functions in regulation with mechanisms other than protein synthesis. However, it should be noted that there is no experimental evidence so far to show that such interactions of Q-containing tRNA with cellular components do occur.

IX. Possible Approaches to the Use of Queuine Analogs for Cancer Diagnosis and Cancer Chemotherapy

As described previously, artificial guanine analogs, such as 8-azaguanine and 6-thioguanine, or queuine precursors, are also incorporated into G-tRNA. For practical purposes, radioactive queuine analogs may be useful as a tool for cancer diagnosis, because they should be specifically incorporated into tRNA in tumor tissues. If some queuine analogs can affect protein synthesis by changing the codon recognition properties of tRNA, or by cross-linking with protein-synthesizing machinery, such as ribosomes, after they are incorporated into tRNA, they may be also useful as anti-cancer reagents. Many queuine analogs have been synthesized chemically for this purpose, and their abilities to be utilized as donors for tRNA-guanine transglycosylase have been tested (N. Shindo-Okada, H. Kasai, H. Akimoto, H. Nomura, and S. Nishimura, unpublished results). Many were active donors. It should be noted that queuine analogs that contain 4-iodobenzyl or 4-hydroxy-3,5-diiodophenyliminomethyl groups in place of

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RNA

the cyclopentenediol were equally well incorporated into G-tRNA. These results suggested that appropriate queuine analogs containing radioactive iodine might be used for the detection of tumor-specific tRNAs or for cancer diagnosis. Such iodine-containing queuine analogs inhibit the growth of tissue culture cells (L5178 Y) at 10 pg/ml, while normal queuine has no effect at 100 pg/ml (N. Shindo-Okada, M. Yoshida, A. Hoshi, and S. Nishimura, unpublished results).

REFERENCES 1 . U. L. RajBhandary, S. H. Chang, H. J. Gross, F. Harada, F. Kimura, and S. Nishimura, FP 28,409 (1969). 2. H. M. Goodman, J. Abelson, A. Landy, S. Breener, and J . D. Smith, Nature (London) 217, 1019 (1968). 3. H. M. Goodman, J. N. Abelson, A. Landy, S. Zadrazil, and J. D. Smith, EJB 13,461 ( 1970). 4. B. P. Doctor, J. E. Loebel, M. A. Sodd, and D. B. Winter, Science 163,693 (1969). 5. H. Rogg and M. Staehelin, BBA 195, 16 (1969). 6. F. Harada and S. Nishimura, Bchem 11,301 (1972). 7. F. Harada, S. Sato, and S. Nishimura, FEBS Lett. 19, 352 (1972). 8. K. Ohashi, F. Harada, Z. Ohashi, S. Nishimura, T. S. Stewart, G. Vogeli, T. McCutchan, and D. SOH, NARes 3,3369 (1976). 9. F. Harada, K. Yamaizumi, and S. Nishimura, BBRC 49, 1605 (1972). 10. M. Sprinzl and D. H. Gauss, NARes 10, r l (1982). 11. R. P. Singhal, E. F. Roberts, and V. N. Vakharia, this volume. 12. H. Kasai, Z. Ohashi, F. Harada, S. Nishimura, N. J . Oppenheimer, P. F. Crain, J. G. Liehr, D. L. von Minden, and J. A. McCloskey, Bchem 14,4198 (1975). 13. J. A. McCloskey and S. Nishimura, Acc. Chem. Res. 10,403 (1977). 14. R. J . Suhadolnik, This Series 22, 193 (1979). 15. N. Tanaka, R. T. Wu, T. Okabe, H. Yamashita, A. Shimizu, and T. Nishimura,J. Antibiot. 35,272 (1982). 16. R. T. Wu, T. Okabe, M. Namikoshi, S . Okuda, T. Nishimura, and N. Tanaka, J . Antibiot. 35, 279 (1982). 17. T. Ohgi, T. Goto, H. Kasai, and S. Nishimura, Tetrahedron Lett. 5, 367 (1976). 18. T. Ohgi, T. Kondo, and T. Goto, Tetrahedron Lett. 46,4051 (1977). 19. T. Ohgi, T. Kondo, and T. Goto,JACS 101,3629 (1979). 20. S. Yokoyama, T. Miyazawa, Y. Iitaka, Z. Yamaizumi, H. Kasai, and S. Nishimura, Nature (London) 282, 107 (1979). 21. B. N. White, G . M. Tener, J. Holden, and D. T. Suzuki,/MB 74,635 (1973). 22. H. Kasai, Y. Kuchino, K. Nihei, and S. Nishimura, NARes 2, 1931 (1975). 22a. T. Walden, J. P. Reyniers, V. Hiatt, and W. R. Farkas, PSEBM 170, 328 (1982). 23. S. Nishimura, This Series 12, 49 (1972). 24. H. Kasai, K. Nakanishi, R. D. Macfarlane, D. F. Torgerson, Z. Ohashi, J. A. McCloskey, H. J. Gross, and S. Nishimura,JACS 98, 5044 (1976). 25. N. Okada, N. Shindo-Okada, and S. Nishimura, NARes 4,415 (1977). 26. M. A. Wosnick and B. N. White, BBRC 81, 1131 (1978). 27. V. N. Vakharia and R. P. Singha],]. Appl. Biochem. 1, 210 (1980). 28. C. M. Garcia and R. P. Singhal, BBRC 86, 697 (1979). 29. T. F. McCutchan, P. T. Gilham, and D. So11, NARes 2,853 (1975).

72

SUSUMU NISHIMURA

30. M. Boisnard, G . Petrissant, and R.-M. Landin, Biochimie 62,61 (1980). 31. C . Vogeli, T. S. Stewart, T. McCutchan, and D. Sol1,JBC 252,2311 (1977). 31a. R. P. Singhal, R. K. Bajaj, C. M. Buess, D. B. Smoll, and V. H. Vakharia, Anal. Biochem. 109, l(1980). 32. Y. Kuchino, H. Kasai, K. Nihei, and S. Nishimura, NARes 3, 393 (1976). 33. T. Uematsu and R. J. Suhadalnik, Bchem 9, 1260 (1970). 34. R. J. Suhadolnik and T. Uematsu, JBC 245,4365 (1970). 35. A. W. Burg and G. M. Brown,JBC 243,2349 (1968). 36. J. J. Yim and G . M. Brown,JBC 251,5087 (1976). 37. J. J. Yim, E. H. Grell, and K. B. Jacobson, Science 198, 1168 (1977). 38. P. A. Marks, E. R. Burka, and D. Schlessinger, PNAS 48,2163 (1962). 39. W. D. Hankins and W. R. Farkas, BBA 213, 77 (1970). 40. W. R. Farkas, W. D. Hankins, and R. D. Singh, BBA 294, 94 (1973). 41. W. R. Farkas and R. D. Singh, JBC 248, 7780 (1973). 42. N. Okada, F. Harada, and S. Nishimura, NARes 3,2593 (1976). 43. W. R. Farkas and D. Chernoff, NARes 3,2521 (1976). 44. E. F. DuBrul and W. R. Farkas, BBA 442,379 (1976). 45. N. Okada and S. Nishimura, JBC 254,3061 (1979). 46. N. Okada, S . Noguchi, S. Nishimura, T. Ohgi, T. Goto, P. F. Crain, and J. A. McCloskey, NARes 5, 2289 (1978). 47. S. Noguchi, Z. Yamaizumi, T. Ohgi, T. Goto, Y. Nishimura, Y. Hirota, and S. Nishimura, NARes 5, 4218 (1975). 48. T. Ohgi, T. Kondo, and T. Goto, Chem. Lett. 10, 1283 (1979). 49. N. Okada, S. Noguchi, H. Kasai, N . Shindo-Okada, T. Ohgi, T. Goto, and S. Nishimura, JBC 254, 3067 (1979). 50. S. Noguchi, Y. Nishimura, Y. Hirota, and S. Nishimura,JBC 257,6544 (1982). 51. N. Shindo-Okada, N. Okada, T. Ohgi, T. Goto, and S. Nishimura, Bchem 19,395 (1980). 52. N. K. Howes and W. R. Farkas,JBC 253,9082 (1978). 53. Y. H. Itoh, T. Itoh, I. Haruna, and I. Watanabe, Nature (London) 267,467 (1977). 54. W. R. Farkas and K. B. Jacobson, Insect Biochem. 10, 183 (1980). 55. J. R. Katze, BBRC 84, 527 (1978). 56. J. Katze and W. R. Farkas, PNAS 76, 3271 (1979). 57. P. F. Crain, S. K. Sethi, J. R. Katze, and J. A. McCloskey,JBC 255, 8405 (1980). 58. W. R. Farkas, JBC 255,6832 (1980). 59. J. P. Reyniers, J. R. Pleasants, B. S. Woshnann, J. R. Katze, and W. R. Farkas, JBC 256, 11591 (1981). 60. J. R. Katze, B. Basile, and J. A. McCloskey, Science 216, 55 (1982). 61. K. Nishikura and E. M. De Robertis,JMB 145,405 (1981). 62. K. B. Jacobson, W. R. Farkas, and J. R. Katze, NARes 9,2351 (1981). 63. W. S. McNutt, Methods Enzymol. 2,464 (1955). 64. W. A. Deutsch and S. Linn, PNAS 76, 144 (1979). 65. W. A. Deutsch and S. Linn, JBC 254, 12099 (1979). 66. Z. Livneh, D. Elad, and J. Sperling, €"AS 76, 1089 (1979). 67. N. Okada, T. Yasuda, and S. Nishimura, NARes 4,4063 (1977). 68. J. R. Katze, M. H. Simonian, and R. D. Mosteller, J . Bact. 132, 174 (1977). 69. N. Okada and S. Nishimura, NARes 4,2931 (1977). 70. N. Sueoka and T. Kano-Sueoka, This Series 10,23 (1970). 71. U. Z. Littauer and H. Inoue, ARB 42,439 (1973). 72. E. Borek and S. J. Kerr, Adu. Cancer Res. 15, 163 (1972).

QUEUOSINE IN TRANSFER

RNA

73

73. S. Nishimura, “Transfer RNA: Structure, Properties, and Recognition” (P. R. Schimmel, D. SOU, and J. N. Abelson, eds.), p. 59. Cold Spring Harbor Laboratory, New York, Cold Spring Harbor, 1979. 74. S. Nishimura and Y. Kuchino, GANN Monogr. Cancer Res. 24, 245 (1979). 75. E. Tsutsui, P. R. Srinivasan, and E. Borek, PNAS 56, 650 (1966). 76. S. H. Blobstein, D. Grunberger, I. B. Weinstein, and K. Nakanishi, Bchem 12, 188 ( 1973). 77. H. Kasai, Z. Yamaizumi, Y. Kuchino, and S. Nishimura, NARes 6, 993 (1979). 78. Z. Yamaizumi, S. Nishimura, K. Limburg, M. Raba, H. J. Gross, P. F. Crain, and J. A. McCloskey, JACS 101,2224 (1979). 79. M . Saneyoshi and S . Nishiniura, BBA 204,389 (1970). 80. M. Saneyoshi and S. Nishimura, BBA 246, 123 (1971). 81. B. N. White, BBA 353, 283 (1974). 82. B. A. Roe, A. F. Stankiewicz, and C . Y. Chen, NARes 4, 2191 (1977). 83. J. R. Katze, BBA 383, 131 (1975). 84. J. R. Katze, NARes 5, 2513 (1978). 84a. V. N. Vakharia and R. P. Singhal, BBRC 105, 1072 (1982). 85. Y. Kuchino, N. Shindo-Okacia, N. Ando, S. Watanabe, and S. Nishimura,JBC 256, 9059 (1981). 86. B. A. Roe, A. F. Stankiewicz, H. L. Rizi, C. Weisz, M. N. DiLauro, D. Pike, C . Y. Chen, and E. Y. Chen, NARes 6,673 (1979). 87. N. Okada, N. Shindo-Okada, S . Sato, Y. H. Itoh, K. Oka, and S. Nishimura, PNAS 75, 4247 (1978). 88. R.-M. Landin, M. Boisnard, and G. Petrissant, NARes 7, 1635 (1979). 89. R. P. Singhal, R. A. Kopper, S . Nishimura, and N. Shindo-Okada, BBRC 99, 120 (1981). 90. N. Shindo-Okada, M. Terada, and S. Nishimura, EJB 115,423 (1981). 91. V. K. Lin, W. R. Farkas, and P. F. Agris, NARes 8,3481 (1980). 92. J. R. Katze and W. T. Beck, BBRC 96,313 (1980). 93. K. B. Jacobson, NARes 5,2391 (1978). 94. K. B. Jacobson, Nature NB 231, 17 (1971). 95. D. Mischke, P. Kloetzel, and M. Schwochau, Nature (London)255, 79 (1975). 96. B. N. White and N. J. Lassani, Insect. Biochern. 9, 375 (1979). 97. H. A. Hosbach and E. Kubli, Mech. Ageing Deo. 10, 141 (1979). 98. R. K. Owenhy, M. P. Stulberg, and K. B. Jacobson, Mech. Ageing Deo. 11, 91 (1979). 99. C. M. Palatnik and E. R. Katz,JBC 252, 694 (1977). 100. R. P. Singhal and R. A. Kopper, F P 40, 1646 (1981). 101. H. J. Grosjean, S. d e Henau, and D. M. Crothers, PNAS 75, 610 (1978). 102. A. L. McNamara and D. W. E. Smith, JBC 253, 5964 (1978). 103. D. W. E. Smith and A. L. McNamara, BBRC 104, 1459 (1982). 104. M. Bienz and E. Kubli, Nature (London)294, 188 (1981). 105. T. Dingermann, A. Ogilvie, F. Pistel, W. Miihlhofer, and H . Kersten, Z p Chem 362,763 (1981).

This Page Intentionally Left Blank

Queuine: An Addendum RAM P. SINGHAL* Laboratory of Cellular and Molecular Biology, Gerontology Research Center, National Institute on Aging, Bethesda, Maryland

I. tRNA-Guanine Transglycosylase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

..........

75

75

C. Substrate Specificity . . . . . . . . . . .

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

80

1. tRNA-Guanine Transglycosylase

The enzyme tRNA-guanine transglycosylase places “free” quanine (or its derivative, queuine or precursors of queuosine’) in position 34 of the tRNA (the “wobble” position in the anticodon) ( 1 , 2). Although breaking and making of a covalent bond and removal of the existing base are involved, no nucleotide triphosphate is required for the reaction (Fig. l),which is limited to the anticodon of four “Qspecific tRNAs”: tRNAAsn,tRNAAsp,tRNAHis,and tRNATyr. The enzyme involved in the posttranscriptional modification of these tRNAs has been purified to homogeneity from several sources. A. Size and Subunits The enzyme from Escherichia coli, rat liver, rat ascites hepatoma cells, rabbit erythrocytes, and wheat germ has been isolated and studied (3-7). Its characteristics vary greatly from one source to another except for the rat liver and ascites hepatoma enzymes, which show a close similarity in specificity (5).The E . coli and rat liver enzymes consist of single polypeptide chains of 46,000 and 80,000 daltons, respectively, whereas the rabbit erythrocyte and wheat germ enzymes consist of two subunits each (Table I). While the rabbit enzyme sub-

* Permanent address: Chemistry Department, Wichita State University, Wichita, Kansas 67208. Precursors of queuosine tested for the E . coli enzyme are 7-cyano-7-deazaguanosine (preQo)and 7-aminomethyl-7-deazaguanosine (preQ I ) . 75 Progress in Nucleic Acid Research and Molecular Biology. VoI. 28

Copyright Q 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-540028-4

76

RAM P. SINGHAL

G’

+

G’

G‘

ampor tANA,

modified tRNA iprcducrl

free sutsnate

replaced residue 8midOfl

FIG. 1. General reaction of tRNA-guanine transglycosylase. Replacement of the wobble base, G2 located in position 37 of the Q-specific tRNAs by the free residue G’ depends on the origin of the transglycosylase and the nature of both residues. In general, G’ and GZare guanine, queuine, or a precursor of queuine such as 7-aminomethyl7-deazaguanine. The acceptor tRNAs are specific to tRNAAs“,tRNAAhp,tRNAHBS, and tRNATyr.Use of radioactive “free” substrate produces a labeled tRNA (the product).

units have different sizes ( M , = 43,000 and 60,000), the wheat germ enzyme appears to have identical subunits (each M , = 68,000)( 7). The rat liver and the rabbit erythrocyte enzymes, although both are mammalian in origin, differ greatly in size and shape.

B. Metal Ion Requirement (Table I) The enzyme preparations from E . coli and wheat germ require magnesium ions for activity, but those from rat liver and rabbit erythrocytes do not. However, the erythrocyte enzyme does require monovalent cations (Na+, K+, or Li+) for full activity.

C. Substrate Specificity (Table I)

The progress of the base replacement reaction depends mainly on (a) the source of the enzyme; ( b )the nature of the reacting “free” base (guanine, G ; prequeuine, preQ; or queuine, Q); and (c) the origin of TABLE I PROPERTIES OF tRNA-GUANINE TRANSGLYCOSYLASE Substrates (K,) Enzyme source

M,

Subunits (M, x

Escherichia

46 None

colib Rat liver

80 None

Rabbit erythrocytes Wheat germ

104 2 (43,60) 140 2 (68,68)

Need Mg2+ Yes

Guanine

Yes (0.053 p M ) No Yes (0.8 p M ) NoC Yes + (0.15p M ) Yes Yes ++ (0.06 pM)

Queuine No Yes (0.3 PM) Yes ++ ( K , = 0.0045 fiM) Yes + (0.095 p M )

Prequeuine‘ Yes

(0.014p M Yes (2 pM) No No

7-Aminomethyl-7-deazaguanine. Among E . coli tRNATYracceptors,G- and preQ-tRNAsTYrareequally good, but preQi- and ( tRNAsTfyrare inactive acceptors. Needs a monovalent cation (Na+, K+ , or Lit). a

77

QUEUINE

TABLE I1 SPECIFICITY OF tRNA-GUANINE TRANSGLYCOSYLASE Minimum guanine nucleus required: the C-2 amino group, the C-6 oxygen, and two electrons on N-7. Substrates: G, Q and their analogs. 7-Methyl-, 7-methyldeaza-, and 7-methyl-7deazaguanines do not function as substrates for the liver enzyme.) The anticancer drugs 8-azaguanine and 6-thioguanine are efficiently incorporated into G-tRNA by the liver enzyme. Inhibitors (of the incorporation of free guanine into G-tRNA): 7-methylguanine M). Neither is (bacterial enzyme); pteridine (competitive inhibitor with KI = 9 x incorporated into C-tRNA.

the accepting Q-specific tRNA (see Table 11).An exception to this is yeast, which fails completely to incorporate exogenous queuine supplied with the growth medium (8). The transglycoslases from different sources react very differently with a given substrate. For example, E . coli, rat, and rabbit enzymes exhibit very different K , values for guanine: 0.053,0.8, and 0.15 pM, respectively (see Table I). Moreover, the enzyme from each source exhibits different specificities for (the bases o f ) G, preQ, and Q. For example, the E . coli enzyme fails to react with Q, but reacts four times faster with preQ than with G. In fact, it has the highest activity (lowest K,,,) for preQ of any enzyme preparation for any of the free bases studied. The liver enzyme reacts with each one of the three substrates, but with greatly altered specificity. Both mammalian enzymes have very high specificities for Q; in addition, each removes Q and places G into E . coli Q-tRNAs, but fails to do so with its homologous tRNA. The liver enzyme inserts G into E . coli tRNATy’with decreasing efficiency in the order G-tRNA > preQo-tRNA > preQ1-tRNA %- Q-tRNA. The mammalian enzymes have a higher affinity for Q than for G; however, the wheat germ enzyme reacts more with G than Q, using homologous G-tRNAs. A significantly high affinity of the bacterial enzyme with preQ and G (none with Q) explains how the bacterium is able to start the posttranscriptional modification of G to Q at a very early stage, i.e., 7aminomethyl-7-deazaguanine (preQ1). Once this preQ is incorporated into G-tRNAs, further modification of preQ to Q appears to occur, while it is part of the tRNA. Both mammalian enzymes react with Q more rapidly than with G (the liver enzyme reacts with preQ, though poorly, while the rabbit enzyme remains inactive). I n addition, Q (“Q factor”), which is present in animal serum and amniotic fluid, causes a G to Q exchange

78

RAM P. SINCHAL

in tRNAs of L-M cells in tissue culture (9-11). In summary, mammals differ from bacteria (perhaps also from wheat germ) in achieving a G to Q posttranscriptional tRNA modification by replacing G with a presynthesized Q (“Q factor”) in their Q-specific tRNAs. At present, it is unclear whether mammals are capable of synthesizing “free” Q as bacteria and plants do, or must depend on an exogenous supply.

11. Q in Neoplastic Cells Nishimura surmises that the presence of G-tRNA in tumor cells is not the result of their fast growth, but is rather a specific property of tumor and undifferentiated cells, perhaps related to some specific metabolism of the tumor cells. A significantly higher concentration of Q must be maintained in order for ascites cells to incorporate Q into their G-tRNAs. Besides the possibilities that insufficient amounts of Q may be available to the tumor cells, or that a potent inhibitor may be present in the host, it may be that the lack of a G to Q exchange in tumor tRNAs arises from a drastically different (lower) affinity of the tumor enzyme for Q than for G. To answer this, the tumor enzyme must be isolated free from the host enzyme and studied carefully for its substrate affinity. Cancerous cells growing in the host and having a common vascular system have access to the “free Q” and also perhaps to the enzyme. Therefore, tumors grown in mice can possibly produce a full complement of Q in their tRNAs. To test this hypothesis, human (adenocarcinoma, ovary and shoulder) tumors were transplanted and grown in athymic “nude” mice. Tumors from such animals were carefully removed, and their tRNAs were isolated. The chromatography of [3H]Asp-tRNA showed no unadsorbed radioactivity (G-tRNA) on concanavalin A-Sepharose columns. The tumor tRNAAsP isoacceptors were indistinguishable from those of the normal rabbit liver tRNAAspby chromatrography on RPC-5 columns. Prior treatment with CNBr also indicated the absence of G-tRNAsAsPin the tumors grown in this manner (R. P. Singhal and V. N. Vakharia, unpublished results). These results suggest that tumors growing in the host can indeed have Q in their Q-specific tRNAs. The tumors may lack the enzyme and require “free” Q to derepress their genome in order to synthesize the transglycosylase. Therefore, the availability of Q and its ability to cross the tumor cell membrane may determine the synthesis of the tumor enzyme and thus the G to Q exchange in tumor tRNAs.

79

QUEUINE

TABLE 111

KINETICPROPERTIES: ASPARTATE Q- AND G-tRNAs AS SUBSTRATES FOR THE AMINOACYLATION REACTION.UNFRACTIONATED tRNAs from L-M Cells(8 Substrate Q-tRNAsA5P G-tRNAA'p

V,, (pmol/min) O.8Oh 0.78' 0.6Zb 0.59'

~~~

K," (nM)

V,,JK, (pmol min-'/pM)

68* 65' 125* 115"

11.8b 12' 5.0h 5.1'

~

Q-tRNAs derived from L-M cells grown in a serum-free medium to which queuine (one Axe,, unit per 100 ml of the medium) was added; G-tRNAs were derived from the L-M cells grown in the absence of queuine. Values derived from Lineweaver-Burk plot. Values derived from Eadie-Hofstee plot.

Thus, the occurrence of G-tRNAs in cancer cells grown in rats or in

cell culture (ascites hepatoma cells, leukemic cells) may arise either from the presence of a transglycosylase with altered kinetic properties or from the presence of a competitive inhibitor in such cells, or from the unavailability of "free" Q in sufficient amounts to derepress the genome to allow synthesis of the tumor enzyme.

111. Function of Q in Q-tRNAs The aminoacylation kinetics studied for G- and Q-tRNAs have produced interesting results. Singhal and Kopper (12)reported the V,,, of aminoacylation of a purified Q-tRNA?,;' sample to be 16 times that of the liver G-tRNAAsp,which had other tRNAs besides Q-tRNAAsp.The K , values for the two tRNA samples were the same. However, unfractionated tRNAs containing G, Q (without mannose), and possibly preQ were compared with pure Q-tRNAsAspthat had manQ; hence, the results can be misleading. Recent work with aspartate G- and Q-tRNAs from L-M cells grown in culture gives a clearer picture (R. P. Singhal and V. N. Vakharia, unpublished results). The results (Table 111) indicate that the QtRNAAspof L-M cells exhibits a higher reaction velocity (V, is 33% greater) and a higher reaction rate ( K , is 55% less) than its counterpart. The heterologous enzyme used here is common to both tRNAs. Q-tRNAAsPappears to be a better substrate than the G-tRNA as determined by the V,IK, ratio. The data strongly suggest an involvement of Q in both the rate and the extent of aminoacyl-tRNA synthesis.

80

RAM P. SINGHAL

REFERENCES 1. N. Okada, S . Noguchi, S. Nishimura, T. Ohgi, T. Goto, P. F. Crain, and J. A. McCloskey, NARes 5,2289 (1978). 2. N. Okada, S . Noguchi, H. Kasai, N. Shindo-Okada, T. Ohgi, T. Goto, and S. Nishimura, JBC 254,3067 (1979). 3. N. Okada and S. Nishimura, JBC 254,3061 (1979). 4. N. Shindo-Okada, N. Okada, T. Ohgi, T. Goto, and S. Nishimura, Bchem 19, 395 (1980). 5. N. Okada, N. Shindo-Okada, S. Sato, Y.H. Itoh, K. I. Oda, and S. Nishimura, FNAS 75,4247 (1978). 6. N. K. Howes and W. R. Farkas,JBC 253,9082 (1978). 7. T. M. Walden, Jr., N. Howes, and W. R. Farkas,JBC 257, 13218 (1982). 8. T. Walden, J. P. Reyniers, V. Hiatt, and W. R. Farkas, FSEBM 170, 328 (1982). 9. J. R. Katze, BBRC 84,527 (1978). 10. J. Katze and W. R. Farkas, PNAS 76,3271 (1979). 1 1 . P. F. Crain, S. K. Sethi, J. R. Katze, and J. A. McCloskey,JBC 255,8405 (1980). 12. R. P. Singhal and R. A. Kopper, FP 40, 1646 (1981).

The Fidelity of Translation

I C. Antibiotics.

ABRAHAMK. ABRAHAM Department of Biochemistry, University of Bergen, Bergen, Norway

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

A. Role of Ribosomes . . . . . . . . . . . . . . B. Ribosomal Mutants. . . . . . . . . . . . . . . . . , ................. C. Role ofppGpp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

B. Temperature . . C. Effect of tRNA. D. Effect o f p H . . . . . . . . . . . . . . . . . . . . . . , . . . . .

............................ A. General

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

D. Temperature Change. . . . . E. Error-Producing Antibiotics. .

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

82 82 83 83 84 84 85 85 86 86 87 88 89 90 90 91 91 91 92 92 92 93 93 94 94 94 96 97

The importance of fidelity in the aminoacylation of tRNAs has long been appreciated (I), though the mechanism by which specificity is maintained during the activation of amino acids was not understood. Ambiguity occurring at the ribosome level during codon recognition by the anticodon in tRNA was first reported in 1962 ( 2 ) . Two other types of translational errors have been reported. One type of error involves a “readthrough” translation of certain termination codons in both prokaryotes and eukaryotes by a normal tRNA (3, 81 Progress in Nucleic Acid Research and Molecular Biology, Vol. 28

Copyright Q 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-540028-4

82

ABRAHAM K. ABRAHAM

4). The other type, demonstrated more recently, is an initiation error taking place in certain in vitro systems deficient in ribosome-releasing factor (5).Such initiation occurs at a codon next to an amber codon (5). In spite of the possiblity that errors in translation can occur at different levels, the frequency of such errors in vivo appears to be rather low (6-8). The frequency of error calculated for valine substitution in the isoleucine positions of rabbit globin is in the range of 2-6 residues per 10,000(7).Since isoleucine and valine are closely related chemically, and both react with isoleucyl-tRNA synthetase (7), this estimate most probably reflects errors at the level of tRNA amino acylation. Errors at the codon/anticodon recognition level most probably are of the order of 1 per 10,000 codons (8).In contrast, most in uitro systems produce errors that are several orders of magnitude higher (9). Codon misreading by isoacceptor tRNAs also occurs in uitro (lo),and most probably also in vivo. Such mistakes that do not result in misincorporation, due to the degeneracy of the genetic code, are discussed at the end of this review.

1. Agents Affecting Translation Fidelity in Vivo

The high accuracy of in vivo translation can be reduced significantly under adverse growth conditions produced by lack of nutrients or by external agents. A. Amino-Acid Starvation Experiments using poly(U) have clearly demonstrated that a high frequency of error occurs if phenylalanine is omitted from the in vitro system, i.e., when only the noncognate aminoacyl-tRNAs are available ( 1 1 ) . It follows that deprivation of any essential amino acid may be expected to increase the error frequency at codons specifying that amino acid, provided no other regulatory mechanism is operating. Nearly a11 proteins made by “relaxed” mutants of Escherichia coli starved for essential amino acids show a striking pattern of electrophoretic heterogeneity (12). Extreme starvation for essential amino acids causes translation errors in eukaryotes also (13).Such errors result in many faulty proteins, which can be detected by two-dimensional polyacrylamide-gel electrophoresis of total proteins. The extent of mistranslation in the case of two specific proteins (EF-G in E . coli, and actin in Chinese hamster ovary cells) during severe histidine starvation, was calculated to be as high as 0.23 per histidine residue (13).Judged from the charge difference in the two-

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dimensional gels of the faulty proteins, these errors seem to occur at the decoding level on the ribosome, and the type of codon misreading in these systems appears to b e the phenylalanine-leucine kind of error (2, 1 1 ) occurring in the poly(U) system, namely, misreading of the third nucleotide of the codon (13). I n bacteria, the accuracy of translation during amino-acid starvation is coupled to the well known phenomenon called “stringent” response. this is discussed further in Section II,B. The stringent response involves a shutdown of the synthesis of rRNA and mRNA for ribosomal proteins in cells deprived of amino acids. Such a stringent response during amino-acid starvation has been reported to take place in yeast also (14).No relaxed mutants have been found in eukaryotes (15).

B. Polyarnines Polyamines increase the fidelity of in vitro translational systems (9,11,16),and there is a correlation between the rate of chain elongation and polyamine content of ribosomes ( 17).Bacterial strains unable to synthesize putrescine when submitted to polyamine starvation, show decreased amino-acid incorporation and growth rate (18).It is most likely that these amines also contribute to the fidelity of translation in vivo. Polyamine-requiring bacterial mutants grown in the absence of these amines produce extensive misreading (18),although direct misreading was not demonstrated.

C. Antibiotics A number of antibiotics, including streptomycin, gentamycin, paramomycin, kanamycin, and neomycin, reduce both the rate and accuracy of translation (19, 20). These antibiotics, at concentrations that inhibit protein synthesis, do not inhibit aminoacylation of tRNAs in vitro (21).Therefore, it is reasonable to assume that they influence translation fidelity only at the ribosome level. Streptomycin-induced errors involve misreading of the first or second position of the codon

(21,22). Several E . coli strains resistant to streptomycin are mutated in the structural gene coding for ribosomal protein S12 (23, 24). A positive effect of the small-subunit proteins S12, S4, and S5 on fidelity is also evident from ribosome reconstitution experiments (25).Since streptomycin binds to the small ribosomal subunit in a 1 : 1 ratio (26,27),it is conceivable that alteration of protein S12 could reduce the affinity of the ribosomes for this antibiotic. On the other hand, mutations affecting ribosomal protein L6 appear to confer increased resistance to all error-producing aminoglycoside antibiotics (28). Several conforma-

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tional alterations of ribosomes can change parameters involved in codon/anticodon recognition (29),so it is not surprising that mutations in several ribosomal proteins of both subunits confer resistance to errorproducing antibiotics (28,30, 31). Since polyamines and aminoglycoside antibiotics affect translational fidelity in opposite directions, polyamine-starved bacteria would be expected to produce more errors in the presence of streptomycin. However, extracts prepared from bacteria starved for polyamines seem to have a higher fidelity than unstarved cells (32).This may be related to the speed of translation, as discussed in Section IV.

D. Bacteriocins The bacteriocins colicin E3 and cloacin DF13 induce a single endonucleolytic cut in the RNA of the small ribosomal subunit of E . coli, and thereby inhibit protein synthesis (33).The fidelity of colicin E-cleaved ribosomes is not altered (34), and the fidelity of cloacin DF13-cleaved ribosomes increases (35). The increased fidelity of cloacin DF13-cleaved ribosomes has been explained (35)on the basis of increased ribosomal GTPase activity of these ribosomes. However, the bulk of evidence from other systems indicates that an increase in GTP hydrolysis in fact reduces the fidelity of translation (see Section 111). Therefore, the mechanism by which cloacin DF13 increases the fidelity of ribosomes is not clear at present. Inhibitors of protein synthesis that reduce ribosomal GTPase activity, such as abrin and ricin (36),have no effect on translational fidelity (37).In fact, several studies show that the enzymic binding of aminoacyl-tRNA to the ribosomal A-site is affected by the bacteriocin (38, 39). However, it is unlikely that the error-producing effect explains the toxicity of these molecules.

E. Biological Aging It has been speculated for some time (40)that the fidelity of translation is related to biological aging. The “error catastrophe” hypothesis predicts that aging will be accompanied by a decrease in the fidelity of protein synthesis ( 4 0 ) .This hypothesis was tested by a number of investigators using different cell types. In certain systems (41) aging resulted in a slight reduction in the efficiency of ribosomes, but no difference in fidelity was detected by most investigators (41-43). A small reduction in the fidelity of liver ribosomes of old rats was reported by one group (44).The bulk of the experimental evidence does not support the idea of decreased fidelity of ribosomes from old animals or from cells that have undergone several passages. Also, when tRNAs from aged and young mice were tested, no significant age-

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related differences in either efficiency or fidelity of translation were detected. The accuracy of protein synthesis in reticulocytes from several mammalian species having short and long life-spans shows no correlation with the life-span of the species (45).Therefore, it is unlikely that ribosomal infidelity is responsible for the accumulation of abnormal enzymes in senescent organisms.

II. Message Decoding

The classical codon reading scheme (46) demands that an anticodon must form stable base-pairs with all three nucleotides of the codon. To account for the base-pairing between inosine or of modified uracil residues in the anticodon with the codon, Crick advanced the “wobble” hypothesis (46). From the difference in energy between a correct and incorrect interaction, it has been calculated that an “incorrect”, aminoacyl-tRNA will bind and incorporate its amino-acid residue into the nascent chain roughly once for every 100 correct incorporations (47), if selection is based merely on codon/anticodon interaction. The difference between actual and predicted error rate has been attributed to GTP hydrolysis in the “kinetic proofreading” model (48).Discovery of an aminoacyl-tRNA binding site other than the A-site on the ribosome (49)has contributed significantly to a better understanding of the “proofreading” events taking place on the ribosome. A. Role of Ribosomes In the “three-site” model of the elongation cycle (50),aminoacyltRNA enters the A-site on the ribosome not by chance, but is flipped to this site after a preliminary contact with the recognition (R) site, which lies on the external face of the small subunit. During this movement of the tRNA, it is held to the ribosomes by no more than the binding of anticodon to codon. The motion of the tRNA therefore constitutes a test of the strength of base-pairing; it is, in a way, a second reading of the anticodon. Energy for the flipping is derived from GTP (50).The movement of aminoacyl-tRNA from the R-site to A-site provides an opportunity to test the correctness of codon/anticodon interaction. According to the model, the anticodon loop of tRNA switches from a 5’-base-stacked, to a 3-base-stacked conformation during this rotation, while the position of the mRNA is unaltered. Prior to this proposal, it was suggested that the movement of mRNA might be effected by a conformational rearrangement of the anticodon loop from one stacking conformation to another (51).The three-site model

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(50)makes it easy to understand the proofreading event taking place on the ribosome. Checking of the “reading frame” is extremely important for accuracy. If there were a gap between successive tRNAs bound to the Psite and the A-site, an incorrect codon would be recognized by the incoming aminoacyl-tRNA. Such mistakes can be prevented if the distance between successive codons recognized by the tRNAs can be checked through a base-pairing mechanism. The proximity of the nucleotide preceding the anticodon, i.e. the thirty-third nucleotide of tRNAs, makes base-pairing between peptidyl-tRNA and aminoacyltRNA possible when the “reading frame” of the codon in the A-site is correct with respect to the preceding codon. The thirty-third nucleotide(U), which is invariant1 in all tRNAs studied so far except for tRNAMetof eukaryotes, may play a significant role in preventing “frameshift error” during translation (50). It is not clear why such a mechanism, designed to prevent frameshifting, is not functional during mistranslation in rel mutants of E . coli (52). Based on the existence of a cluster of seven tRNA-binding-proteins on the outer surface of the small subunit (50),and the fact that mutations affecting some of these proteins interfere with the fidelity of translation (25), this external region of the small subunit has been suggested to determine aminoacyl-tRNA recognition (50).It should be realized, however, that other regions of the small subunit involved in mRNA binding, as well as proteins of the large subunit (28, 53) contribute to the fidelity of translation. B. Ribosomal Mutants Most of the mutants that contain alterations in ribosomal components are those resistant to or dependent on antibiotics such as streptomycin, spectinomycin, erythromycin (54). Few alterations of ribosomal protein have been observed in these mutants, and the altered proteins were further confirmed in some mutants by reconstitution experiments (55). Mutants possessing altered translation fidelity are those resistant to or dependent on aminoglycoside antibiotics as discussed in Section 1,C.

C. Role of ppGpp Mistranslation in relaxed mutants can be prevented if the level of ppGpp is elevated in one way or another (57-59). This nucleotide inhibits the irreversible partial reactions of protein synthesis involvSee Singhal et al. in this volume and Singhal and Fallis in Vol. 23. [Ed.]

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ing GTP hydrolysis (52).Improvement in the accuracy of tRNA selection by ppGpp has been attributed to the delayed catalytic step, facilitating the dissociation of incorrect substrates (52). An error-reducing effect, operating through a similar mechanism, has also been reported for fusidic acid (52).If this argument is correct, one should also expect inhibitors of ribosomal GTPase activity to contribute to the accuracy of translation. This point is discussed further in Section IV,C. It has been suggested that some mistranslation in relaxed mutants is a result of a spontaneous reading frameshift (52). Two types of evidence have been presented to support this postulate (52). First, mistranslation in relaxed mutants occurs not only at the amino-acid codons, but also at the termination codons, resulting in readthrough translation (52). Second, limitation of a particular amino acid also produces enhanced phenotypic suppression at codons specifying other amino acids (52).The spontaneous frameshifting has been ascribed to aberrant translocation (52, 56).

D. Readthrough Translation Peptide-chain synthesis normally terminates when one of the three “stop” codons UAA (ochre), UAG (amber), or UGA (opal) is encountered by a translating ribosome (60).These codons function as a termination signal at the end of a cistron, and also within the cistron, when they are generated by nonsense mutations. The termination process is mediated by the termination factors resulting in the hydrolysis of the peptidyl-tRNA linkage and release of the peptide (61). Most often the termination process is very efficient. However, in many systems, termination is suppressed or bypassed, allowing chain elongation to continue beyond the termination site. This phenomenon is known as “readthrough translation.” Readthrough translation can take place at mutationally generated internal termination codons as well as at normal termination codons. Several instances of readthrough translation consist of a mutated tRNA having a base change in its anticodon (62). Such tRNAs are genetically defined as “suppressor” tRNAs. Normal tRNAs can also misread a termination codon as an amino acid codon. A few examples of this type of misreading have already been discussed (Section 11,B). High Mg2+concentration in in vitro translation systems (63) and low temperature (64) facilitate readthrough. Naturally occurring readthrough translation in a mammalian system was first demonstrated in reticulocytes (3). In this system, suppression of termination at the normal termination codon of p-globin mRNA is effected by tRNATv. (3) It is, however, not clear whether suppression

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is caused by misreading. The best-known example of readthrough translation involves the Qp coat-protein cistron (65). As in the reticulocyte system, the amber codon is suppressed by tRNATv.The mechanism for the misreading of the UGA codon by tRNATQis not yet clear. Suppressor tRNAs do not adversely affect cells that contain them, and the efficiency of termination suppression of different amber codons by the same tRNA varies significantly (66).It has been postulated that the sequence of mRNA adjacent to the termination codon determines the efficiency of suppression (64, 67). A suppressor tRNA possessing a base change outside the anticodon region has been reported (68). Methylation or base-change outside the anticodon arm can alter the codon-reading ability of tRNAs (69). An unmodified tRNATyrreads through a termination signal in TMV RNA, while a corresponding isoacceptor tRNA, in which the first base of the anticodon is highly modified, does not act as a suppressor

(70). Are all readthrough proteins a product of translational error? At least in the QP system, it appears that the coat readthrough protein synthesis is essential for infection by QP (71).No essential function has been attributed to readthrough translation products from TMV RNA or p-globin mRNA (3). In avian sarcoma and murine leukemia viruses, the gag gene and the pol gene are translated together by readthrough translation and then processed to produce the two proteins (72, 73).

111. Accuracy of in Vitro Translation Systems Poly(U)-directed polyphenylalanine synthesis is widely used to measure the fidelity of in vitro translation systems. Using leucine of high specific acitivity, error frequencies of the order of one per 1000 translated codons can easily be detected. Further, the error rate can be elevated artificially by limiting the availability of the correct amino acid, phenylalanine, in the system. On the other hand, the greater solubility of polyleucine- and leucine-containing peptides, compared to polyphenylalanine, makes detection of incorporated leucine difficult. Since leucine and phenylalanine are chemically quite different, the error originating at the level of amino-acid activation is minimal in this system, and the observed values give a direct measure of decoding error. Finally, the absence of termination codons in homopolymers and some copolymers makes it possible to test the misreading of amino-acid codons as termination codons with such copolymers by measuring the release of polypeptide from peptidyl-tRNA.

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A. Polyvalent Cations The fidelity of translation is very much dependent on the concentrations of polyvalent cations used in the in vitru protein synthesis system (11, 16). Earlier systems used high concentrations of M 2 + as the only polyvalent cation (22, 74). But what is the optimal Mg2+concentration? This is a difficult question to answer for several reasons. First, with identical poly(U) and ribosome concentrations, prokaryotic and eukaryotic ribosomes exhibit different Mg2+ optima and error frequencies (75). In a poly(U)-programmed translation system at 13 mM Mg2+,and in the presence of streptomycin, error frequencies of 25% and 63% were reported for rabbit reticulocyte ribosomes and E . coli ribosomes, respectively (75). This is an extremely high error frequency, at least one order of magnitude higher than usual. Second, with synthetic messengers, the optimal Mg2+concentration is dependent on the concentrations and nature of polynucleotide and the amino acid tested (44). Generally speaking, ribosomes are more inaccurate at high Mg2+.In translational systems using synthetic messengers, where it is easy to measure the fidelity, a polyvalent cation concentration yielding highest fidelity, not highest synthesis, should be considered as optimal. More recent in uitro translation systems are optimized with polyamines (11)and other divalent cations (9). These polycations complement each other (I]), and therefore the optimal concentration of polyamine is very dependent on the concentration of the other ions. Several laboratories, using different translation systems, have clearly demonstrated that polyamines increase the fidelity of messenger decoding by the ribosomes (9,11,16).As pointed out above, polyamines complement Mg2+ in translation systems, and therefore their effect should be tested at low concentrations of Mg2+. In this context, it is interesting to note that polyamines cause increased infidelity of poly(U) translation at high concentration of Mg2+ (20). Several intermediate steps of protein synthesis in uitro are stimulated by polyamines, the extent of stimulation depending on assay conditions. A combination of Mg2+and polyamines in vitru results in higher incorporation of amino acids than does optimal Mg2+ alone (11).Non physiological concentrations of polyamines can induce profound conformational changes of nucleic acid components in the system, resulting in aggregation and precipitation (76). The mechanism by which polyamines increase the fidelity of translation is not clear. The ability of these amines to reduce leucine incorporation in the poly(U) system, also when aminoacyl-tRNAs are

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used instead of free amino acids, indicates that the observed effect is exerted at the level of codon/anticodon recognition on the ribosome. Like Mg2+, polyamines influence the conformation of ribosomes. Such conformational changes of ribosomes have been demonstrated by altered susceptibility of individual ribosomal proteins to lactoperoxidase-catalyzed iodination (77).The influence of polycations on the speed and accuracy of the elongation process is discussed in Section IV.

B. Temperature An increase in temperature up to 37°C has a positive effect on the transIation fidelity of most in vitro systems (64, 78).The wheat germ system, however, is an exception to this general rule (13), since it works optimally at temperatures below 30°C. For very short incubation periods, however, amino-acid incorporation can be slightly higher at 37°C (13).We have also observed that the temperature optimum for fidelity of this system is dependent on the concentration of polycations and on ionic strength (37).At higher temperatures, the optimal concentration of polycations is shifted toward higher values, as has been reported for the E . coli system (64,78).Since high temperature is known to destabilize the 30 S-poly(U)-tRNA complex (79), temperature increase might facilitate the rejection of incorrectly bound tRNA, and thereby increase fidelity. The effect of temperature on the accuracy of translation is not restricted to synthetic messengers, as has been clearly demonstrated for galactosidase synthesized in vitro at 20°C and 37°C (64).

C. Effect of tRNA Addition of a large excess of tRNA reduces the ambiguities in the decoding of synthetic messengers (80). Although this was demonstrated with crude tRNA preparations, the effect is obviously due to an increase in the ratio between cognate and noncognate tRNAs, thereby facilitating the selection process. The interesting question is whether physiological variations in the concentration of tRNA have a similar effect in vivo? There is a general feeling that one of the many roles of ribosomes is to equalize all codon/anticodon interactions, so that a differential rate of decoding does not occur (81).There are, however, several lines of evidence indicating that peptide chain elongation is nonuniform (82-84). The relative duration of discontinuity or pause in the elongation can be experimentally modulated by tRNA (83) and polyamines (84).Such discontinuities in elongation can influence fidelity (84).This is discussed in more detail in Section IV.

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D. Effect of pH The effect of a pH change on fidelity is more complex than usually pictured (85, 86). I n bacterial systems, poly(U) translation is more accurate at neutral or slightly acid p H (80).In a later communication from the same group, an opposite effect of pH change was reported for reticulocyte ribosomes (87). It should be pointed out that in most systems the endogenous messenger-dependent activity is reduced at neutral or slightly acid pH, compared to that of synthetic messengers. Since activity without poly(U) was not reported in these experiments, it is difficult to draw a general conclusion. The buffer most commonly used in in vitro translation systems, TrisC1, has a high temperature coefficient. Therefore, some of the observed effects of temperature on the translation fidelity might be an indirect effect of p H change.

E. Energy Regeneration Very convincing evidence as to the importance of an efficient energy-regenerating system for accurate translation has been presented (9). It appears that it is not the concentration of nucleoside triphosphate, but the ratio of triphosphates to their hydrolysis products that is important for high fidelity. Efficient energy regeneration increases the fidelity of translation in the wheat germ system also (37). However, in another study it was found that addition of elongation factors and GTP (without a GTP-regenerating system) to a system lacking these components increased the extent of poly( U) translation, but decreased the accuracy (88).This observation is difficult to explain. It may well be that phenylalanine tRNA has become the ratelimiting factor in the system, where elongation factors and/or GTP were rate-limiting beforehand.

F. Soluble Factors Are ribosomes self-maintained with molecular mechanisms for messenger-dependent tRNA selection, transpeptidation, and translocation? If so, elongation factors and GTP do not create new mechanisms, but only impart increased power to the existing mechanism of elongation (89). It is clear that no external energy is required for the transpeptidation reaction. However, it is difficult to envisage a mechanism for tRNA selection and mRNA movement without conformational changes. As pointed out above, the actual error rate of messenger decoding is at least two orders of magnitude lower than predicted (47). Elongation factors and GTP play a very important role in this selection process (48) and in the movement of messenger.

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Also, release factor RF-1 appears to reduce readthrough translation of the amber codon (64). Release factor RF-1 also prevents incorrect reinitiation at a codon immediately following an amber codon of R17 phage (90).

IV. Relationship between Speed and Accuracy A. General In early experiments with synthetic messengers (19,20,91) it was observed that the frequency of translational errors is a function of the Mg2+ concentration and that the concentration of Mg2+required for maximal activity is higher than that required for maximal accuracy. Observations of this nature led to the belief that speed and accuracy of protein synthesis might be inversely related (92). This assumption was supported by the observation that addition of elongation factors and GTP to a system free of these components increased the extent of poly(U) translation but decreased the accuracy (88).The validity of this thesis has been critically reviewed (86). The bulk of evidence indicates that most of the fast systems tend to be more accurate. The main problem is the calculation of the speed of synthesis from total amino acids incorporated, since in almost all in vitro systems only a fraction of ribosomes are active (93). Furthermore, as stated earlier, accuracy is very much dependent on the ratio between cognate and noncognate aminoacyl-tRNAs in the system. Such data are not given in most reports. A number of factors that influence the speed of synthesis are discussed below.

B. Inhibitors of the Elongation Process Several translation inhibitors reduce the elongation rate by inactivating either ribosomes or elongation factors. Others inhibit the rate of elongation by interfering with the formation of complexes between elongation factors, GTP, and ribosomes. These inhibitors are very useful in studying the relationship between the speed and accuracy of translation. A large number of protein toxins are known to block the elongation cycle by catalytically inactivating the 60 S subunit of eukaryotic ribosomes (94,95). The most thoroughly investigated toxins of this group are abrin and ricin. Others, like a-sarcin, mitogillin, and restrictocin, cleave a 320-nucleotide-long piece of RNA from the 3' end of the large ribosomal RNA (95, 96). In all cases, the catalytic inactivation of the large ribosomal subunit results in a decrease in the

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affinity of elongation factors for ribosomes. Toxin-inactivated ribosomes possess reduced factor-dependent GTPase activity and aminoacyl-tRNA binding to the ribosomes (36). Diphtheria toxin inhibits the translocation process by catalytically inactivating EF-2. This inactivation requires NAD+, and the inactivated EF-2 contains ADP-ribose, covalently attached to the protein (97, 98). The (ADP-ribosyl) (EF-2) * GTP complex can interact with ribosomes with low affinity. The rate of GTP hydrolysis and translocation by ADP-ribosylated EF-2 is inhibited (99). We have studied the effect of abrin, ricin, and diphtheria toxin on the speed and accuracy of poly(U) translation in nuclease-treated rabbit reticulocyte lysates (100).All three toxins strongly inhibit the elongation rate but do not reduce the accuracy of translation (100). Cycloheximide, another inhibitor of the translocation process, has been reported to increase the fidelity of poly(U) translation (101).No data were given on the speed of the poly(U) translation. Cycloheximide in the range used (1 to 4 x lO-'M) in our laboratory, did not inhibit poly(U) translation, but inhibited globin mRNA translation by more than 50%. A possible explanation for the reciprocal relationship between speed and accuracy observed in the absence of elongation factors and GTP (88)is discussed in Sections III,E and F.

-

C. Ribosomal GTPase Activity An increased fidelity of bacterial ribosomes cleaved by cloacin D F 13 was reported by one group (35),whereas no effect on fidelity was observed by another (33).Both groups observed a decrease in the rate of translation by toxin-inactivated ribosomes. The increased fidelity was explained on the basis of an elevated GTPase activity of the inactivated ribosomes (35).This explanation is difficult to understand. First, cloacin inactivates the small ribosomal subunit, and the GTPase site is located on the large subunit (102).Second, it is not the ribosomal GTPase activity, but the regeneration of GTP, that is important for accuracy (9). Therefore, increased GTPase activity of the ribosomes should be expected to reduce the accuracy of translation by reducing the ratio between triphosphate and its hydrolysis products (9).

D. Temperature Change In almost all in vitru and in vivu systems, amino-acid incorporation is higher when the temperature is raised. As pointed out in Section III,B, a moderate increase in temperature also increases the fidelity of translation. In most poly(U) systems, only a small fraction of the ribo-

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somes are active (93).Therefore, a higher rate of amino-acid incorporation could be the result of increased initiation, not of elongation. However, noninitiating systems containing natural messengers also show an increased rate of elongation when the temperature is raised (64).Changes in temperature influence the pH (see Section II1,D) and the Mg2+ optima of in vitro systems (78) in a manner that increases accuracy. These indirect effects alone are not sufficient to explain the significant effect of temperature on the fidelity of translation. Temperatures above 37°C increase the accuracy of poly(U) translation without increasing the total amino-acid incorporation (78,75).As pointed out above, there is no a priori reason to believe that an increase in elongation rate should result in increased amino-acid incorporation. A high temperature could make initiation in poly(U) systems inefficient

(103).

E. Error-Producing Antibiotics Error-producing antibiotics, such as streptomycin, neomycin, and kanamycin, have been reported to increase the rate and decrease the accuracy of translation (19, 92, 104), whereas other reports indicate that these antibiotics reduce the rate as well as the accuracy (105109).The bulk of recent evidence favors the view that under physiological conditions they reduce both the speed and the accuracy of protein synthesis (86).It has been pointed out that the mechanism of action of these antibiotics is complicated (86, 110), and the errorproducing effect is perhaps not the only mechanism for their action

(110).

F. Summary Accuracy of translation in vivo offers the most convincing evidence in favor of the view that speed and fidelity are not opposing parameters of translation. The error frequency in vivo is at least two orders of magnitude lower (7, 8 ) , while the rate of elongation is severalfold higher, than in most in uitro systems (93).

V. Codon Misreading by lsoacceptor tRNAs The very high fidelity of protein synthesis in vivo is not incompatible with the idea that a significant misreading of certain codons can take place also in uiuo, without affecting the amino-acid sequence of proteins synthesized. For instance, in a codon family (i.e., groups of four codons specifying the same amino acid), the third nucleotide could be misread by the first nucleotide of the anticodon, without any

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adverse effect on translation fidelity (10).In a protein-synthesizing system from E . coli, programmed with MS2 phage RNA, misreading of this type took place in vitro (111-114). When two isoacceptor tRNAs with different anticodons competed for the same codon, the tRNA able to read the codon by the classic base-pairing scheme was only one order of magnitude more efficient than the competing tRNA misreading the wobble base (113, 114). The misreading frequency observed in this system under conditions of competition was very high (lo%), compared to the translational error frequency (0.03%)of in vitro systems (9). Misreading of the third base was not observed for the Iysine codon AAA by lysine tRNA having the anticodon CUU (115), while there was considerable misreading of the glutamine codon CAA by glutamine tRNA having the anticodon CUG (113).Misreading of the glutamine codon CAA was observed even in the presence of the glutamine tRNA having the correct anticodon ( 113). To account for these results, a two-out-of-three misreading hypothesis was invented (10). It is assumed that the probability of such misreading for any codon is a function of the strength of interaction between anticodon and the first two nucleotides of the codon. Thus, the misreading probability is greatest for codons making G-C interactions in the first two positions, intermediate for codons having one A-U and one G-C interaction, and minimal for codons making only A-U interactions. This model explains the lack of misreading of lysine codon AAA, and the misreading of glutamine codon GAA (113, 115). These experiments were not conducted under conditions yielding optima1 fidelity in uitro ( 9 , I I ) .Therefore, its relevance under in viuo conditions is not clear. When this system was optimized with polyamines, the misreading of glutamine codon CAA by glutamyl-tRNA with the anticodon CUG was significantly reduced ( 1 16). Demonstration of codon misreading by isoacceptor tRNAs (19)has emphasized the problems involved in message decoding. Base-pairing between codons and anticodons alone does not account for the high accuracy observed. The deviation from the classical base-pairing rules (117 ) during the interaction of the third base of the codon with the wobble position of the anticodon is even more clear in the mitochondria1 genetic system (118,119).According to the wobble hypothesis (117)a minimum of 31 tRNAs is required to read the 61 possible amino acid codons. Based on an almost complete sequence analysis of human mitochondrial DNA (118) it has been concluded that, for each codon family, there is only one tRNA in the mitochondrion, and that this tRNA

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always contains a U or modified U in the wobble position. The twoout-of-three hypothesis ( 10) was suggested as a possible explanation for these results.

VI. Summary and Future Perspectives Several factors that influence the fidelity of in vivo and in vitro translation are discussed. The in vitro systems, which are more errorprone, can be improved by controlling factors such as temperature, pH, ionic strength, energy regeneration, and composition of the system with respect to polyvalent cations, tRNAs, and protein factors. The notion that speed and accuracy of protein synthesis are opposing parameters is questioned on the basis of a Iarge number of data indicating that fast systems are more accurate. An important factor that is relevant for accurate translation is perhaps the coordinated movement of messenger bound to several ribosomes in a polysome. A nonsynchronized movement could affect codonlanticodon interaction adversely. Such nonsynchronized movement might occur during poly( U) translation, where initiation can take place virtually at any point along the messenger. Therefore, the reading frame with respect to one ribosome is most probably different from that of another ribosome translating the same poly(U) molecule. However, since Watson-Crick base-pairing requires antiparallel orientation of codon and anticodon, translation will still be unidirectional. Are nonsynchronized translational systems more errorprone? Cycloheximide can be a useful tool to answer some of these questions, since a concentration of this inhibitor that inhibits translation of natural messenger almost completely, does not inhibit poly(U) translation under certain conditions (100). The mechamism of message decoding as we know it at present, does not offer a satisfactory explanation for the high fidelity of this process. The stability of interaction between codon and anticodon by itself is far too weak to account for the precision of message decoding. The interactions of cognate triplets are only ten times more stable than corresponding interactions of triplets containing a single mismatched pair of nucleotides (120, 121). Judged from the error frequency of message decoding (8),it is clear that, in practice, the advantage of cognate tRNA interaction is several orders of magnitude over noncognate interaction. Ribosomes, tRNA, and factors contribute significantly to the stability of codon/anticodon interaction (122). A critical aspect of the tRNA selection mechanism is that it must provide for the rapid release of noncognate tRNA molecules. There-

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fore, the theoretical models (40,123) should take into account the fact that selection must take place before the tRNA is fixed into the ribosomal A-site. If selection is achieved after such fixing, clearly, any perturbation of the system that slows down the release of noncognate tRNA molecules could affect the fidelity of the process. The three-site model of the elongation cycle attempts to solve some of these problems (50),and provides an explanation for the weaker interaction of tRNA with the R-site compared to the A-site. Furthermore, the switching of tRNA from the R-site to the A-site provides an opportunity to test the stability of codon/anticodon interaction during a conformational change of tRNA structure, independent of other forces acting on the tRNA bound to the ribosomal A-site.

ACKNOWLEDGMENTS I wish to thank Professor A. Pihl for comments and criticism and Miss Teresa Cieplinska and Mrs. Torill Sage for valuable assistance in the preparation of this paper. The Norwegian Cancer Society is thanked for financial aid.

REFERENCES L. Pauling, Festchr. Prof. Dr. Arthur Stoll Siebzigsten Geburttstag 1957, p. 597. 0. W. Jones and W. Nirenberg, PNAS 48,2115 (1962). A. I. Geller and A. Rich, Nature (London) 283, 41 (1980). [See also 70, Ed.] H. Engelberg-Kulka, NARes 9, 3 (1981). [See also 70, Ed.] 5. M. Royoji, R. Berland, and A. Kaji, PNAS 78, 597, 3 (1981). 6. R. B. Loftfield, BJ 89, 82 (1963). 7. R. B. Loftfield and D. Vanderjagt, BJ 128, 1353 (1972). 8. P. Edelmann and J. Gallent, Cell 10, 131 (1977). 9. P. C. Jelenc and C. G. Kurland, PNAS 76, 3174 (1979). 10. U. Lagerkvist, PNAS 75, 1759 (1978). 11. A. K. Abraham, S. Olsnes, and A. Pihl, FEBS Lett. 101,93 (1979). 12. P. H. O’Farrell, Cell 14, 545 (1978). 13. J. Parker, J. W. Pullard, J. D. Friesen, and C. P. Stanners, PNAS 75, 1091 (1978). 14. K. G. Gross and A. 0. Pogo,JBC 249,568 (1974). 15. J. R. Warner, R. J. Tushinski and P. J. Wejksnora, in “Ribosomes” (G. Chambliss, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds. ), p. 207. Univ. Park Press, Baltimore, Maryland, 1980. 16. K. Igarashi, K. Kaskiwagi, R. Aoki, M. Kojima, and S. Hirose, BBRC 91,440 (1979). 17. K. Igarashi, K. Hara, Y. Wantanabe, S. Hirose, and Y. Takeda, BBRC 64,897 (1975). 18. S. H. Goldemberg and I. D. Algranati, Med. B i d . 59, 360 (1981). 19. J. Davies, W. Gilbert, and L. Gorini, PNAS 51, 883 (1964). 20. S. M. Friedman and I. B. Weinstein, PNAS 52,988 (1964). 21. J. Davies, L. Gorini, and B. Davies, J . Mol. Phartnacol. 93, 1 (1965). 22. J. Davies, D. S. Jones, and H. B. Khorana, J M B 18,48 (1966). 23. P. Strigini and L. Gorini, JMB 47, 517 (1970). 24. G. Funatsu and H. G. Wittmann, J M B 68,547 (1972). 25. M. Ozaki, M. Mizushima, and M. Nomura, Nature (London)222,333 (1969). 1. 2. 3. 4.

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26. G. Schreiner and K. Nierhaus, JMB 81,71 (1973). 27. D. Lado, M. A. Cousin, T. Ojasoo, and J. P. Raynaud, EJB 66,597 (1976). 28. R. Kiihberger, W. Pipersberg, A. Petzet, P. Buckel, and A. Bock, Echem 18, 187

(1979). 29. C. G. Kurland, R. Rigler, M. Ehrenberg, and C. Blomberg, PNAS 72,4248 (1975). 30. W. Pieperberg, V. N. Oseda, and Bock, Mol. Gen. Genet. 171, 23 (1979).

31. L. Topisirovic, R. Villarroel, M. De WiIde, A. Herzog, T. Cobezar, and A. Bollen, Mol. Gen. Genet. 28, 89 (1977). 32. S. H. Goldemberg and I. D. Algranati, EJB 117, 251 (1981). 33. T. Boon, PNAS 69,549 (1972). 34. P. C. Tai and B. D. Davies, PNAS 71, 1021 (1974). 35. J. C. Twilt, G. P. Overbeck, and J. Van Duin, EJE 94,484 (1979). 36. S. Benson, S. Olsnes, A. Pihl, J. Skorve, and A. K. Abraham, EJB 59,573 (1975). 37. A. K. Abraham, unpublished observations. 38. F. Turnovsky and G. Hognauer, BBRC 55, 1246 (1973). 39. G. Sander, EJB 75,523 (1977). 40. L. E. Orgel, PNAS 49, 517 (1963). 41. N. Mori, D. Minzuno, and S. Goto, Mech. Ageing Deu. 10,379 (1979). 42. J. M. Buchanan, C. L. Bunn, R. I. Lappin, and A. Stevens, Mech. Ageing Deu. 12, 339 (1980). 43. R. I. Wojtyk and S . Goldsterin,]. Cell. Physiol. 103,299 (1980). 44. J. J. Butzow, M. G. McCool, and G. L. Eichhorn, Mech. Ageing Dev. 15,203 (1981). 45. G. P. Hirch, R. A. Popp, M. C. Francis, B. S. Bradshaw, and E. G. Bailiff, Ado. Pathobiol. 7, 142 (1980). 46. F. H. C. Crick, J M B 19,548 (1966). 47. J. J. Hopfield and T. Yamane, in “Ribosomes” (G. Chambliss, G . R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 585. Univ. Park Press, BaItimore, Maryland, 1980. 48. J. J. Hopfield, PNAS 71, 4135 (1974). 49. J. A. Lake, in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 207. Univ. Park Press, Baltimore Maryland, 1980. 50. J. A. Lake, PNAS 74, 1903 (1977). 51. C. Woese, Nature (London)226,817 (1970). 52. J. Gallent and D. Foley in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 615. Univ. Park Press, Baltimore, Maryland, 1980. 53. C. G. Kurland, R. Rigler, M. Ehrenberg, and C. Blomberg, PNAS 72,4248 (1975). 54. J. Davies and M. Nomura, Annu. Reu. Genet. 6,203 (1972). 55. M. Nomura, E. A. Morgan, and S. R. Jaskunas, Annu. Rev. Genet. 11,297 (1977). 56. C. G. Kurland, in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L.Kahan, and M. Nomura, eds.), p. 597. Univ. Park Press, Baltimore, Maryland, 1980. 57. N. P. Fiil, B. M. Willumsen, J. D. Friesen, and K. Von Meyenberg, Mol. Gen. Genet. 150,87 (1977). 58. J. D. Friesen, G. An, and N. P. Fiil, Cell 15, 1187 (1978). 59. J . Gallant and R. Lazzarini, in “Protein Synthesis” (E. H. McConkey, ed.), p. 309. Dekker, New York, 1976. 60. S. Brenner, W. Stretton, and S. Kaplan, Nature (London) 206,994 (1965). 61. J. Lucas-Lenard and F. Lipmann, ARB 40,409 (1971).

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62. H. Goodman, J. Abelson, A. Landy, S. Brenner, and J. Smith, Nature (London) 217, 1019 (1968). 63. M. Capecchi, JMB 30,213 (1967). 64. J. L. Manley and R. F. Gesteland,JMB 125, 433 (1978). 65. A. M. Weiner and K. Weber, JMB 80,837 (1973). 66. H. Yahata, Y. Ocada, and A. Tusgita, Mol. Gen. Genet. 106,208 (1970). 67. H. Engelberg-Kulka, NARes 9, 983 (1981). 68. D. Hirsh,JMB 58,439 (1971). 69. A. M. Korner, S. I. Feinstein, and S. Altman, in “Transfer RNA” (S. Altman, ed.), p. 105. MIT Press, Cambridge, Massachusetts, 1979. 70. M. Bienz and E. Kubli, Nature (London) 294,188 (1981). 71. H. Hofstetter, H. J. Monstein, and C. Weissman, BBA 374,238 (1974). 72. K. Beemon and J. Hunter, PNAS 74,3302 (1977). 73. H. Oppermann, J. M. Bishop, H. E. Vermus, and L. Livintow, Cell 12,993 (1977). 74. 0. W. Jones and M. W. Nirenberg, PNAS 48,2115 (1962). 75. S. M. Friedman, R. Breezney, and I. B. Weinstein,JBC 243,5044 (1968). 76. A. K. Abraham and A. Pihl Trends Biochem. Sci. 6, 106 (1981). 77. C. J. Michalski, S. M. Boyle and B. H. Sells, C a n . J .Biochem. 57, 250 (1979). 78. W. Szer and S. Ochoa, JMB 8, 823 (1964). 79. M. A. Glukhova, N. A. Belitsina, and A. S. Spirin, EJB 52, 197 (1975). 80. M. Grunberg-Manago and J. Dondaon, BBRC 18,517 (1965). 81. B. F. C. Clark, in “Ribosomes” (G. Chambliss, G . R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 413. Univ. Park Press, Baltimore, Maryland, 1980. 82. A. Protzel and A. J . Morris, JBC 249,4594 (1974). 83. P. M. Lijardi, V. Mahdavi, D. Shields, and A. G . Candelas, PNAS 76,6211 (1979). 84. A. K. Abraham and A. Pihl, EJB 106,257 (1980). 85. M. Yarus, This Series 23, 195 (1979). 86. M. Laughrea, Biochimie 63, 145 (1981). 87. H. Lamform and M. Grunberg-Manago, BBRC 27, 1 (1967). 88. L. P. Gavrilova, 0. E. Kostiashkina, V. E. Koteliansky, N. M. Rutkevitch, and A. S. Spirin, JMB 101, 537 (1976). 89. A. S. Spirin, This Series 21, 39 (1978). 90. M. Ryoje, R. Berland, and A. Kaji, PNAS 78, 5973 (1981). 91. A. G . So, J. W. Bodley, and E. W. Davie, Bchem 3, 1977 (1964). 92. C. G . Kurland, ARB 46, 173 (1977). 93. E. G . H. Wagner, P. E. Jelec, M. Ehrenberg, and C. G. Kurland, EGJ 122, 193 (1982). 94. S. Olsnes and A. Pihl, Nature (London) 238,459 (1972). 95. D. Vazquez and A. JimBnez, in “Ribosomes” (G. Chambliss, G. R. Craven, J. Davies, K. Davis, L. Kahan, and M. Nomura, eds.), p. 847. Univ. Park Press, Baltimore, Maryland, 1980. 96. F. P. Conde, C. Fernadez-Puentes, M. T. V. Montero, and D. Vazquez, FEMS MicrobioE. Lett. 4, 349 (1978). 97. D. M. Gill and L. L. Dinius, JBC 248,654 (1973). 98. A. M. Pappenheimer and D. M. Gill, Science 182,353 (1973). 99. A. M. Pappenheimer, ARB 46,69 (1977). 100. A. K. Abraham and A. Pihl, EMBOJ., submitted. 101. M. Kurkinen, FEBS Lett. 124, 79 (1981).

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102. D. Donner, R. Villems, A. Liljas, and G. G. Kurland, PNAS 75,3192 (1978). 103. T. Nakamoto and D. Kolakofsky, PNAS 55,606 (1966). 104. A. S. Spirin, 0. E. Kostiashkina, and J. Joniik,JMB 101,553 (1976). 105. B. J. Wallace, P. C. Tai, E. L. Herzog, and B. D. Davis, PNAS 70, 1234 (1973). 106. B. D. Davis, P. C. Tai, and B. J. Wallace, in “Ribosomes” (M. Nomura, A. Tissi&res,and P. Lengyel, eds.), p. 771. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1974. 107. H. R. V.Amstein, R. A. Cox, and J. A. Hunt, Nature (London) 194, 1042 (1962). 108. J. Davies, and B. D. Davis,/BC 243,3312 (1968). 109. P. C. Tai and B. D. Davis, Bchem 18, 193 (1979). 110. J. R. Menninger, JBC 251,3392 (1976). 111. S. K. Mitra, F. Lustig, B. Akesson, U. Lagerkvist, and L. Strid,JBC 252,471 (1977). 112. E. Goldman, W. H. Holmes, and G. W. Hatfield,JMB 129,567 (1979). 113. T. Samuelsson, P. Elias, F. Lustig, T. Auxberg, G. F@lsch,B. Akesson, and U. Lagerkvist, JBC 255,4583 (1980). 114. S. K. Mitra, F. Lustig, B. Akesson, T. Auxberg, P. Elias, and U. Lagerkvist,JBC 254,6393 (1979). 115. P. Elias, F. Lustig, T. Auxberg, B. Akesson, and U. Lagerkvist, FEBS Lett. 98, 145 (1979). 116. A. K. Abraham, Med. Biol. 59, 368 (1981). 117. F. H. Crick,/MB 19 548 (1966). 118. B. G. Barrel, S. Anderson, A. T. Bankier, M. H. L. de Bruijn, E. Chen, A. R. Coulson, J. Drouin, T. C. Eperon, D. P. Nierlich, B. A. Roe, F. Sanger, P. H. Schreier, A. J. H. Smith, R. Staden, and I. G. Young, PNAS 77,3164 (1980). 119. S. G.Bonitz, R. Berlani, G. Curuzzi, M. Li, F. G. Nobrega, M. P. Nobrega, B. E. Thalenfeld, and A. Tzagoloff, PNAS 77,3167 (1980). 120. 0. C. Ulenbeck, J. Baller, and P. Doty, Nature (London) 225,508 (1970). 121. J. Eisinger, BBRC 43,854 (1971). 122. H. Grosjean, D. G. SoI1, and D. M. Crothers,JMB 103,499 (1976). 123. J. Nino, J M B 84, 297 (1974).

Structure qnd Functions of Ribosomal Protein S1 ALAP-RAMANSUBRAMANIAN Max-Planck-lnstitut f u r Molekulare Cenetik, Abteilung Wittmann, Berlin-Dahlem, Federal Republic of Germany

I. Isolation, Physical Properties, and Shape.. . . . . . . . . . . . . . . . . . . 102 A. Isolation Procedure. . . . . . . . 104 B. General Properties , C. Physical Constants . D. Probable Shape. . . . 11. Structure. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 A. Amino-Acid Sequences. . . . . . . . . . 109 B. Four Repeating Homologous Stretches . . . . . . . . . . . . . . . . 111 C. Secondary Structure Elements . . . . . . . . . . . . . . . . . . . . . . . 113 111. Discernible Structural and Functional Domains. . . . . . . . . . . . 113 A. M u t a n t F o r m M l S l . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114 B. Stable Fragment by Limited Proteolysis . . . . . . . . C. Ribosome-Binding Fragment. . . . . . . . . . . . . . . . . . D. Nucleic-Acid-Binding Fragment from the Central Region. 118 E. The -SH Groups . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 F. Evidence for an Interdomain Hinge Region . . . . . . . . . . . . 119 IV. The S1 G e n e . . . . . A. Interactions with Nucleic Aci

. . . . . . . . . . . . . . . . . 123 . . . . . . . . . . . . . . . . . . . 123

VII. Functions of S1

B. S1 Function in Protein Synthesis . . . . . . . . . . . . . . . . . . . . . VIII. Mode of Action of S1 . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

130 134 137

The ribosomes of bacteria and of the chloroplasts in plant and algal cells contain between 50 and 60 different proteins. Ribosomes of the eukaryotic cell cytoplasm contain a still greater number (i.e., about 80) of individual proteins. The most studied and the best characterized of all ribosomes is the Escherichia coli ribosome (I),which contains one copy each,of 51 proteins and four copies of a dimorphic protein in the form of a tetramer on the large subunit. The E . coli r-proteins' range in

'

Abbreviations. r-protein, ribosomal protein; rRNA, ribosomal RNA. Prefixes S and L designate, respectively, r-proteins of the small subunit and large subunit. 101 Progress in Nucleic Acid Research and Molecular Biology, Vol. 28

Copyright 0 1983 by Academic Press, Inc. All rights of reproduction in any form reserved.

ISBN 0-12-5400284

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A L A P - W A N SUBRAMANIAN

M, from 5000 to 61,000; they also cover an equally broad range in isoelectric character, from acidic proteins of pZ 4.5 to the most highly basic proteins in the cell. [The proteins of the ribosome constitute less than half of its macromolecular mass (36%in the case of E . coli ribosome), the remainder being carried by either three or four RNA molecules. An article on the structure of rRNA is included in this volume (21.1 The evolution of such a complex and distinct set of proteins in both the 70 S type (bacteria, plastids, mitochondria) and the 80 S type (eukaryotic cytoplasm) of ribosomes would indicate the performance of important, specific functions by these individual proteins. Yet there are very few r-proteins to which a clear-cut functional role has been assigned. The difficulty is attributed to the cooperative nature of the interactions of r-proteins in both ribosome assembly and ribosome function (see reference 3 for a review on this topic) and to the constraints in r-protein evolution, because they form a part of a particle that performs an essential, vital function for the organism. There are two r-proteins that have clear functional roles in protein biosynthesis: S1 on the small ribosomal subunit, and L7/L12 (the dimorphic protein) on the large subunit. Together on the ribosome they amount to 110,000 daltons, or over one-eight of the r-protein mass. They are also the best studied of all r-proteins. The structure, function and evolution of L7/L12 have been described in recent review articles (4, 5); and the structure and homology relationships of a chloroplast L12 protein have been published (6).In this article, I summarize and interrelate the results of studies on the structure and functions of S1.

1. Isolation, Physical Properties, and Shape S1 is a prominent protein of E . coli ribosome (Fig. 1).Although this statement would appear to be self-evident v i s - h i s Fig. 1, there were, in the early years of ribosome research, serious questions on both the molar stoichiometry of S1 and of its relation to the ribosome (7-9). With a binding constant of 2 x lo8 M-' for its attachment to the ribosome (10,11), S1 is one of the most weakly held of all ribosomal proteins. It is therefore partly removed when ribosomes are purified by centrifugation through a high-salt (e.g., 1 M NH4C1) buffer (a traditionally used procedure to purify ribosomes). Because S1 is easily removed, ribosome-free fractions of the cell extract always contain appreciable amounts of free S1. Immunological assays (12)indicate that 10% and 20% of the cellular content of S1 are present, respectively, in postribosomal supernatant and 1 M NH&l washes of ribo-

RIBOSOMAL PROTEIN

s1

103

FIG.1. Ribosomal proteins of‘ Escherichia coli separated by two-dimensional gel electrophorsis. S 1 and a few other ribosomal proteins mentioned in the text are indicated. At the right is shown purified S 1 prepared by the poly(U)-Sepharose procedure (Section I). Details of electrophoresis and spot identifications are given in references 191 and 192.

somes. Since the various assays for the translational function of the ribosome require supplementation by one or both of these fractions, earlier such assays with S1-lacking ribisomes did not reveal a requirement for S1. That requirement was revealed later when methods were developed to remove free S1 by specific procedures (e.g., use of antiS l IgG or binding to poly(U)-Sepharose; see Section VII). S1 remains bound to polyribosomes more firmly than it does to free ribosomes. I n an experiment done to detect possible variations in the stoichiometry of r-proteins in the cell (13),the ratio of S1 (polysome) to S1 (free ribosome) was 1.9, while the corresponding ratio for all other ribosomal proteins (except S21) was almost exactly 1, indicating a greater loss of S1 from the free ribosome. S1 is also exchanged between ribosomes, as was shown by the “heavy isotope transfer” experiment wherein E . coEi was first grown in DzO on deuterated glucose and l5NH4C1 and then transferred to normal medium and allowed further growth (14).A ready exchange of S1 bound to the 30 S subunit with that added free is also easily observed (15). The molar stoichiometry of S1 in early experiments was as low as 0.1, evidently as a consequence of its loss during the isolation and analysis steps (8,9).A stoichiometry of 0.7 to 0.9 was later obtained b y using endogenous or in uitro-generated polysomes for the analysis

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(16). Since then, I have obtained stoichiometry values of 0.7 to 1.2 in unwashed ribosomes isolated from different strains of E . coli grown under different growth conditions (17). S1 is thus a unit protein, like all E. coli r-proteins other than L7/L12 (18-20). Salmonella typhimurium grown in a rich medium that permits 2.4 doublings per hour contain 250 mg of rRNA per gram dry weight of cells (21). If these numbers are valid for E. coli, then, at 1 mol of S1 (M,= 6.1 X lo4)per mole of rRNA ( Z M , = 1.48 x lo6),there would be 10 mg of S1 per gram dry weight, or 2 mg of S1 per gram wet weight (22) of E . coli cells. The soluble proteins are estimated to be 50% of the bacterial dry mass (22), and therefore S1 would constitute, according to this analysis, 2% of the soluble proteins in maximally growing E. coli. A. isolation Procedure Several methods are described in the literature for the purification of S1 from either 70 S ribosomes or isolated 30 S subunits (12,23-26). We have previously published (27) a procedure for the large-scale purification of both S1 and the groEprotein (28), but afterward have exclusively used an affinity chromatography procedure using commercially availabIe poly(U)-Sepharose. This method is based on the previous findings that S1 binds so strongly to poly(U)- or poly(C)cellulose columns that 6 M urea is required to elute it (29),and that S1-free ribosomes can be prepared by passing a ribosome solution through poly(C)-cellulose (30).When we repeated the latter procedure (but using poly(U)-Sepharose) we found that 80% of the input ribosomes did not emerge from the column. In an effort to elute the bound ribosomes we washed the column with 1 M NH4C1-containing buffer and observed that the ribosomes now eluted totally free of S1. The bound S1 was next eluted with 6 M urea. We immediately realized that a method of choice was in our hands, suitable both for isolation of S 1from ribosome and for preparing S l-free ribosomal particles. The method can be used on a small scale to prepare S1-free 70 S ribosomes or 30 S subunits needed for in vitro assays, e.g., a 5-ml column for 15 mg of 30 S or 45 mg of 70 S; or it can be used on a large scale to isolate S1, e.g., a 50-ml column with 5 g of ribosome per passage. The ribosome or 30 S subunit is dissolved in 10 mM TrisC1, pH 7.6,20 mM Mg acetate, 6 mM 2-mercaptoethanol, 1M NH4C1. The ribosome concentration may be as high as 50 mg/ml. The S1 eluted from the column by 6 M urea is generally contaminated by RNA and traces of r-proteins, which are removed by chromatography on a column of DEAE-cellulose (in the presence of 6 M urea, 10 mM TrisC1, pH 7.6,6 mM 2-mercaptoethanol). The protein is eluted with a

RIBOSOMAL PROTEIN

s1

105

0 to 0.15 M NaCl gradient, e.g., 2 x 400 ml for a 50-ml column. The peak of S1 appears at about 0.07 M NaCI. The S1 is very pure at this stage as determined by electrophoresis (Fig. 1)or sedimentation analysis, and, after removal of urea by dialysis, it is functionally active in poly(U)- or phage-RNA-dependent protein synthesis with S l-free ribosomes (31). The saturating level of activity in such assays is reached by the addition of 1 mol of S1 per mole of ribosome; i.e., the isolated S1 is homogeneous in its functional properties (see Fig. 9, Section VII). Some additional details of this purification and a procedure to synthesize poly(U)-Sepharose of higher than commercially available capacity are included in a recent publication (32).

B. General Properties The absorption spectrum of S1 is that of a typical protein with a peak at 280 nm and a tryptophan shoulder at 285 nm. Purified S1 shows an A280 :A260 ratio of 1.6, and the protein concentration may be estimated from the 280 nm absorption using the conversion factor A&., = 7.5 (33).The true concentration of S1 can also be estimated by the usual Lowry procedure (using as standard bovine serum albumin) because we have observed that the concentration so determined is only about 10% higher than that determined by the absolute procedure of acid hydrolysis and amino-acid analysis. Free S1 is a very soluble protein. Solutions containing 20-30 mg of protein per milliliter have been prepared for the study of its smallangle X-ray scattering behavior (15,27,34).The isoelectric point of S1 is 4.7 (12); below this pH, S1 is only sparingly soluble. S1 has a tendency to bind to plastics and to Millipore and Diaflo filters; the loss by such means could be substantial at low (<0.2 mg/ml) protein concentration. Such loss (e.g., during protein concentration by Diaflo filtration) is prevented by the presence of 6 M urea. Purified S1 is rapidly inactivated by heat above 50°C (12,26). Investigators working with S 1have been concerned with the question of possible irreversible effects of urea on S1. Therefore, apart from taking the usual precautions in working with urea (i.e., fresh solutions of pure grade urea, working at 0-4°C and keeping the urea exposure time to a minimum, adding 2-mercaptoethanol as a scavenger), they have compared the properties of S1 prepared with and without urea. In molecular mass, sedimentation coefficient, radius of gyration, frictional coefficient, poly( U)-binding ability and in protein synthesis activity with synthetic or natural mRNA, no detectable differences have been observed in such comparisons (12,15,27,31,34). The NMR spectrum of S1 was also found to be independent of the method of preparation (35).

106

ALAP-RAMAN SUBRAMANIAN

C. Physical Constants 1. MOLECULARMASS The M, of S l has been determined by many investigators using a variety of methods, but, for reasons not clear, these values show a large variation. The M , values yielded by different methods are: 63,000-83,000, by sedimentation equilibrium (e.g., 12,15,36), 67,700 by sedimentation velocity-diffusion constant (36), 75,000-80,000 by small-angle X-ray scattering (27, 34), and 65,000-76,000 by dodecyl sulfate-gel eletrophoresis (12,26, 36-39). The median value from the above survey is 72,000, which is about 20% greater than the chemical molar mass of S1, i.e., 61,159, determined from the amino-acid sequence (33).We have found that the central region of S1 gives an anomalously high M,in dodecyl sulfate gels (32),but this would not explain the high M , values by the other physical methods. However, the experiments on the M, of S1 have clearly established S1 as a monomeric protein that shows no tendency to form dimers or higher aggregates even at high concentration (15, 36).

2. RADIUSOF GYRATION This parameter, which provides information on the shape characteristics of a particle, is obtained accurately from the small-angle X-ray scattering curve. It is by definition the root-mean-square distance of all the atoms in the particle from the particle’s center of gravity. The RG of S1 has been determined from the innermost linear portion of the Guinier plot of the scattering curve by three groups, yielding values of 58 & 72 A, and 80 A (15,27,34).It is not clear why there is such a large spread in these values. The 72 value was obtained using S1 prepared without urea (34),and the other two values came from S1 prepared in the presence of urea. One of the groups has also determined the Rc from the total scattering curve using the distance distribution function and derived 80 %, (27)..A calculation of Rc from the hydrodynamic parameters of S1 yielded the value 65-70 A (36). Thus an average value of 70 A would appear to be close to the Rc of

s1.

For a protein of its M , , S1 has an unusually large Rc. For comparison, the Rc of oxyhemoglobin (M, = 64,500) is 26 (40).The relative high Rc value of S1 reflects a very asymmetric shape, discussed later in this section. In a recent experiment, the Rc of S1 in situ on the 30 S subunit was determined by a neutron-scattering procedure (41).It was necessary

A

RIBOSOMAL PROTEIN

s1

107 TABLE I PHYSICAL CONSTANTS OF S1

Parameter

M,(amino acid sequence) M , (physical measurements) Radius of gyration, RG Sedimentation coefficient, S ) ] O , ~ Diffusion constant, D20.w Partial specific volume (measured), 6 Intrinsic viscosity, [q] Hydration (H2O per gram of protein) Isoelectric point, pl Frictional ratio, j7fo Stokes radius Length a

Value

References

61,159 -72,000 -70 .& 3.2 S 4.5 x crnzisec 0.74 ml/gd 9.8 ml/g 0.3 g 4.7 1.7 -45 8, -230 .&

33,44 See text 15, 27,34, 36 12, 15, 36 36 36 36 27 12 12,36 12,36 12, 15, 27, 36

Partial specific volume calculated (193)from the amino-acid composition in Table

I1 is also 0.74 ml/g.

for these measurements to have the 30 S subunits fixed by glutaraldehyde treatment (42),and therefore the measurements were not on the native form of the protein. With this proviso, the value of 60-65 A is so close to both the average RG given above and the Rc determined in the same laboratory earlier (58 %I) that we may assume that no gross changes in the shape of S1 occur when the protein binds to the ribosome.

3. OTHERPHYSICAL PARAMETERS Several other physical constants of S1 have been determined, often in more than one laboratory. These include sedimentation coefficient, diffusion constant, partial specific volume, intrinsic viscosity, degree of hydration, and Stokes radius. Table I lists the values for these constants, as well as the values derived therefrom for the frictional coefficient and the maximum length of S1.

D. Probable Shape The actual shape of a protein may be known definitively only when the protein is crystallized and the X-ray diffraction pattern of the crystal is analyzed and solved. (In preliminary experiments, preparations of S1 have not shown a readiness to crystallize). However, the physical measurements on S 1 have given some definitive indications of its shape in solution.

108

ALAP-RAMAN SUBRAMANIAN

FIG.2. Distance-distribution function p ( r ) of S1 from its small-angle X-ray scattering data. The function gives the probability of a given distance r within the S1 structure. Reproduced, with permission, from European Journal of Biochemistry (27).

Without going into details, all the physical constants of S1 are consistent with a highly asymmetric shape for this protein. If it were a prolate ellipsoid, the hydrodynamic data would have suggested an axial ratio of about 1 : 10 (12, 15, 36). The maximum length of S1 calculated from the data of different authors are in the range 210-280 hi (12,15,27, 3 4 , 3 6 ) with an average value of 230 A. The maximum length of a particle can also be derived from the distance-distribution function of its small-angle X-ray scattering curve. In the case of S1 (27),this analysis yields a value of280 A (Fig. 2). The maximum dimension of the E . coli ribosome has been determined by many investi ators by different procedures. These values range from 190 to 250 (43). Thus the length of S1 is of the same magnitude as, or slightly larger than, the maximum size of the ribosomal particle. This conclusion and the inference of a hinge region between ribosome-binding and mRNA-binding domains of S 1 will have a strong bearing on the proposed specific function of S1 (Section VII). The small-angle-scattering curve of a particle at the region of the higher angles carries information on the actual shape of the particle, although the data are resolvable only for simple objects. The scattering curve of S1 is not that of a simple triaxial body (27, 34). More complex shapes, e.g., two V-shaped triaxial bodies (34),or a long cylinder with an attached short cylinder at one end (H. Labischinski, personal communication) have been proposed, but the experimental data cannot at present be solved unambiguously.

x

RIBOSOMAL PROTEIN

s1

109

II. Structure A. Amino-Acid Sequence

The primary structure of S1 was determined both b y amino-acid sequencing of the protein and b y nucleotide sequencing of the S1 gene ( 3 3 , 4 4 , 4 5 ) and is shown in Fig. 3. S1 from E . coli strain MRE 600 is a single chain of 557 amino-acid residues, with an M,of 61,159. The most striking feature of the primary structure of S1 is the presence of four homologous internal repeats in the central and C-

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110

ALAP-RAMAN SUBRAMANIAN

terminal regions of the protein, discussed in the next section. Some of the other features of the structure are the following. As in the case of other r-proteins, a third of the residues of S1 are of basic and acidic amino acids, and these are distributed throughout the chain, often in small clusters of two to six residues. The next largest group is the aliphatic hydrophobic amino acids, valine, leucine, and isoleucine, which account for 26% of the total residues and are also distributed throughout the chain (except in positions 347 to 366 and 535 to 557), again often in small clusters of two to four residues. Alanine shows a heavier distribution in the C-terminal third of the chain while being sparse in the central region. Tryptophan, tyrosine, histidine, and methionine are, on the other hand, preferentially distributed in the central third of the chain; this region also contains the two -SH groups (Cys-292 and Cys-349) of S1. The amino-acid composition of S1 (Table 11) shows some distinctive features when compared to the total E . coli r-protein composition (33,46).S1 has a lower content of histidine, methionine, proline, and tyrosine and a higher content of valine than the average r-protein. It is TABLE I1 AMINO-ACID COMPOSITION OF S 1 FROM ITS PRIMARY STRUCTURE^ Amino acid (symbol)

Number of residues per mole

Mole-percent of residues

Alanine (A) Arginine (R) Asparagine (N) Aspartic acid (D) Cysteine (C) Glutamine ( Q ) Glutamic acid (E) Glycine (G) Histidine (H) Isoleucine (I) Leucine (L) Lysine (K) Methionine (M) Phenylalanine (F) Proline (P) Serine (S) Threonine (T) Tryptophan (W) Tyrosine (Y) Valine (V)

48 30 23 43

8.6 5.4 4.1 7.7 0.4 2.5 10.8 8.6 1.4 5.4 8.1 7.7 1.1 3.1 1.8 4.5 4.5 1.3 1.1 12.0

2 14 60 48 8 30 45 43 6 17 10 25 25 7 6 67

Total number of residues, 557; M , = 61,159; modified amino acids, none.

RIBOSOMAL PROTEIN

s1

111

richer in the acidic amino acids and poorer in the basic amino acids (in particular arginine) as would be expected for an acidic protein. The sum of the percentage of acidic and basic amino acids in S1 (32.8%)is almost exactly the same as in the total 30 S proteins (32.6%). With the amino acid sequence of S 1, the primary structure data on all E . coli r-proteins are complete (1,47). The small ribosomal subunit contains 21 proteins, of which the smallest is S21 (70 residues) and the second largest is S2 (240 residues). The total number of amino-acid residues used in the construction of this subunit of E . coli MRE 600 ribosome is 3,084, i.e., an average of 147 residues per protein chain. The large subunit contains 34 protein molecules, and these have a total of 4,232 amino-acid residues, i.e., an average of 124 residues per protein chain. Evidently S1, with its 557 residues, is an unusually large r-protein. This large size presumably reflects Sl’s function of binding and holding mRNA during translation (Section VII) and its evolution by fusion of multiple functional domains (Section 111).

B. Four Repeating Homologous Stretches Unique among ribosomal proteins, the primary structure of S1 contains four repeating homologous stretches in the central and C-terminal region of the molecule. The lengths of these repeats are quite constant, ranging from 73 to 75 residues. I have designated them R1, R2, R3, and R4. Their positions in the sequence are R1 = 195-267, R2 = 280-354, R3 = 367-441, and R4 = 454-527. If stretches R1 and R2 are aligned, one under the other (Fig. 4) 55% of the amino-acid residues in the corresponding positions are identical and, in addition, 16% of the residues in the corresponding positions are conservative replacements, i.e., substitution of one amino acid by another of very similar physicochemical properties. Such a degree of homology is well beyond a random occurrence (48).It is of the same degree as the homology found among the phylogenetically conserved L12 proteins (436). Alignment of R1 and R3 show 40% identical amino acid residues and 13%conservative replacements. The stretch R4 from near the Cterminus contains the lowest homology. When aligned with R1, it shows only 33% identical amino-acid residues and 19% conservative replacements. This last homology is still well beyond a random occurrence (48). The degree of homology between the stretches R1, R2, R3, and R4 are such that they could most likely have been evolved by repeated duplication and fusion of a single short gene. This inference is supported by the DNA sequence of the S1 gene (44,45) which shows a

112

ALAP-RAMAN SUBRAMANIAN

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Lys ArgtValLL s His-Pro-Ser Glu+Ile-Val-Asn-Val-Gly-Asp 1 2 0 '

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Ser-Leu-Gly-Leu-Lys-Gln Leu-Gly-Glu-Asp Pro-Trp 350

Ser-Leu-Gly-Leu-Lys-Gln Cys-Lys-Ala-Asn Pro-Trp

FIG.4.Two of the repeating sequences in the primary structure of S1. Identical and conservatively replaced amino-acid residues are boxed, respectively, by solid and dashed lines. There are two more stretches of homologous sequences at positions 367441 and 454-527 (see Section 11,B).

high degree of nucleotide homology in these regions (see Section IV). The N-terminal region of S1 from positions 1 to 194 does not contain any, even short, homologous segments. As described in Section 111, the N-terminal region of S1 contains its ribosome-binding domain, whereas the central and C-terminal regions, which carry the homologous stretches, contain the RNA-binding domain of S1. What is the significance of the four homologous stretches in the S 1 molecule? Many nucleic-acid-binding proteins (mainly DNA-binding) have been isolated from normal E . coli cells and from cells infected by coliphages. These proteins cover a wide range in M , , but the simplest among them are proteins with M , under 10,000, made of 7090 amino-acid residues. Often they are dimers or tetramers in their native state, and this state of aggregation is important for their physiological function. Some examples are the gene-5 protein of bacteriophage fd (49) and NS1 and NS2 ofE. coli (50,51).The primary physiological role of the gene-5 protein is considered (52) to be to stabilize and protect the single-strand daughter virion DNA. If the RNA-binding domain of S1 were evolved in such a manner as to mimic such

RIBOSOMAL PROTEIN

s1

113

DNA-binding proteins but without recourse to self-aggregation, we should expect to find either two or four homologous stretches in its nucleic-acid-binding domain. The present finding that there are four homologous stretches in S1 [rather than the three we observed previously (44)] is therefore significant. This point is further elaborated in Section VII.

C. Secondary Structure Elements The secondary structure of a protein is definitively determined when the protein's three-dimensional structure is solved at high resolution. Barring that, there are optical and computational methods to estimate the secondary structure elements in the protein, and these methods have been applied for the determination of the secondary structure of S1(12,15,33,53).These data show that between 30%and 40% of the amino-acid residues of S1 are involved in forming the ahelix, and a similar number in forming the @-sheet. The computer method has, in addition, predicted that 20-30% of Sl's residues are involved in the @-turn, i.e., the 180" turn between the formed elements of a-helix and @-sheet, which permits the creation of compact domains in a protein. The mix of a-helix and @-sheetin S 1 is like that predicted for most other ribosomal proteins (47),but the distribution of a-helices and /Isheets in the different regions of the S1 chain does not appear to be at random (Fig. 5 ) . The N-terminal, nonhomologous region (positions 1194),with its ribosome binding domain, contains slightly over half the a-helical structures in S1, but only about a sixth of the total @-sheets. Thus the predicted @-sheet structures are concentrated in the four homologous regions that carry the nucleic-acid-binding domain. Here again, S1 shows a resemblance to gene 5 protein whose structure, determined at 2.3 A resolution, is composed entirely of antiparallel @sheets (49). Figure 5 shows also that the four homologous regions of S1 do not all have the same predicted secondary structure: while R2 is made of only @-sheet,RI, R3, and R 4 contain both a-helix and @-sheet.

111. Discernible Structural and Functional Domains With its large size and dual roles in protein biosynthesis and Q@ RNA replication, S 1 has invited studies probing the existence of distinct structural and/or functional domains within the molecule. These studies have employed physical and biochemical procedures: e.g., analysis of the small-angle scattering curve for evidence of distinct scattering domains, fluorescence polarization analysis of S 1 labeled

114

I

ALAP-RAMAN SUBRAMANIAN

Trypsin I

80

120

40

500

5 51

-R4FIG.5. Secondary structure of S 1 predicted by a computer program with Chou and Fasman parameters. Loops and zigzags represent a-helix and p-sheet, respectively. The position of the facile trypsin-sensitive site (arrow) and of the four homologous, repeating stretches are indicated. Adapted, with permission, from European Journal of Biochemistry (33).

with fluorescent groups, or isolation of discrete fragments of S1 and examination of functions retained by them. These studies, summarized in this section, show that S1 is organized into at least two distinct domains.

A. Mutant Form MlSl While isolating E . coli mutants resistant to antibiotics, S. Mizushima obtained one that did not show the usual S1 spot in its twodimensional gel pattern of ribosomes. Further experiments revealed the presence of an anti-S1 cross-reacting protein in the mutant ribosome (54). Using the cross-reaction for monitoring protein purification, a pure protein was isolated (Fig. 6) that was able to restore to S1lacking ribosomes about 75% of the activity for translating synthetic or natural mRNA (54) and to restore full functional activity to Sl-lacking QP replicase (55). Detailed structural work has shown that the mutant S1 (MlS1) is a truncated form lacking the C-terminal 120 amino-acid residues of S 1 (unpubIished results of K. Foulaki, B. Wittmann-Liebold, and the au-

RIBOSOMAL PROTEIN

s1

115

FIG.6. Immunological homology of purified S 1 and mutant S1. (A) The proteins electrophoresed in dodecylsulfate gel and stained with Coomassie blue. (B) Double immunodiffusion with anti41 in the center well, S1 and mutant S1 in peripheral (alternate) wells: spurless precipitin lines are formed. Reproduced, with permission, from Journal of Biological Chemistry (54).

thor, quoted in references 33 and 44). Thus M l S l (calc. M, = 48,404) contains 437 amino-acid residues. Analysis of its small-angle X-ray scattering shows M l S l to be a slightly less elongated molecule than S1, i.e., of radius of gyration 60 A and of maximum length 205 8, (see Table I for data on S 1). The shape characteristics of M 1 s 1’s scattering curve is, however, very similar to that of S1 (unpublished results of H. Labischinski and the author). Since the M l S l molecule ends at position 437, it evidently lacks the repeating homologous segment R4 (see Fig. 4).Thus the loss of this segment apparently does not affect the QP-replicase activity of the molecule (55), but it results in reduced effectiveness (75%) in protein synthesis function (54). In addition, whereas a second molecule of S1 can bind to “inactive” 30 S subunits (see Section VI), such was not observed in the case of M l S l (56). Another consequence of the loss of the C-terminal region is in the increased solvent exposure of Cys-292; for, while only one SH group (Cys-349) is reactive in S1, both S H groups are reactive in M l S l (32).

116

ALAP-RAMAN SUBRAMANIAN

6. Stable Fragment by limited Proteolysis When S 1is treated at 0°C with a small amount of trypsin (1-2 pg of enzyme per milligram of Sl), a large fragment is produced in nearquantitative yield (Fig, 7). The fragment (SlF1)has been isolated and characterized (31), and the sequencing of S1 has provided its exact location (33). The cleavage site is at Arg(170)-Arg(171)-Ala(172). About 80% of the fragment is due to cleavage at the Arg-Ala bond, and the remainder is due to cleavage at the Arg-Arg bond. (This was inferred from the pattern of appearance of a minor amino-acid residue after each Edman degration step.) The major component of S l F l therefore contains the sequence from positions 172 to 557, i.e., 386 amino acid residues and M,41,330. S l F l is totally inactive in restoring function to S1-lacking ribosomes (31)and to S1-lacking QP replicase (55). In our early experiments, a binding between S l F l and 30s subunits was indicated (31),but later experiments, with radioactively labeled SlF1, showed that such binding is very weak (57), about 5% of that due to intact S1 (unpublished results of Z. Ali and the author). S l F l is also unable to bind to Sl-lacking QP replicase (55).Thus the loss of the 170 N-terminal residues from S1 results in the inability to bind to ribosomes or to Qp replicase, and, in consequence, a complete loss of the biological activity. S 1F1, however, retains a characteristic property of S 1, namely, the ability to bind to nucleic acids (31,56).It binds to poly (U)-Sepharose so strongly that it is not released by 1M salt, but only by 6 M urea (31). Thus, the region of S1 contained in S l F l should be expected to have the nucleic-acid-binding domain of S 1. From its primary structure, S 1F1 contains 50 trypsin-sensitive peptide bonds, yet it is very resistant to further cleavage, indicating a very compact folded structure. As noted in Section 11, this region of S1 is rich in P-sheet structure and contains all four repeating homologous segments (Fig. 8). The small-angle X-ray scattering curve of S l F l indicates it to be an elongated molecule of maximum length 210 A and radius of gyration 61 A. However, unlike that of S 1 and MlS1, the scattering curve of S l F l can be interpreted as due to a simple triaxial body, e.g., a cylinder of 200 length (unpublished results of H. Labischinski and the author). The characteristic cleavage of S1 by trypsin occurs not only in free S1, but also in S1 bound to 30 S subunits. It is inhibited when either S1 or the 30 S subunit is treated with poly(U) [observations of the author and Rummel and Noller (58)l.The latter result shows that the conformation of S1 near its facile trypsin-sensitive bonds is modified when the RNA binding site of S1 is occupied.

RIBOSOMAL PROTEIN

s1

117

FIG. 7. Facile trypsin cleavage at a specific region in S1. Slot 1 shows S 1 before treatment, and slots 2-10 show it after cleavage with trypsin at 0°C ( 1 , 2 , 5 ,10,20,30,40, 60, or 90 min of reaction). The stable fragment (SIF1) contains the nucleic acid binding domain of S1. Reproduced, with permission, from Journal of Molecular Biology ( 3 1 ) .

C. Ribosome-Binding Fragment Yet another method used to produce functionally active fragments of S1 is chemical cleavage at the methionine residues in the S l chain. By this means, three large and three small fragments are produced, and two of them contain functional domains of S1 (32, 57).

118

A L A P - W A N SUBRAMANIAN 7

b

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SH-2

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7

1

S1 F1 SH-1

SH-1

SH-2

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FIG.8. Linear representation of S1 and its functional domains. The four repeating, homologous stretches are shown stippled, and their beginning and ending positions are given. Fragment F2a contains the ribosome binding domain of S1, while fragment S l F l contains the nucleic-acid-binding domain. The small fragment F3 can bind to nucleic acids strongly whereas F2b does not. The mutant protein (MlS1) is 75% as active as S1 in protein synthesis and fully active in QP RNA replication. SH-2 is the “reactive” -SH group of S1, accessible both in free protein and in S 1 bound to the 30 S subunit. It becomes unreactive (like SH-1) in the S 1 .RNA complex (see Section 111).

The most interesting of this group of fragments is the one (F2a) from the N-terminal region of S l that contains the sequence from positions 1to 193 (calculated M , = 21,304.) Fragment F2a binds to S1lacking 30 S subunits, and this binding shows about the same affinity as binding by S1. In assays where either F2a or S1 is labeled, unlabeled S 1competes out the binding by F2a and, conversely, unlabeled F2a competes out the binding by S1 (57).Thus S1 and F2a seek the same site on the 30 S subunit for their binding. From the sequence position of F2a (Fig. 8), it is evident that it contains the region of S1 degraded by trypsin and therefore missing from SlF1. By isolating this region by a chemical procedure and demonstrating its specific binding to 30 S subunits, we were able to show that S1 binds to the ribosome by means of a specific sequence of its structure in the N-terminal third of the molecule. D. Nucleic-Acid-Binding Fragment from the Central Region

The second functional fragment (F3)produced by methionyl bond cleavage in S1 has the sequence from position 224 to 309; i.e., it contains 86 amino-acid residues ( M , = 9725). F3 was found to bind to poly(U) quite well by Millipore binding assay (32). F3 also binds to poly(U)-Sepharose, and about 50% of the bound protein is not re-

RIBOSOMAL PROTEIN

s1

119

leased by 1 M salt but only by 6 M urea (unpublished observation of the author). The binding between poly(U) and F3 is abolished b y low concentrations of aurintricarboxylate, which also inhibits the binding between poly(U) and S1 (32).Thus, this fragment of S1 from the central region of the molecule must have the core of Sl’s nucleic-acidbinding domain. Neither the ribosome-binding fragment F2a nor the C-terminal fragment F2b (which contains two of the homologous domains) have any noticeable ability to bind to poly(U) (32). A peculiar property of F3 is its relatively slow electrophoretic migration in dodecyl-sulfate gels. Thus, it gives the high apparent M,of 16,000 when analyzed by this procedure (32).This central region of S1 is therefore responsible for the high apparent M , of S 1 when the latter is determined by gel electrophoresis (see Section I).

E. The -SH Groups There are two -SH groups in S1, of which only one is reactive in the native protein, while neither is reactive in the S1 . RNA complex. The position of the reactive -SH group was determined by an analysis of the fragments produced when S l , MlS1, and S l F l are treated with a cysteine-specific cleavage reagent (59). The result was later confirmed by autoradiographic analysis of the CNBr fragments from N-ethyl[ 14C]maleimide-labeledS 1 (32). The reactive -SH group of S1 (Cys-349) is exposed in the free protein (23, 59-61) as well as in 30 S subunits (62-66), but becomes inaccessible to reaction when either is treated with poly(U) or other poly- or oligonucleotides (64,67). The two -SH groups are both reactive in free MlS1, as noted earlier (32). The mono N-ethylmaleimide derivative of S1 (MalN-S1) is able to bind to nucleic acids, apparently as efficiently as S1, but is not able to unwind them (60,68,69). MalN-S1 binds to ribosomes with the same binding constant as S l (10) as expected from the remoteness of the -SH group to the N-terminal region. The 30 S subunits containing MalN-S1 have been reported to be inactive in binding Met-tRNA when programmed by MS2 RNA (60, 70). MalN-S1 is, however, fully active in QP RNA replication (71) and in poly(U) translation (72)and about 50% as active as S1 in MS2 RNA translation (unpublished experiments of T. Suryanarayana and A. R. Subramanian). The significance of the last result will be discussed in Section VII.

F.

Evidence for an lnterdomain Hinge Region

The small-angle X-ray scattering data of S 1, plotted by the Guinier procedure for cross-section factor, show two linear, regions, which

120

ALAP-RAMAN SUBRAMANIAN

indicate two distinct scattering domains in the molecule (27). In fluorescent polarization measurements, S 1 with an attached fluorescent group has a rotational correlation time of 26 nsec (70). For a rigid molecule the size of S1, a much larger correlation time, i.e., 60 nsec, is expected; therefore, a construction of S1 into two domains, each of which can rotate freely against the other, was proposed (70). The proton NMR spectrum of S1 has also been interpreted as suggesting a greater degree of structural flexibility in S1 than is normal in a globular protein the size of S1 (35). The two structural domains of S1 indicated by the physical measurements can be correlated with the functional domains of S1 from the biochemical studies described earlier. One of the two domains is composed of the N-terminal region extending from position 1 to 170 and contains the ribosome-binding site of S1. This region must have a loosely ordered structure, since it is easily fragmented by the protease trypsin. The second domain extends from position 171 to the C terminus of S1 and contains the nucleic acid binding site. This region is extremely resistant to the action of trypsin and therefore must have a tight organization. It contains the two SH groups and the four repeating homologous segments of S1. There are two peptide bonds between the two domains that are subject to very facile cleavage by trypsin. The interdomain region must therefore contain a short, highly exposed length of polypeptide chain permitting free movement of the domains relative to each other. As discussed later (Section VII) a flexible region between its two domains is an important aspect of the specific functional role of S1.

IV. The S1 Gene

The chance discovery of a mutant form of S1 (the mutation later proved to be unconnected with any antibiotic resistance) permitted the mapping of the S1 gene on E . coEi chromosome (73).Like other rproteins, S l is carried by E . coli on a single gene, which is located at 20.5 min on its chromosomal map. This segment of E. coli DNA has been transduced into A phage, and the S1 gene therein has been subcloned into plasmids (74, 75). The nucleotide sequence of the S1 gene and its flanking regions have been determined (44,45). The S1 gene contains 1668 base pairs in its amino-acid coding region, corresponding to 556 amino-acid residues, i.e., one residue less than that obtained by the protein-chemical procedure. There are also two other differences between the deduced and the protein-chemical1 y determined sequences (44).For practical

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reasons, the protein and the DNA were isolated from two different and independent strains E . coli. The minor differences in the sequences are therefore thought to arise from strain-specific differences between the MRE 600 and JS6.5 strains used in these studies. The promoter of the S1 gene has tentatively been located in the upstream” sequence, which, interestingly enough, contains an open reading frame that may code for a 116-residue protein (45).There are two ribosome recognition sequences neighboring each other in the upstream region of the S1 gene, but both are farther away from the initiating AUG codon than is generally observed in other genes (76). The codon usage in the S1 gene follows very closely the pattern observed in the case of all highly expressed genes of E . coli (77, 78). Among the individual codons, highly preferred ones are: ACC (Asn), 100%; ATC (Ile), 97%; CAG (Gln), 93%; and CCG (Pro), 90%; while the codon pairs CGY, GGY, ATY, and CCY (Y = a pyrimidine nucleoside) were 100% used for the amino acids arginine, glycine, isoleucine, and proline, respectively. The most interesting feature of the sequence of the S1 gene is undoubtedly the presence therein of several segments of extensive nucleotide homology (44,45).All these segments lie in the central and C-terminal regions of S1 (see Section 111). Such homology has not been observed in other r-protein genes or in the sequences of the rRNA molecules (45). What regulates the expression of the S 1 gene in the bacterial cell? Experiments with E . coli cells that carry multicopy plasmids containing the S1 gene show the gene expression to be mostly regulated at the translational level: these cells contain a huge amount of S1 mRNA but only a relatively modest excess of S 1 protein (75). Free S 1 added to in vitro systems expressing the S1 gene show a relative repression (79).However, since S1 binds to RNA independently of its nucleotide sequence (see Section VI), a mechanism of autogenous translational control (80)of the S1 gene would be rather provocative. Some experiments indicate that the regulation of S1 synthesis is achieved in a manner different from that of other r-protein genes (79,81). An unusually short half-life (40 sec) for S1 mRNA in the cell has also been reported (82). “

V. location of S1 within the Ribosome Several methods have been developed to gain information on the relative positions of individual ribosomal components in situ and thus to obtain the molecular topography of the ribosomal particle. These

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methods include: ( a ) cross-linking of neighboring components by means of chemical cross-linking agents or by photoactivation; ( b )immunoelectron microscopy, i.e., visualization of a component on the image of the ribosome by means of its specific antibody; and (c) the neutron scattering procedure using ribosomes containing deuterated components. All these procedures have been applied in the case of S1, but the results have not been fully satisfactory. Cross-linking of 30 S subunits with a bifunctional reagent has yielded dimeric products that contain S 1 covalently linked to three other ribosomal proteins, namely, S2, S10, and S18; no other Sl-containing cross-links were detected (83). The reagent used has a span length of 14.5 when fully extended (84); therefore, these results mean that part of S2, S 10, and S 18 must lie within this distance from a cross-linking group (Lys or Cys residue) in S1. It is interesting that a molecule 230 long with 45 such cross-linkable groups formed only three detectable cross-linked products: it supports the view that only a small region of the S1 molecule is in actual contact with the ribosome (see Section VII). In earlier experiments, the same research group showed that cross-linking of initiating 70 S ribosomes yields products containing S1 and the initiation factors IF-1, IF-2, and IF-3, but did not determine whether the linkage was direct or bridged by another protein, e.g., S2 (84).No cross-link between S1 and IF-3 was reported in an experiment where the latter was photochemically cross-linked in the 30 S subunit by UV irradiation, i.e., when the linkable distance was close to zero (85). Attempts to cross-link S1 and 16 S rRNA have also yielded results. The 3' end of 16 S rRNA was modified to make it capable of crosslinking to protein, and the cross-linked RNA was then analyzed for covalently bound protein; S1 (and S21) was detected in this experiment (86). However, there are potential artifacts in this procedure (2, 87). In a more recent experiment, a bifunctional reagent was used to achieve the S1-16 S rRNA cross-linking in 30 S subunits (88). The cross-linked 16 S rRNA contained S1, and the position of the linkage on the RNA chain was deduced to be at the region 861-889 (16 S rRNA contains 1541 nucleotides). This result indicates a location of S1 near the central region of 16 S rRNA rather than at its 3' end (see Section VI). Location of S1 on the ribosome by immunoelectron microscopy has presented problems because, having a weak affinity constant, S1 is removed from the ribosome by the binding of the antibody molecule. Nevertheless, two of the research groups specializing in this area have assigned positions to S1. According to one group, S1 is located at the

A

A

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distal portion of a platform-like structure on the 30 S subunit, not far removed from its “head” (89).According to the other group, two locations of S1 could be discerned, one on the headlike structure and the other on the right shoulder of the particle (90). Evidently, these two assignments are not in perfect agreement, but we must note that there is agreement on the general location. Using neutron scattering, an approximate position for the center of gravity of S1 has been assigned and this location is in the general area suggested by the immunoelectron microscopy (41). Neutron scattering data have also provided distances between the centers of gravity of S1 and four other ribosomal proteins (S3, S4, S5, and S7), and all four distances are in the range 50-65 A. I have earlier noted (Section I) that neutron scattering measurements indicate strongly that the shape of S1 in the 30 S subunit may not be different from its shape when free in the isolated state.

VI. Interactions of S1 There are three interactions of importance in understanding the function and evolution of S1. These are Sl’s interactions with the ribosome and QP replicase and its interaction with external nucleic acids. The last one has apparently determined the evolution of S1 as a natural component of the two macromolecular assemblies. QP replicase contains no nucleic acid in its structure; therefore, the association of S1 therein is evidently due to protein-protein interaction. In the case of the ribosome, however, the association could be through protein-protein interaction, protein-RNA interaction, or a combination of both. In this section, I evaluate the experimental results relating to these questions. A. Interactions with Nucleic Acid

Many aspects of the interactions of S1 with nucleic acids have been covered by two articles in this series dealing with ligand-induced conformational changes in RNA (91) and helix-destablizing proteins (69). I shall therefore restrict myself to only those essential features of S1-nucleic acid interactions that are pertinent to the role of S1 in protein biosynthesis (Section VII). The ability of S1 to bind to polynucleotides was one of the earliest of its properties noticed by investigators in the field (92, 93). For the quantitative analysis of this interaction, a truly wide range of physical techniques have been used, including: ( a ) Millipore filter binding,

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useful for both qualitative and quantitative analysis of the binding by oligo- and polynucleotides (94,95); ( b )equilibrium dialysis, useful for quantitative anaIysis of oligonucleotide binding (67); (c) assay by the loss of reactivity of Cys-349, which occurs when S1 binds to poly(oligo-)nucleotides (67); ( d )analysis of the fluorescence quenching of tryptophan residues when S1 binds to poly-(oligo-)nucleotides (96); ( e ) high-resolution electron-microscopic imaging of S1 and S1-nucleic-acid complexes (68); and (f) temperature-jump relaxation procedures (97,98). In addition, other properties of S1, such as its ability to lower the melting temperature (99), alter the circular dichroic spectrum (99, loo), or change the fluorescence properties of (free or ethidium bromide-bound) polynucleotides (60, IOO), have also been used. Still another means of detecting S1.RNA binding is to pass S1 through a short column of poly(U)- Sepharose as described in Section I.

1. S1 AND THE STRONGRNA BINDINGSITE S1 can bind single-stranded DNA and RNA, but it exerts its strong, characteristic binding only toward RNA. Thus this binding site of S I must distinguish between the ribose and deoxyribose residue of a polynucleotide chain. Above 0.1 M salt, the binding between S1 and RNA is insensitive (67) or slightly enhanced (101) by further addition of salt. Therefore, a strong hydrophobic component exists in the S1.RNA interaction. The size of the RNA binding site of S1 was determined by means of the experiments with oligonucleotides of increasing chain length. The binding constant increased with increasing chain length and then became constant; by plotting the data, the saturation affinity was found at r] = 13.6 (67). Using the quenching of tryptophan fluorescence, a single-site size of 10 residues was inferred (101). More than one oligonucleotide molecule of smaller size can bind to an S1; and at greater chain length, more than one S1 can bind to a single polynucleotide chain (67). The latter phenomenon was also observed in electron miscrocopy studies on S1 and nucleic acids (68)where, at saturation, a binding of one S l per 10-15 nucleotides was observed. Thus a site size of 10-12 nucleotides would appear to be reasonable estiyate. This length of oligonucleotide, if stretched, would be about 40 A. Values reported for the binding constant of S 1 to oligo- or polynucleotides of saturating chain length show a large divergence, i.e., from 4 X lo5 M-' to 3 X lo7 M-' (67, 95, 97, 98, 101).Some divergence should be expected because different groups have used different procedures (which have different degrees of uncertainty), and measure-

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ments were made at different ionic conditions and temperatures. A median value of about 3 x lo6 M-’ would appear to be a fair estimate of the binding constant. Does S 1 show preferences among the homopolynucleotides used in these studies? Whereas S 1 bound to poly(A)-cellulose is released by 1 M salt, unfolding by urea is necessary for its release from poly(C)or poly( U)-cellulose. Thus a stronger binding to polypyrimidine is observed. A threefold greater binding constant for S l * poly(C) over Sl-poly(A) (101) and a 10-fold greater binding constant for S1.oligo(U) over Sl*olgio(A)(67) is reported, but the opposite, namely a threefold greater binding for S1. olgio(A) over S 1 * olgio(U) has also been reported (98). Draper and von Hippel (101) have deduced the existence of a strong cooperativity (parameter o,approximately 31) for poly(C) binding and its absence in the case of poly(A) binding. These results therefore indicate that insofar as its basic binding affinity toward RNA is concerned, S1 does not make a strong distinction with respect to nucleotide composition or base sequence, but it exerts a stronger hydrophobic component in its interaction with pyrimidines.

2. DOESS1 HAVEA SECOND POLYNUCLEOTIDE BINDINGSITE? S1 has been shown to bind to the denatured DNAs of phage A, Simian virus 40, and calf thymus, to the single-stranded DNA of 4x174 phage, and to synthetic polydeoxynucleotides (56,68,96,102). This binding is salt-sensitive. From a study of the quenching of tryptophan fluorescence of S 1 by polydeoxynucleotides, a second nucleic acid binding site on S1 was proposed. Some of the properties of this site are: site size, 5 nucleotides; binding constant, 3 x lo6 M-I; cooperativity, nil; binding constant for double-stranded DNA, times that for single-stranded; involvement of charged groups, two basic residues in S1 in charge-charge interaction with DNA phosphate groups; specificity for DNA versus RNA, none (102). A good critique of the two-site model is given in this series (69). At present there is no independent evidence for a second DNA (or RNA) binding site on S1, but I wish to add a speculative thought. The discovery of four homologous stretches of sequences in S1 is intriguing in this connection. The gene 5 protein of phage fd (see Section 11) in its native state is a dimer (87 residues per monomer) with two binding sites for oligodeoxynucleotides (49). If each of S 1’s homologous segments were an oligonucleotide binding site, two such segments would suffice to generate a nucleic-acid-binding protein. However, S 1 actually contains four homologous segments. It is likely that a rationale

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for this structural feature of S1 may resolve the real number of nucleicacid-binding sites in S1 and provide further insight into S1 function.

3. RNA HELIX-DESTABILIZING PROPERTY This topic has been reviewed recently (69) and therefore is not discussed here.

B. Interactions within the Ribosome and

G2/3 Replicase

1. INTERACTIONSWITHIN THE RIBOSOME Although itself a nucleic-acid-binding protein, S 1 plays no apparent role in the assembly of the ribosome (103).This may appear surprising at first sight, but is understandable in terms of two of its RNA binding characteristics, namely, the lack of sequence specificity and the relatively low (3 x lo6 M - l ) binding constant. The thermodynamic constant for the association of S1 with the 30 S subunit has been determined by sedimentation methods (10,11). By applying the equations developed for band sedimentation equilibrium (104),a value of 1.7 X lo8 M-’ was obtained. Thus the affinity of S1 for 30 S subunits is about 50 times stronger than its affinity for “naked” RNA. The association constant is not affected by raising the temperature from 6°C to 18°C or by the presence of 50 S subunits in the reaction mixture (11).Since the ribosome concentration in an E . coEi cell is about 20 X M (calculated from the ribosome content, water content, and cell volume), given the above association constant, there will not be an appreciable concentration of S l-free ribosomes in the cell. Through what forces does S1 bind to the 30 S ribosomal subunit? There are several experiments supporting the thesis that this binding is mediated through protein-protein interaction rather than by protein-RNA interaction. Whereas S1 can bind to 30 S subunits subjected to a mild nucleolytic cleavage, it cannot bind to 30 S subunits subjected to mild proteolysis (105).In other experiments, the binding of S 1 to 30 S subunits lacking defined lengths of 16 S rRNA has been examined. Removal of 49 nucleotides (10) or 160 nucleotides (206) from the 3’ end of the 16 S rRNA had no adverse effect on the binding of S1. The more critical result on this point has come from our work with S1 fragments (Section 111).An N-terminal fragment of S1 containing residues 1-195 (F2a) is able both to bind strongly to 30 S subunits and to compete with S1 for this binding, but has no appreciable nu-

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cleic-acid-binding ability. On the other hand, a fragment containing the central and C-terminal regions of S1 (SlF1) is able to bind RNA very strongly but has only a weak affinity for the ribosome. Thus, S1 apparently binds to 30 S subunits essentially through the proteinprotein interactions of its N-terminal domain. Which are the r-proteins involved in the binding of S1 to the 30 S subunit? By partial reconstitution experiments, S9 has been found to be very important in this binding (107).Whether other r-proteins are also involved in this binding, as well as the question of whether free S9 and S1 will interact with each other in the absence of the ribosomal structure, have not been examined. Two of the cross-linkable neighbors of S1 (S2, S10) showed only a small stimulatory effect on the binding between 30 S subunits and S1 (107). An analytically useful effect of the attachment of S1 to 30 S subunits is the distinct decrease in the particle’s electrophoretic mobility. This allows for a quick detection of the binding between S1 and S1lacking 30 S subunits (108, 109).

2. S 1 AND INTERACTIONS WITH rRNA S1 binds to isolated, free 16 S and 23 S rRNA (110) and to tRNA ( 1 1 1 ) just as it binds to any synthetic polynucleotide. In the case of 23 S rRNA, the strong binding was in a pyrimidine-rich region. However, S1 is never found associated with the 50 S subunit, and the dissociation of the ribosome into two subunits (either by IF-3 or by lowering Mg2+)is indifferent to the presence or the absence of S 1 (26, 112, and my observation). S 1 also binds to the 49 nucleotide long, 3’end-fragment of 16 S rRNA (113).The binding site of S1 on this fragment was in a dodecamer subfragment (113)that contained the phylogenetically conserved, prokaryotic ribosome-mRNA recognition sequence. This observed possibility of the participation of S1 in the ribosome-mRNA recognition step of protein synthesis initiation has intrigued many investigators (see Section VII). However, the binding constant of S1 with the 49-mer is 5 x lo6 M-’ (97),the same magnitude as S 1’s interaction with any polynucleotide, indicating the absence of a special affinity. As noted (Section V), the reliability of the observed S1-16 S rRNA 3’-end cross-link has been questioned. More important, the presence or absence of S 1or the addition of S1 antibodies showed no effect on the binding of a synthetic octanucleotide with complementary base sequence to the 3’ end of 16 S rRNA, while another r-protein (S21) showed a strong effect (114, 115). Thus we must conclude that although an interaction between S 1and the 3’-end of 16 S rRNA is feasible, it remains to be proved.

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3. S1 AND “INACTIVE” 30 S SUBUNITS When 30 S subunits are exposed to low concentrations of Mg2+ and/or to NaC1, they are converted into an inactive form; this can, however, be readily reactivated by raising [Mg2+],replacing NaCl by KCl or NH4C1, and warming to 40°C (116,117).A second molecule of S1 binds to such inactive 30 S subunits (10).The affinity of S1 for this second site is 20-fold lower than for the first site. More important, the second site is absent in inactive 30 S subunits that lack the 3’-end 49-mer in their 16 S rRNA (10). Thus the second S1 site appears to arise from the 3‘ end of 16 S RNA. The octanucleotide probe has been applied to test the accessibility of the 3’ end in inactive 30 S subunits; the results indicate, however, that whereas the 3‘ end is accessible in active 30 S subunits, it is inaccessible in the inactive form (114). I have noted above (Section 111) that M l S l does not show evidence of binding at this second site. 4. S1 AND Qj3 REPLICASE Isolated from Qj3 phage-infected E . coli, Qj3 replicase contains one molecule each of SI, EF-Tu, EF-Ts, and synthetase; the last one is the only phage-coded polypeptide (39, 118-121). The calculated M,’s of the four components are: S1 = 61,159 (Section I); EF-Tu = 43,225 (122);EF-Ts = 30,257( 123);synthetase = 65,317( 124).Thus, contrary to the generally believed earlier inference (39, 121), S1 is not the largest component of this enzyme. The binding constant of S1 toward Sl-lacking Qj3 replicase is 1 X lo8M-’ (55,73).Since QP replicase does not contain nucleic acid, the binding affinity of S l toward it must be due entirely to protein-protein interaction. The phage-coded polypeptide appears to be the component with which S1 interacts in the replicase, because a stable subcomplex of these two molecules has been isolated (121,125).It is striking that the binding constant of S1 with QP replicase is almost exactly the same as its binding constant with 30 S subunits; perhaps it indicates that the same surface of S1 is involved in both these interactions.

VII. Functions of S1 The determination of the function of an individual r-protein has generally been a vexing problem because ribosomes lacking a single r-protein (made by in vitro reconstitution or by mutation and selection Drocedures) are often functionally active, as if the protein in question

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had no clear function (126-128). S1 is one of the few exceptions to this rule. Ribosomes lacking S1 show only a very low level of activity (see Section VI1,B). An E . coli mutant lacking S1 has not been isolated, although one with a shorter S1 (Section 111) and another with a reduced affinity for the ribosome (E. Dabbs, personal communication), have been obtained. An amber mutation (i.e., one that would cause premature chain termination and release) was induced in the S1 gene, but this mutation was lethal in the absence of an amber suppressor (129).It appears therefore that S1 performs an essential function for E . coli. The question of the occurrence of structural and/or functional analogs of S1 in organisms other than E . coli has not received adequate attention. For example, a comparative examination of prokaryotic, eukaryotic, and organellar ribosomes for the presence or absence of S 1 analogs therein has not been reported. An “Sl-like” protein has been observed in Mycobacterium smegmatis ribosomes ( 130), and we have evidence (unpublished) for an S l-like, poly( U)-binding protein in spinach chloroplast ribosomes. Bacillus stearothermophilus ribosomes or cell extracts do not give immunological cross-reaction with S1-antibody (131);and an S1-like protein has not been detected in Bacillus subtilis ribosomes (132). Evidence for certain unique features in the mRNA of B . subtilis and other gram-positive bacteria that would enable the homologous ribosomes to translate them more efficiently has been reported; the generally observed inefficient translation of gram-negative bacterial mRNA by gram-positive ribosomes is then ascribed to the absence of such features in the latter class of mRNA ( 1 3 3 , 1 3 3 ~ ) . Returning to E . coli, what can be the essential function of S1 in this organism? Because S1 occurs predominantly on the ribosome, its major function is evidently connected with protein biosynthesis. However, because S1 can easily be detached from the ribosome, it can have other functions, either as a component of other macromolecular assemblies, or by itself. The former possibility is realized in QP replicase. Free S1 has been shown to stimulate DNA transcription in uitro, but its physiological significance there remains uncertain.

A. S1 Function in QP Replication and DNA Transcription 1.

QP REPLICATION

The function of S 1 in QP replicase is discussed in detail in review articles (121, 125). Its presence in QP replicase is required for the specific binding of this enzyme onto QP RNA so that transcripts of the

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latter, called minus strands, can be synthesized. The transcription of the minus strands themselves later on to produce the manyfold replication of the original phage RNA apparently does not require S1. The astonishing specificity of QP replicase in its own biological milieu (134)is generated by the subcomplex of S1 and the phage-coded polypeptide (125). We have already seen that M l S l is fully reactive in QP replicase function and its binding constant therein is the same as that of S1 (55). S l F l is inactive in substituting for S1, because it does not, apparently, bind to the S1-lacking complex (55). The mono N-ethylmaleimide derivative of S1 is as active as S1 in QP replicase function (71): the unwinding property of S1 is evidently not important in this function (Section 111).

2. DNA TRANSCRIPTION It has often been noticed that the addition of ribosomes to an

E . coli extract containing the DNA * RNA polymerase complex, or to a purified system containing RNA polymerase and DNA, stimulates RNA synthesis (135, 13$6).The stimulation was later observed with purified ribosomal subunits, and, in the case of 30 S subunits, it was found to be a property of the acidic r-proteins (137,138).Fractionation indicated S 1as the major stimulatory component (137).Other workers have added purified S1 to defined in vitro transcription systems and noticed stimulation of both phage and bacterial DNA transcription

(139, 140).

The effect of S1 on transcription has also been examined using the monovalent antibody fragment Fab. The specific Fabs against 20 small subunit r-proteins were each added to a system transcribing A plac DNA in the presence of 30 S subunits. The antibody fragment against S1 (and also those against a few other r-proteins) showed a complete inhibition of the ribosomal stimulation ( 1 4 1 ) . Since the stimulation of transcription by S1 could simply be due to its RNA binding (and therefore sequestering) property, the above results in themselves do not prove a participation of S1 in transcription. There is also the proposed role of ribosomes in the attenuation of mRNA transcription, but here the ribosome is believed to participate purely passively and, therefore, a role therein for an RNA-binding r-protein is not envisaged (142). B. S1 Function in Protein Synthesis When S 1-free ribosomes are tested for their ability to translate synthetic or natural mRNA in an S1-free system (30,31,143,144),only a very low level of activity is observed. The addition of purified S1

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stimulates this intrinsic activity to the level observed with S l-containing ribosomes (Fig. 9). Similarly, when antibodies (or the monovalent Fab) specific for S1 are added to an Sl-containing system (140, 141, 145,146) there is a virtually total inhibition, which can be reversed by neutralizing the antibody with purified S1 (Fig. 9). Thus S1, like L12

jliA 1 150

-

.U 3

= 50 -

4

yl

E

L d E Z r z 2 26

--

10

14 18 22 mM Mg'*

12

14 16 18 mM Mg+'

21

FIG.9. Translation ofmRNA by S1-lacking ribosomes in an S1-free system with and without added purified S1. The upper panels show the translation of synthetic messengers: panel A, poly(U), and panel B, ploy(A); the lower panels show the translation of phage MS2 RNA, where panel D illustrates the effect of S 1 antibody and its neutralization by added S1. The incubation mixture (for A, 9, and C ) contained 44 pmol of S1-free ribosomes, the indicated (radioactive) amino acids, 2 pg of poly(U) ( M , -lo5) or 2 p g of poly(A) ( M , -lo5) or 20 p g of MS2 RNA ( M , -lo6)),and other ingreidients required for in uitro protein synthesis as given in reference 31. The nominal Mg2+concentration at each point is given (the actual, free Mgi+ concentration will be lower because of chelation by ATP and P-enolpyruvate present in the system). Where indicated (+S1), 65 pmol of purified S 1 were also present. After 20 minutes of reaction at 3 T C , the polypeptides synthesized were precipitated with boiling trichloroacetic acid (supplemented with 0.25% tungstic acid in the case of polylysine), filtered, and counted. The MS2 phage coat protein (the major synthetic product in panels C and D) contains 10.9 mol-% valine and 10.9 mob% alanine; hence the actrtaf amount of amino acids polymerized in C and D are ninefold greater than the ordinate values. Panel D is reproduced, with permission, from European Journal of Biochemistry ( 1 64), and the reader should refer to it for further details. Panels A, 9, and C are from the unpublished results of T. Suryanarayana and the author (see Section VI1,B).

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(147, 148), but unlike L1 (128),produces a striking stimulation of the protein synthetic capability of the ribosome. The results with specific mRNAs used in these in vitro systems are discussed in some detail below. 1. SYNTHETICMESSENGERS Escherichia coli ribosomes translate the classical synthetic mRNA poly(U) quite well (149),but the translation is strongly dependent on the presence of S1 (Fig. 9). The S1-dependence of this system, first demonstrated by van Duin and van Knippenberg (143),has since been confirmed by other workers (30,31,72,144).The dependence is greatest at low amounts of poly(U); it becomes progressively less as the amount of poly(U) added to the system is increased. I n our assay procedure, shown in Fig. 9, using 40 pmol of ribosomes, there is a 20fold or greater stimulation by S1 when 4 p g (nominally 40 pmol) or less of poly(U) are used. At 50 p g of poly(U), the system is independent of S1. The amount of S1 necessary to produce the saturating level of activity is 1 mol of S1 per mole of ribosome. A large excess of S1 is inhibitory (146,150),but this inhibition depends'on the poly(U) :ribosome ratio; there is strong inhibition when this ratio is low, but none when there is a large excess of poly(U) in the system (81,143)(observations of T. Suryanarayana and the author). Thus the inhibitory effect observed with excess S1 can b e ascribed to the binding and consequent sequestering of poly(U) molecules by free S1. Synthetic messenger poly(A) is also translated by ribosomes, and S1 has been reported to stimulate (143, 144) or be indifferent to this translation (72).A recent analysis of poly(A) translation (by T. Suryanarayana and the author) shows it to be essentially like the poly(U) system in having a strong dependence on S1 at low mRNA : ribosome ratios (Fig. 9) and no dependence when this ratio is high. Ribosomes have, in the absence of S1, a stronger affinity for poly(A) than for poly(U) (72).Therefore, the dependence of poly(A) translation on S1 is observed only when the poly(A) : ribosome molar ratio in the translation systemis well below one. At these low poly(A) :ribosome ratios, the system becomes also sensitive to inhibition by excess free S1. 2. NATURALMESSENGERS The genomic RNA of bacteriophage MS2, or its close relatives f2 and R17, is the natural mRNA predominantly used in the study of S1. This RNA contains four genes (151-153), but the translation of only

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the earlier known three genes, namely those of A-protein, coat protein, and synthetase (154), have been analyzed in these studies. The translation of M S 2 RNA is strongly dependent on the presence of S1 (Fig. 9). The translation also requires the initiation factors (155,156). All three of the phage proteins are made in vitro but, like in the infected cell, the coat protein is by far the major product of synthesis (157-159). The effect of S1 on the individual steps in the translation of MS2 RNA has been examined. S1 has no influence on ribosome dissociation (Section VI), the first step in the initiation of protein synthesis (160,161). The presence of S1 on the 30 S subunit is strictly necessary for the ribosome to bind M S 2 RNA (162, 163),i.e., the second step of initiation. S 1 is not necessary for the subsequent step of binding M e t tRNA: ribosomes programmed with AUG bind met-tRNA; and this binding is not influenced by the presence or the absence of S1 (144, 163). Ribosomes programmed with M S 2 RNA are S1-dependent for their Met-tRNA binding (144,163) and are inhibited by S1 antibodies (164),but this merely reflects the S 1dependence of the earlier mRNA binding step. When S1 antibodies are added to a system after the initial mRNA binding step, they do not cause inhibition of subsequent translation (165). This could mean either that S1 is not necessary for the elongation step or that the functional domains of S1 in the newly formed initiation complex are no longer accessible to the antibodies. The fact that S1 is a normal r-protein rather than one that, like the initiation factors, binds to ribosomes only at the initiation step, argues for a continued role of S 1 in protein synthesis. The antibody-insensitive stage is already reached when M S 2 RNA and 30 S subunits in the presence of 1 2 mM Mg2+ are incubated at 37-40°C for a few minutes. Thereafter the complex is resistant both to exchange with subsequently added mRNA and to initiation-specific inhibitors (165-167). Future experiments should reveal the precise nature of the interactions in this mRNA. 30 S complex.

3. INHIBITORSOF S1 FUNCTION An inhibitor of protein-nucleic acid interaction, and hence a potent inhibitor of protein synthesis initation, is the dye aurintricarboxylic acid (168, 169). On E . coli ribosomes, its main target is S1. Aurintricarboxylate (ATA) inhibits S 1’s interactions with RNA, presumably by itself binding to the RNA binding domains of S1 (170). The commercial ATA consists mostly of a heterogeneous mixture of phenolformaldehyde-type cross-linked polymers; the reactive species are only the polymeric ones (170).

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When S1 interacts with poly(U) or oligo(U), a conformational change occurs so that the complex is no longer cleaved by trypsin (see Section 111). The same resistance to trypsin is observed when S1 is treated with ATA (unpublished observations of the author), giving a further indication that the polymeric, anionic ATA inhibits S1 function by itself occupying the RNA-binding domain.

VIII. Mode of Action of S1 Results summarized so far make it abundantly clear that the major function of S1 in protein biosynthesis is at the mRNA binding step. There are also experiments in which a direct chemical link between mRNA and S1 has been made (171-174). Thus, when a ribosome * poly(U) . Phe-tRNA complex is ultraviolet irradiated, S 1cross-links to poly(U) (173).S1 is one of the several r-proteins that cross-link to a ribosome-bound photoaffinity analog of AUG ( 1 74),but since, as we noted, S1 is not necessary to bind AUG onto the ribosome, it is difficult to evaluate this result. Are ribosomes lacking S 1 totally incapable of binding mRNA? That does not appear to be the case. Both poly(U) and poly(A) bind to S l-free ribosomes (poly(A) somewhat more strongly than polyfU)), but S 1 considerably enhances the ribosome’s affinity for either molecule (72, 95, 175, and observations in my laboratory). Quantitative measurements have been reported in the case of poly(U) * ribosome interaction (95,175).Using poly(U) of defined length, a binding constant of 1 X lo6 M-’ has been observed with S1-free ribosomes, and 5 X lo9 M-’ with S1-containing ribosomes (95).In the case of MS2 RNA, as noted (162, 163, and my observations), the presence of S1 makes a very considerable enhancement of its binding to the ribosome. Thus, although it does not introduce a novel function into the ribosomal structure, S1 is able to amplify manyfold the preexisting but weak ribosomal affinity for mRNA. Are there some basic differences between the mechanisms employed by S1 and the ribosome for binding mRNA? Polj nucleotides carrying a fluorescent label bind to the ribosome, whereupon a fluorescence enhancement is observed ( 1 76).Free S 1and S l-lacking ribosomes also bind to the labeled polynucleotides, but only S1 produces the fluorescence enhancement ( 1 76),indicating differences in mechanisms of interaction.

1. MODE OF ACTION:PASTPROPOSALS Two proposals are in the literature to explain the mode of action of S1. According to one of them (113, 177, 178), S1 facilitates the ribo-

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some . mRNA recognition step that involves hydrogen bonding between the 3’-end sequence of 16 S rRNA and its complementary sequence on the mRNA. According to the second view (69, 164), S1 functions by unwinding double-strand regions in mRNA and thereby facilitates mRNA . ribosome interaction. The idea (179, 180)2of a second RNA RNA recognition step in initiation (other than the classical mRNA * (fMet-tRNA) interaction) is supported by many lines of evidence (76,181),but does S1 participate in this step? S1 clearly has a strong influence on the overall degree of initiation, and it also has a differential influence on the level of initiation at the three MS2 RNA genes (159,182). Therefore the possibility of S1 participating in the RNA * RNA recognition process was considered and suggested (177, 178). However, experimental support (97, 183,184) for the suggestion has never been strong. Evidence, pro and con, regarding the putative interaction between S1 and the 3’ end of 16 S rRNA is considered in Section VI, with the conclusion that such an interaction, though feasible, has yet to be proved. The 3’ end of 16 S rRNA itself (in the form of a 49-mer fragment) has been subjected to high-resolution NMR and temperature-jumprelaxation studies (97,185,186). These studies show a stable, helical “hairpin” structure in the fragment, but this structure does not include the nucleotide sequence believed to be involved in the ribosome . mRNA recognition. Only below 21°C may the latter form a “bulge-loop” on the stable hairpin; this bulge-loop is then melted by the addition of S i (97). Thus, at the growth temperatures ofE. coZi, the recognition sequence at the 3’ end of 16 S rRNA is likely to be singlestranded and free, as was observed by Backendorf et al. (114, 115) with their octadeoxynucleotide probe. The second proposal, i.e., the unwinding model, has two lines of experimental support. S 1 is capable of unwinding double-strand regions in RNA (68,69,99,100);and it has a 1000-fold greater affinity for the single-strand than it does for the double-strand structure (102). Second, with MS2 RNA as messenger, the requirement for S1 is minimized or abolished when the RNA is treated with formaldehyde (164). There are reasons to believe that the observed unwinding property of S1 may not be crucial for its function in protein synthesis. We have seen that S l is necessary for the efficient translation of poly(U) even though poly(U) exists at the assay temperature as a random coli, with-

-

Two other novel ideas of the same authors, namely, base-pairing between the 3’ end of 16 S rRNA and the termination codons (179) and determination of the cistron specificity in MS2 RNA translation by the 3’ end of 16 S rRNA (180)have not found experimental support.

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out secondary structure (187, 188). Thus the removal of secondary structure of MS2 RNA (the presumed consequence of formaldehyde treatment) in itself cannot explain the observed behavior of the formaldehyde-treated RNA. Formaldehyde treatment is known to introduce additional, nonspecific initiation sites in MS2 RNA (189),which could in part mask the S1 effect. Moreover, from the conflicting results on S1 versus poly(A) translation (see Section VII,B,l) it has become clear that the S1 effect, to be firmly established, has to be studied over a wide range of RNA concentration. A characteristic of the unwinding of nucleic acids by S1 is its antagonism by Mg2+ (69), while protein synthesis requires this ion. But the critical argument against the unwinding model comes from the effect of sulfhydryl group blocking on unwinding versus protein synthesis. When S1 is treated with N-ethylmaleimide, it yields a monomaleimide derivative (Section 111). The modified S 1 binds to nucleic acid as well as S1 does, but is unable to unwind the secondary structure (60, 68, 69). Recent experiments show that maleimidetreated S1 is fully reactive in Q/3 replicase function (71),fully reactive in poly(U)-directed protein synthesis (72 and my observations), and 50% active in MS2-RNA-directed protein synthesis (observations of Suryanarayana and the author). Thus, although probably interesting in its own right (69), the unwinding property of S1 does not hold the key to the biological functions of S1. 2. MODEOF ACTIONOF S1

S1 has many striking structural features that ought to be incorporated into any proposal regarding its mode of action. There are (to summarize sections 1-111) its highly elongated shape, its bidomain structure with a hinge (or freely rotatable) region in between, and four structurally homologous segments in the RNA-binding domain. In addition, its binding to RNA is base-sequence-independent, and its affinity to ribosome is weak. The second critical finding, which should also be incorporated in any proposal, is one that has emerged from studies using ribosomes completely free of S1. Such ribosomes still possess an intrinsic capacity to bind and to translate mRNA. Given high concentrations of mRNA, Sl-free ribosomes are often as efficient as S1-containing ribosomes, but the latter become clearly superior when the mRNA concentration is reduced. We are proposing a model for the function and mode of action of S1 incorporating these two principles. According to this model, the func-

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tion of S1, which has a large perimeter of action due to its elongated, bidomain shape, is TO ACT AS AN RNA BINDING PROTEIN AND TO BRING mRNA TO THE PROXIMITY OF THE RIBOSOME. This action of S I would reduce the concentration of mRNA that would otherwise be necessary to saturate the intrinsic mRNA binding site on the ribosome. It would also aid the subsequent mRNA * (16 S rRNA) and mRNA . (Met-tRNA) interactions that take place on the ribosomal surface, but without S1 being directly involved. The fact that one molecule of S1 is always associated with the ribosome could indicate that the S1 . mRNA attachment is maintained throughout translation and therefore that S 1 acts additionally as an “mRNA HOLDING” protein. There are certain features of this model that can be tested experimentally. The known characteristics of the interactions of S1 with the ribosome and with nucleic acids, as well as most of the properties of S1 itself, are in accord with this model. There are two properties of S1, however, on which the model is silent. One is the differential specificity that S1 so clearly exhibits in the translation of the different genes on phage MS2 RNA (157-159,182). The other is the specific function (and hence the evolutionary pressure for its formation) of the four homologous segments in the RNA binding domain of S1 (Section 111). I would like to end this article on an evolutionary note. It has been suggested that the ribosome, with its high speed and accuracy, must have evolved from a simpler, yet translationally competent, particle by means of a series of refinements (e.g., see 128,190).The proposed model of Sl’s mode of action would suggest that S1 was such an evolutionary refinement in E . coli. What its counterpart in other families of organisms might be is an interesting topic for future research.

ACKNOWLEDGMENTS I thank Professor H. G. Wittmann for helpful comments on the manuscript and Dr. T. Suryanarayana for useful discussions.

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126. W. A. Held, S. Mizushima, and M. Nomura,JBC 248,5720 (1973). 127. E. Dabbs,J. Bact. 140,734 (1979). 128. A. R. Subramanian and E. Dabbs, EJB 112,425 (1980). 129. M. Kitakawa and K. Isono, Mol. Gen. Genet. 185, 445 (1982). 130. T. Yamada, FEBS Lett. 142,267 (1982). 131. K. Isono and S . Isono, PNAS 73, 767 (1976). 132. K. Higo, E. Otaka, and S. Osawa, Mol. Gen. Genet. 185,239 (1982). 133. W. J. Sharrock, B. M. Gold, and J. C. Rabinowitz,JMB 135,627 (1979). 133a. J. R. McLaughlin, C. L. Murray, and J. C. Rabinowitz,JBC 256, 11283 (1981). 134. C. Weissmann and S. Ochoa, This Series 6,353-399 (1967). 135. D. Shin and K. Moldave,JMB 21,231 (1966). 136. M. Revel and F. Gros, BBRC 27, 12 (1967). 137. J. C. Leavitt, K. Moldave, and D. Nakada, J M B 70,15 (1972). 138. J. C. Leavitt, R. H. Hayashi, and D. Nakada, ABB 161,705 (1974). 139. H. F. Kung, J. Morrissey, M. Revel, C. Spears, and H. Weissbach, JBC 250, 8780 (1975). 140. G . van Dieijen, P. H. van Knippenberg, J. van Duin, B. Koekman, and P. H. Pouwels, Mol. Gen. Genet. 153, 75 (1977). 141. J. C. Lelong, R. Maschler, M. CrBpin, C. Jeantet, G. W. Tischendorf, and F. Gros, Biochimie 61, 881 (1979). 142. C. Yanofsky, Nature (London)289,751 (1981). 143. J. van Duin and P. H. van Knippenberg, J M B 84, 185 (1974). 144. J. E. Sobura, M. R. Chowdhury, D. A. Hawley, and A. J. Wahba, NARes, 4, 17 (1977). 145. J. C. Lelong, D. Gros, F. Gros, A. Bollen, R. Maschler, and G. Stoffler, PNAS 71, 248 (1974). 146. G. van Dieijen, C. J. van der Laken, P. H. van Knippenberg, and J. van Duin,JMB 93, 351 (1975). 147. E. Hamel, M. Koka, and T. Nakamoto, JBC 247,805 (1972). 148. I. Pettersson and C. G. Kurland, PNAS 77,4007 (1980). 149. M. W. Nirenberg and J. H. Matthaei, PNAS 47,1588 (1961). 150. M. J. Miller and A. J. Wahba,JBC 249,3808 (1974). 151. P. Model, R. E. Webster, and N. D. Zinder, Cell 18,235 (1979). 152. J. F. Atkins, J. A. Steitz, C. W. Anderson, and P. Model, Cell 18, 247 (1979). 153. M . N. Beremand and T. Blumenthal, Cell 18,257 (1979). 154. W. Fiers, R. Contreras, F. Duerinck, G. Haegeman, D. Iserentant, J. Merregaert, W. Min Jou, F. Molemans, A. Raeymaekers, A. Van den Berghe, G. Volckaert, and M. Ysebaert, Nature (London) 260, 500 (1976). 155. S. Ochoa and R. Mazumder, in “The Enzymes” (P. D. Boyer, ed.), 3rd ed., VoI. 10, pp. 1-51. Academic Press, New York, 1974. 156. M. Grunberg-Manago and F. Gros, This Series 20, 209-287 (1977). 157. M. Kozak and D. Nathans, Bacteriol. Rezj. 36, 109 (1972). 158. H. F. Lodish, in “RNA Phages” (N. D. Zinder, ed.), pp. 301-318. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1975. 159. S. Isono and K. Isono, EJB 56, 15 (1975). 160. A. R. Subramanian, E. Z. Ron, and B. D. Davis, PNAS 61, 761 (1968). 161. A. R. Subramanian and B. D. Davis,JMB 74, 45 (1973). 162. W. Szer and S . Leffler, PNAS 71,3611 (1974). 163. W. Szer, J. M. Hermoso, and S. Leffler, PNAS 72,2325 (1975). 164. G. van Dieijen, P. H. van Knippenberg, and J. van Duin, EJB 64, 511 (1976).

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G. van Dieijen, P. Zipori, W. van Prooijen, and J. van Duin, EJB 90,571 (1978). P. Zipori, L. Bosch, G. van Dieijen, and G. van der Hofstad, EJB 92,225 (1978). J. van Duin, G. P. Overbeek, and C. Backendorf, EJB 110,593 (1980). A. P. Grollmann and M. L. Stewart, PNAS 61,719 (1968). A. P. Grollman and M. T. Huang, in “Protein Synthesis (E. H. McConkey, ed.), Vol. 11, pp. 125-134. Dekker, New York, 1976. R. G. Gonzhlez, B. J. Blackburn, and T. Schleich, BBA 562,534 (1979). I. Fiser, K. H. Scheit, G. Stoffler, and E. Kuechler, BBRC 60, 1112 (1974). I. Fiser, P. Margaritella, and E. Kuechler, FEBS Lett. 52,281 (1975). P. Margaritella and E. Kuechler, FEBS Lett. 88, 131 (1978). 0. Pongs, H. U. Petersen, M. Grunberg-Manago, E. Lanka, R. Bald, and G. Stoffler, JMB 134,329 (1979). S. V. Kirillov, V. I. Makhno, and Y. P. Semenkov, EJB 89, 297 (1978). D. E. Draper and L. Gold, Bchem 19, 1774 (1980). J. A. Steitz, K. U. Sprague, D. A. Steege, R. C. Yuan, M. Laughrea, P. B. Moore, and A. J. Wahba, in “Nucleic Acid Protein Recognition” (H. J. Vogel, ed.), pp. 493508. Academic Press, New York, 1977. J. A. Steitz, in “Biological Regulation and Development” (R. F. Goldberger, ed.), pp. 349-399. Plenum, New York, 1979. J. Shine and L, Dalgarno, PNAS 71, 1342 (1974). J. Shine and L. Dalgarno, Nature (London)254, 34 (1975). M. Gmnberg-Manago, in “Ribosomes,” 1980; pp. 445-477. J. A. Steitz,JMB 73, 1 (1973). J. A. Steitz, A. J. Wahba, M. Laughrea, and P. B. Moore, NARes 4, 1 (1977). S. Goelz and J. A. Steitz,JBC 252,5177 (1977). R. A. Baan, C. W. Hilbers, R. van Charldorp, E. van Leerdam, P. H. van Knippenberg, and A. L. Bosch, PNAS 74, 1028 (1977). R. A. Baan, Ph.D. Thesis, State University of Leiden, 1977. M. N. Lipsett, PNAS 46,445 (1960). W. Szer,JMB 16,585 (1966). H. F. Lodish and H. D. Robertson, CSHSQB 34,655 (1969). C. Woese, in “Ribosomes,” 1980,3pp. 357-373. A. R. Subramanian, EJB 45,541 (1974). A. Kyriakopoulos and A. R. Subramanian, BBA 474,308 (1977). E. J. Cohn and J. T. Edsall, “Proteins, Amino Acids and Peptides.” Hafner, New York, 1943.

The Yeast Cell Cycle: Coordination of Growth and Division Rates STEVEN G. ELLIOTT~ AND CALVINS. MCLAUCHLIN Department of Biological Chemistry, California College of Medicine, University of Calijornia, Imine, California

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................. C. Metabolic Activities D. DNA Synthesis.. . . . . . . . .............. .......... E. RNA Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Proteins and Enzymic Activitie . . . 111. Control of Cell Division. . . . . . . . . . . . . . . . . . . . . . .

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I. Methods for Cell-Cycle Analysis A. Volume, Cell Mass, and Density

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One of the central problems for cell biology in this decade is the biochemical elucidation of the mechanisms of cell division in eukaryotic cells. A thorough molecular description of the cell cycle is an essential component of our understanding of aging and development. The abnormal cell division associated with cancer indicates that an important goal of cancer research is the biochemical and genetic description of the nature of this abnormal cell division process. The fungi, and especially the yeasts, have an important role to play in the development of the biochemistry of the cell cycle. It is clear that the basic processes in eukaryotic cell division are well represented in the yeasts. Thus, an examination of the current status of cellcycle studies in yeast may be useful to researchers interested in mammalian and plant cell-division, as well as those interested in yeast. This review is timely because the rapid advances in the molecular biology of yeasts, including but not limited to recombinant DNA technology, promise exciting advances in this decade.

'

Present address: Department of Biology, University of California, Santa Barbara, California 93106. 143 Progress in Nucleic Acid Research and Molecular Biology. Vol. 28

Copyright 0 1983 by Academic Press, Inc. A11 rights of reproduction in any form reserved. ISBN 0-12-540028-4

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Two essential, distinct processes take place during the cell-division cycle. Growth occurs to increase the cellular components and to duplicate the genetic material. Second, a defined sequence of morphological events distributes the duplicated genetic material equally between the mother and daughter cell and ensures an adequately equitable distribution of the other cellular components. Under certain conditions, cell division can occur without growth, but generally the two processes are interrelated. For yeasts in balanced growth, the two processes must be interrelated, because if cell division occurs faster than cell growth, the average cell size will decrease until a cell is no longer competent to divide at the initial rate. Conversely, if cell growth is more rapid than cell division, the average cell size will increase until the cell is no longer capable of growing at the initial rate. Three basic patterns of cellular growth have been considered (I). These are idealized in Fig. 1. Step accumulation (periodic synthesis) can cause a doubling in cellular content by a short burst of synthesis (Fig. lA, 1A’). This is a common method of accumulation for DNA (I). The increase in amount of a component can occur through an exponential rate of synthesis, which leads to an exponential rate of accumulation (Fig. lB, 1B’). Finally, the rate of synthesis can be held constant, resulting in a linear pattern of accumulation. In this case the rate of synthesis must double at some point in the cycle, or the resulting progeny will grow at half the rate of the parent. This results in a characteristic “step” pattern of synthesis (Fig. lC, 1C’). Each of these growth patterns can be related to specific types of metabolic control.

tl t 2

t

Time or cell c y c l e position

FIG. 1. Idealized accumulation and rate curves during the cell cycle: (A) step accumulation,(Bf exponential accumulation, (C) linear increase with doubling in rate of accumulation, (A’) periodic synthesis, (B’) exponential synthesis, (C’) linear increase with doubling in rate of synthesis.

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Periodic variations in the rate of synthesis require that both the initiation of synthesis and the termination of synthesis be under some sort of specific metabolic control. The exponential increase in the rate of synthesis could be accomplished by several types of autoregulatory control. Step variations in the rate of synthesis could be most simply accomplished if the substance examined is under promoter control and the inflection or step point is at S (Fig. 2), as the number of copies of a given gene and its promoters are doubled at that point. The patterns of accumulation and synthesis of a number of different cellular components have been studied in yeast. I n many ways yeast is an ideal organism for studies of cell growth and division. This organism progresses through a number of well-defined morphological and genetic steps during cell division that allow a precise delineation of the cell-cycle position of individual cells. Photomicroscopy of growing cells is possible because yeast grows rapidly on solid media. Large quantities of synchronously growing cells can be generated by a variety of methods. Mutants defective in cell division are available for study. The yeast cell cycle is similar to that of higher eukaryotes; it can be divided into four phases: G1, S, G2, and M. Two types of yeast have been studied extensively; their cell cycles are shown in Figs. 2 and 3. Budding yeasts, such as Saccharomyces cerevisiae (Fig. 2) are

FIG.2. Cell cycle of Saccharomyces cereuisiae. Abbreviations: ids, initiation of DNA synthesis; be, bud emergence; cds, completion of DNA synthesis; nm, nuclear migration; mnd, medial nuclear division; se, spindle elongation; Ind, late nuclear division; ck, cytokinesis; cs, cell separation.

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FIG.3. Cell cycle of Schizosacchuromyces pombe. Abbreviations: nd, nuclear division; cpf, cell plate formation; ids, initiation of DNA synthesis; cpm, cell plate maturation; cs, cell separation; cds, completion of DNA synthesis.

characterized by bud initiation early in the cycle. The bud grows in size throughout the cycle. When the bud approximates three-fourths of the size of the maternal cell, the nucleus migrates into the neck of the bud, a spindle is formed, and nuclear division and cytokinesis follow (2). Fission yeasts such as Schixosaccharomyces pombe (Fig. 3 ) divide by a different process. These cells grow primarily in length throughout the cell cycle. When the length doubles, the centrally located nucleus divides and migrates to opposite ends of the cell. A plate then forms in the center of the cell, and cell separation occurs. These two yeasts differ in the relative proportion of time spent in the different cell-cycle phases ( 3 ) .In S . pombe, a major proportion of time is spent in G2 with a relatively short G1; in S. cerevisiae, the major portion of the cell cycle is spent in G1.

1. Methods for Cell-Cycle Analysis

The methods used to study cell division are, perhaps, the most important and controversial aspect of cell-cycle research. An ideal method would allow the cells to grow and divide in a constant, unchanging natural environment. The method of analysis should in no way perturb this “normal” pattern of the cell cycle. Yeast cells as they are naturally found rarely exist under conditions of constant environ-

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ment. Therefore, there is difficulty in defining a normal cell cycle. However, the ideal cell cycle described above is a useful concept because it is clear that cells exposed to a changing environment can show periodic events that are not an intrinsic part of the normal cell cycle. The concept of “balanced growth” was established in the late 1950s and early 1960s ( 4 , 5 ) .Balanced growth exists when every cellular component precisely doubles in amount with each division (6). Interestingly, according to the above definition, cells in batch culture are not in balanced growth (7, 8). This concept was later refined to suggest that cells are, for practical purposes, in balanced growth when the population size increases exponentially and the cell ages are stably distributed (9). Another method of defining cells in balanced growth is through the measurement of mitotic cells. If the length of mitosis is independent of the rest of the life cycle, and the number of mitotic cells increases exponentially, cells will be in balanced growth. This definition does not apply to synchronously growing cultures, and other criteria must be used for determining normalcy, such as length of the cell cycle before and after treatment, size at division in successive cell cycles, and the presence of a “normal” decay of synchrony during growth (6). Methods used to study the cell cycle can be divided into induction synchrony, selection synchrony, cell selection, and single cell methods. Cultures can be induced to divide synchronously by imposing a cell-cycle block that allows all the cells in the culture to collect at the same stage in the cell cycle. After removal of the block, cells divide synchronously for up to three cell cycles. Such blocks include: C1 arrest due to nutrient limitation followed by addition of fresh media (feed-starve synchrony) (10); chemical inhibition of DNA synthesis (e.g., deoxyadenosine, A23187,2 hydroxyurea, fluorodeoxyuridine) (11-13); temperature-sensitive cdc mutants ( 1 4 ) ;and the pheromone a factor (15). Synchronous division can also be induced by periodic heat shocks (16, 17) and periodic addition of a limiting nutrient (17, 18). Each of these methods can produce large quantities of cells in synchronous growth, which is a decided advantage over other methods. If, during the cell-cycle block, cells continue to grow, they are by definition in unbalanced growth. There is concern that the inducing agent may alter the cell growth characteristics and result in an abnormal cell cycle. DNA blocks, cdc mutants, and heat-shock-induced synA23187 is calcimycin (CA 52665-69-7).[Ed.]

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chronous cultures all show shortened cycles and an increase in size at division for at least the first cell cycle (12, 19, 20). An analysis of theoretical and observed synchronous cultures prepared by the feed/ starve method established, on the basis of the natural variability in generation time, that the degree of synchrony obtained is better than expected (21).This suggests that synchronous cultures prepared by this method may be perturbed. Indeed, a number of components accumulate in a different pattern during the cell cycle induced by feedstarve techniques compared to methods that are more likely to result in balanced growth. Selection synchrony involves the physical separation of cells into cell-cycle fractions. The earliest method involved the removal of small G 1 cells from sucrose gradients. These divide synchronously when inoculated in fresh medium (22).This method can be scaled up by using a zonal rotor (23, 24). Cells can also be separated into cellcycle stages in density gradients (25-27). Both methods can cause perturbations in some cellular properties in subsequent growth (28, 29). The perturbations may be due to either the osmotic changes, nutrient or oxygen starvation during harvesting and centrifugation, or in some cases cold shock. A less stressful method of selection synchrony involves continuous-flow centrifugation ( 3 0 ) or centrifugal elutriation (31), in which cells are continuously bathed in medium at growth temperature. Cell harvesting becomes unnecessary because the cells in culture are loaded directly into the rotor. In this method, liquid is pumped through a separation chamber while the rotor is spinning. Two forces, a centrifugal force and an opposing flow of liquid, act on the cells. Small cells can be separated from an exponentially growing culture and used to form a synchronous culture by choosing flow rates and centrifuge speeds at which small cells are flushed out (29).Perturbations observed in synchronous cultures prepared by sucrose-gradient centrifugation are eliminated (29).However, this method is not perfect because some enzymes show periodicities in asynchronous control cultures treated to the continuous flow method (32). Complete fractionation of a culture for cell selection avoids the problem of perturbation. A culture in balanced growth can be fractionated under conditions where metabolism is stopped. Rates of synthesis for many components can be determined by pulse-labeling the culture prior to fractionation. A good correlation exists between cell size and cell-cycle position (23,31).Assigning a cell-cycle position to a particular fraction and defining the ends of a cell cycle in the fractions may be difficult when size varies as much as fourfold in a popula-

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tion; under these conditions, it is difficult to determine which fractions represent one complete cell cycle. The ends of the cell cycle can be estimated by using only those fractions that include a doubling in cell volume. The volume range used is estimated from the mean cell volume of the original culture (33). The number of variables tend to prevent an ideal cell-cycle separation by size selection methods. Cell-cycle fractionation based on size is influenced by cell-to-cell size variation and the discrepancy between cell size and cell-cycle position. Volume increase is approximately exponential during the cycle, whereas size fractionation results in a more linear separation. In cell-cycle fractions, the beginning of the cycle may be better resolved owing to a tendency of the larger sized cells to be contaminated with small cells, but not vice versa. Because older cells tend to be larger than younger cells (3 4 ),a different relationship exists between size and cell-cycle position for each cell age. However, on balance, this is the method of choice for the biochemical analysis of the cell cycle, with minimum perturbations. Single-cell methods overcome this problem of cell size variation to some extent because the abnormally large or small cells need not be analyzed. A large variety of cellular parameters can be measured on single cells. Visual characteristics can be examined by time-lapse photomicrography (35,36),and dry mass can be measured by an interference microscope or Cartesian diver ( 1 , 3 7 , 3 8 ) .The synthesis of many macromolecules can be measured by autoradiography (3 9 ),and fluorescent enzyme assays have been applied to individual cells (4 0 ). Data from a number of cells can be collected and plotted against the cell-cycle position. However, it is not yet possible to determine cellcycle position for each cell precisely, so cell size is usually used as a measure of cell-cycle position. Despite this disadvantage, shared with the cell-selection method, the single-cell method may provide unambiguous answers to many unanswered questions about the cell cycle.

II. Patterns of Growth A. Volume, Cell Mass, and Density In budding yeast, total cell mass increases continuously through the cell cycle (18,37,38,41).The data suggest that the rate of increase may be linear, although an exponential rate of increase cannot be eliminated. Volume increase is sigmoidal in budding yeasts throughout the cell cycle (35,37,38).Volume increases rapidly following bud

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emergence, with most of the volume increase due to the growth in the size of the bud, not the mother cell (35, 37, 38, 4 2 , 4 3 ) , although the mother cell does increase in size prior to bud emergence ( 4 4 ) .The volume changes are primarily due to the rate of formation of new cellwall material, since the cell wall is a rigid structure ( 4 5 ) . The fact that volume and dry mass increase at different rates indicates that, because the wall is rigid, cell density will change periodically. This density change may be partialIy offset by changes in cellular water content. The volume of water that flows in and out of the cell has not been determined, although some results suggest that cells lose water during bud emergence ( 2 7 ) .It is clear that density does change periodically with a maximum at the start of the cell cycle ( 2 5 , 2 7 ) .The changes in density are correlated with periodic fragmentation and reformation of the vacuoles. The vacuoles decrease in size and fragment at bud emergence; then, as the cells grow, the vacuoles enlarge and fuse ( 2 7 ) . The rate of mass accumulation in the fission yeast S. pombe has been thoroughly studied. Initial studies suggested that mass accumulates linearly with a doubling in rate during the cell cycle ( 4 6 ) .Subsequent studies in different media suggest that the rate of increase in mass is exponential ( 4 7 ) .It is not clear that the difference between linear and exponential growth can be determined accurately under these conditions, although a statistical analysis of volume distribution suggests that the exponential increase of cell mass is unlikely ( 7 , 4 8 ) . At lower temperatures, the rate of increase of dry mass falls at the end of the cycle ( 4 6 ) . Cell volume increases exponentially for three-fourths of the cycle and then remains constant ( 7 , 3 6 , 4 6 , 4 8 , 4 9 ) .The increase in volume is accounted for by the increase in cell length. As in the budding yeast, the differing rate of volume and cell mass accumulation throughout the cell cycle of the fission yeast dictate a variation in cell density through the cell cycle. The maximum density occurs at the end of the constant-volume phase, while the minimum density is observed at the beginning of the cell cycle (I). The observed density changes are not mediated by vacuolar changes, because in normal growth media S . pombe does not have a vacuole ( 3 ) .

B. Cell Walls, Septa, and Membranes All yeasts have a rigid outer cell wall composed of polysaccharide and protein; however, the structure of the cell wall differs between budding and fission yeast. The wall of S. cerevisiae is composed primarily of glucan, mannan, chitin, protein, and lipid ( 4 5 , 5 0 , 5 1 ) .Mannan is the major carbohydrate, although a significant constituent re-

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sponsible for rigidity is a P-glucan in Pl-3 and p1-6 linkages (45,50). The cell wall of S . pombe is composed mainly of a P-glucan and an aglucan with some galactomannan. There is very little mannan and chitin (52, 53). Schizosaccharom yces pombe grows primarily in length during the cell cycle. The shape and the size of the cell is affected by the cell wall; the rate of wall growth must correlate with the rate of accumulation of other cellular components. Wall extension at the tips occurs during three-fourths of the cycle (48,49, 54-59). There is some wall synthesis outside the growth regions at the tip, probably due to reorganization and thickening of the wall (56, 59). Length extension appears to increase exponentially, which correlates with the exponential volume increase (49,56,57). The rate of wall extension is not equal at both ends of the cell. The end opposite the cell plate (the original end) grows first and is responsible for three-fourths of the length increase. The new end grows for a short period of time late in the cycle (49,56, 60). During the constant-volume phase at the end of the cycle, wall synthesis is primarily at the cell plate ( 1 ). The cell plate appears 10 minutes after the initiation of the constant volume phase (0.85 of the cycle) (49). The septum is biased toward the newer end according to a length parameter such that the volumes of the resultant cells are equal (611. In S. cereoisiae, most of the wall synthesis is in the bud. Glucan and mannan wall components increase continuously throughout the cell cycle (43, 59, 62-65). Biely (65) analyzed in detail the rate of synthesis of wall components and observed a reduction in the prebudding phase and at cell division. The continuous synthesis of glucan can be explained partly by the stability of glucan synthase and its mRNA. Mannan synthase, however, is unstable, as is its mRNA (66). There is very little synthesis in the mother cell (59, 67); the small amount observed is probably due to the reorganization of wall material. During the initial stages, synthesis is over the entire surface of the emerging bud. Later synthesis is at the tip (67-70). When the bud is approximately two-thirds the size of the mother cell, the entire bud again shows incorporation, presumably due to final maturation (67, 71 ). Synthesis is apparently directed by small vesicles, derived from the rough endoplasmic reticulum, which fuse with the cell wall to release their contents (wall precursors and synthases) onto the growing area of the bud (71-75). Later thickening of the wall occurs by depositing another layer inside the first by growing into the cytoplasm (71). Finally, a septum grows inward from the wall and thickens.

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Vesicles are also involved in this process. The cross wall is composed of two layers. The layer closer to the bud, which becomes the bud scar, is composed of a chitin ring and material made of mannan with small amounts of glucosamine and glucan. The outer layer is composed primarily of glucan (2, 62, 76). Following cell separation, a permanent bud scar remains on the surface of the bud and a birth scar remains on the mother cell (2, 76). Inside the cell wall is a three-layer membrane that is essential to the uptake of small molecules. This membrane is composed of lipid, protein, polysaccharide, and sterols (51, 77, 78). A number of membrane components accumulate exponentially in budding yeasts, including free and esterified sterols, phospholipid, and unesterified fatty acids (79,80).Incorporation of leucine and phosphate into membranes also increases exponentially (81). Unlike wall growth, membrane growth appears to be the same in both mothers and buds because of the random migration of membrane components (81).

C. Metabolic Activities The rate of oxygen uptake and carbon dioxide evolution are convenient measures of the metabolic activity. Oxygen uptake measures the energy production and metabolism in organisms dependent primarily on respiration for ATP production. Yeast can grow fermentatively, and, in the absence of oxygen, grows exclusively by converting glucose into ethanol, C02, and energy. Usually, yeast cells grow by a combination of respiration and fermentation. Although glycolytic activity during the cell cycle has been studied extensively, there is some confusion about the pattern of increase. In S . cerevisiae, four patterns of oxygen uptake have been reported: step increases (38,82, 83); periodic increases and decreases one or more times per cycle (27, 63,84,85); exponential increases (86);and linear increases with doublings in rate (18). In S. pombe, there are different patterns of oxygen uptake, including step increases once or twice per cycle (87-go), exponential increases with a maximum (91),and linear increase with a doubling in rate (85, 88, 92). The evolution of C02 has also been examined in S . pombe. Both step and linear increases are observed

(92, 93). Although differing results have been observed for glycolytic activity during the cycle, it is important to note that control cultures, to determine the effect of the synchrony procedure on the culture, were incorporated in only a few of the studies (88, 92, 93). As discussed above, many synchrony procedures induce artifacts. Two groups compared selection synchrony, heat-shock synchrony and measurements on single cells on the pattern of glycolytic activity,

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using the gradient diver method (85,92). These three methods gave a linear doubling increase. This pattern was observed in both complex and minimal media. Oxygen uptake and carbon dioxide evolution were continuous by deoxyadenosine and hydroxyurea synchrony ( 88, 93). A linear doubling increase of COZ evolution was observed by selection synchrony in complex medium, but step increases occurred in minimal media (93).The situation remains unresolved, and a final answer may require the determination of which patterns are artifacts, and whether the strain or the history of the culture is important in the observed pattern. Resolution of these questions will establish which methods are most able to reflect the true nature of the cell cycle.

D. DNA Synthesis 1. NUCLEAR DNA A number of researchers have shown in S. ceretiisiae through a variety of methods that DNA is synthesized periodically in S. ceretiisiae. Early studies utilized synchronous cultures prepared by feedstarve synchrony with resulting perturbations ( 94,951. Williamson in 1965 avoided perturbations arising in earlier synchronization procedures by pulse-labeling asynchronous cells in balanced growth ( 96). Autoradiography of single cells showed that DNA was synthesized for a brief portion (one quarter) of the cell cycle starting at the time of bud emergence. These results were confirmed later by size fractionations in zonal centrifugation (23,97) and elutriator centrifugation (98).The observation that nitrosoguanidine causes mutations at the replication point during DNA synthesis in E . coli (99)suggested a simiIar experiment in yeast. An increase in the mutation frequency was observed for a short period of the cell cycle following bud emergence (100-103), consistent with periodic DNA synthesis. DNA synthesis is also periodic in other yeasts, including C. albicans (142) and S. pombe (104, 105). In fission yeast, the S period is very short (10 to 15 minutes), about 10% of the cycle (104,105) compared with 30 minutes (25% of the cycle) in S. ceretiisiae (96). In addition, the S period for fission yeast occurs earlier in the cell cycle and spans the period between the end of mitosis and a point after cell division. In slow-growing cells and abnormally small cells, the S phase is delayed. Under these conditions, the time between S phase and cell separation is constant (105-107). In fast-growing cells, the S phase begins immediately after mitosis. The time from the end of S to the following cell division varies according to the growth rate (105,

106,108). In S. cereuisiae, DNA synthesis is initiated at or near the time of

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bud emergence. The time required to complete S is relatively constant at moderate growth rates (109-112). Therefore, as the length of the cell cycle increases owing to a decrease in growth rate, the time of S phase is delayed as in fission yeast. At slow growth rates, the time from initiation of S to cell separation also increases (112), as does the length of S itself ( 1 11-113). The duration of S appears to be a function of the medium (114). The increase in S can be accounted for by changes in the replication-fork rate, not by changes in the number of replicons (113,114). DNA replication itself is semiconservative (115-11 7) and is initiated internally (14,113,118,119).A single strand of DNA observed in the electron microscope often has several replication bubbles (14, 118, 119). By measuring the center-to-center distance between bubbles, it has been estimated that the average replicons are 10-40 p m apart. A similar result (10-60 p m ) was obtained by fiber autoradiography of pulse-labeled DNA (113,119).Assuming that replication origins are evenly distributed, yeast DNA has 140. This estimate may be low, because in any one stretch of DNA all the origins may not b e simultaneously active. The bubbles observed are replication origins, because both bubbles and DNA fragments the size of replicons accumulate following blocks in DNA synthesis (120, 121). Certain sequences of DNA, “autonomously replicating sequences” (ARS; 121a), are associated with increased transformation frequencies and allow autonomous replication of plasmid DNA. These sequences are presumed to be replication origins (122, 123). The frequency of ARS sequences in random clones indicates that there are 150 to 300 replication origins, or about 10 to 20 per chromosome (122, 123). Several ARS sequences have been sequenced (124,125).,Comparisonof two out of three such sequences indicated some limited homology; however, there was no significant homology with the third sequence (124). This suggests either that conformation of the DNA is important in DNA synthesis, or that only limited regions are recognized by DNA initiation factors. Consecutive pulses of different specific activity and fiber autoradiography can give the direction and rate of DNA synthesis. Experiments based on this method established that replication is bidirectional and extends at a rate of 0.7 Fm/min (119). A second group obtained a similar result and observed that the rate decreased at slower growth rates (113).At doubling times ranging from 105 to 380 minutes, the rate of fork movement dropped from 2.04 pm/min to 0.56 pm/min. Assuming the haploid nucleus has 9.2 x lo9 daltons of DNA (about 14,000 kilobases), the replication rate is 1 pm/m in (2.8 kilobases per minute) and proceeds bidirectionally. The minimum number

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of replication origins required to complete DNA synthesis in the observed 30 minutes is approximately 80. Therefore, there are 2 to 3 times more replicons present than required to complete DNA synthesis during S. While most adjacent replicons initiate within a few minutes of each other, the more distant ones do not and there are in fact activations throughout S phase (113).Several workers have reported that the induction of mutations by nitrosoguanidine for different genetic markers have peak mutation frequencies during different intervals in S phase. This suggests some temporal order in DNA synthesis ( 1 0 0 , 101, 103, 126). In contrast, ribosomal DNA, of which there are approximately 140 copies each replicated once per S phase, is synthesized throughout (127, 128). Protein synthesis is required about 10 minutes prior to the initiation of the S phase in S . cerevisiae (129-132) and in S . pombe (129). However, elongation of DNA does not require protein synthesis. RNA synthesis is also required prior to DNA synthesis (130). These required protein and RNA synthesizing steps are apparently associated with the start region of the cell division cycle as defined by the cdc28 gene product and the (Y factor-sensitive step (130). Studies on in vitro DNA synthesis indicate that initiation of DNA may occur via synthesis of an RNA primer (133).Such a mechanism can generate Okazaki fragments that can be observed under certain conditions (121, 134). The fragments are later joined by DNA ligase (134, 136). Some RNA synthesis mutants show secondary defects in DNA synthesis, and in many DNA synthesis mutants RNA synthesis is blocked (137).In addition, a 37,000-dalton protein factor associated with RNA polymerase that may stimulate DNA synthesis has been identified (138). A number of temperature-sensitive mutants that show defects in DNA synthesis have been isolated and identified in three different yeasts, S . cerevisiae (135, 137, 139-141), S . pombe (143, 144), and Ustilago maydis (145,146).These DNA-synthesis mutants have been characterized as to whether they are defective in initiation or elongation of DNA synthesis. Seven genes in S . pombe that show defects in DNA synthesis have been identified (143).Three of these show apparent defects in the initiation of DNA synthesis (cdclO, cdc21, cdc20). The other genes displayed defects in chain elongation or in the joining of DNA fragments. Gene cdcl7 has been identified as a DNA-ligase mutant (136), and cdc22 is defective in nucleoside diphosphokinase (147). In S . cerevisiae, five mutants (cdc'i, 8, 9, 21, and 40) showing defects in the replication of DNA have been identified (139-141, 148). An additional seven genes involved in the temperature-sensitive de-

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pression of DNA synthesis have been identified. These exhibit either a rapid total cessation or a reduced rate of DNA synthesis at the restrictive temperature (137).The function of two cdc genes is known. Mutant cdc9 is defective in DNA ligase (135),and cdc2l is defective in thymidylate synthetase (148,149). A DNA polymerase mutant has not yet been identified in either S . cerevisiae or S . pombe. However, in Ustilago maydis, one of the five temperature-sensitive mutants defective in DNA synthesis is a temperature-sensitive DNA-polymerase I mutant (145). In S . cerevisiae, two DNA polymerases (termed PolA and PolB) have been identified and characterized (150-152). The major fraction (PolA) has a molecular weight of approximately 150,000, as does the minor fraction (PolB) (150,152).Neither protein cross-reacts immunologically. PolA [Pol1 according to Chang (150)lis similar to the higher eukaryotic DNA polymerase a (153) while PolB (PolII) is similar to the prokaryotic polymerases I1 and I11 (150). Unlike other higher eukaryotes, a third low-molecular-weight polymerase (Polp) is not found in yeast (150,151). DNA polymerase activity oscillates during the cell cycle, with a maximum just before S phase (154,155). Homogenate mixing experiments from different stages of the cell cycle lead to the conclusion that small molecules are not responsible for the activity changes (154). There was no indication of changes in activity due to membrane attachment. An examination of the rate of sedimentation of DNA polymerase in sucrose gradients during the cell cycle indicates that there are no apparent changes in the size of the polymerase (155).The changes in activity are not due to degradation and resynthesis, because DNA polymerase is stable in the presence of cycloheximide and the periodic oscillations in activity continue in synchronous cultures (155). These results suggest that the activity changes are due to periodic covalent attachment of small effector molecules. The periodic variation of enzyme activity can be dissociated from DNA synthesis by X-rays, which delay DNA synthesis, but not the peak in polymerase activity. While these studies provide information about the regulation of DNA polymerase activity, it is not clear that the activity is indeed periodic. Both studies involved synchronous cultures prepared by the feed-starve synchrony method, which may cause oscillations in activity by nutrient changes, not by cell cycle regulation.

2. PLASMIDS Small circular DNAs about 2 pm in circumference are present in many strains of S . cerevisiae (119,156-158). A similar 2 p m plasmid is

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present in S. italicus (159,160)and in S . pombe (161).A survey of 70 yeast strains, including 17 different species, indicated that DNA plasmids occur only in S. cerevisiae (9 of 10 strains), S. ita2icus (2 of 3 strains), and one strain of Kluyveromyces lactis (159).The K . lactis strain, unlike the others, had two linear DNA plasmids, one of which was associated with the killer phenotype. The function of the plasmids is not clear at present. The existence of strains that lack the plasmid (158, 161) and the ability to cure the plasmids by transformation indicate that they are not essential for growth. The plasmids have been implicated in resistance to drugs affecting mitochondrial function ( 159, 162, 163). These observations suggest that the 2-pm DNA plays an informational role in mitochondrial biogenesis or in the function of mitochondrial membranes. Another possibility is that 2-pm DNA plays a role in chromosome structure. This hypothesis is based on the observation that 2-pm DNA cosediments with folded chromosomes from dividing cells, but only 10% as much cosediments with cells arrested in G 1 (164). The structure of the plasmids has been examined in detail. They are present in multiple copies, the number varying from 24 to 108 copies per cell (156, 163). They are similar in density (1.701 g/cm3) and nucleosome structure to nuclear DNA (156, 157, 165), and are 6318 base-pairs in length with two inverted repeats of 599 base-pairs situated opposite each other (158,166-168).These regions can apparently recombine, resulting in two forms depending on the orientation of the noninverted regions (169-1 7 1 ) .There are three protein-coding regions capable of coding for proteins of M , 48,600,43,200, and 33,200 (167).The second of those may encode replication functions, and the third catalyzes recombination between the inverted repeats (169). Genetic evidence suggests that 2-pm DNA has a nuclear location ( 1 72-1 74).This is consistent with the observation that replication is controlled by the same nuclear gene products as nuclear D N A (160,

175). Replication is via a single origin located adjacent to and partly including one of the inverted repeats ( 1 69,176).Replication intermediates ( 8 structures) have been observed in the electron microscope

(119, 160, 176). Replication during the cell cycle has also been examined in synchronous cultures induced by ‘‘a factor,” a mating pheromone ( 117, 185).The results indicate that synthesis is periodic with a peak in the first third of S phase that leads to a doubling in amount by the end of the cycle. Owing to its small size, a single molecule replicating bidirectionally at 1pmlmin could be replicated in 30 seconds. In theory,

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a single plasmid could be replicated many times during S phase. Density transfer experiments indicate that this is not the case (117). One generation after a density shift, nearly all the DNA was of hybrid density. This indicates that replication is semiconservative, and that each molecule is replicated once in a cell cycle. If each 2-pm DNA is replicated only once per cycle, how can transformations by plasmids containing 2-pm DNA result in such high copy numbers? The answer may be that cells take up large numbers of plasmids during transformation, or that the high copy number may be due to unequal segregation of the plasmids during mitosis resulting in one nonviable cell lacking 2-pm DNA and one viable cell with all the copies. Alternatively, the number of copies may be directly regulated. In addition to 2-pm DNA, many S. cerevisiae strains often contain two double-standard RNA plasmids of M , 3.4 X lo6 and 1.2 to 1.7 X lo6, called L and M, or P1 and P2, respectively (177-179). These RNAs are not required for growth (180).Proteins are associated with the RNA, forming a virus-like particle (181,182).However, the dsRNA is not infectious (183).The RNA of lower M, apparently encodes two proteins, a toxin (killer protein) of M , 30,000, which kills sensitive yeast strains lacking this RNA (180,182), and a second protein of M , 32,000. The larger RNA codes for the major capsid protein and two smaller proteins (181).mRNA requires nuclear genes for maintenance and killing (180,182). Replication of the plasmids requires some of the same genes as nuclear and 2-pm DNA (184).However, mutants involved in the initiation step of DNA, required for nuclear DNA synthesis, are not essential for dsRNA synthesis (184,185).When the mode of replication of the large dsRNA was examined by density-shift experiments, very little RNA of hybrid density was formed and fully light-density RNA appeared within a half-generation of the density transfer. These results indicate that replication is either conservative or semiconservative, with only a small portion of the molecules undergoing replication (186).The rate of synthesis of the L dsRNA in a size-fractionated culture apparently increases exponentially through the entire cell cycle (186), with no indications that replication is confined to or absent from any cell cycle stage. Another study, which utilized synchronous cultures prepared by both induction and selection synchrony methods, indicated that dsRNA synthesis is periodic with a peak during S phase (185).It is not clear why these different results should have been obtained, although it is possible that they reflect the different synchrony methods used and thus may be of use in establishing proper procedures for the study of periodic events.

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3. MITOCHONDRIALDNA Mitochondrial DNA exists as a circular DNA varying from a molecular weight in S . pombe of 1 3 , X lo6 (6 pm) to 40 to 50 x lo6 (25 pm) in S. cerevisiae and S . carlsbergensis (187). It can be separated from nuclear and 2-pm DNA through its lower density (188, 189). The amount of mtDNA is not tightly coupled to the amount of nuclear DNA. The mtDNA/nDNA ratio varies with media composition and growth phase (189-191). In S. cerevisiae, if protein synthesis is blocked by temperature-sensitive protein-synthesis mutants, cycloheximide, or amino-acid starvation, mtDNA synthesis continues whereas nDNA synthesis stops (192,193). A similar result occurs in afactor-blocked cells (194).This difference between mitochondrial and nuclear DNA synthesis is not universal. In S . pombe, cycloheximide inhibits both (129). Mitochondrial DNA synthesis may require some nuclear genes for replication because the genes cdc8 and 21, which block nuclear DNA synthesis, also block mtDNA (193, 195).However, mtDNA synthesis appears to be less affected than nDNA synthesis (196).Several genes involved in the initiation of nDNA synthesis are not required for mtDNA synthesis. A number of different approaches have been used to determine whether mtDNA synthesis is periodic through the cell cycle. A study of the rate of appearance of mitochondrial mutations following mutagenesis in synchronous or size fractionated cultures demonstrated a periodicity in “petite” induction. The maximum occurred during or just before S (100). Another study suggests that the mutation frequency is the same throughout the cell cycle (197). The observation that mtDNA exhibits a high degree of recombination and repair (116, 198), which is increased during S phase (1981, could explain an increased mutation frequency during S (199). In addition, there is a positive correlation between cell size and the amount of mtDNA (191).Since daughter cells tend to b e smaller than mother cells, they are more likely to become petites. Thus, size fractionation may cause a periodicity that is related to cell age, not to cell-cycle position. These experiments may better define the subtle experimental problems in this ‘field than the periodicity of mtDNA replications. Studies on the rate of synthesis of mtDNA through the cell cycle also provide conflicting results. Experiments with adenine labeling of synchronous cultures of S . pombe prepared by selection of small cells from sucrose gradients indicated that synthesis is periodic for both mtDNA and nDNA, although the maxima occurred at different times

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(200).Periodic synthesis was also observed in S . cerevisiae during S phase in size-fractionated cells, a method that should eliminate synchrony-induced perturbations as an explanation (201). Periodic synthesis after S phase has been observed in synchronous cultures of S . Zactis (202). However, continuous, nonperiodic synthesis has been reported ( 117,203). It has been suggested that methods based on labeling may not be valid (204).In synchronous cultures prepared by a modification of Williamson and Scopes feed-starve synchrony method ( l o ) ,the maximum incorporation of radioactive adenine into mitochondria1 DNA was observed during S phase. In contrast, the amount of mtDNA apparently increased continuously. A similar discrepancy between labeling rates and amount of mtDNA was observed in a size-fractionated culture (201). When an isotope-dilution analysis was performed on labeled cells, a continuous pattern was observed (204).This suggests that the observed periodicity could be an artifact of the synchrony procedure caused by selective uptake at a particular point in the cycle, or an artifact of pool metabolism. The latter explanation may be most probable because during S phase the precursor pools may be depleted, thereby artificially increasing the specific activity of the pulse-labeled pool. This could result in increased incorporation during and after S phase. This is subject to test. Traditional density-shift experiments suggest that replication is dispersive (115, 116, 205). However, a semiconservative replication pattern was observed in vitro (206).The dispersive pattern is apparently a result of the extensive recombination that occurs during S phase; it has been suggested that in vivo replication is semiconservative, but that redistribution by recombination gives rise to an apparent dispersive pattern (205).

E. RNA Synthesis Total RNA synthesis increases exponentially during the cell cycle in S . cerevisiae in both synchronous and size-fractionated cultures (23,98,207,208).Accumulation is exponential in the budding yeasts Candida albicans and Candida utilis, although a linear doubling model cannot be eliminated (18,142).Different RNA species, including ribosomal (18 S, 26 S, and 5 S) transfer (4 S) and messenger RNA, are also synthesized exponentially (207-209). One report suggests that mRNA and rRNA are synthesized in a stepwise manner during the cell cycle (210),and another report suggests the rate of synthesis is constant (211). This latter conclusion is unlikely because, unless the rate doubles at some point in the cycle, the rate per cell will be

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reduced by half after each division. In addition both conclusions are not consistent with the observed exponential increase in protein during the cycle (Section 11,F). The reasons for these experimental discrepancies are not immediately apparent. On balance, the data suggest that an exponential increase in the rate of RNA synthesis is typical of the normal cell cycle. The periodicity of RNA polymerase activity in S. cerevisiue has been examined. RNA polymerase 11, which is responsible for mRNA synthesis activity, varies periodically, whereas RNA polymerase I activity increases continuously through the cell cycle (212, 232). The rate of synthesis of several mRNAs throughout the cell cycle has been determined. The synthesis of the mRNAs for histones H2A and H2B has been studied in a-factor-synchronized and size-fractionated cultures, using cloned histone genes for hybridization assays (213). Both histone mRNAs exhibit periodic synthesis with a maximum prior to S phase. This result is in agreement with the observed periodic synthesis of histone proteins (214,215).In contrast, approximately 100 other mRNAs coding for basic proteins appear to be synthesized continuously (213).Translation in vitro of total mRNA from different stages of the cell cycle followed by two-dimensional gel electrophoresis indicates that most of the resolved proteins (400-500), and therefore their mRNAs, were present in approximately constant relative amounts throughout the cell cycle (216).Because of the short half-life for most mRNAs (217), it is unlikely that many are synthesized periodically and yet are present continuously. Individual tRNAs have also been examined during the cell cycle. Of thirty-three different tRNAs examined in a size-fractionated culture, none showed evidence of periodic synthesis; all appeared to increase exponentially

(218). The situation in S. pombe appears to be different. In synchronous cultures prepared by zonal centrifugation and elutriation, and in single cells, total RNA increases continuously during the cell cycle ( 4 7 , 219-221 ). However, an analysis of the rate of synthesis indicated that it increases in a stepwise manner with a linear doubling mode of increase. There is no sharp transition from the lower rate to the higher rate (220).Some results suggest that the time of the rate doubling is correlated with the attainment of a particular size, and that this “size control” is important in the coordination between the rate of growth and the rate of cell division (210, 222, 223). These conclusions were based on the observation that mutants of different average cell size exhibit rate steps at times in the cell cycle that correlate with cell size, not with cell-cycle position. A more recent study suggests that in

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synchronous cultures prepared by elutriation, where perturbations induced by the gradient technique used previously are eliminated, no correlation between the time of the step and size exists ( 1 1 ) . Instead, the step showed a strong correlation with cell-cycle position that is in agreement with the results on protein synthesis (Section 11,F). The earlier result may be an artifact due to the synchrony method. RNA synthesis and cell division appear to be coordinated by changes in rate at specified points in the cell cycle.

F. Proteins and Enzymic Activities In budding yeast, total protein increases exponentially during the cell cycle (27, 86, 98, 224-226). The early work was not accurate enough to distinguish between an exponential and linear doubling mode of increase, but later rate measurements confirmed the exponential increase. This result is consistent with the observed exponential increases in ribosomal protein and ribosomal RNA (207,209,227). Under the same conditions, the fraction of total ribosomes actively transcribing mRNA is constant through the cell cycle (Fig. 4)which is consistent with exponential protein synthesis. The amount of protein also increases continuously in S . pombe, with an increasing rate through the cell cycle (16,20,39,47).These results are not accurate enough to distinguish between linear and exponential synthesis. Recently, a detailed analysis of the rate of protein synthesis in nonperturbed synchronous cultures prepared by centrifugal elutriation indicated that the rate increases for most of the cycle with a slowing of the increase starting at about midcycle, reaching a constant rate by the end. This was followed by an abrupt increase in rate at the time of cell-plate formation (aceeleration point), which resulted in a step-like pattern during the cell cycle (228).Analysis of the rate pattern indicated the accumulation was neither linear with a doubling in rate nor exponential but somewhere in between. Asynchronous control cultures subject to the same separating procedures did not show this pattern, so it is unlikely that these events are artifacts of the synchronizing procedure. The timing of the acceleration point does not appear to be regulated by a size control (Section 111) but instead is associated with cell cycle position, because in mutants that divide at different sizes the timing in the cell cycle is the same (228). Many biologists have suggested that the continuous increases in total protein content may not necessarily mean that individual proteins and enzymes increase in amount in the same way. Individual proteins could be synthesized periodically and total protein could

,

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75

FLOW R A T E

1,3

-

*

I:

.

l,9

l,7

*

.

*

2:

2.3

.

2,5

*

w Lo

z

si 2

50

-

2 II

25

-

0

0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9

1.0

C E L L C Y C L E POSITION

FIG.4. Percentage of ribosomes present as polysomes during the cell cycle of Sacchnromyces cereoisiae. Cells were fractionated according to size in an elutriator rotor at 4°C in the presence of cycloheximide. Polysomes were then centrifuged through a sucrose gradient. The ratio of ribosomes present as monosomes versus polysomes is plotted vs cell cycle position.

increase continuously if the synthesis of different proteins was dispersed throughout the cell cycle. In 1959, periodicities were reported in the activities of peptidase and catheptic activities in synchronous cultures of S. cerevisiae (229).This set off a flurry of research on enzyme activities through the cell cycle in yeast and other organisms, with similar results. In general, most of the enzymes showed either stepwise or periodic increases and decreases ( 1 , 230, 231). By 1971, all of28 enzymes examined in S. cerevisiae, 5 of 9 in S. pombe, and 7 of 7 in other yeasts were periodic (230).Steps were observed for some of these enzymes, including RNA polymerase 11, in cultures fractionated by size in a zonal rotor (41,112,212,231,232),where perturbing effects should be eliminated. The only exception found at that time was in RNA polymerase I activity, which was continuous under these conditions (232).The activity of P-galactosidase, determined by a fluorescence assay on single cells in balanced asynchronous culture, was also periodic in S. Zactis (40). A number of models have been advanced to explain the periodic enzyme behavior including oscillatory repression and linear tran-

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scription (208,231,233).All the reports on enzyme periodicities have been based on measurements of enzyme activity. Two basic assumptions in the above models are that enzyme activity and enzyme synthesis are tightly coordinated, and that regulation is at the level of transcription. Stated differently, the final activity is due to the periodic synthesis of an unstable mRNA that is faithfully translated into protein, the amount of this determining the final activity. While in general many mRNAs are unstable with short half-lives (on the order of 20 to 40 minutes) (217,234),some mRNAs are considerably more stable (217).In addition, there is a lag between inhibition of protein synthesis and inhibition of enzyme activity (234). Studies on DNA polymerase show that oscillations in activity continue in the absence of protein synthesis (155).These results indicate that enzyme activity and enzyme synthesis are not necessarily coupled through the cell cycle. Some of the experiments have been criticized because it was rarely demonstrated that the periodicities were not induced by the synchronizing procedure. When 19 enzymes, many of which were thought to be periodic, were examined with careful controls in S. pombe, very few showed clear indications of periodicity (2 8 ).In addition, some of the asynchronous controls for the selection synchrony technique indicated that perturbations that resembled the periodicities in synchronous culture could be induced (2 8 ).Similar perturbations in enzyme activity were observed in asynchronous control cultures of several different yeasts including C. albicans (2 8 ),S. pombe (32),and S. Zactis (Fig. 5 ) . More recently, the activities of P-galactosidase in S. lactis, of acid phosphatase and a-glucosidase in S. cerevisiae, and of acid phosphatase in S. pombe were analyzed in synchronous cultures, where perturbations were absent. All showed continuous increases (235).A similar result was obtained for a-glucosidase in size-fractionated S. cerevisiae (236). The rate of synthesis of many proteins during the cell cycle has been determined. In some cases, these allow a comparison between enzyme activity and enzyme synthesis. In S. pombe, total ribosomal protein showed step synthesis (237),as did a pH-2.1-extractable protein (238).Similar steps in acid-extractable proteins were observed in Kluyveromyces fragilis (239,240).In S . pombe and S . cerevisiae, protein bands resolved in dodecyl sulfate on one-dimensional gels were also synthesized continuously (241-243). In all these studies, proteins may not have been sufficiently resolved to eliminate the possibility that periodicities were obliterated by the multiplicity of proteins in the fractions and gel bands examined. For this reason, a study of the synthesis of individual proteins from S. cerevisiae resolved on

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0.5

> k

20.1 I-

0

a

0.05

0

1

2

3

4

HOURS

FIG. 5. A perturbed asynchronous control culture of Saccharomyces lactis. Cells growing in 0.5%yeast extract plus 2%lactose were collected in the separation chamber of an elutriator rotor. After 15 minutes, they were flushed out and allowed to grow asynchronously.P-Galactosidase activity was measured b y a colorimetric method with p-nitrophenol-P-D-galactopyranoside as substrate.

two-dimensional gels was undertaken. Cells in exponential growth were pulse-labeled, then size-fractionated on an elutriator rotor. Over 200 different proteins were assayed in detail, including 41 ribosomal proteins, 8 basic proteins, and over 150 other cellular proteins (98, 216, 227, 236, 244). The polypeptides from several enzymes were identified on the gels, including a-glucosidase, which shows periodic variations in activity under some conditions. All the proteins showed exponential synthesis during the cell cycle. The only exception observed was for histone proteins, which were synthesized periodically with a peak at the beginning of S phase (216).This result is in agreement with an earlier report that utilized the feed-starve synchrony technique (214),and with the observed periodic synthesis of histone mRNA (213).An exhaustive examination of about 900 proteins identified eight others that appear to be synthesized periodically throughout the cell cycle (272).These results suggest that periodic synthesis is rare for proteins, and suggest that the exceptional periodic proteins play a direct role in the cell division process. Exponential synthesis of most proteins during the cell cycle has significant implications with regard to cell-cycle regulation. If cell-

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AND CALVIN

s. MCLAUGHLIN

cycle proteins are synthesized continuously, regulation could be modulated through one or more of the following: periodic changes in activity, periodic degradation, or periodic assembly processes that use size controls to trigger periodic events in the cell cycle. Protein degradation is very low in yeast. The average half-life is 18 days in cells dividing every 90 minutes (245).Proteins associated with DNA synthesis are stable ( 1 2 ) , as are the gene products associated with most cdc mutants (246).Protein degradation was examined during the cell cycle by two-dimensional gel electrophoresis on a sizefractionated culture; only 2 of 111 cellular proteins showed apparent periodic increases in degradation, and none showed changes in size or isoelectric point during the cell cycle (244).A survey of 900 proteins identified nine that may be periodically synthesized, modified, or degraded. Five may be related to each other (272). Additional studies are required to determine if the two proteins we identified in earlier studies (244) are a subset of the nine proteins recently identified (272).These results establish that periodic degradation or modification of cellular proteins is rare. This suggests that these rare, periodically degraded proteins may be important components of the cell division process. In fact, a labile protein is required for the initiation of the cell cycle (247),and protease activation of a zymogen has been proposed as a model for chitin synthesis in bud morphogenesis ( 7 6 ) . Protein synthesis is also required to initiate the cell cycle (Section II,D), which suggests that a specific protein or class of proteins is not present in excess and must be synthesized prior to cell cycle initiation. It was demonstrated by an indirect method that a thermolabile protein(s) is required for a function associated with mitosis and that this protein(s) is accumulated during the cell cycle. When cells are heated or pulsed with UV or cycloheximide at different stages in the cell cycle, cell division is delayed, but the delay at some stages of the cycle is longer than the length of the pulse. The delay increases during the cycle to a maximum during mitosis (17,248).This phenomenon is called “set-back” and is apparently the result of inactivation of a labile protein. A certain amount of this protein is required to perform its function; when it is inactivated, it must be resynthesized. Cells late in the cell cycle accumulate more than cells early in the cycle, and both kinds of cells are “set back” to the same point in the cycle when the accumulated protein is inactivated. Other results suggest that many proteins involved in the cell cycle are present in excess or are synthesized continuously. Almost all S. cerevisiae cdc mutants are able to traverse several cell cycles in the absence of new cdc gene-

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product synthesis (246). This indicates that the gene products are present in excess, and it is unlikely that periodic synthesis of any of them would affect their action. These proteins may be subject to periodic activations, participate in periodic assembly processes, or simply be passive elements in the cell-division process. Consistent with the continuous exponential synthesis of the vast majority of cellular proteins is the observation that in S. cereuisiae, cdc mutants continue to make most cellular proteins at the nonpermissive temperature (220).In S. pombe, there are no qualitative differences on two-dimensional O’Farrell gels between arrested mutants and wild-type cells (249).Therefore, most proteins are synthesized independently of cell-cycle controls.

111. Control of Cell Division

The results discussed above indicate that most cellular components accumulate continuously under a variety of growth conditions during the cell cycle. Dividing cells in balanced growth maintain a constant average cell size. However, when the growth rate changes due to changes in the environment, the division time must also adjust to maintain growth competence. The result of changes in growth rate is usually an equal change in the rate of division. As growth slows, “phase expansion” takes place. In both budding and fission yeast, most of the increase in cell-cycle length at low growth-rates is due to an increase in the G1 phase ( 1 , 2 , 112). If the growth rate is reduced far enough, cells stop in a resting state, called GO, distinct from G1 (250, 251). Transition from this resting state to proliferation is regulated at a point termed “start” ( 2 ) , which spans the GUS boundary. After traversal of this point, cells are irreversibly committed to cell division. If some protein synthesis occurs, cells can continue to division, with little net growth (251, 252). After completion of the cycle, these cells then arrest in G1 as small cells. This indicates that growth is not required after a cell cycle is initiated, but is required for initiation. Some attempts have been made to determine what component cells monitor to determine whether they should undergo a cell cycle or arrest in G1. All the intermediates in the sulfate assimilation pathway, including methionine, are required to prevent G1 arrest (253). Therefore, the signal is farther down the pathway than formation of amino acids. I n a temperature-sensitive methionyl-tRNA-synthetase mutant at the restrictive temperature, G1 arrest occurs even when there is methionine in the cells (254).This suggests that the G1 arrest

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STEVEN G . ELLIOTT AND CALVIN S. MCLAUGHLIN

signal is a deficiency of charged tRNA or a product made from it, such as a specific protein. The G1 arrest signal may be distinct from the signal that tells cells to start. This suggestion comes from mutant studies indicating that G1 arrest may be regulated by different gene products than is initiation of a cell cycle (255, 256). Several temperature-sensitive mutants defective in the initiation of the cell cycle have been isolated (2,257,258). These are characterized by continued growth and traversal of the cycle, after which they arrest as unbudded cells in a G1 state. These cells continue to grow even when they arrest, which suggests that there is no cell-cycle control over growth. The biological basis for start is not known, although there are some indications it may involve the construction of a spindle plaque (2,259, 260). In S . pombe, in addition to a control over the GUS boundary, there is a second control at the G2fM boundary (19, 105, 107, 108, 261). Mutants defective in the regulation over both steps have been isolated (261-263). Genes defective in start in S. pombe act in a similar way to those in S. cereuisiae, i.e., G1 arrest, but continued growth. However, some of those defective in the G2/M regulatory step can divide normally, but they are abnormally small. This result has been interpreted to mean that these gene products act to delay mitosis until a certain cell size is reached, after which mitosis occurs. There is also one gene product, cdc2, that acts at both the GlfS boundary and the G2fM boundary; some alleles can cause G1 or G2 arrest, and others are small

(263). Two basic models have been proposed to explain how cells coordinate growth and division rates. The first is a probabilistic model called “transition probability.” It was first proposed to explain the variation in intermitotic times in mammalian cells (264) and later applied to yeast (265, 266). In its simplest form the model is as follows. Cells exist in two states, A and B. A is a resting state equivalent to GI, and B is a proliferating state equivalent to S, G2, and M. Cells passing from A to B are irreversibly committed to completing B and returning to A. However, the transition from A to B is the rate-limiting step in cell division, and is random. A cell has a certain probability of leaving A; this depends on environment. The rate of traversal from A to B therefore follows first-order kinetics. This model is supported by an array of data suggesting that there is variation in the time required for cells to enter S phase after release from blocks in G1, and that the rate of entry approximates first-order kinetics (265, 266). The second model (size control) is a deterministic one, which states that transition to proliferation is not random. Cells can monitor

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their instantaneous cell size, and at a certain threshold size, the cell cycle is initiated. Several mechanisms to monitor cell size have been proposed (1,267). The practical result of the model is that cells coordinate growth and division by altering, delaying, or accelerating cycle time. If the cells are small, the size control causes a delay until growth catches up. If the cells are too large, the cells immediately pass the control point and proceed to division, where size is reduced by half, which tends to correct the inbalance. Convincing data have been presented in support of the model. In S. cerevisiae, there is a strong correlation between cell size and bud emergence or the start step as defined by start mutant or a-factor arrest points. Second, daughter cells, which are smaller than mother cells at division, take longer to initiate their cell cycle ( 2 , 112). Further evidence comes from studies on S. pombe. Oversized cells divide faster than smaller cells, and smaller cells grow proportionately more in size in one cycle than larger cells (7,19,48,106,108).Mutants presumably defective in this control have been isolated (261-263). However, unlike S . cerevisiae, two size controls may be at play in S. pombe, one at the G2/M boundary and one at the GUS boundary (19,106,108). Neither model can explain all the data (268).Therefore, composites of the two models have been proposed, such as the tandem model where cells grow to a certain cell size and then traverse start in a probabilistic manner. A better fit of the data is obtained by the “sloppy size control” model (268-270).The model includes a size control but suggests that there is variation in the traversal of start such that the probability of initiating a cell cycle is greater for large cells than small ones. A variation of this model is called the “G1 rate model” (271), which states that a size control governs traversal of start, but variations in rates of metabolic reactions cause variations in start and other essential steps in the cell cycle. Definitive proof for the various models awaits the molecular basis of the control of cell division. Cloning of cell-cycle genes identified from mutant studies or from studies on periodic events, especially periodically synthesized or modified proteins, and a detailed analysis of the mechanism of action of such gene products, will go a long way toward achieving a biochemistry of cell division.

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J. R. Ludwig and C. S. McLaughlin, in press. J. R. Ludwig, Ph.D. Thesis, University of California, Irvine, 1981. L. Chia and C. S. McLaughlin, Mol. Gen. Genet. 170, 137 (1979). P. Bower, Personal communication. J. M. Mitchison, J. Cummins, P. R. Gross, and J. Creanor, Erp. Cell Res. 57, 411 (1969). S. G. Elliott, unpublished. J. M. Mitchison and P. Walker, E r p . Cell Res. 16,49 (1959). R. S. S. Fraser and P. Nurse, Nature (London) 271, 726 (1978). R. S. S. Fraser and P. Nurse, ]. Cell Sci. 35, 25 (1979). H. 0. Halvorson, J. Gorman, P. Tauro, R. Epstein, and M. Le Berge, F P 23, 100 (1964). H. Hilz and H. Eckstein, Bchern 7, 351 (1964). P. Nurse and A. Wiemken,j. Bact. 117, 1108 (1974). S. G. Elliott, J. R. Warner, and C. S. McLaughlin,]. Bact. 137, 1048 (1979). J. Creanor and J. Mitchison,]. Cell. Sci., in press. B. Sylven, C. A. Tobias, H. Malmgren, R. Ottoson, and B. Thorell, Erp. Cell Res. 16,75 (1959). H. 0 .Halvorson, B. L. A. Carter, and P. Tauro, Adu. Microbiol. Phys. 6,47 (1971). H. 0. Halvorson, in “Cell Differentiation in Microorganisms, Higher Plants and Animals.” Fischer, Jena, 1976. B. L. A. Carter and I. W. Dawes, E x p . Cell Res. 92,253 (1975). P. Tauro, H. 0. Halvorson, and R. Epstein, PNAS 59,277 (1968). J. Creanor, J. W. May, and J. M. Mitchison, EJB 60,487 (1975). J. Creanor, S. G. Elliott, and J. M. Mitchison, in press. J. R. Ludwig, J. J. Foy, S. G. Elliott, and C. S. McLaughlin, Mol. Cell. Biol. 2, 117 (1982). W. H. Wain and W. D. Staatz, Erp. Cell Res. 81,269 (1973). J. H. Duffus, BBA 228,627 (1971). J. H. Duffus and C. S. Penman, J. Gen. Microbiol. 79, 189 (1973). C. S. Penman and J. H. Duffus, Z. Allg. Mikrobiol. 16, 361 (1976). W. H. Wain, E x p . Cell Res. 69, 49 (1971). R. W. Shulman, L. H. Hartwell, and J. R. Warner,JMB 73,513 (1973). D. P. Dickinson, Ph.D. Thesis, University of Edinburgh, 1979. S. G. Elliott and C. S. McLaughlin, J. Bact. 137, 1185 (1979). H. 0. Halvorson, BBA 27, 267 (1958). B. Byers and L. Sowder, J . Cell Biol. 87,70 (1980). B. Shilo, V. G. H. Riddle, and A. B. Pardee, Exp. Cell Res. 123,221 (1949). J.G. Bullock and W. T. Coakley, Erp. Cell Res. 103,447 (1976). D. P. Dickinson, J. Cell Sci. 51, 203 (1981). R. Pinon, Chromosome 70,337 (1979). C. P. Milne, Masters Thesis, University of Washington, Seattle, 1972. J. C. Johnston, R. A. Singer, and E. S. McFarlane, J . Bact. 132,723 (1977). M. W. Unger and L. H. Hartwell, PNAS 73, 1664 (1976). M. W. Unger,]. Bact. 130, 11 (1977). B. L. A. Carter and P. Sudbery, Genetics 96,561 (1980). P. Sudbery, A. R. Goodey, and B. L. A. Carter, Nature (London) 288,401 (1980). S. I. Reed, Genetics 95, 561 (1980). L. H. Hartwell,]. Cell Biol. 85, 811 (1980). B. Byers and L. Goetsch, CSHSQB 38, 123 (1973).

176

STEVEN G. ELLIOTT AND CALVIN S. MCLAUGHLIN

260. 261. 262. 263. 264. 265. 266. 267.

B. Byers and L. Goetsch, J. Bact. 124, 511 (1975). P. Fantes and P. Nurse, E x p . Cell Res. 115, 317 (1978). P. Nurse and P. Thuriaux, Genetics 96,627 (1980). P. Nurse and Y. Bisset, Nature (London)292,558 (1981). J. A. Smith and L. Martin, PNAS 70, 1263 (1973). B. Shilo, V. Shilo, and G. Simchen, Nature (London)264,767 (1976). B. Shilo, G . Simchen, and A. B. Pardee, J . Cell. Physiol. 97, 177 (1978). P. Fantes, W. D. Grant, R. H. Pritchard, P. E. Sudbery, and A. E. Wheals,J. Theor. Biol. 50,213 (1975). P. G . Lord and A. W. Wheals,J. Cell Sci. 50,301 (1981). L. Alberghina and L. Mariani, in “Biomathematics and Cell Kinetics Developments in Cell Biology” (A. J. Valleron and P. M. MacDonald, eds.), pp. 89-102. Elsevier/North Holland Biomedical Press, New York, 1978. L. Alberghina, L. Mariani, and R. Zippel, Differentiation 15, 135 (1979). L. N. Castor, Nature (London)287,857 (1980). A. T. Lorinez, M. J. Miller, N. Xuong, and E. P. Geiduschek, Mol. Cell B i d . 2, 1532 (1982).

268. 269. 270. 271. 272.

Prokaryotic and Eukaryotic 5 S RNAs: Primary Sequences and Proposed Secondary Structures’

I

I

RAM P. SINGHAL AND JONI K. SHAW Department of Chemistry, Wichita State Uniuersity, Wichita, Kansas

I. Structure of Prokaryotic 5 S RNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Homology in Primary Structures.. . . . . . . . . . . ............. B. Invariant Segments of the Structure. . . . . . . . . ............. C. The Base-Paired Region ......................... 11. Structure of Eukaryotic 5 S A. Homology in Primary S B. Invariant Segments of the Structure. ................... C. The Base-Paired Regions.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Conformation of Prokaryotic 5 S RNA . . . . . . . . . . . . . . . . . . . . . . . . . . A. Reactivity for the Chemicals. . . . .... .... B. Exchangeable Protons . . . . . . . . . .................... C. Reactivity with Various Nucleases . . . . . . . . . . . . . . . . . . . . . . . . . . D. Binding with Complementary Oligonucleotides. . . . . . . E. Reconstitution of Ribosomes from 5 S RNA . . . . . . . . . . F. Physical Measurements.. . . .......................... IV. Summary.. . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix: Sequences of 5 S RNAs .................. A. Notes . . . . . . . . . . . . . . . . . . . B. References and Footnotes for Sequence List . . . . . . . . . . . . . . . . . C. 5 S Sequences . . . . . . . . . . . . . . . . . . Prokaryotes . . . . . . . . . . . . . . . . . . . . Eukaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chloroplasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ............ Archaebacterial . . . . . . . . . . . . . . . . . . . . . . Addendum: References to 32 Additional Sequences. . . . . . . . . . . . .

178 178 178 183

185 190 191 193 195 195 196 197 198 198

206 208 208 208 251

Although the structure of at least one 5 S RNA has been known since 1967 (1), it is only recently that the sequences of the 5 S RNAs The authors plan to update the 5 S RNA sequence list (Appendix) periodically. Researchers are requested to send new 5 S (and 5.8 S) RNA sequences and changes in the known structures to the first author (R. P. S.). 177 Progress in Nucleic Acid Research and Molecular Biology, Vol. 28

Copyright 0 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-5400284

RAM P. SINGHAL AND JON1 K. SHAW

178

from a wide variety of sources has become available. Excellent review articles on the structures and function of 5 S (and 5.8 S) RNAs have been published (2-4). In this paper, we list all the 5 S RNA sequences known, excluding those of the “precursor” molecules (see the Appendix). The secondary structures of both prokaryotic and eukaryotic 5 S RNAs are postulated from their primary sequences. Chemical and enzymic reactivities and conformational studies are discussed here with regard to the proposed secondary structure. For other sources of 5 S (and 5.8 S) sequences, the reader is referred to other publications in which residues are numbered sequentially without “subpositions” (see below) (5-7).

1. Structure of Prokaryotic 5 S RNAs

A. Homology in Primary Structures Mature 5 S RNA in prokaryotes is derived by posttranscriptional processing of the direct transcript. Direct comparisons of the sequences from different bacteria indicate that each structure has a similar chain length and contains no minor or modified nucleotides. To characterize invariant purine and pyrimidine nucleotides in various positions of the 5 S RNA structure, 29 prokaryotic sequences available at the time of this study were tabulated in a manner to yield a most common secondary structure and to maximize regions of homology (see sequences P1 to P29, Appendix). Thus, 119 nucleotide “positions” and 12 “subpositions” (10 : 1and 10 :2; 16 : 1; 29 : 1;33 : 1; 70 : 1 and 70 :2; 54 : 1; 86 : 1; 101 : 1; 109 : 1; 108 :2) were established. Although a residue in each subposition was not found for each 5 S RNA sequence, inclusion of the subpositions in the sequential numbering produced constant helical segments and invariant residues. The nature and the frequency with which a given nucleotide occurs in a nucleotide position of the 5 S RNA structure are listed in Table I. The data indicate the presence of 29 “invariant” nucleotides (“conserved positions”) in more than 27 sequences. In addition, 10 positions were found to have one specific nucleotide in more than 24 (out of 29) sequences. Among 26 positions, in one position G or U, in six positions C or G, in eight positions C or U (pyrimidine, Y), and in eleven positions A or G (purine, R) residues appeared in most sequences analyzed.

B. Invariant Segments of the Structure From this analysis and sequence comparison there appear two kinds of structural conservations, one in which specific residues have

PROKARYOTICAND EUKARYOTIC 5

s RNAs

179

TABLE I NATUREOF RESIDUES IN 29 PROMYOTIC5 S RNAs" Nature of the conserved nucleotideb

A

G

C

U

1 1 1 5 1 1 5

3 13

2 9 10 9 4 2 1 20 10 4 1 10 8 1

23

3

3 24

22 1 1

-

1 7 11 6 11 1 11 8 17

-

9 15 13 26 10 13 21 1 1 3 1

7 28

1 25 6 12 22 5 29 27

-

-

-

27

2

-

12

-

3 1 1

-

-

-

28 3

-

1

-

-

27 1 2 10 2

1

-

17 10 29

29

28 15 1

-

8 26 27 28

5 17 6 9 14 1 4 6 1

2 3 2 28 -

1 1 6 4 1 12

-

1 18 4

-

1 1 14 14 1 18 2

-

1

Nucleotide position

Specific nucleotide

Group-specific nucleotide

1 2" 3" 4 5 6 7 8 9 10 10: 1 10:2 1I d 12d 13 14 15 16 16: 1 17 18 19 20 21 22 23 24 25 26 27 28 29 29: 1 30 31 32 33 33: 1 34 35 36 37 (continued)

180

RAM P. SINGHAL AND JON1 K. SHAW

TABLE I (Continued) ~~

~

~~~

Nature of the conserved nucleotideb A

G

C

28

-

-

7

1 13

29 29 -

29 -

-

-

-

-

29 3 28

15

-

14

1 15 29

26 25

1

-

29 29 13

-

1 1 3 6 1 25 2

-

3 28 1 2 1 1

6 28 29

-

2 29 26 13

-

16 1 11

3 9 2

26

-

-

1 1

2

-

28 5 21 11 7 13 1

28

29 1 1

-

26 1

-

-

26 1 27

27 26 2 2

1 1

-

U

Nucleotide position 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 70: 1 70 : 2 71 72 73 74 74 : 1 75

Specific nucleotide

Group-specific nucleotide

PROKARYOTIC AND EUKARYOTIC

5

181

s RNAS

TABLE I (Continued) Nature of the conserved nucleotideb

-

A

G

C

U

28 1 -

23 15 5 22 24 25 25 8 3 3 5 6 8

1 5 3

28 1

-

1 3 3 1 1 2

3 4 5

1

-

10 -

4 29 2 26 2 3

26 1

-

13 8 9

-

3 3

-

3 2 2 3 1 28 6 22

-

27 3 24

5 2 2 1 3 5 2 5 12 2 20 24 24 22 14 3

-

14 1

-

1 27 26 9 1 2

1 2 1 2 5 1

3 12 11 3

24 5 4 18

-

-

-

-

26 4 18 1 2 1

-

18 8 16 1 2 2 5 2 25 1 9 1

-

1 1 27 2 1 7

-

1 1 8 11 8

Nucleotide position

Specific nucleotide

Group-specific nucleotide

76 77 78 79 80 81 82" 83 84 85 86 86: 1 87 88 89 90 91 92 93 94 95 96 97 98 99 100

101 101 : 1 102 103 104 105 106 107 108 108: 1 108 : 2 109 110 111 112 (continued)

RAM P. SINGHAL AND JON1 K. SHAW

182

TABLE I (Continued) Nature of the conserved nucleotideb ~~

A

G

C

U

1 9 3 14 4 11 1

15 4 11 14 10 14

13 5 9

-

-

-

11 2 13

2 1 14

-

1 5

Nucleotide position

Specific nucleotide

Group-specific nucleotide

113 114 115 1 16c 117" 118 119

-

C or G

-

-

-

R

-

Y

a See the Appendix for the list and origins of 5 S RNA sequences used for the study (sequences P1 to P29). Residues in parentheses indicate the nature of the predominant nucleotide in the 29 RNAs examined. A residue in this position in one of the sequences was not characterized. In E . coli, two different strains were analyzed. Strain CA 265 contains A in place of C in position 11, and strain MRE 600 contains U in place of G in position 12.

*

been conserved in specific positions of the structure?and one in which several nucleotides in a sequence have been conserved (8).One region of the homology contains the sequence of residues from 1 to 10 that pairs with the sequence of residues from 110 to 119. Although the sequence is not highly conserved? pairing between the residues of two sections is maintained in most structures. This section, which contains both the beginning and the end of the molecule, contains predominantly G C pairs. Another polynucleotide segment of 13 nucleotides from residue 17 to 29 is conserved without any deletion or insertion of the nucleotides. Several conserved segments?for example residues 30 to 33 and residues 47 to 50, exhibit potential sites for basepairing in most sequences examined. The longest conserved section of the molecule appears to include residues 34 to 70 with very little change from one sequence to another (see the Appendix). Within this region of homology, the hexanucleotide -C-C-G-A-A-C- (residues 41 to 46) is perhaps the longest conserved segment of the 5 S RNA structure. Although there are other sections that contain a larger number of invariant residues, this particularly long section is unique in being conserved in every prokaryote examined without deletion or addition of a single nucleotide. For example, the nanonucleotide -R-A-U-R-GU-A-G-Y- (residues 71 to 79) is conserved, but with occasional

P R O W Y O T I C AND EUKARYOTIC

5

s RNAS

183

changes in the kind of purine or pyrimidine nucleotides in various sequences. The segment of 16 nucleotides from residues 83 to 96 appears to contain the largest number of G and C residues. This is followed by a decanucleotide (residues 99 to 109) that, though not conserved, contains two smaller conserved sections within itself, namely -A-G-A-(residues 99, 100, and 101)and -U-A-G-G- (residues 103 through 106).

C. The Base-Paired Regions There are four sections in the molecule that exhibit potential sites for hydrogen bonding, thus probably producing four stem regions in the structure. ( a ) Stem 1:As mentioned earlier, tail and end regions of the molecule, residues 1 to 10 pairing with residues 108 to 119. ( b ) Stem 2: Residues 15 to 22 pairing with residues 67 to 59; these segments contain predominantly G and C residues, which form more stable base-pairs. (c) Stem 3: Two base-pairing segments reside within a section of the residues 27 to 33 pairing with residues 55 to 47, respectively. Another region of four pairs is conserved in most sequences, residues 29 to 33 pairing with residues 53 to 47, respectively. ( d ) Stem 4: Residues 76 to 85 pair with residues 99 to 90, respectively. The probable base-pairings in each stem region of the structure are listed in Table 11. It appears that most sequences permit base-pairs in each stem region, with G * C base-pairs outnumbering all other types of pairing for each stem. The A . U type of base-pairs appear to be the next in frequency of occurrence. The stem 4 region can exist only if one admits a greater majority of other kinds of base-pairs. For example, the pairing of residues 79,80, and 81 with 96,95, and 94, respectively, involves other than the usual G . C and A . U base-pairs. Numerous nucleotide positions in the 119-chain structure exhibit predominance for purine or pyrimidine nucleotides as a class (in the Appendix, compare the nature of residues located in positions 3,7, 12 to 17,20,22,23,27 to 32,34 to 39,41 to 46,48 to 59,61,66 to 79,83 to 85,90 to 93,95,96,99 to 106, 109, 113, 116, 119).The homologies in the sequences of prokaryotic 5 S RNAs support the concept of evolutionary conservation. They support the theory of point mutations, especially for analogous bases. The comparison of the sequences indicates the presence of conserved and semiconserved residues in the structure. Moreover, large sections, especially residues 35 to 70 and 71 to 85, are conserved in most species with only very few deletions or additions of nucleotides and with a high number of invariant residues.

TABLE I1 PROBABLE BASE-PAIRED REGIONSOF PROKARYOTIC 5 S RNAs Nature of the base-pair* Region of base-paired residues"

G .C

A .U

Stem 1 1 . 118

4

10

21 10 18 19 14 20 14 14 24

4 14 8 10 1 1 8 2 1

28 26 15 14 13 21 4 29

1 2 13 14 7 12 -

U . U(1), A . A(l ), C . A ( l ) G . U( 12) u ' C(1), c ' C(1) G . U(l), G G(1) G ' U(1) G . U(12), C . C(l)

13 2 29 28 15 29 3

26

C . U( 16) A . C(l)

14 26

u . C(1)

28 1

A . C(l) G . A( l),A . A(3), A . G(25) G . U(9), A . G(1) G . U(25), U . U(1) G . U(14), U . U(3), U . C(l) G . G(l), C . U(9), C . C(2) G . U(l), G . G(1), U . U(l) A . C(l), A . G(1) G . U(2), A . G(1), U . C(l) G . U(1), G . G(3), A . G(l)

2 . 117 3 . 116 4 . 115 5 . 114 6 . 113 7 . 112 8 . 111 9 . 110 1 0 . 109 Stem 2 15 . 6 7 16 . 66 1 7 . 64 18 . 6 3 19.62 20.61 21 . 6 0 22 ' 59 Stem 3 27 .55 28 . 5 4 29 ' 53 30 . 5 0 31 . 4 9 32 . 4 8 33 . 4 7 Stem 4 76 ' 99 77 ' 9 8 78 ' 97 79 ' 96 80 . 9 5 81 . 94 82 . 93 83 .92 84.91 85 * 90

19 3 1 6 22 25 25 24

-

-

11 3 2

-

Other pairings noted

G . U(11), U . U(1), U . C(1), A A(1) G . U(1), A . G(l) G ' U(4) G . U(2) G U(13), C . C(l) G . U(8) G . U(7) G . U(11), G . G(l) A . C(2), U . C(l)

-

I

-

4 When a residue is characterized only as a purine (R) or a pyrimidine (Y), or else no residue is present in a particular position (deletion), the nature of possible base-pairing for that particular residue in the 5 S RNA structure cannot be stated. Hence, the sum of the different types of base-pairs having such residues (R, Y)does not equal 29 (the total number of sequences examined). b The nature of the base-pairing was determined for 29 prokaryotic structures (see sequences P1 to P29, Appendix). The numbers in the G . C and A . U columns and the numbers in parentheses appearing after the other base-pairs indicate the number of occurrences of that particular type of base-pair in the 5 S RNA sequences.

PROKARYOTIC AND EUKARYOTIC5

s RNAs

185

From these considerations and other studies of the prokaryotic 5 S RNA (see Section IV), a secondary structure for this molecule is proposed in Fig. 1. The structure suggests that certain sections of the structure may be responsible for specific functions, such as interaction of 5 S RNA with proteins and other RNAs.

II. Structure of Eukaryotic 5 S RNAs A. Homology in Primary Structures In this study of the structures of eukaryotic 5 S RNAs, those derived from mitochondria and chloroplasts were not included. To characterize the invariant residues, 35 eukaryotic 5 S RNAs whose structures were available were arranged in a manner yielding the maximum sequence homology. This resulted in the establishment of 119 nucleotide positions and eight subpositions (10 : 2, 46 : 1, 70 : 1, 73: 1, 74 : 1, 86: 1, 101 : 1, and 119 : 1). The subposition 70: 1 of prokaryote 5 S RNA was occupied in each eukaryote 5 S RNA examined. The frequency of occurrence of nucleotides in various positions of the 35 structures examined is shown in Table 111. The results indicate the presence of “invariant” or “conserved” nucleotides in 41 positions in over 32 sequences (i.e., with greater than 90% probability): 13 A, 12 G, 10 C, and 6 U residues. Another 19 “semivariant” residues can be located in over 24 sequences (i.e., with greater than 70% probability): 1 A, 7 G, 5 C, and 6 U residues. There appear to be more C and G residues than A and U residues in variant and semivariant positions (34 C and G vs 26 A and U). Five positions have either C or G, two positions have C or U (Y), and eight positions have A or G (R). The structure of 5 S RNA from eukaryotes is similar to that from prokaryotes. Although both structures appear to be polynucleotides of 119 nucleotides, subpositions in the structures are somewhat different. In addition, most yeast 5 S RNAs contain a pseudouridine in position 51 (see sequences E27 to E30, Appendix). While prokaryotes contain up to 12 subpositions, eukaryotes in this arrangement have up to 10. However, both have 5 subpositions in common (10 : 2, 70 : 1, 74: 1 , 8 6 : 1, and 101: 1).

B.

invariant Segments of the Structure

Similar to the prokaryotic structure, eukaryotic 5 S RNAs also exhibit “invariant” sections among various sequences. For example, the invariant segment of prokaryotes, -C-C-G-A-A-C-, is present as -C-NG-A(or C)-U(or A)-C- in positions 41 to 46 (where N is A, C, or U).

PROKARYOTIC

5s RNA

._

86 I

FIG.1. A generalized structure of prokaryotic 5 S RNAs derived from a comparative study of 29 sequences from different organisms. A residue in a bold-line circle is an “invariant” nucleotide (A, C, G, or U), and one in a thin-line circle indicates one of two nucleotides [C or G, C or U (Y), A or G (R)]. These “invariant” nucleotides are found in the same “positions” in most 5 S RNA sequences examined (see the text and Table I for details). A dashed-line circle indicates a “subposition” that is not filled (“deletion”) in most prokaryotic sequences. For the nature of the base-pairing in stem regions, see Table 11.

TABLE 111 NATUREOF RESIDUES IN 35 EUKARYOTIC 5 S RNAsO Nature of the conserved nucleotideb

A

G

C

U

Position

4 1 4 3 3 18

3 31 14 4 7 7 7 27 18 24

-

-

11 13

3 5 15 24 3 1

1 2 3" 4

7 7 7

1 26

-

24 7 1 11 7

-

32 32 22 12

-

22 33 1 1

-

25 -

-

-

1 1 1 11 8 2 2 26 3 1

-

7 27 1 1 -

7 34 35 2 9 22 22 1 12 25 17 9

-

-

1 6 1 30

10 1 2 20 24

32 12 4

22

-

-

-

-

2 32 16 35

12

23

-

33

11

12 35

33

-

-

2

9 4

1

-

32 5 3 9 5 1 5 16 7 1 7

-

5 3 1

-

2 7 10 33 2 14 -

35 2 10

-

Specific nucleotide

Group-specific nucleotide

9 6

7 8 9 10 10:2 11 12 13 14 15 16 17 18 19 20 21d 22d 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 41 42 43 44 (continued)

187

TABLE I11 (Continued) Nature of the conserved nucleotideb

A

G

C

12 1 8 8

-

-

-

2 1 23 35 1 4 1 35 32 2

10 4

3 12 33

-

10 3 18

-

35

12 3 6 21 1

-

4

1 10 2

3

32

-

-

-

-

33

1 34

-

1 32 14

-

-

4 9 9 3 6

-

-

16 18 26 13 1 1 31 28 35 1 9 3 31 16 8

-

34

6

35

10

-

8 22 29 12

3

34 3 2 9 10 2 14 1 1

34 20 3

1 9 8

-

-

-

25 2 1 2

-

-

-

33

2

U

Position 45 46 46: 1 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 70: 1 71 72 73 73: 1 74 74 : 1 75 76 77 78 79 80 81 82 83 84 85 188

Specific nucleotide

Group-specific nucleotide

TABLE 111 (Continued) Nature of the conserved nucleotideb ~~

G

A

1

34 33 5 35 1 2 12

2 12

35 -

1 13

-

-

8 1

2 1 35 24 35 1 10 1

1 34 34 6 15 1 2 12

-

9 21 27 7 28 7 28 7

5

-

3

-

3 24 15 2

3 14 3 1

1

-

-

C

U

Position

-

17 -

-

-

32 31 2 29 22 7

2 1 8 6 3 26

10

1 15 8 6 5 3 6 1 27 7 10 27 7 18 1 3 13 1 22 22

86 86: 1 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 101 : 1 102 103 104 105 106 107 109 110 111 112 113 114‘ 115 116” 117 118 119 119: 1

1

-

4 11 28 28 4 3 7 1

-

15 8

7 9 4

5 31 8 1 1

-

8

Specific nucleotide

Group-specific nucleotide

~

a See the Appendix for the list and origins of the 5 S RNA sequences used for the study (sequences E 1 to E35). b Residues in parentheses indicate the nature of the predominant nucleotide found in over 24 sequences. c A residue in this position in one of the sequences was not characterized beyond being a purine or a pyrimidine nucleotide. Since two different sequences have been reported for positions 21 and 22, they were omitted from this analysis.

*

189

RAM P. SINGHAL AND JON1 K . SHAW

190

However, consistency regarding which parts of the prokaryotic 5 S RNAs are conserved is not observed in the eukaryotic structure (for example, compare segments 71-79,99-109, and 103-106 between the 5 S RNAs from the two sources).

C. The Base-Paired Regions The base-pairing scheme in various stem regions of the structure is shown in Table IV. As opposed to the prokaryotic 5 S RNA (four stems), this structure appears to have five base-paired regions (stems). The specificity of the nucleotides involved in yielding stem 1, stem 2, and stem 3 is identical between the two structures, and similar residues constitute stem 4. Stem 5 consists of five base-pairs and is unique to the eukaryotic structure. There is a preponderance of G C basepairs in the structures derived from each kind of organism. Figure 2 illustrates a proposed secondary structure for eukaryotic 5 S RNA based on the above results and also on the proposed structure for prokaryotic 5 S RNAs (see Fig, 1).

-

TABLE IV PROBABLEBASE-PAIRED REGIONSOF EUKARYOTIC 5 S RNAs Frequency of the base-pairb Region of base-paired residues0

G C

A*U

24 14 7 27 14 7 17 16 31 3

7 6 7

Other possibilities

Stem 1

1 . 118 2 ' 117 3 , 116 4 . 115 5 . 114 6 . 113 7.112 8 . 111 9 . 110 10 . 109 Stem 2 15 . 6 7 16 .66 17 .64 18 . 6 3 19 .62 20 ' 61 21 . 6 0 22 .59

22 23 2 21 33 19 11 25

21 25 18 5 1 3

G G G G

'

U(4)

. U(12), A . C(3) . U(20), c . C(1) . U(8) A . C(2) G . U(13) G ' U(3) A . G(1)

5 4 31 11

A . C(l)

15

G . U(1) G . U(2), C . A(1) G . U(1), c ' U(7)

-

20

-

-

A . A(l), C . U(1)

u . U(1), G . G(1) C . U(l), C . A(1)

PROKARYOTIC AND EUKARYOTIC

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TABLE IV (Continued) Frequency of the base-paiP Region of base-paired residuesD

G .C

A .U

10 34 32 26 28

24 1 1 3

Other possibilities

Stem 3

28 .54 29 .53 30 . 50 31 . 4 9 32 . 4 8 33 . 4 7

-

-

32

-

G G G G

. A(1), G . G(1)

. U(6) . U(1), C . U(6) . U(1), C . A(1), C . U(1)

Stem 4

78 . 97 79 ' 9 6 80 . 95 81 ' 9 4 82 . 9 3 83 . 92

34 2 8 24 29 -

85 ' 9 1 86 . 9 0 Stem 5 68 . 107 69 . 106 7 0 . 105

33 33

1 1

1 28 23

-

13 11 6 9

3

C . A(1)

G . U(33) G U(1), U . U(13)

-

A . C(2), A G(22), G G(1) G . U(1) G , U(1)

G . U(7), G . G(21) C . C(6), C . U ( l ) G . U(7), C . U(l), G

.

A(1)

70: 1 . 104 71 . 103

5

28

15 4

G . U(7), C . C(1) G . U(2), G . A( 1)

a When a residue was characterized only as a purine or a pyrimidine, or else was not present in a particular position (deletion), the nature of the base-pairing for that particular position is indeterminate. Hence, the sum of the different types of base-pairs having such residues (R, Y) does not equal 35 (the total number of sequences examined). The nature of the base-pairing was determined for 35 eukaryotic structures (see sequences E l to E35, Appendix). The numbers in the G . C and A . U columns and the numbers in parentheses appearing after the alternate base-pairs indicate the number of occurrences of that particular type of base-pair in the 5 S RNA sequences.

111. Conformation of Prokaryotic 5 S RNAs

Over 80 sequences of 5 S RNAs from various sources have been determined. The results of chemical, biological, and physical studies indicate that 60-70% of the molcule is base-paired. The molecule

I

-,

87

FIG.2. A generalized structure of eukaryotic 5 S RNAs derived from a comparative study of 35 sequences of different organisms. A residue in a bold-line circle is an “invariant” characteristic nucleotide (A, C, G, or U), and one in a thin-line circle indicates one of two nucieotides [C or G; C or U (Y);A or G (R)]. These “invariant” nucleotides are found in the same “positions” of most 5 S RNA sequences examined (see the text and Table 111 for details). The dashed-line circle represents a “subposition” that is not filled (“deletion”)in most eukaryotic sequences. For the nature of the base-pairing in the stem regions, see Table IV.

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s RNAS

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binds with ribosomal proteins L5, L18, L25, and L22, in addition to the T-JI-C stem of the tRNAs (2,8a). Structural integrity appears to be essential for its function in protein synthesis. Two conformers, native (“A”) and denatured (“B”), have been described for this RNA (8b). A wide variety of chemicals have been employed to study the reactive regions of this molecule. These include monoperphthalic acid, carbodiimide, Kethoxal, chloracetaldehyde, L4-phenyldiglyoxal, and hydrogen exchange. In addition, the several nucleases, including T1, T2, IVYpancreatic and sheep kidney ribonucleases on 5 S RNA have been employed. Other studies include binding of the complementary oligonucleotides with the 5 S RNA and “reconstitution” of the ribosome subunit using 5 S RNA from E. coli and ribosome proteins from Bacillus sp. (see below). The use of a chemical or an enzyme for studying structure depends not only on the actual reaction or the method employed, but also on the accessibility of the residues to the reagent. (Certain reactive sites may be inaccessible owing to three-dimensional folding or masking by a protein.) In this section, results of various studies dealing with the conformation of 5 S RNA that support the proposed secondary structure are discussed.

A. Reactivity for the Chemicals a. Monoperphthalic Acid.The reactivity of monoperphthalic acid with only non-hydrogen-bonded A residues o f 5 S RNA (to yield adenosine l-oxide) indicates areas of A U base-pairs in the structure (9). The results indicate that 10 (out of 23) A residues are not hydrogenbonded in E . coli 5 S RNA. Binding of 5 S RNA to the T+-C stem of tRNA appears to be abolished after specific modification of the A residues in positions244 and 45 by monoperphthalic acid (10).Hence, the bases in this area probably do not participate in base-pairing (see loop 111, Fig. 1).

b. N-Cyclohexyl-N’-[2-(4-methylmorpholinio)ethyl]carbodiimide. In general, carbodiimides react with U, JI, G , and I residues in the non-hydrogen-bonded regions of the molecule ( 3 , 11 ). Under appropriate reaction conditions, carbodiimides react selectively with U res2 The residue numbers cited in this text differ from those given by the author in the original literature. Here, the residues are referred to by our numbering scheme, which allows for “subpositions” in the structure. See, for example, E . coli 5 S RNA sequence P18 in the Appendix. There is a difference of one number between the two numbering methods, since we locate the eleventh residue in position 10 : 2, but this difference disappears after residue 86, as the E . coli structure lacks a residue in position 86.

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RAM P. SINGHAL AND JON1 K. SHAW

idues in positions 13,39,64,76,87,89,and 103 of E . coli 5 S RNA (3). Most reactive residues appear to be located in positions 87 and 89. These results suggest that the molecule has a compact structure and a large portion of it is not readily reactive. c. 3-Ethoxy-2-oxobutyraldehyde (Kethoxal). The action of Kethoxal on 5 S RNA ( E . coli) indicates differences in reactivity between conformer A (“native”) and conformer B (denatured), both quantitatively and qualitatively ( 1 2 , 1 3 ) .For example, in conformer A, G:40 is much more reactive than those G residues located in other positions, where they react (in decreasing order) G : 12 > G : 6 8 > G:23 > G : 85 > G : 107 > G : 15, G : 22, G : 43. In conformer B , G :43 exhibits the greatest reactivity. The reactivity in G : 4 3 only in the denatured conformer suggests protection of this reaction site by the protein L5 in the native form. However G :40 was reactive in both reactive native and denatured 5 S RNAs. When intact ribosomes are treated with Kethoxal, not only do the exposed residues of the RNA react, but also 90%of the 5 S RNAs dissociate from the ribosomes (14). G : 40 reacts immediately, but G : 43 starts reacting only after this dissociation. The authors suggest that G :43 is protected by a protein, and that the residue perhaps serves as a recognition (binding) site for that particular protein. Furthermore, the interaction of this area of the 5 S RNA with the T-+C stem of the tRNA may occur through a ribosomal protein. d . Chloracetaldehyde. The fluorescence-yielding modification with chloracetyldehyde (alkylation of the 6-amino group of adenine under specific reaction conditions), results in modification of 16 different adenine residues in the E. coli 5 S RNA, the greatest reactivity being in positions 28,33, 38, 56,57, and 58 (cf. sequence P-18 in the Appendix) (15).Among these, only residues in positions 28 and 33 are base-paired (stem 3) in our proposed secondary structure (see Fig. 1). This segment of a six-base-paired region (stem 3) surrounded by two loops (I1 and 111) can easily undergo conformational change during reaction conditions. The authors indeed observe that this region undergoes a conformational change as judged by fluorescence measurements (15). e. Cross-Linking Agent. The cross-linking agent 1,Cphenyldiglyoxal has been employed to link guanine residues in the base-paired regions (16).There is evidence for cross-linking between G : 2 and G : 111 in the E . coli structure (see sequence E-18),but to achieve this, extensive perturbation and unstacking of the bases located in stem 1 are necessary. Hence, the results are inconclusive.

PROKARYOTIC AND EUURYOTIC 5

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B. Exchangeable Protons Tritium-hydrogen exchange identifies those protons in a doublehelical structure that are free from tertiary constraints (17). The results indicate that 125-170 hydrogens are exchangeable, which agrees with the number expected from several proposed models of 5 S RNA ( 1 7).

C. Reactivity with Various Nucleases The study of RNA reactivity with nucleases has the disadvantage of not being able to account for possible conformational changes in the RNA structure brought about by the protein (enzyme). However, the reaction of 5 S RNA with RNase T1 yields five large fragments (18). This does not necessarily indicate uninterrupted stretches of basepairs. The results from RNase T1 together with those from pancreatic RNase digestions indicate the presence of four single-stranded regions, which may involve both stems and loops: residues 10 to 12 (loop I), 24 to 30 (loop 11, stem 3), 39 to 43 (loop 111),and 57 to 63 (loop 11, stem 2). Another study using RNase T2 confirmed the presence of a highly exposed single-stranded region in the vicinity of residue 39 located in loop I11 (19). In a separate study, using sheep kidney RNase and RNase IV, the results indicate that the most accessible region lies in the vicinity of G : 40 of the E . coli 5 S RNA (20). The authors conclude from this work that the G residue in position 40 is located in an accessible single-stranded region that extends from residue 33 through 49. These studies support the notion of a highly conserved, single-stranded area appropriate for interacting with the T-$-C stem of the tRNA structure.

D. Binding with Complementary Oligonucleotides The binding of complementary oligonucleotides with a macromolecule indicates accessibility and “exposed” areas of the molecule. Similarly, a failure to bind with the molecule indicates unavailability or involvement of the residues in secondary and tertiary structures. Experiments with 5 S RNA indicate the following that fail to bind (base-pair); residues 9 to 11 (stem 1 region); 24 to 31 (loop 11, stem 3); 57 to 64 (stem 2); 95 to 98 (stem 4). These results support the existence of the base-paired regions (stems 1 to 4) in the proposed 5 S RNA model. Failure to react with residues 11, 24 to 26, 57, and 58 can be due to tertiary interactions or strong stacking of the purine bases in that area of the molecule (20).

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RAM P. SINGHAL AND JON1 K. SHAW

E. Reconstitution of Ribosomes from 5 S RNA The reconstitution of ribosomes using E . coli 5 S RNA and Bacillus ribosomal proteins yields ribosomes that can function normally in protein synthesis (22).In a separate study, mismatching of the residues located in the stem 1 region of B . lichenifomis was induced artificially by melting and reannealing two forms (“major” and “minor”) of 5 S RNA each with ribosomal proteins from the same organism (23). This modified 5 S RNA reassociated with the ribosomal proteins more slowly, but the reconstituted ribosome did function normally. The results indicate a need for a specific stem 1structure for proper protein interaction with the 5 S RNA.

F. Physical Measurements From various physical studies (25-27) involving NMR, small-angle X-ray scattering, sedimentation velocity, sedimentation equilib rium, viscosity measurements, and ultraviolet spectroscopy of 5 S RNA, it can be concluded about this molecule that ( a )it has a structure that includes helices aligned along the long axis; ( b )the tertiary interactions yield a Y-shaped structure; (c) it is more elongated and asymmetric; ( d )from hydrodynamic studies, it has a prolated, ellipsoidal shape; ( e )as suggested by the migration of purified 5 S RNA in two bands in polyacrylamide gel electrophoresis, it probably exists in two conformations.

IV. Summary Sequences of 5 S RNAs from both prokaryotic and eukaryotic origins have been arranged in the Appendix in a manner that yields the maximum homology among various structures. To achieve this, 119 nucleotide “positions” and 11 “subpositions” are established for a “standard” prokaryotic 5 S RNA structure. Similarly, 119 “positions” and 8 “subpositions” are established for a standard eukaryotic structure. From comparisons among the 29 prokaryotic and 35 eukaryotic sequences in the Appendix, several “invariant” (conserved C, U, A, or G ) and “semiconserved” (C or G, purine or pyrimidine) nucleotides have been derived for each structure. Homologies among sequences are also discussed. The probability of base-pairing in different parts of both prokaryotic and eukaryotic 5 S RNA molecules is evaluated. Four basepaired “stem” regions and four single-stranded “loop” regions appear

PROKARYOTIC AND EUKARYOTIC

5 s RNAs

197

in the prokaryotic structures. An additional base-paired section (stem

5 ) is suggested for the eukaryotic structure. On the basis of these results, a structural model for each prokaryotic and eukaryotic 5 S RNA is proposed (Figs. 1 and 2). Various studies of bacterial 5 S RNAs using chemical reactivity, enzymic hydrolysis, proton exchangeability, binding of complementary oligonucleotides, reconstitution of ribosome subunits, and different physical measurements, the results of which support the proposed model in general, are briefly discussed. (See Appendix, p. 198, and Addendum, p. 251.)

REFERENCES 1. G . G. Brownlee, F. Sanger, and B. G. Barrell, Nature (London) 215,735 (1967). 2. V. A. Erdmann, This Series 18,45 (1976). 3. V. A. Monier, in “Ribosomes” (M. Nomura, A. Tissi&res,and P. Lengyel, eds.), p. 141. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1974. 4. J. T. Madison, ARB 37, 131 (1968). 5. V. A. Erdmann, NARes 9, r25 (1981); 10, r93 (1982). 6. B. G . Barrell and B. F. C. Clark, “Handbook of Nucleic Acid Sequences,” p. 68. Joynson-Bruvvers, Oxford, England, 1974. 7. M. 0. Dayhoff, “Atlas of Protein Sequences and Structures,’’ p. 351. National Biomedical Research Foundation, Washington, D.C., 1972. 8. P. Wrede, D. Pong, and V. A. Erdmann, J . Med. Gen. 120,83 (1976). 80. P. Spierer, A. A. Bogdanov, and R. A. Zimmerman, Bchem 17,5394 (1978). 8b. M. Aubert, J. F. Scott, M. Reynier, R. Monier, PNAS 61, 292 (1968). Also see reference 12. 9. F. Cramer and V. A. Erdmann, Nature (London) 218,92 (1969). 10. V. A. Erdmann, M. Sprinzl, and 0. Pongs, BBRC 54,942 (1973). 11. J. C. Lee and V. M. Ingram,JMB 41,431 (1969). 12. H. E. Noller and R. A. Garrett,JMB 132,621 (1979). 13. W. Herr and H. F. Noller,JMB 130,421 (1979). 14. I. Larrinua and N. Delihas, PNAS 76, 4400 (1979). 15. M. Digweed and V. A. Erdmann, NARes 9,3187 (1981). 16. R. Wagner and R. A. Garrett, NARes 5,4065 (1978). 17. J. Ramstein and V. A. Erdmann, NARes 9,4081 (1981). 18. B. J. Jordan, ] M B 55,423 (1971). 19. R. Vigne, B. R. Jordan, and R. J. Monier,JMB 76, 303 (1972). 20. C . Bellemare, B. R. Jordan, and R. J. Monier,JMB 71,307 (1972). 21. J. B. Lewis and P. Doty, Nature (London) 225,510 (1970). 22. P. Wrede, 0 . Pongs, and V. A. Erdmann,JMB 120,83 (1978). 23. V. A. Erdmann, PNAS 18,45 (1976). 24. H. A. Raue, S. Lorenz, V. A. Erdmann, and R. J. Planta, NARes 9, 1263 (1981). 25. G. E. Fox and C. R. Woese, J . M o l . Euol. 6, 61 (1975). 26. R. Osterberg, B. Sjoberg, and R. A. Garrett, EJB 68,481 (1976). 27. J. W. Fox and K. P. Wong,JBC 254,10139 (1979).

198

RAM P. SINGHAL AND JON1 K. SHAW

APPENDIX: Sequences of 5 S RNAs A. Notes

1. Several “subpositions” have been created to achieve maxi-

2. 3.

4.

5.

mum homology among various structures. The prokaryotic and eukaryotic 5 S RNA sequences are listed here with 119 “positions” and with one or more “subpositions” of residues 10, 29, 33, 46, 70, 73, 86, 101, 108, and 119. The use of subpositions makes it possible to preserve a common numbering method for 5 S RNAs without regard to origin. For characteristics of the nucleotide positions involved in various stem and loop regions of the molecule, the reader is referred to Figs. 1 and 2 and Tables I1 and IV. The residues underlined in Appendix Section C were found in Iess than 1 mole per mole of the 5 S RNA by the authors sequencing the structure (see the reference for that particular sequence). * and Y indicate either an A or G (R) and either C or U (Y), respectively, in that particular position. For details, see the note cited with the reference for t b t particular sequence. In the E . coli 5 S RXA structure (P18), C indicates either C or A in position 11 and U indicates either U or G in position 12. The authors employed the two-dimensional electrophoretic method of Sanger et a2. for characterizing oligonucleotides derived from RNase T1 and pancreatic RNase digestions. The sequence of this particular 5 S RNA is calculated on the basis of a small number of “orientation points,” derived from the general properties of the 5 S rRNA structure.

fi

B. References and Footnotes for Sequence List PROKARYOTES P1. Anacystis nidulans (blue-green algae, 1405/1 Katz/Allen):M . J. Corry, P. I. Payne, and T. A. Dyer, FEBS Lett. 46,63 (1974). P2. Bacillus breois (ATCC 8185): C. R. Woese, K. R. Luehrsen, C. P. Pribula, and G. E. Fox, J. Mol. Eool. 8, 143 (1976). The authors derived this structure from oligomer catalogs (see note 5). P3. Bacillusfimtus (ATCC 14575): C. R. Woese, K. R. Luehrsen, C. P. Pribula, and G. E. Fox,]. MoZ. E d . 8,143 (1976). The residue at position 3 was a pyrimidine and those at positions 82 and 116 were purines. P4. Bacillus lichenijormis (S 244): H. A. Rauk, T. J. Stoof, and R. J. Planta, EJB 59,35 (1975).

PROKARYOTIC AND EUKARYOTIC

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s RNAs

199

P5. Bacillus megaterium (KM, a University of Illinois strain): C. P. Pribula, G. E. Fox, and C. R. Woese, FEBS Lett. 44,322 (1974). P6. Bacillus pasteurii (ATCC 11859): C. R. Woese, K. R. Luehrsen, C. P. Pribula, and G. E. Fox,]. Mol. Euol. 8, 143 (1976). The residue at position 2 was a pyrimidine and that at position 117 a purine. P7. Bacillus Q: H. A. Raue, A. Rosner, and R. J. Planta, Mol. Gen. Genet. 156, 185 (1977). P8. Bacillus stearothermophilus (1439, FV, “a”): C. A. Marotta, F. Varricchio, I. Smith, S. M. Weissman, M. L. Sogin, and W. R. PaceJBC 251,3122 (1976).This is a thermophilic strain derived from B. stearothermophilus 1420R after repeated growth in a low-phosphate medium at 62°C. P9. Bacillus stearothermophilus (799, “b”): C. A. Marotta, F. Varricchio, I. Smith, S. M. Weissman, M. L. Sogin, and N. R. Pace,JBC 251,3122 (1976). P10. Bacillus subtilis (168): H. A. €tau&,T. J. Stoof, and R. J. Planta, EJB 59,35 (1975); C. A. Marotta, F. Varricchio, I. Smith, S. M. Weissman, M. L. Sogin, and W. R. Pace, JBC 251,3122 (1976). P11. Lactobacillus breuis (X2, a University of Illinois strain): C. R. Woese, K. R. Luehrsen, C. P. Pribula, and G. E. Fox, J . Mol. Euol. 8, 143 (1976). P12. Lactobacillus uiridescens: L. J. Alexander and T. S. Stewart, NARes 8,979 (1980). P13. Micrococcus lysodeikticus (ATCC 4698): H. Hori, D. Syozo, K. Murao, and H. Ishikura, NARes 8,5423 (1980).This organism has the highest GC content of DNA among all living organisms (72%)according to the authors. P14. Streptococcus faecalis (a Microbiology Teaching Laboratory of the University of Illinois strain): C. R. Woese, K. R. Luehrsen, C. P. Pribula, and G . E. Fox,]. Mol. Euol. 8, 143 (1976). P15. Streptomyces griseus: A. Simoncsits, NARes 8, 4111 (1980). This organism is a fungus-like prokaryote. P16. Clostridium pasteurianium (ATCC 6013): C. P. Pribula, G. E. Fox, and C. R. Woese, FEBS Lett. 64,350 (1976). P17. Mycobacterium smegmatis (SN2): V. A. Erdmann, NARes 9, r25 (1981). P18. Escherichia coli (MRE 600 and CA 265): G. G. Brownlee, F. Sanger, and B. G. Barrell, Nature (London)215,735 (1967); G . G. Brownlee, and F. Sanger,JMB 23, 337 (1967);G. G. Brownlee, F. Sanger, and B. G. Barrel1,JMB 34,379 (1968). In E . coli, two different strains of bacteria were analyzed. Strain MRE 600 differs from the other strain CA 265 in having C instead of A at position 11 and U instead of G at position 12. P19. Halobacterium cutirubrum (NRC 34001): R. N. Nazar, A. T. Matheson, and G. Bellemare, JBC 253, 5464 (1978). P20. Photobacter (8265): C. R. Woese, C. P. Pribula, G. E. Fox, and L. B. Zablen, J . Mol. Euol. 5,35 (1975). P21. Pseudomonasfluorescens (ATCC 13430): B. Dubuy and S. M. WeissmanJBC 246, 747 (1971). P22. Thennus ahuaticus (ATCC 25104): R. N. Nazar and A. T. Matheson, JBC 252, 4256 (1977).This organism inhabits thermal environments and has unusual “thermal” enzymes, ribosomes, tRNA, and plasma membranes. The A residue at position 1 and the U residue at position 119 appear to occur in less than 1 mole per mole of 5 S RNA. P23. Proteus vulgaris: V. A. Erdmann, NARes 9, r25 (1981). Residues appearing at positions 8-38, 47, 48, and 56-98 were identified tentatively by the author. P24. Aerobacter aerogenes: S. J. Sogin, M. L. Sogin, and C. R. Woese, J. Mol. Euol. 1,

RAM P. SINGHAL AND JON1 K. SHAW

200

P25.

P26.

P27.

P28.

P29.

P30. P31. P32.

173 (1972);H. Hori, Mol. Gen. Genet. 145, 119 (1976). The authors derived this sequence by the “oligomer catalog” technique (see note 5). Residues in positions 8, 11, 12, 16, 18, 62, 63, 66, and 97 were identified tentatively. Erwinia aeroidiae (NRRLB 138):S. J. Sogin, M. L. Sogin, and C. R. Woese, J. Mol. Euol. 1,173 (1972);H. Hori, Mol. Gen. Genet. 145, 119 (1976).The authors have derived this sequence by the “oligomer catalog” technique (see note 5). Residues in positions 10:2-12, and 61-63 were identified tentatively. Proteus mirabilis: S. J. Sogin, M. L. Sogin, and C. R. Woese, J . Mol. Evol. 1, 173 (1972); H. Hori, Mol. Gen. Genet. 145,119 (1976).The authors have derived this sequence by “oligomer catalog” technique (see note 5). Residues in positions 8, 12, 18, 63, and 68 were identified tentatively. Serratia marcescens: S. J. Sogin, M. L. Sogin, and C. R. Woese, J . Mol. Evol. 1,173 (1972); H. Hori, Mol. Gen. Genet. 145, 119 (1976). The authors derived this sequence by “oligomer catalog” technique (see note 5). Residues in positions l l , 12, 62, and 63 were identified tentatively. Salmonella typhimurium: S. J. Sogin, M. L. Sogin, and C. R. Woese, J . Mol. Evol. 1,173 (1972);H. Hori, Mol, Gen. Genet. 145,119 (1976).The authors derived this sequence by “oligomer catalog” technique (see note 5). Residues in positions 11, 12, 62, and 63 were identified tentatively. Yersinia pestis: S. J. Sogin, M. L. Sogin, and C. R. Woese, J . Mol. Eool. 1, 173 (1972); H. Hori, Mol. Gen. Genet. 145, 119 (1976). The authors derived this sequence by “oligomer catalog” technique (see note 5). Residues in positions 12 and 63 were identified tentatively. Beneckea harueyi (392):V. A. Erdmann, NARes 10, r93 (1982). Thermus thermophilus (HB8): V. A. Erdmann, NARes 10, r93 (1982). Mycoplasma capricolum (ATCC 27 343): H. How, M. Sawada, S. Osawa, K. Murao, and H. Ishikura, NARes 9, 5407 (1981).

EUKARYOTES El. Chlamydomonas reinhardii (green alga, “I”): J. L. Darlix and J. D. Rochaix, NARes 9,1291 (1981).Two species of cytoplasmic 5 S RNA were isolated. This is the major species of 122 nucleotides, which differs from the minor species in 16 residues at various positions. E2. Chlamydomonas reinhardii (green alga, “11”): J. L. Darlix and J. D. Rochaix, NARes 9,1291 (1981).This is a minor species of 121nucleotides isolated from the cytoplasm. E3. Chtorella pyrenoidosa (alga): V. A. Erdmann, NARes 9, r25 (1981). E4. Helianthus annuus (sunflower): V. A. Erdmann, NARes 9, r25 (1981). E5. Lemna minor (duckweed): T. A. Dyer and C. M. Bowman, in “Genetics and Biogenesis of Chloroplasts and Mitbchondria” (M. Bucher et al., eds.), p. 645. Elsevier/North-Holland, Amsterdam, 1976. E6. Lycopersicon esculentum (tomato): V. A. Erdmann, NARes 9, r25 (1981). E7. Phaseolus oulgaris (dwarf bean): V. A. Erdmann, NARes 9, r25 (1981). The U residue at position 119 : 1appears to occur in less than 1mol per mole of 5 S RNA. E8. Secale cereale, C.V. ‘Lovasz patonai’ (rye):V. A. Erdmann, NARes 9, r25 (1981). E9. Spinacia oleracea (spinach): N. Delihas, J. Anderson, H. M. Sprouse, M. Kashdam, and B. Dudock, JBC 256,7515 (1981). E10. Triticum aestioum (wheat embryo): V. A. Erdmann, NARes 9, r25 (1981). E l l . Viciafaba (broad bean): V. A. Erdmann, NARes 9, r25 (1981).

P R O M Y O T I C AND EUKARYOTIC

5

s RNAS

20 1

E12. Aspergillus nidulans (ascomycetes): B. Piechulla, U. Hahn, L. W. McLaughlin, and H. Kuntzel, NARes 9, 1445 (1981). E13. Crithidia fasciculata (trypanosome): R. M. MacKay, M. W. Gray, and W. F. Doolittle, NARes 8, 4911 (1980). E14. Dictyostelium discoideum (slime mold): H. Hori, S. Osawa, and M. Iwahuchi, NARes 8,5535 (1980). E15. Drosophila melanogaster (F6 of KC, fly): V. A. Erdmann, NARes 9, r25 (1981). E16. Callus gallus (chicken): V. A. Erdrnann, NARes 9, r25 (1981). Two different sequences of chicken embryo fibroblast cultured cells have been published. They differ in positions 20 and 21: position 20 has been reported to have C or U residue and 21 to have U or G. The yield of U in position 119 : 1 was less than expected. E17. Homo sapiens (mammalian cultured cells): This is a common structure found in HeLa cells and KB cells. V. A. Erdmann, NARes 9, r25 (1981).The yield of uridine in position 119 : 1 was less than expected. KB cell and HeLa cell 5 S RNA structures contain one and three phosphate groups in the 5‘ position, respectively. E18. Homo sapiens (mammals): This is a common structure reported for mouse, marsupial, rat, and rabbit. R. Williamson and G. G. Brownlee, FEBS Lett. 3,306 (1969), mouse; M. J. Averner and N. R. Pace,JEC 247,4491 (1972), marsupial; F. Labrie and F. Sanger, EJ 114, 29 (1969), rat and rabbit. E19. Iguana iguana (leguan): V. A. Erdrnann, NARes 9, r25 (1981). E20. Lingula anatina (brachiopoda): H. Komiya, N. Shimizu, M. Kawakami, and S. Takemura, J . Eiochem. (Tokyo) 88, 1449 (1980). E21. Lytechinus oariegatus (sea urchin): V. A. Erdmann, NARes 9, r25 (1981). E22. Neurospora crassa (ascomycetes): B. Piechulla, U. Hahn, L. W. McLaughlin, and H. Kuntzel, NARes 9, 1445 (1981). E23. Salmo gairdneri (rainbow trout): H. Komiya and S. Takemura,J. Biochem. (Tokyo) 86, 1067 (1979). E24. Terrupene Carolina (turtle): K. L. Roy, FEBS Lett. 80,266 (1977). The U at position 119 : 1 was found in less than 1 mole per mole of 5 S RNA. E25. Tetrahymena themophilus (ciliated protozoa): V. A. Erdmann, NARes 9, r25 (1981). Residues appearing at positions 3, 5, 114, and 116 were identified tentatively by the author. E26. Saccharomyces carlsbergensis (yeast, “a”): V. A. Erdmann, NARes 9, r25 (1981). Residues appearing at positions 14-20 were identified tentatively by the author. E27. Saccharomyces carlsbergensis (yeast, “b”): V. A. Erdmann, NARes 9, r25 (1981). E28. Saccharomyces cereoisiae (yeast): V. A. Erdmann, NARes 9, r25 (1981). E29. Kluyoeromyces lactis (yeast): V. A. Erdmann, NARes 9, r25 (1981). E30. Pichia membranaefaciens (yeast): V. A. Erdmann, NARes 9, r25 (1981). Residues at positions 81-87 were identified tentatively by the author. E31. Tomlopsis utilis (yeast): K. Nishimura and S. Takemura, J. Biochem. (Tokyo)76, 935 (1974). E32. Xenopus laeois (toad, somatic cells from kidney): V. A. Erdmann, NARes 9, r25 (1981). The U at position 119 : 1was found in less than 1mol per mole of 5 S RNA. E33. Xenopus laeois (toad, oocyte cells): V. A. Erdmann, NARes 9, r25 (1981). Residues at positions 79 and 80 were identified tentatively by the author. The U at position 119 : 1 was found in less than 1 mole per mole of 5 S RNA. E34. Xenopus mulleri (toad, somatic cells): V. A. Erdmann, NARes 9, r25 (1981). E35. Xenopus mulleri (toad, oocyte cells): V. A. Erdrnann, NARes 9, r25 (1981). E36. Acanthamoebu castellanii (amoeboid protist): R. M.MacKay and W. F. Doolittle, NARes 9,3321 (1981).

RAM P. SINGHAL AND JON1 K. SHAW

202 E37. E38. E39. E40.

Misgurnus fossilis (oocyte): V. A. Erdmann, NARes 10,193 (1982). Misgurnus fossilis (somatocyte): V. A. Erdmann, NARes 10, r93 (1982). Crypthecodinium cohnii: V. A. Erdmann, NARes 10, r93 (1982). Schizosaccharomyces pombe (yeast I F 0 No. 0345):V. A. Erdmann, NARes 10,193 (1982). E41. Euglena gracilis (photosynthetic protozoa): N. Delihas, J. Anderson, W. Andresini, L. Kaufman, and H. Lyman, NARes 9,6627 (1981). E42. Artemia salina (brine shrimp): L. Diels, R. DeBaue, A. Vandenberghe, and R. DeWachter, NARes 9,5141 (1981).

CHLOROPLASTS C1. C2. C3. C4.

Lemna minor (duckweed): T. A. Dyer and C. W. Bowman, B] 183,595 (1979). Nicotiana tabacum (tobacco): T. A. Dyer and C. W. Bowman, BJ 183,595(1979). Phaseolus oulgaris (dwarf bean): T. A. Dyer and C. W. Bowman, BJ 183,595 (1979). Spinacia oleracia (spinach): N. Delihas, J. Anderson, H. M. Spouse, and B. Dudock, NARes 9,2801 (1981). C5. Vicia fabo (broad bean): T. A. Dyer and C. W. Bowman, Biocham. ]. 183, 595 (1979).

MITOCHONDRIA M1. Triticum sp. (wheat): M. W. Gray and D. F. Spencer, NARes 9,3523 (1981).

ARCHAEBACTERIAL Al. Halococcus morrhuae (ATCC 17082): V. A. Erdmann, NARes 10, r93 (1982). A2. Sulfolobus acidocaldarius: V. A. Erdmann, NARes 10, r93 (1982). A3. Thernoplasma acidophilum (122-1B2 or 122-1B3):K. Luehrsen, G. E. Fox, M. W. Kilpatrick, R. T. Walker, H. Domdey, G. Krupp, and H. J. Gross, NARes 9, 965 (1981).

C. 5 S Sequences See pages 204-209 (prokaryotes, pp. 204-205; eukaryotes, pp. 206-209; chloroplasts, pp. 208-209; mitochondria, pp. 208-209; archaebacterial, pp. 208-209).

ADDENDUM: References to 32 Additional Sequences See Addendum on p. 251.

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Structure of Transfer RNAs: Listing of 150 Additional Sequences’ RAMP. SINGHAL, EDDAF. ROBERTS, AND VIKRAMN. VAKHARIA Department of Chemistry, Wichita State Uniuersity, Wichita, Kansas

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

11. Abbreviations and Definitions. . . . . . . . . . . . .................. A. tRNA Code Numbers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. References and Footnotes for Se ................... IV. Sequences of 126 tRNAs . , , , ................... Addendum: Additional tRNA Sequences. .......................

. .

211 215 215 216 219 227 227

1. Organization

The structure of various tRNAs can be organized in a manner to yield common nucleotide “positions” in the cloverleaf structui e of most tRNAs. To achieve this, “subpositions” of residues 17 (17: 1, 17 : 2 ) and 20 (20 : 1, 20 : 2, 20 : 3) in the dihydrouridine loop and residue 47 (47 : 1, 47 : 2, etc.) in the variable loop are introduced. Using this numbering scheme, each tRNA yields a structure with common “ invariable” or “conserved” nucleotide positions ( 1 ) . Figure 1 indicates, in addition to “subpositions,” “invariable“ (A, C, G, or U), and “semiconserved” (R, i.e., A or G; Y, i.e., C or U; G or C) positions found in most tRNA sequences. The structures of 178 tRNAs were organized in this manner and listed in this series over 2 years ago ( 1 ) . Here, we update this list by

‘

The authors plan to update the tRNA sequence list periodically. Researchers are requested to send new tRNA sequences and changes in the known structures to the first author (R. P. S.). A program in FORTRAN language has been developed for storing, updating, and retrieving the tRNA sequences and for drawing the secondary structure of tRNAs by computer. [Singhal et al., Comput. Programs Biomed. 14,277 (1982)l. Address inquiries for this to the first author (R. P. S.). 211 Progress in Nucleic Acid Research and Molecular Biology, Vol. 28

Copyright 0 1983 by Academic Press, Inc. All rights of reproduction in any form reserved. ISBN 0-12-5400284

212

RAM P. SINGHAL

et al.

FIG.1. A generalized structure of tRNAs. The stippled boxes indicate the nucleotide positions that differ in the eukaryotic initiator tRNAs. Dashed-line circles indicate a “subposition” not necessarily “occupied” in every tRNA (for example 17, 17 : 1 of the a region of the D loop; 20,20 : 1,20 : 2 of the @ region of the D loop; 47 : 1,47 :2, etc., of the V loop). The structure contains 16 invariant (conserved) and 20 semiconserved (R = purine; Y = pyrimidine; G/C = G or C ; A/C = A or C ) nucleotides. Residues in bold-line circles are the invariant nucleotides present in most prokaryotic and eukaryotic tRNAs.

listing another 126 tRNA sequences available to us (this number includes revision of several of the sequences listed previously). Although we have used the same format as before, mitochondria1 tRNAs (especially those derived from their gene analysis) cannot be accommodated in this numbering scheme. Hence, their residues are numbered as they appear in the original literature (a sequence extending beyond 76 residues is completed with a note in the reference section; such a sequence bears an asterisk in position 76) (see Addendum, p. 227).

STRUCTURE OF TRANSFER

213

RNAS

In Table I, tRNA sequences have been grouped according to their origins (sequences published in the earlier listing are indicated by sequence code numbers in italics). (See Section II,A below for explanation of the sequence code numbers.) An Arabic number in parentheses after the sequence code indicates the number of different organisms having that common tRNA structure-as referred to by the sequence code. For example, i M 1 0 ( 7 ) under mammals indicates the common sequence for the initiator-methionine tRNA determined from seven normal, mammalian sources and cited in the previous sequence collection; similarly, R11 under yeast mitochondria indicates a sequence of tRNAArgfrom that source listed in this collection. TABLE I SEQUENCES OF tRNAs FROM VARIOUS SOURCES“ Origin

1. Asteroidea (starfish) 2. Bacteria (1) B. stearothermophilus (2) B . subtilis

5.

6.

1 3 11

iM13

5 2 4 2 3 3 2 1 6

D5, V11 R8(2), P2(2) L15, S9 R9, D6, L19 iM10, M4, V11 N3,’ W6 K9 E6, H4, K5, iM12, F16, V17

58

(4) H. volcanii

13

(6) Salmonella (7) Staphylococcus ( 8 ) Thermoplasma acidophilum (9) Thermus thermophilus Blue-green algae Bombyr mori Cancer cells (1) Ascites hepatoma (2) Leukemia (3) Morns hepatoma Mitochondria (4) Myeloma ( 5 ) Sarcoma (6) Cells in culture Drosophila

Sequence code numbersb

2F, l Y , 4V 8A, 7R, 16G, 2K, 3iM, 5M, 3F, 2T, 7Y(2), 14V l A , 9A, l R , 2 R , l N , l D , l C , l Q , 2Q, I E , 2E, 3E, 2G, 3C(2), 4G(3), 15G(2), lH(2),lZ, 41, lL(2),215, 3L, IK, liM(5),l M , 2M, lF, 1S,2S, l T , l W , 2 W , 3W, 4W, 8W,” 12W(4),2Y(3), 1V(2),2V(2),3V(2) 10R, 7D, 5 4 , 7E, 51,20L, 10K, 22iM, 4P, 15S, 16S, 8Y, 18V 14G, 4iM, 4F 1C(2),1H(2), 1L(2),5V 5G(3),6G(3) 8M 2iM, 15iM(2) F8 A 5 , A 6 , G10, G11, F18

(3) Escherichia

(5) Mycoplasma

3. 4.

Total no.

3 7 6 1 3 1

(continued)

214

RAM P. SINGHAL et

al.

TABLE I (Continued) Origin

Total no.

Sequence code numbersb

7. Euglena gracilis (1) Chloroplast (2) Cytoplasm 8. Mammals

1 1 2 34

Mitochondria 9. Neurospora crassa (1) Cytoplasm (2) Mitochondria 10. Plants (1) Chloroplast

4

D9 F6 F7, F20 N2(3), D4, D5, G12, G13, H5, L14, L16, K6, K7, K8, iM10(7), M4, F14(4) F19(2), W7, S9, S10, V11, V12(2), V13 M10, S12(2), S13

2 7 7 12

(2) Cytoplasm 11. Rhodosprillum rubrum 12. Scenedesmus obliquus Chloroplast 13. Tetrahymena thermophila Cytoplasm 14. Towlopsis utilis 15. Virus and phage

2 6 24

16. Yeasts

38

Mitochondria

1 2 1 2

12

iM7, F17 A7, L12, L13, iM6, T6, W9, Y5 G9, iM5, iM8, F12, F13, F15(2) L9, L10, L11, iM16, iM19, M7, F5, F22, P5, T5, W11, V15 iM20 L23, F21 iMll iM17, M6 iM21, M9 A2, 12, L7, iM14, Y4, V10 W 2 ) , 4N, 8D, 3Q(3), 4Q, 6Q, 7G(3), 6H, 31(4), 4L, 21L, l P , 3P, 3S, 4S, 3T A3, A4, R4, R5, R6,02, C2, E4, E5, G8, L5,L6, L8,L17, L18, K3, K4(2), iM9, M3, F9, F11, S5, S6, S7(2), S8, S11, S14, 2'4, W5, Y3(2), Y6, V6 to V9 R11, C3, G17, H2, H3, 16, L22, iM18, F10, S17, S18, W10

A total of 293 sequences are listed in this table. This number includes 167 sequences described in the earlier compilation [This Series 23, 227 (1979)l. Although only 108 sequences appear here, another 18tRNAs have structures identical with or very similar to one or another of the 108 sequences (see Section 111 for details). Thus, this compilation deals with 124 newly elucidated tRNA structures. (The 126 sequences include eight listed in the Addendum section of the earlier compilation, but not those described in the Addendum section of this compilation.) For definitions of the sequence code numbers, see Section 11, below. The number in parentheses indicates the number of organisms (species) having this tRNA structure. The tRNA structures listed in italics appear in the earlier compilation [This Series 23, 227 (1979)l. See the text for details. The sequence of tRNAAS"from mammals (code N2) differs from mammary carcinoma (code N3); that of tRNATQfrom E . coli (code 1W) differs from a temperaturesensitive mutant (code 8W) only in one base position. See Section I11 for details.

STRUCTURE OF TRANSFER

215

RNAS

When the structures derived from two (or more) organisms for the same or different isoacceptors represent essentially a common sequence, then minor sequence differences, if any, are indicated by an asterisk above the affected residue. Details are given in the note appearing under that sequence code in Section I11 (references and notes for sequence list). The references are limited to the citation of the major or the latest publication if several publications deal with the same tRNA structure. Differences between mutant and the wild-type structures are indicated by asterisks in the sequence and explained in Section 111. Additional information may be found in other articles on this subject (1-3). Sequences of tRNA genes and those of mutant suppressor tRNAs are not included in this list, but appear in other compilations (4-6). REFERENCES 1 . R. P. Singhal and P. A. M. Fallis, Structure, Function, and Evolution of Transfer RNAs (with Appendix Giving Complete Sequences of 178 tRNAs), This Series 23,

227 (1979). 2. M. Sprinzl and D. H. Gauss, Compilation of tRNA Sequences, NARes 10, r l (1982). 3. D. H. Gauss, F. Gruter, and M. Sprinzl, Compilation of tRNA sequences, in “Transfer RNA: Structure Properties and Recognition” (R. R. Schimmel, D. Soll, and J. N. Abelson, eds.), p. 520. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1979. 4. M. Sprinzl and D. H. Gauss, Compilation of Sequences of tRNA Genes, NARes 10, r57 (1982). 5. J. E. Celis and P. W. Piper, Compilation of Mutant Suppressor tRNA Sequences, NARes 10, r83 (1982). 6. J. E. CeIis, Collection of Mutant tRNA Sequences, in “Transfer RNA: Structure, Properties and Recognition” (P. R. Schimmel, D. Soll, and J. N. Abelson, eds.), p. 539. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1979.

II. Abbreviations and Definitions A. tRNA Code Numbers Each sequence is numbered arbitrarily and carries, in addition, a single letter indicating the usual amino-acid acceptance (the letters are those recommended by IUPAC-IUB CBN (see, e.g., JBC 243, 3557). A number preceding the letter indicates prokaryotic origin; a numberfdlowing the letter indicates eukaryotic origin. Thus 1A represents an alanine tRNA from a prokaryote; R4 represents an arginine tRNA from a eukaryote. The one-letter code is as follows (Zoc. cit.):

RAM P. SINGHAL

2 16 A R N D C Q E G H I L

alanine arginine asparagine aspartic acid cysteine glutamine glutamic acid glycine histidine isoleucine leucine

fZt d.

K lysine M methionine iM F P S T W Y V

initiator methionine phenylalanine proline serine threonine tryptophan tyrosine valine

B. Abbreviations The symbols in parentheses are those recommended for use in text; those outside the parentheses, with notations above or beneath the primary symbol, are in the form recommended for use in long sequences (the aim being to reduce overall length to one character).

1. SPECIAL NOTATIONS N Unidentified residue Special comment for an uncharacterized residue; see the note cited with the reference for that sequence

R, Y A purine, a pyrimidine nucleoside p

Change occurs at this nucleotide position in the mutant tRNA structure (see note following the reference for this sequence).

2. URIDINEDERIVATIVES

A U

An unidentified modification of uridine

D

(hU) 5,6-dihydrouridine

T d U

(m5U) ribothymidine or 5-methyluridine uridine or dihydrouridine

f

(s2T)2-thioribothymidine (5-methyl-2-thiouridine)

T

(Tm) 2'-O-ribothymidine (2'-O-methyl-5-methyluridine)

U

(mo5U)5-methoxyuridine

m a

STRUCTURE OF TRANSFER

RNAS

217

(cmo5U) 5-carboxymethoxyuridine (also called uridin-5oxyacetic acid) (mcm5U) 5-(methoxycarbonylmethyl)uridine (also called uridin-5-oxyacetic acid methyl ester) (s4U)4-thiouridine

( s W )2-thiouridine (mcm5s2U)5-(methoxycarbonylmethyl)-2-thiouridine (mnm5s2U)5-methylaminomethyl-2-thiouridine (mnm5U) 5-methylaminomethyluridine (acp3U) 3-(3-amino-3-carboxypropyl)uridine (cmnm5U)5-carboxymethylaminomethyluridine

(q) pseudouridine (m “P) 1-methylpseudouridine

(qm) 2’-O-methylpseudouridine (Urn)2’-O-methyluridine Special comment for this residue, see the note cited with

the reference for that sequence.

3. CYTIDINE DERIVATIVES A C 3 C 5 C

E

An unidentified modification of cytidine (m3C) 3-methylcytidine (m5C)5-methylcytidine (ac4C) N4-acetylcytidine

(s2C) 2-thiocytidine

C

m

6

(Cm)2’-O-methylcytidine Special comments for this residue; see the note cited with

the reference for that sequence

218

RAM P. SINCHAL

et al.

4. ADENOSINE DERIVATIVES A An unidentified modification of adenosine A m A (mlA) 1-methyladenosine 2 (m2A)2-methyladenosine A 6 (m6A) N6-methyladenosine A

A i x K b

(i6A)N6-isopentenyladenosine (ms2i6A)N6-isopentenyl-2-methylthioadenosine N6-(threoninocarbony1)adenosine; (tc6A) N-[N-(B-P-D-ribofuranosylpurin-6-yl)carbamoyl]threonine

N6-Methyl-N6-(glycinocarbony1)adenosine; (mt6A)N-[N-(9~-~-ribofuranosylpurin-6-yl)-N-methylcarbamoyl] threonine

A

(ms2tfiA)N6-Methylthio-IP(threoninocarbony1)adenosine

R

N 6 - [Tris(hydroxymethyl)methylamidothreoninocarbonyl] adenosine; (@A) N-[N-(9-P-~-ribofuranosylpurin-6-yl)carbamoyllthreoninetris(hydroxymethyl)methylamide. (This compound is an artifact, being produced by enzymic amide bond formation between Tris buffer and t6A during isolation.)

A m

(A") 2'-O-methyladenosine

lt

Special comments for this residue; see the note cited with the reference for that sequence.

5. GUANOSINE DERIVATIVES A An unidentified modification of guanosine G m G (m'G) 1-methylguanosine 2 G (m2G)N2-methylguanosine

E

(miG)N2,N2-dimethylguanosine(not G as indicated earlier)

G

(m7G)7-methylguanosine

G m

(G") 2'-0-methylguanosine

+

Ly

STRUCTURE OF TRANSFER

6

219

RNAS

Special comments for this residue; see the note cited with the reference for that sequence.

6. OTHERMODIFIEDNUCLEOSIDES m I (m' I) 1-methylinosine W

(W) wyosine; 3-P-~-ribofuranosylwye; 4,9-dihydro-4,6-dimethyl-9-oxo-3-/3-~-ribofuranosyl-lH-imidazo[ 1,2-a]purine

Y

(yW) wybutosine; a-(carboxyamino)wyosine-7-butyricacid dimethyl ester

W

%

(oyW) peroxywybutosine; a-(carboxyamin0)-P-hydroperoxywyosine-7-butyric acid dimethyl ester (the natural product is probably hydroxybutosine rather than the peroxy compound; Kasai et aZ., NARes 6, 993, 1979).

(Q) queuosine; ribosylqueuine; 7-{[(cis-4,5-dihydroxy-2-cyclopenten- l-yl)amino]methyl}-7-deazaguanosine

Q

8 d

(manQ) P-D-mannosylqueuosine, found in mammalian tRNAAsP,residue 34

N

m

(galQ) P-D-galactosylqueuosine, tRNATyr,residue 34

found

in

mammalian

(N") 2'-O-methylribo-purine or -pyrimidine nucleoside

111. References and Footnotes for Sequence List ALANINE 8A. Bacillus subtilis: H. Ishikura, K. Murao, and Y. Yamada, Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980. 9A. Escherichia coli (1B): E. Lund and J. E. Dahlberg, Cell 11,247 (1977).

ARCININE R6. Yeast: G. Keith and G. Dirheimer, BBRC 92, 116 (1980). These authors have revised this tRNA sequence (cf. sequence R6, This Series 23,227). Both C and U residues and C and G residues have been found in positions 4 and 73, respectively. 7R. Bacillus subtilis: H. Ishikura, K. Murao and Y. Yamada, Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg (1980). R8. Mouse leukemia (1 and 2): F. Harada and S. Nishimura, Biochem. Int. 1, 539

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RAM P. SINGHAL

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(1980). The difference between isoacceptors tRNAPg and tRNAge is in position 50: P and C, respectively. R9. Morris hepatoma 5123 (mitochondrial): H. P. Ag;awal, R. C. Gupta, K. Randerath, and E. Randerath, FEBS Lett. 130,287 (1981). U-45: both U and G are found in this position. 10R. H . uolcanii (Halobacterium sp.): R. Gupta, doctoral dissertation, University of Illinois, Urbana, 1981; also see NARes 10, r l (1982). R11. Yeast (mitochondrial): G. Dirheimer, personal communication as cited in NARes 10, r l (1982).

ASPARACINE N3. Walker 256 mammary carcinosarcoma: B. A. Roe, A. F. Stankiewicz, H. L. Rizi, C. Weisz, M. N. DiLauro, D. Pike, C. Y. Chen, and E. Y. Chen, NARes 6,673 (1979). This sequence differs from that of the typical mammalian tRNAAS” (N2) in only the wobble position: Q34 + G34. (see earlier compilation for sequences of N2 and N3). 4N. Phage T5:V. M. Kryukov, M. G. Schlyapnikov, S. I. Kazantsev, A. V. Kaliman, V. N. Ksenzenko, and A. A. Bayev, Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980.

ASPARTATE D3. Beef liver: a tentative structure proposed by V. N. Vakharia, FP 40, 1753 (1981). m D4. Rabbit liver (2): V.N. Vakharia and R. P. Singhal, BBRC 105,1072, (1982). A-58: 1methyladenosine (40%) and adenosine (60%) are found in this position. D5. Rat liver and ascites hepatoma: Y. Kuchino, N. Shindo-Okada, N. Ando, S. Watanabe, and S. Nishimura,]BC 256,9059 (1981). 4-34: Rat liver contains mannosylqueuosine (manQ), but the tumor contains an unmodified G. D6. Morris hepatoma 5123D (mitochondrial-GUG): H. P. Agrawal, K. Randerath, and E. Randerath, NARes 9, 2535 (1981). 7D. H . oolcanii (Halobacterium sp.): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, r l (1982). 6-54: This residue is present mostly as 1-methylpseudouridine and in part as pseudouridine. 8D. Phage T5: V. M. Kryukov, M. G. Schlyapnikov, S. I. Kazantsev, A. V. Kaliman, V. N. Ksenzenko, and A. A. Bayev, Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980. 6-55: Present as 80% Prd and 20% U. 8-65: present as q r d (30%)and Urd (70%). D9. Euglena gracilis: W. G. Farmerie, S. H. Chang, and W. E. Barnett, FP 39, 2022 (1980).

CYSTEINE C3. Saccharomyces cereuisiae (mitochondrial). This sequence was deduced from gene analysis of mt DNA of tRNACys: J. L. Bos, K. A. Osinga, G. Van der Horst, and P. Borst, NARes 6,3255 (1979). (See earlier compilation for this sequence.)

GLUTAMATE E5. Schizosaccharornyces pombe: T. W. Wong, T. McCutchan, J. Kohli and D. Soll, NARes 6,2057 (1979). E6. Drosophila melanogaster (4): M. Altwegg and E. Kubli, NARes 8,215 (1980 .&32: 2’-0-methylcytidine is reported in less than 1 mol per mole of the tRNA. (3-49,& 50: A 5-MeCyd is found in position 49 and/or position 50.

STRUCTURE OF TRANSFER

221

RNAS

7E. H . oolcanii (Halobacterium sp.): R. Gupta, Ph.D. thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, r l (1982).

GLUTAMINE 5Q. H. uolcanii (Halobacterium sp.): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, r l (1982). 6Q. Phage T5: V. M. Kryukov et al., Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980. *-54,5-55: Urd (30%)was found in addition to the modified residue (70%)in each position.

GLYCINE 14G. Mycoplasma mycoides sp. Capri: M. W. Kilpatrick and R. T. Walker, NARes 8, 2783 (1980). 15G. Escherichia coli (two mutants: EMS-9 and SP-11) UGA sup mutant: N. E. Prather, E. J. Murgola, and B. H. Mims, NARes 9,6421 (1981). A-36: A is replaced by C in the wild type. 1-37: this residue is often ms2i6Ain the suppressor mutants. 16G. Bacillus subtilis (1): H. Ishikura, K. Murao, and Y. Yamada, Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980. G17. Yeast (mto): G. Dirheimer, personal communication as cited in NARes 10, r l (1982). A-37: This can exist as either i6A or msZi6A.

HISTIDINE H2. Saccharomyces cereuisiae (mitochondrial): *This sequence was deduced from the gene analysis of the mitochondrial DNA; M. Boisnard and G. Petrissant, FEBS Lett. 129, 180 (1981). H3. Yeast (mitochondrial): A. P. Sibler, R. P. Martin, and G. Dirheimer, FEBS Lett. 107, 182 (1979). H4. Drosophila melanogaster tRNA:'' (mutant): M. Altwegg and E. Kubli, NARes 8, 3259 (1980). $-54: is replaced by a U in the mutants; an additional, unpaired G is located at the 5' end. H5. Sheep liver: M. Boisnard and G. Petrissant, F E B S Lett. 129, 180 (1981). 6H. Phage T5: V. M. Kryokov et al., Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980.

ISOLEUCINE 31. Phage T4: C. Guthrie and W. H. McClain, Bchem 18, 3786 (1979). Differences between wild type and mutants are tabulated below.

Changes in the residue Strain Residue position:

4

69

70

71

72

Wild type HA1 (mutant) HA101 (revertant) HA103 (revertant)

C U U U

G G G

G G G A

U

U U C U

A

U

C U

RAM P. SINGHAL et al.

222

41. Escherichia coli (minor): Y. Kuchino, S. Watanabe, F. Harada, and S. Nishimura, Bchem 19,2085 (1980). 51. H. uolcanii (Halobacterium sp.): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, r l (1982). 16. Yeast (mitochondria): G. Dirheimer, personal communication, 1981, as cited in NARes 10, r l (1982).

LEUCINE L8. Schizosaccharomyces pombe sup 8-e mutant: R. Wetzel, J. Kohli, F. Altruda, and D. Soll, Mol. Gen. Genet. 172,221 (1979). L9. Bean (chloroplast-1): M. L. Osorio-Almeida, P. Guillemaut, G. Keith, J. Canaday, and J. H. Weil, BBRC 92,102 (1980). Residue 37 is either i6A or zeatin [N6-(trans4-hydroxyisopentenyl)adenine].

L10. Bean (chloroplast-2): M. L. Osorio-Almeida, P. Guillemaut, G. Keith, J. Canaday, and J. H. Weil, BBRC 92, 102 (1980). A-37: A can be PAA,ms2i6A,zeatin, or ms2zeatin [zeatin is N6-( truns-4-hydroxyisopentenyl)adenine]. L11. Bean and spinach (chloroplast-3): M. L. Osorio-Almeida, P. Guillemaut, C. Keith, J. Canaday, and J. H. Weil, BBRC 92, 102 (1980); J. P n a d a y , P. Guillemaut, R. Gloeckler, and J. Weil, Plant Sci. Lett. 20,57 (1980). U-47: Bean contains U, but spinach (chloroplast) contains G in this position. L12. Neurosporu crassa (mitochondria-1): J . E. Heckman, J. Sarnoff, B. Alzner-DeWeerd, S. Yin, and U. L. RajBhandary, PNAS 77, 3159 (1980). L13. Neurospora crassa (mitochondria-2): J. E. Heckman, J. Sarnoff, B. Alzner-DeWeerd, S. Yin, and U. L. RajBhandary, PNAS 77,3159 (1980). L14. Bovine liver: R. Pirtle, M. Kashdan, I. Pirtle, and B. Dudock, NARes 8,805 (1980). L15. Rat tumor Morris hepatoma 5123D: E. P n d e r a t h , R. C. Gupta, H. P. Morris, and K. Randerath, Bchem 19,3476 (1980). N-34: symbolized by the authors as “Mm” (residue*No. 35 in their numbering scGme) and considered to be a modification of Cm. N-37: A modification of mlG. N-44: symbolized by the authors as “L” (residue 45 in their numbering scheme) and considered to be a hypermodified residue-an unusual modification in this position of the tRNAs. This tRNA from two different preparations was found to differ in the position 49 (60% U and 40% A) and in positions 10, 12,34, and 47 : 2 (partially modified residues mZG, ac4C,Yr and A, respectively). L16. Cow mammary gland (2): M. A. Tukalo, I. G. Vasilchenko, V. V. Vlasov, and G. K. Matsuka, Ukr. Biokhim. Zh. 52,547 (1980). 5-12: partially exists as ac4C. L17 and L18. Anacystis niduluns (CAG and CAA); B. LaRue, N. Newhouse, K. Nicoghosian, and R. J. Cedergren, JBC 256, 1539 (1981). L19. Morris hepatoma 5123D (mitochondria-UAG): K. Randerath, H. P. Agrawal, and E. Randerath, BBRC 100,732 (1981).Position 5 contains a mixture of both U and G. 20L. H. uolcunii (Halobacterium sp.): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, r l (1982). 21L. Phage T5: V. M. Kryukov, M. G. Schlyapnikov, S. I. Kazantsev, A. V. Kaliman, V. N. Ksenzenko, and A. A. Bayev, Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980. L22. Yeast (mitochondria); G. Dirheimer, personal communication (1981) as cited in NARes 10, r l (1982). L23. Rhodosprillum rubrum: N. Newhouse, K. Nicoghosian, and R. L. Cedergren, Can. J . Biochem. 59,921 (1981). ‘This sequence extends beyond residue 76 with AAGGCACCA in positions 77 to 85.

STRUCTURE OF TRANSFER

223

RNAs

LYSINE K5. Drosophila melanogasier (2): S. Silverman, I. C. Gillam, G. M. Tener, and D. SOH, NARes 6,435 (1979). N-54 is probably Tm. K6, K7, and K8. Rabbit liver (1, 2, and 3, respectively); M. Raba, K. Limburg, M. Burghagen, J. R. Katze, M. Simsek, J. E. Heckman, U. L. RajBhandary, and H. J. Gross, EJB 97,305 (1979). Three isoacceptors differ only in 4 positions:

Residues in positions Isoacceptors of tRNALYs

Sequence code no.

29

34

37

41

1 2 3

K6 K7 K8

G A A

C C mcm5s2U

tc6A tc6A ms2t6A

C U

C

K9. Simian virus 40-transformed BALB/3T3 mouse fibroblasts: M. Raba, K. Limburg, M. Burghagen, J. R. Katze, M; Simsek, J. E. Heckman, V. L. RajBhandary, and H. J. Gross, EJB 97,305 (1979). U-27 is a mixture of U and JI; A-37 was not characterized, but was not tc6A. &-54 is a mixture of Tm, T, $, and U in the ratio of 60 :30 : 10 : 3, respectively. 10K. H. uolcanii (Halobacterium sp.): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, rl(l982). Residues 34,52, and 54 show partial modifications.

INITIATOR METHIONINE iM11. Scenedesmus obliquus (cytoplasm): P. 0. Olins and D. S. Jones, NARes 8, 715 (1980). For sequence, see previous listing of tRNAs [ThisSeries 23,227 (1979)], under iM11, where it is cited as a personal communication. iM12. Drosophila m e hogaste r: S. Silverman, J. Heckman, G. J. CowIing, A. D. Delaney, R. M. Dunn, I. C. Gillam, G. M. Tener, D. SOU, and U. L. RajBhandary, NARes 6,421 (1979). iM13. Asterina amurensis (starfish ovary): Y. Kuchino, M. Kato, H. Sugisaki, and S. Nishimura, NARes 6,3459 (1979). This sequence differs from that of the mammalian initiator tRNA (iM10) in nine nucleotide positions: 16,26,49-51,55,63,64, and 65. iM14. Tomlopsis utilis: S. Yamashiro-Matsumura and S. Takemura, J. Biochem. (ToA kyo) 86, 335 (1979). G-64 is a derivative of G or Gm. 15iM. Thennus thennophilus (1,2): K. Watanabe, Y. Kuchino, Z. Yamaizumi, M. Kato, T. Oshima, and S. Nishimura,J. Biochem. (Tokyo) 86,893 (197). 6-51and 6-63: The second isoacceptor contains G and C in these positions. iM16. Spinach chloroplast: J. L. Calagon, R. M. Pirtle, I. L. Pirtle, M. A. Kashdan, H. J. Vreman, and B. S. Dudock, JBC 255,9981 (1980). iM17. Scenedesmus obliquus (chloroplast): J. M. McCoy and D. S. Jones, NARes 8, 5089 (1980). iM18. Yeast (mitochondria): J. Canaday, G. Dirheimer, and R. P. Martin, NARes 8,1445 (1980). U-72 contains a mixture of U and $. iM19. Bean (chloroplast), and iM20, bean (cytoplasm): J. Canaday, P. Guillemaut, and J. H. Weil, NARes 8,999 (1980).

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iM21. Tetrahymena thermophila (cytoplasm): Y. Kuchino, M. Takashi, and S. Nishimura, NARes 9, 4557 (1981). 22iM. H . uolcanii (Halobacterium sp.): R.Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, rl (1982).

METHIONINE 5M. Bacillus subtilis W 168: Y. Yamada and H. Ishikura, NARes 8,4517 (1980). M6. Scenedesmus obliquus (chloroplast): D. S. Jones, Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980. M7. Spinach chloroplast: R. Pirtle, J. Calagan, M. Kashdan, H. Vreman, and B. Dudock, NARes 9, 183 (1981). 8M. Thermoplasma acidophilum: M. W. Kilpatrick and R. T. Walker, NARes 9,4387 (1981). M9. Tetrahymena thermophila (cytoplasm): Y. Kuchino, M. Takashi, and S. Nishimura, NARes 9, 4557 (1981). M10. Hat (mitochondria): G. Dirheimer, personal communication (1981) as cited in NARes 10, rl(l982). 6-49: there is an additional nucleoside (uridine) between G49 and A-50 of this sequence.

PHENYLALANINE F12 and F15. Pea: G. A. Everett and J. T. Madison, Bchem 15, 1016 (1976). This sequence is virtually identical to tRNAphefrom wheat germ (F12) and barley embryo (F15); see earlier listing of references for F12 and F15 [This Series 23, 227 (1979)l. F16. Drosgphila melanogaster (2):M. Altwegg and E. Kubli, NARes 7,93 (1979). 6-32 and U-47: partial modifications of the listed sequence of isoacceptor 2, C-32 Cm and/or U-47 ++ D modification yields four isoacceptors. F17. Neurospora crassa (cytoplasm): B. Alzner-DeWeerd, L. I. Hecker, W. E. Barnett, and U. L. RajBhandary, NARes 8,1023 (1980). F18. Bombyx mori: G. Keith and G. Dirheimer, BBRC 92, 109 (1980). 6-32 is sometimes found as Cm. 6 -4 7 is sometimes found as U. C-48 occurs more frequently as m5C than C in this position. e -4 9 occurs more frequently as C than m5C. G-57 is sometimes an A. F19. Bovine lens (1,2):F. K. Lin, T. D. Fun, S. H. Chang, J. Honvitz, P. F. Agris, and B. J. Ortwerth, JBC 255,6020 (1980). 6-57: Isoacceptor 1 differs from isoacceptor 2 in having A instead of G; note that isoacceptor 2 is identical to beef liver tRNAPhe (see sequence code No. F14). F20. Euglena gracilis (cytoplasm): S. H. Chang, L. I. Hecker, C. K. Brum, J. J. Schnabel, J. E. Heckman, M. Silberklang, U. L. RajBhandary, and W. E. Barnett, NARes 9,3199 (1981).This sequence differs from a previously published structure in four positions: 13, 14, 37, and 47. F21. Rhodospirillum rubrum: N. Newhouse, K. Nicoghosian, and R. J. Cedergren, Can.J . Biochem. 59, 921 (1981). F22. Bean and spinach (chloroplasts): J. Canaday, P. Guillemaut, R. Guillemaut, R. Gloeckler, and J. Weil, Plant Sci. Lett. 20, 57 (1980). The bean sequence is the same as the one published by P. Guillemaut and G. Keith, FEBS Lett. 84, 351 (1977), but contains G instead of C in position 44. There are differences between bean and spinach sequences in three positions, of which those changes at positions 17 and 39 are due to posttranscriptional modifications.

-

STRUCTURE OF TRANSFER

22s

RNAS Residues in positions

tRNAPhefrom: Bean chloroplasts Spinach chloroplasts

16 U C

17 D U

39

4 U

PROLINE P2. tRNA primer (1,2) for Moloney murine leukemia virus in mouse and chicken cells: JBC 254,10979 (1979). In positions 32 and 34, while isoacceptor 1 contains Urn and I, the second isoacceptor contains 4 and an unidentified derivative of U, respectively. 3P. Phage T5: V. M. Kryukov et al., Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980. 4P. H. uolcanii (Halobacterium sp.): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, rl (1982). P5. Spinach (chloroplast): M. A. Kashdan, H. Sprouse, L. Otis, and B. Dudock, FP 40, 1646 (1981).

SERINE S9. Hepatoma: E. Randerath, R. C. Gupta, R. J. Rhines, and K. Randerath, F P 39,2023 (Abstract No. 2199). This structure is identical to the one from rat liver tRNAISer (see sequence code No. S9 in the earlier compilation), except that it lacks the ribose methylation in position 18 (G instead of Gm) (1980). S11. Schizosaccharcomyces pombe spppressor 3-e: A. Rafalski, J. Kohli, P. Agris, and D. So11, NARes 6,2683 (1979). U-34 is a mixture of s2T and mcmW. S12. Beef heart (mitochondria) and bovine (mitochondria): P. Arcari and G. G. Brownlee, NARes 8,5207 (1980); M. H. deBruijn, P. H. Schreier, I. C. Eperon, B. G. Barrel], E. Y. Chen, P. W. Armstrong, J. F. H. Wong, and B. A. Roe, NARes 8, 5213 (1980). Each of 19 U residues of the beef heart are modified to m5U (T)in the bovine mitochondria; in addition, an unmodified A exists in position 37, and no CCA 3' end is found in the bovine mitochondrial structure. A-60: An additional G (insertion) occurs between residues 60 and 61. This sequence was deduced from gene analysis of mitochondrial DNA of tRNASer. S13. Human (mitochondria): M. H. deBruijn, P. H. Schreier, I. C. Eperon, B. G. Barrel], E. Y. Chen, P. W. Armstrong, J. F. H. Wong, and B. A. Roe, NARes 8,5213 (1980). A-60: An additional A (insertion) occurs between residues 60 and 61. This sequence was deduced from gene analysis of mitochondrial DNA of tRNASer. S14. Sacchoromyces cereuisiae (suppressor SUQ5-01): C. Waldron, B. S. Cox, N. Wilk, R. F. Gesteland, P. W. Piper, D. Colby, and C. Guthrie, NARes 9,3077 (1981).U35: U is replaced by G in the wild type. 15s and 16s. H. uolcanii (Halobacterium sp.) (1, 2): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, rl (1982). S17 and S18. Yeast (mitochondria) (1, 2): G. Dirheimer, personal communication (1981) as cited in NARes 10, rl(l982).The sequence No. S17 contains an insertion of G between positions 26 and 27 as indicated by an asterisk on 6-26.

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THREONINE T5. Spinach (chloroplast): M. A. Kashdan, R. M. Pirtle, I. J. Pirtle, J. L. Calagon, H. J. Vreman, and B. S. Dudock,JBC 255,8831 (1980). T6. Neurospora crassa (mitochondria): J. E. Heckman, J. Sarnoff, B. Alzner-DeWeerd, S. Yin, and U. L. RajBhandary, PNAS 77,3159 (1980).

TRYPTOPHAN W6. Avian sarcoma virus (primer for avian retrovirus DNA synthesis): B. Cordell, F. M. DeNoto, J. F. Atkins, R.F. Gesteland, J. M. Bishop, and H. M. Goodman,JBC 255, 9358 (1980). This sequence is identical to that present in uninfected avian cells [this common sequence was described as code No. W6 in the earlier compilation (ThisSeries 23, p. 288)], yet studies indicate that two conformations may exist influencing biological activities of the molecule. 8W. Esherichia coli temperature sensitive due to a lesion in the gene for tRNATrp:S. P. Eisenberg, L. SOU, and M. Yarus,JBC 254,5562 (1979). This mutant varies in only one base change, G-7 + A-7. For the sequence of E . coli tRNATv (code No. lW), see the earlier compilation (ThisSeries 23, p. 288). W9. Neurospora crassa (mitochondrial): J. E. Heckman, J. Sarnoff, B. Alzner-DeWeerd, S. Yin, and U. L. RajBhandary, PNAS 77,3159 (1980). W10. Yeast (mitochondrial): A. P. Sibler, R. Bordonne, G . Dirheimer, and R. Martin, C. R. Hebd. Seances Acad. Sci. Ser. D 290,695 (1980). W11. Spinach (chloroplast): J. Canaday, P. Guillemaut, R. Gloeckler, and J. H. Weil, NARes 9,47 (1981). 1-37 is a mixture of i6A or ms2ifiA. 12W. Escherichia cold (Su+7mutants): M. Yarus and L. Breeden, Cell 25, 815 (1981). The differences among various mutants including the one listed earlier (3W) are tabulated below.

~~

~~~

~~~

~

Residue in position E . coli tRNATv mutants (plasmids) pMY3” pMY31 pMY34 pMY313 pMY315

Sequencecode

22

29

56

3w 12w 12w 12w 12w

G A G G A

G G G

C C

A G

A C

C

Listed as pSU+7am in our previous listing.

TYROSINE Y6. Schizosaccharomyces pombe: G. Vogeli, NARes 7, 1059 (1979). 7Y. Bacillus subtilus (1,2): B. Menichi, H. H. Arnold, T. Heyman, G . Dirheimer, and G . Keith, BBRC 95,461 (1980). A-37: Isoacceptor 2 differs from isoacceptor 1in having either i6A or its modified residue ms2ifiAA. 8Y. H. uolcanii (Halobacterium sp.): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, rl (1982).

STRUCTURE OF TRANSFER

RNAS

227

Y9. Yeast (mitochondria): G. Dirheimer, personal communication, 1981, as cited in NARes 10, rl (1982). A-37: This residue exists as P A or ms2i6A.

VALINE V13. Rat liver (1): N. Shindo-Okada, Y. Kuchino, F. Harada, N. Okada, and S. Nishimura, J . Biochem. (Tokyo) 90,535 (1981). This sequence lacks A-76. Rat ascites hepatoma (2) was also published with this sequence and found to be identical to tRNAValfrom mouse myeloma and rabbit liver (see sequence code No. V11). 14V. Bacillus subtilis (1): H. Ishikura, K. Murao, and Y. Yamada, Abstracts cited in proceedings of EMBO-FEBS tRNA Workshop, Strasbourg, 1980. V15. Spinach (chloroplast): H. M. Sprouse, M. Kashdan, L. Otis, and B. Dudock, NARes 9,2543 (1981). V16. Neurospora crussu (mitochondrial): J. E. Heckman, J. Sarnoff, B. Alzner-DeWeerd, S. Yin, and U. L. RajBhandary, PNAS 77,3159 (1980). V17. Drosophila mela.ogaster (4):W. R. Addison, I. C. Gillam, and G. M. Tener, JBC 257,674 (1982).U-20 is partially modified to acpW in this sequence. It is probable that a cytidine residue between residues 47 and 51 is modified to m5C. 18V. H. oolcanii (Halobacterium sp.): R. Gupta, doctoral thesis, University of Illinois, Urbana, 1981, as cited in NARes 10, rl (1982).

IV. Sequences of 126 tRNAs See pages 228-241. Sequences for Ala, Ars, Asn, and Asp tRNAs, pp. 228-229; Glu, Gln, Gly, and His tRNAs, pp. 230-231; Ile and Leu (contd.) tRNAs, pp. 232-233; Leu, Lys, and Met-i (contd.) tRNAs, pp. 234-235; Met-i (contd.), Met, and Phe tRNAs, pp. 236-237; Pro, Ser, and Thr tRNAs, pp. 238-239; Trp, Tyr, and Val tRNAs, pp. 240-241.

ADDENDUM: Additional tRNA Sequences 1 . Structures of an additional 24 tRNA sequences included in proof. ( a ) References and footnotes, see p. 243. ( b )Sequences of 24 tRNAs. 2. Availability of a total of 318 tRNA sequences. A total of 293 sequences are listed in Table I (see p. 213), which includes the 126 sequences listed here. An additional 24 sequences are described on pp. 243 and following. A total of 318 tRNA sequences are thus available in the literature (as of Dec. 1982) including the one sequence that has just appeared: tRNA Code #F26: A. Mazabraud, The nucleotide sequence of tRNAPheof Xenopus lavis. Biochimie 64, 955 (1982). (The Addendum continues on p. 243.)

IsoNo. -

acceptor

Organism

Aminoacyl Stem

D Stem 910111213i4151617~1

D Loop

Anticodon Stem Anticodon Loop

D Stem -

2181920:1

2 3

22 24 26 28 30 32 34 36 38 2 1 23 25 27 29 3 1 33 35 37

ALANINE 8A 9A

1B

B. subtitis

pG-G-A-G-C-C-U-U-A-G-C-U-C-A-G-C-D

-G-G-G

a -A-G-A-G-C-G-C-C-U-G-C-U-U-U-G-C-A-C

6

E. coli

pG-G-G-G-C-U-A-U-A-G-C-U-G-A-G-C-D

4-G-G

-A-G-A-G-C-G-C-C-U-G-C-U-U-U-G-C-A-C

Yeast

pG-C-U-C-G-C-G-U-G-G-C-G-U-A-A-D

4-G-C

-A-A-C-G-C-G-$-C-U-G-A-C-U-U-C-U-A-A

B. subtilis

pG-C-G-C-C-C-G-U-A-G-C-U-C-A-A-U

4-G-A-D

-A-G-A-G-C-G-U-U-U-G-A-C-U-I-C-G-G-A

Mouse leukemia

pG-G-G-C-C-A-G-U-E-8-C-G-C-A-A-D

-G-G-A-D

-A-A-C-G-C-&-C-U-G-A-C-U-I-C-G-?-A

-A-A-A

-A-U-U-A-A-U-G-A-$-U-U-C-G-A-C-$-C-A

V

ARGININE R6

3

7R R8

1,2

R9

Hepatoma (mto)

*

m 2

m

pU-G-G-U-A-A-U-U-A-G-U-U-U-A-A-A-U

U

U

B

t m

m

10R

H. voZcanii

pG-U-C-C-U-G-A-U-A-G-G-G-$-A-G-U

4-G-A-C-U

-A-U-C-C-U-C-C-U-G-G-C-U-U-G-C-G-G-A

R11

Yeast (mto)

pG-C-U-C-U-C-U-U-A-G-C-U-U-A-A-D

-G-G-D-U

A -A-A-A-G-C-A-a-A-A-U-A-C-U-U-C-U-A-A

Phage T5

pG-G-U-U-C-C-U-U-A-G-C-U-C-U-A-A-U

-G-G-U-U

-A-G-A-G-C-C-G-C-A-C-C-U-U-G-U-U-A-A

Rabbit L i v e r

pU-C-C-C-G-U-C-U-A-G-U-A-a-A-G-U

-G-G-D-G

-A-G-U-A-U-A-U-C-C-G-C-C-U-Q-U-C-A-C

Rat L i v e r , Tumor

pU-C-C-U-C-G-U-U-A-G-U-A-$-A-G-U

-G-G-D-G

-A-G-U-A-U-C-C-C-C-G-C-U-C-Q-U-C-A-C

Hepatoma (mto)

pG-A-G-A-U-A-U-U-A-G-U-A-A-A-U

t

ASPARAGINE 4N

A

ASPARTATE

D4 D5 D6

2 "major"

m

-A

U

rl

*

5

-A-U-U-A-C-A-I$-A-A-C-C-U-U-G-U-C-A-A

7D

H. volcanii

pG-C-C-C-G-G-G-U-G-G-U-G-$-A-G-U

-G-G-C-C-C

-A-U-C-A-U-A-C-G-A-C-C-C-U-G-U-C-A-C

8D

Phage T5

pG-C-G-A-C-C-G-G-G-G-C-U-G-G-C-U-U

-G-G-U-A

-A-U-G-G-U-U-C-U-C-C-C-C-U-G-U-C-A-C

D9

E. gracilis

pU-C-U-U-C-G-G-U-A-G-U-A-$-A-G-D

-G-G-D-A

-A-G-U-A-U-G-$-C-C-G-C-C-U-G-U-C-A-C

rn

n

A

Isoac- Anticodon

No. ceptor -

Stem

V a r i a b l e Loop

W C Stem

T$C L o o p

T$C Stem

Aminoacyl Stem

3g4041424344454647: 1 2 3 4 5 6 7 8 91011121314151648495~515~53545556575~5g6~616263~4656667686g70717273747576 ALAN INE

+ 8A 9A

-C-A-G-C-G-G-T-JI-C-G-A-U-C-C-C-G-C-U-A-G-G-C-U-C-C-A-C-C-A

4-C-A-G-G-A-G-G-U

19

+ 4-C-A-G-G-A-G-G-U

-C-U-G-C-G-G-T-JI-C-G-A-U-C-C-C-G-C-A-U-A-G-C-U-C-C-A-C-C-A

-$I-C-A-G-A-A-G-A-D

-U-A-U-G-G-G-T-O-C-G-A-C-C-C-C-C-A-U-C-G-U-G-A-G-U-G-C-C-A

ARGININE R6

3

7R

R8 M

M (D

-U-C-A-A-A-A-G-G-U

1.2

m

+

*

-U-A-G-G-G-G-T-JI-C-G-A-C-U-C-C-C-C-U-C-G-G-G-C-G-C-G-C-C-A

5 * m -U-C-O-A-G-G-O-O-C-G-A-C-U-C-C-U-G-G-C-U-G-G-C-U-C-G-C-C-A

-$-C-A-G-A-A-G-A-D

*

R9

-U-U-A-G-A-U-U-A-U

-G-A-U-A-A-U-A-A-U-C-A-U-A-A-U-U-A-C-C-A-A-C-C-A

10R

-G-C-C- .'.-G-G-G-A

-C-C-G-G-A-G

R11

-$I-A-U-U-A-A-U-A-U

-U-C-C-A-U-G-T-O-C-A-A-A-U-C-A-U-G-G-A-G-A-G-A-G-U-A-C-C-A

2

5

-9-C-G-A-A-U-C-U-C-C-G-U-C-A-G-G-A-C-G-C-C-A

ASPARAGINE 4N

-G-@-U-G-A-G-G-G-U

+

ASPARTATE

5 5

m

-G-C-G-G-G-A-G-G-A

-C-C-G-G-G-G-T-O-C-G-A-A-U-C-C-C-C-G-G-A-C-G-G-G-A-G-C-C-A

D 5 "major"

-G-C-G-G-G-A-G-A

~-C-G-G-G-G-T-JI-C-G-A-U-U-C-C-C-C-G-A-C-G-G-G-G-A-G-C-C-A

D6

-G-G-U-U-A-A-G-U

-U-A-U-A-G-A-C-U-U-A-A-A

D4

2

5 5

*

-U-C-U-A-U-A-U-A-U-C-U-U-A-C-C-A

2

7D

4-G-U-C-G-U-G-A

C-G-C-G-G-G-O-O-C-G-A-A-U-C-C-C-G-C-C-U-C-G-G-G-C-G-C-C-A m

8D

4-G-G-A-G-A-G-A-A

D9

-G-C-G-G-A-A-G-A-C

-U-G-U-G-G-G-T-O-C-A-A-A-U-C-C-C-A-U-C-G-G-U-C-G-C-G-C-C-A A A n

*

A

*

~-C-C-G-G-G-T-~-C-A-A-U-U-C-C-C-G-G-C-C-G-G-A-G-A-G-C-C-A

No.

Isoacceptor Organism

Anticodon D Stem D Loop D Stem Stem Anticodon Loop 91011121314151617~1 2181920:1 2 3 22 24 26 28 30 32 34 36 38 21 23 25 27 29 31 33 35 37

Aminoacyl Stem

GLUTAMATE E5

m pU-C-C-G-U-U-G-U-G-G-U-C-C-A-A-C

S. pombe

4

-G-G-C-D

rl

-A-G-G-A-U-U-C-G-U-C-G-C-U-U-U-C-A-C t

2

Prosophila

pU-C-C-C-A-U-A-U-G-G-U-C-$-A-G-D

H. volcanii

PG-C-U-C-U-G-U-U-G-G-U-G-$-A-G-U-C-C

5Q

H. votcanii

PA-G-U-C-C-C-A-U-G-G-G-G-$-A-G-U

~m -G-G-C-C-A -A-U-C-C-U-G-U-U-G-C-C-U-U-C-U-G-G-G

69

Phage T5

pU-G-G-G-G-A-U-U-A-G-C-U-U-A-G-C-U-U

-G-G-C-U-C -A-A-A-G-C-U-U-C-G-G-C-C-U-U-U-G-A-A

14G

M. mycoides

pG-C-A-G-G-U-G-U-A-G-U-V-U-A-A-U

-G-G-C

-A-G-A-A-C-U-U-C-A-G-C-C-U-U-C-C-A-A

15G

E. coZi(UGA

pG-C-G-G-G-C-A-U-C-G-U-A-U-A-A-U

-G-G-C-U

-A-U-U-A-C-C-U-C-A-G-C-C-U-U-C-A-A-A

E6

7E

U

-A-G-G-A-U-A-U-C-U-G-G-C-U-U-U-C-A-C m A -G-G-C-C-A -A-U-C-A-U-A-U-C-A-C-C-C-U-N-U-C-A-C -G-G-C-D

GLUTAMINE U

GLYCINE

16G

1

G17

S

SUP)

A

6 t s

C

B. subtilis

pG-C-G-G-G-U-G-U-A-G-U-U-U-A-G-U

-G-G-D

-A-A-A-A-C-C-U-C-A-G-C-C-U-U-C-C-A-A

Yeast (mto)

PA-U-A-G-A-U-A-U-A-A-G-U-U-A-A-U-D

-G-G-D

-A-A-A-C-U-G-G-A-U-G-$-C-U-U-C-C-A-A

-A-G-A-A-A-A-U-A-C-G-C-U-U-G-U-G-G-U -A-G-A-A-A-A-$-A-C-G-C-$-U-G-U-G-A-$ m

U

t

HISTIDINE H2

S. cerewisiae

pG-G-U-G-A-A-U-A-U-A-U-U-U-C-A-A-U

-G-G-U

H3

Yeast (mto)

pG-G-U-G-A-A-U-A-U-A-U-U-U-C-A-A-D

-G-G-D

H4

pG-G-C-C-G-U-G-A-U-C-G-U-C-$-A-G-D 2 d pG-G-C-C-G-U-G-A-U-C-G-U-A-$-A-G-U

-G-G-D-D

H5

Drosophila (mutant) Sheep Liver

6H

Phage T5

pU-G-U-G-G-C-U-A-U-A-U-C-A-U-A-A-U-U

-G-G-U-U

(mto)*

-G-G-D-D

-A-G-G-A-C-C-C-C-A-C-G-$-U-G-U-G-G-C

m

-A-G-U-A-C-U-C-U-G-C-G-JI-U-Q-U-G-G-C

m

-A-A-U-G-G-U-C-C-U-G-A-U-U-G-U-G-A-A

Isoac- Anticodon Stem

No. c e p t o r. -

TJlc Stem

T$C Loop

T$C Stem

Aminoacyl Stem

394041424344454647: 1 2 3 4 5 6 7 8 9101112131415164849S0s1S2~3S4ssS6~758sg60616263646~6667686g70717273747576 GLUTAMATE

5

-C-G-A-C-G-G-G-A

-G-C-G-G-G-G-T-$-C-G-A-C-U-C-C-C-C-G-C-A-A-C-G-G-A-G-C-C-A

-C-C-A-G-A-A-G-G

-C-C-C-G-G-G-T-$-C-G-A-U-U-C-C-C-G-G-U-A-U-G-G-G-A-A-C-C-A

-G-G-U-G-A-U-G-A

5 m -C-C-A-G-G-G-$-$-~-G-A-A-U-C-C-C-U-G-A-C-G-G-A-G-C-A-C-C-A

SQ

-G-G-C-A-A-C-G-A

-C-C-C-A-G-G-$-$-I-G-A-A-U-C-C-U-G-G-U-G-G-G-A-C-U-A-C-C-A

69

4-$-C-G-A-G-A-U

-C-A-U-U-G-G-T-$-C-A-A-A-U-C-C-A-A-U-A-U-C-C-C-C-U-G-C-C-A

14G

4-C-U-G-A-U-U-G

-U-G-A-G-G-G-U-$-C-G-A-U-U-C-C-C-U-U-C-A-C-C-U-G-C-U-C-C-A

1SG

4-C-U-G-A-U-G-A

-U-G-C-G-G-G-T-~-C-G-A-U-U-C-C-C-G-C-U-G-C-C-C-G-C-U-C-C-A

4-C-U-G-A-U-G-U

-C-G-U-G-A-G-T-$-C-G-A-U-U-C-U-C-A-U-C-A-C-C-C-G-C-U-C-C-A

-A-C-A-U-U-G-A-A

-U-G-C-G-A-G-T-~-C-G-A-U-U-C-U-C-G-C-U-A-U-C-U-A-U-A-C-C-A

H2

4-C-G-U-U-A-A-A

-U-C-U-G-A-G-U-U-C-G-A-U-U-C-U-C-A-G-U-A-U-U-C-A-C-C-C-C-A

n3

-G-C-G-U-U-A-A-A

-U-C-U-G-A-G-T-$-C-G-A-U-U-C-U-C-A-G-U-A-U-U-C-A-C-C-C-C-A

H4

-C-G-U-G-G-U-A-A

-~-C-C-A-G-G-~~-$-C-G-~-A-U-C-C-U-G-G-U-C-A-C-G-G-C-A-C-C-A

HS

-C-G-C-A-G-C-A-A

5 m -C-C-U-C-G-G-U-$-C-G-A-A-U-C-C-G-A-G-U-C-A-C-G-G-C-A-C-C-A

6H

-$-C-A-G-G-C-C-U-A

E5 E6

4

7E

* *

GLUTAMINE

tQ

z

* *

GLYCINE

16G

1

G17 HISTIDINE

A

5

5 5

-U-G-U-G-G-A-T-$-C-G-A-A-U-U-C-U-A-C-U-A-G-C-C-A-C-A-C-C-A

No.

Isoacceptor

Organism

Aminoacyl Stem

D Stem

D Loop

91011121314151617~1

2181920:1

A n t icodon Stem Anticodon Loop

D Stem 2 3

22 24 26 28 30 32 34 36 38 21 23 25 27 29 3 1 33 35 37

ISOLEUCINE 31

Phage T4

pG-G-C-C-C-U-G-U-A-G-C-U-C-A-A-U S

-G-G-D-D-A

-G-C-A-G-C-A-G-U-C-C-G-C-U-N-A-U-N-A

m

t

E. c o l i

pG-G-C-C-C-C-U-U-A-G-C-U-C-A-G-U

-G-G-D-D

-A-G-A-G-C-A-A-G-C-G-A-C-U-N-A-U-A-A

51

H. volcanii

pG-G-G-C-C-A-A-U-A-G-C-U-C-A-G-U-C-A

-G-G-U

-u-G-A-G-C-&C-$-C-G-G-C-U-G-A-U-N-A

I6

Yeast ( m t o )

pG-A-A-A-C-U-A-U-A-A-U-U-C-A-A-D-D

-G-G-D-U

-A-G-A-A-U-A-G-U-A-U-$-U-U-G-A-U-A-A

-G-G-D-G-D

-A-A-G-G-G-G-G-C-A-G-A-$-U-N-C-A-G-C

41

2

LEUCINE S.

pombe(sup 8-e) pG-C-G-G-C-U-A-U-G-C-C-C-G-A-G-D

A

m

A

*

L9

1

Bean (chl)

PG-G-G-G-A-U-A-U-G-G-C-G-A-A-A-U-U

-G-G-D-A

-G-A-C-G-C-I$-A-C-G-G-A-C-U-N-A-A-A-A

L10

2

Bean (chl)

pG-G-C-U-U-G-A-U-G-G-U-G-A-A-A-U-U

-G-G-D-A m

-G-A-C-A-C-G-C-G-A-G-A-C-U-C-A-A-A-A

L11

3

Bean (chl)

pG-C-C-G-C-U-A-U-G-G-U-G-A-A-A-U-U

-G-G -D -A

-G-A-C-A-C-G-C-U-G-C-U-C-U-U-A-G-G-A

L12

1

N. c r a m (mto)

PA-U-C-C-G-A-G-U-G-A-U-G-G-A-A-D

-G-G-D-A

-G-A-C-A-U-A-A-C-A-U-G-C-U-U-A-A-A-A

L13

2

N.

PA-U-A-G-G-U-G-U-G-C-U-G-G-A-A-D-U

-G-G-D-A

4-A-C-A-G-G-U-U-C-C-G-$-U-U-A-G-G-C

-G-G-D-C-$

-A-A-G-G-C-G-C-U-G-G-A-$-U-I-A-G-G-C

-G-G-D-C-$

U * -A-A-G-G-C-G-C-C-A-G-A-C-U-N-A-A-N-$ m U

E3

w t o

U

K

L8

CPQSSU (rnto)

2

K

L14

Bovine L i v e r

pG-G-U-A-G-C-G-U-G-G-C-C-G-A-G-C

L15

Hepatoma

pG-U-C-A-G-G-A-U-G-G-C-C-G-A-G-U

2 2

2

2

K *

pG-U-C-A-G-G-A-U-G-G-C-C-G-A-G-C

m

m

+ m

A

U

m m

m

* m

L17

A. niduzam (CAG) pG-C-G-G-A-A-C-U-G-G-C-G-G-A-A-U-D

-G LG-D -A

-A-A-G-G-C-G-C-U-G-C-G-U-U-C-A-G-G-N m m -G-A-C-G-C-G-C-U-A-G-A-U-U-C-A-G-G-$

L18

-G-G-D-A

-G-A-C-G-C-A-G-C-A-G-A-C-U-C-A-A-A-A

L19

A. nidulans (CAA) pG-G-G-C-A-A-G-U-G-G-C-G-G-A-A-U-D * m2 Hepatoma (mto) PA-C-U-U-U-U-A-U-A-G-G-A-U-A-G-A-A

-A-G-D

-A-A-U-C-C-A-$-U-G-G-U-C-U-U-A-G-G-A

20L

H. volcanii

-G-G-C-C-A

L16

2

Cow Mam. Gland

pG-C-G-U-G-G-G-U-A-G-C-C-A-A-N-C-C-A

-G-G-D-D-C

-A-C-G-G-C-:-C-A-G-C-G-U-U-G-A-G-G-G

A m

m

Isoac- Anticodon No. ceptor -

Variable Loon

Stem

T$C Stem ~ _

T$C Loo? _ _

TQC Stem _ _

_Aminoacyl Stem

394041424344454647: 1 2 3 4 5 6 7 8 910111213141516484950~15253545~565758596061626364656667686g70717273747576 ISOLEUCINE

+ 31 41

-G-G-G-A-A-A-G-G-U

2

-$-C-G-C-U-U-G-G-U

+71

5

* * * *

-U-A-C-C-A-G-T-JI-C-A-A-A-U-C-U-G-G-U-C-U-G-G-G-U-C-A-C-C-.4 -C-G-C-U-G-G-T-*-C-A-A-G-U-C-C-A-G-C-A-G-G-G-G-C-C-A-C-C-A

5

2

51

-C-C-G-G-G-A-G-G-C

-C-C-G-C-G-G

16

4-G-U-A-C-A-A-A

-U-A-U-A-G-G-T-*-C-A-A-U-C-C-C-U-G-U-UA-G-U-U-U-C-A-C-C-A

-C-C-U-G-C-U-G-U-U-G-U-A-A-A-A-C-G

-C-G-A-G-A-G-T-$-C-G-A-A-C-C-U-C-U-C-U-G-G-C-C-G-C-A-C-C-A

-$-C-G-A-A-U-C-C-G-C-G-U-U-G-G-C-C-C-A-C-C-A

LEUCINE

m

5

L8

L9

1

-$-C-C-G-U-C-G-A-C-U-U-A-A-U-A-A-A-U-C-A

-U-G-A-G-G-G-T-*-C-A-A-G-U-C-C-C-C-U-C-U-A-U-C-C-C-C-A-C-C-A

L10

2

-U-C-U-C-G-U-G-C-U-A-A-A-G-A-G-C-G

-U-G-G-A-C-G-T-~-C-G-A-G-U-C-C-U-C-C-U-C-U-U-C-A-A-G-U-C-A-C-C-A

L11

3

-A-G-C-A-G-U-G-C-U-A-G-A-G-C-A

-U-C-U-C-G-G-T-~-C-G-A-G-U-C-C-G-A-G-U-A-G-C-G-G-C-A-C-C-.4

L12

1

-C-A-U-G-U-G-G-G-C-U-U-C-A-A-G-C-U-G

-U-G-A-A-G-G-T-*-C-A-A-G-U-C-C-U-U-C-U-U-C-G-G-A-U-A-C-C-A

L13

2

-C-G-G-A-A-U-G-G-U-U-U-A-A-A-A-A-C-U-G

-U-A-C-A-A-G-T-*-C-A-.~-G-U-C-U-U-G-U-C-A-U-C-U-A-U-A-C-C-A 5 m

ta

0 0

-$-C-C-A-G-U-C-$-C-$-U-C-G-G-G-G-G * A *

L14

-C-G-U-G-G-G-T-JI-C-G-A-A-U-C-C-C-A-C-C-G-C-U-G-C-C-A-C-C-A

-$-C-U-G-G-N-U-C-C-G-N-A-U-G-G-A-G

5 m -C-G-U-G-G-G-T-JI-C-G-A-A-U-C-C-C-A-C-U-U-C-U-G-A-C-A-C-C-A

-C-G-C-A-G-ll-C-$-C-C-C-U-G-G-A-G-G

5 -C-G-U-G-G-G-T-*-C-G-~-A-U-C-C-C-A-C-U-U-C-U-G-A-C-A-C-C-A

L17

-$I-C-U-A-G-U-G-G-U-U-U-C-A-C-G-A-C-U-G

-U-C-C-G-G-G-T-$-C-A-A-G-U-C-C-C-G-G-G-G-U-U-C-C-G-C-A-C-C-A

L18

-$-C-U-G-C-C-G-C-U-A-G-C-G-A-U-A-G-U-G

-U-G-U-G-G-G-T-*-C-G-A-G-U-C-C-C-A-C-C-U-U-G-C-C-C-A-C-C-A 5

L19

-A-C-C-A-A-A-A-A

-C-C-U-U-G-G-U-G-C-A-A-C-U-C-C-A-A-A-U-A-A-A-A-G-U-A-C-C-A

20L

5 -C-G-C-U-G-U-C-C-U-G-U-A-G-A-G-G-U-C

-C-G-C-C-G-G

L15 L16

2

m

5

2

-$-C-G-A-A-U-C-C-G-G-U-C-C-C-A-C-G-C-A-C-C-A

Anticodon D Stem D Loo~ D Stem Stem Anticodon Loop 91011121314151617~1 2181920:1 2 3 22 24 26 28 30 32 34 36 38 21 23 25 27 29 31 33 35 37

IsoNo.

acceptor Organism

Aminoacyl Stem

21L

Phage T5

pG-G-G-G-C-U-A-U-G-C-U-G-G-A-A-C-U

L22

Yeast (mto)

pG-C-U-A-U-U-U-U-G-G-U-G-G-A-A-D-D

L23

R. rubrun:

pG-C-C-U-U-U-G-U-A-G-C-G-G-A-A-D

A

-G-G-D-A

-G-A-C-A-A-U-A-C-G-G-C-C-U-U-A-G-A-U

-G-G-D-A

-G-A-C-A-C-G-A-U-A-C-$-C-U-U-A-A-G-A

- G-G -D-A

-A-C-G-C-G-G-C-A-G-A-C-U-C-A-A-A-A-JI

-G-G-D

-A-G-A-G-C-A-$-G-A-G-A-C-U-C-U-U-A-A

-G-G-D

-A-G-A-G-C-A-$-G-G-G-A-C-U-C-U-U-A-A

-G-G-D

-A-G-A-G-C-A-$-C-A-G-A-C-U-U-U-U-A-A

A

U

6

m

LYSINE L

K5

2

Drosophila

pG-C-C-C-G-G-C-U-A-G-C-U-C-A-G-D-C

K6

1

Rabbit Liver

pG-C-C-C-G-G-C-U-A-G-C-U-C-A-G-D-C

K8

3

Rabbit Liver

pG-C-C-C-G-G-A-U-A-G-C-U-C-A-G-D-C

L

L

234

L

t

a

t

b

*

K9

SV40 fibroblasts pG-C-C-C-G-G-C-U-A-G-C-U-C-A-G-D-C

-G-G-U

-A-G-A-G-C-A-U-G-A-G-A-C-U-C-U-U-A-A

10K

W. uo lcanii

-G-G-C

-A-G-A-G-C-G-U-C-U-G-A-C-U-C-U-U-N-A m

-G-G-A

-A-G-C-G-U-G-C-U-G-G-G-C-C-C-A-U-A-A

-G-G-A

-A-G-C-G-U-G-C-U-G-G-G-C-C-C-A-U-A-A

-G-G-A

-A-G-C-G-C-G-C-A-G-G-G-C-U-C-A-U-A-A

-G-G-D m -G-G-D

-A-G-C-U-C-G-U-C-G-G-G-C-U-C-A-U-A-A m

-G-G-D

-A-G-C-U-C-G-$-G-G-G-G-C-U-C-A-U-A-A

pG-G-G-C-C-G-G-U-A-G-C-U-C-A-N-U-U-A

5

A

METHION INE-INITIATOR

iM12

Drosophila

iM3

A . amurensis

iM14

T.

1534

1,2

m 2

PA-G-C-A-G-A-G-U-G-G-C-G-C-A-G-U

m 2 PA-G-C-A-G-A-G-U-G-G-C-G-C-A-G-U m 2 pA-G-C-G-U-C-U-U-G-G-C-G-C-A-G-D

is

l l t i1

S

2

U

t t t

T. t h e r m o p h i h s

pC-G-C-G-G-G-G-U-G-G-A-G-C-A-G-C-C-U

iN16

Spinach (chl)

pC-G-C-G-G-G-G-U-A-G-A-G-C-A-G-U-U-U

it117

S. obZi,Tms

iM18

Yeast (mto)

pU-G-C-A-A-U-A-U-G-A-U-G-U-A-A-D-U

-G-G-D-U

m -A-A-C-A-U-U-U-U-A-G-G-G-U-C-A-U-G-A

iM19

Bean (chl)

pC-G-C-G-G-A-G-U-A-G-A-G-C-A-A-C-U-U

-G-G-D

-A-G-C-U-C-G-C-A-A-G-G-C-U-C-A-U-A-A

(chl) pC-G-C-A-G-G-A-U-A-G-A-G-C-A-G-U-C-U

-A-G-C-U-C-G-C-A-A-G-G-C-U-C-A-U-A-A U

Isoac- Anticodon Stem

No. c e p t o r -

V a r i a b l e Loop

TW Stem

T$C Loop

TJlC S t e m

Aminoacyl S t e m

3 ~ ~ ~ 4 1 ~ ~ 4 3 1~ 2~ 34 45 5~ 6~ 74 87 910111213141516484950~1525354~~56~758~g60616263646~6667686g70717273747576 : 21L

-$-C-C-G-U-A-G-C-U-U-A-A-A-U-G-C-G

-U-G-G-G-A-G-T-$-C-G-A-G-U-C-U-C-C-C-U-A-G-C-C-C-C-A-C-C-A

L22

-$-G-U-A-U-U-A-C-U-U-U-A-C-A-G-U-A

-U-G-A-A-G-G-T-Jl-C-A-A-G-U-C-C-U-U-U-A-A-A-U-A-G-C-A-C-C-A

L23

-C-U-C-C-U-U-U-G-T,

-U-A-A-C-C-C-A-G-G-U-G-G-U-A-G-T-*-C-G-A-C-U-C-U-C-C-C-C-A

LYSINE 2

-$-C-U-C-A-G-G-G-D

K6

1

-$-C-C-C-A-G-G-G-D

K8 @a

CJl 0

+

K5

3

-$-C-U-G-A-G-G-C-0

m

* A

-C-G-U-G-G-G-N-U-C-G-A-G-C-C-C-C-A-C-G-U-U-G-G-G-C-G-C-C-A

5 m -C-G-U-G-G-G-T-$-C-G-A-G-C-C-C-C-A-C-G-U-U-~-G-G-C-G-C-C-A

+ +

K9

-$-C-U-C-A-G-G-G-D

1 OK

-U-C- A-G- A-C - G-G-U

m

5 5

-C-C-A-G-G-G-T-$-C-A-A-G-U-C-C-C-U-G-U-U-C-G-G-G-C-G-C-C-A 5 m

-C-G-U-G-G-G-T-$-C-G-A-G-C-C-C-C-A-C-G-U-U-G-G-G-C-G-C-C-A

5

A

X

2

-C-G-C-G-$-G-U-$-C-G-A-A-U-C-G-C-G-U-C-C-G-G-C-C-C-A-C-C-A

METHIONE INITIATOR -C-C-C-A-G-A-G-E-D

ibll3

-C-C-C-A-G-A-G-G-D

iM14

-C-C-C-U-G-A-U-G-D

15iM 1,2

-C-C-C-G-A-A-G-G-U

iM16

-C-C-U-U-G-A-G-G-U

iM17

-q-C-C-C-A-A-U-G-D

iM18

-C-C-U-A-A-U-U-A

iM19

m

5

iF.112

-C-C-G-A-G-G-A-U-C-G-A-A-A-C-C-U-U-G-C-U-C-U-G-C-U-A-C-C-A

+ + +

m

5 5

a

-C-C-C-U-G-G-A-U-C-G-A-A-A-C-C-A-G-G-A-G-A-C-G-C-U-A-C-C-A *

s

m

*

-C-G-C-C-G-G-T-*-C-A-A-A-U-C-C-G-G-C-C-C-C-C-G-C-A-A-C-C-A

+

-C-A-C-G-G-G-T-Jl-C-A-A-A-U-C-C-U-G-U-C-U-C-C-G-C-A-A-C-C-A -C-G-C-A-G-G-T-$-C-A-A-A-U-C-C-U-G-C-U-C-C-U-G-C-A-A-C-C-A

-U-A-U-A-C-G-T-Jl-C-A-A-A-U-C-G-U-A-U-U-A-U-U-G-C-U-A-C-C-A +TI

-C-C-U-U-G-A-A-G-U

5 m -C-C-G-A-G-G-A-~-C-G-A-A-A-C-C-U-C-G-C-U-C-U-G-C-U-A-C-C-A

-U-A-C-G-G-G-T-IL-C-A-A-A-U-C-C-C-G-U-C-U-C-C-G-C-A-A-C-C-A

IsoNo. a c c e p t o r -

Organism

D Stem

Am inoacyl Stem

Anticodon Stem A n t i c o d o n Loop

D Stem

91011121314151617~1 2181920:1 2 3 22 24 26 28 30 32 34 36 38 21 23 25 27 29 31 33 35 37 U

m 2

t

iM20

Bean ( c y t )

PA-U-C-A-G-A-G-U-G-G-C-G-C-A-G-C

-G-G-A

-A-G-C-G-U-G-G-U-G-G-G-C-C-C-A-U-A-A

iM21

T . thermophita

m PA-G-C-A-G-G-G-U-G-G-C-G-A-A-A-D

-G-G-A

-A-U-C-G-C-G-U-$-G-G-G-C-U-C-A-U-A-A

22iM

H . volcnnii

PA-G-C-G-G-G-A-U-G-G-G-A-$-A-N-C-C-A

-G-G-A-G

-A-U-U-C-C-G-C-C-G

5M

R. s u b t i l i s

pG-G-C-G-G-U-G-U-A-G-C-U-C-A-G-C

-G-G-C-D

-A-G-A-G-C-G-U-A-C-G-G-U-U-C-A-A-C

M6

s.

-G-G-C-C

-A-G-A-G-C-A-N-C-C-G-$-$-U-C-A-U-A-C

M7

Spinach (chl)

-G-G-D-D

-A-G-A-G-$-A-$-$-G-C-$-U-U-C-A-U-A-C

-G-G-A

-G-G-A-G-C-G-C-C-G-G-A-,$-U-C-A-U-A-A

t

(CYt)

-G-G-G-C-U-C-A-U

METHIONTNE

Obtiquus ( c h l ) pG-C-C-U-G-C-U-~-A-G-C-U-C-A-G-U-D

pA-C-C-U-A-C-U-U-A-A-C-U-C-A-G-C

L

U

S

8M

T. acidophitum

p~-C-C-G-G-G-G-U-G-G-C-U-C-A-N-C-u

M9

T. thermophita

pA-G-C-A-G-G-G-U-G-G-C-G-A-A-A-D

-G-G-A

-A-U-C-G-C-G-U-$-G-G-G-C-U-C-A-U-A-A

M10

(CYt) R a t (mto)

pG-C-U-U-G-U-A-U-A-G-U-U-U-A-A-D-D

-G-G-D-U

-A-A-A-A-C-A-U-U-U-G-$J-C-U-C-A-U-A-A

Wheat,barley,pea

pG-C-G-G-G-G-A-U-A-G-C-U-C-A-G-D-D

-G-G-G

-A-G-A-G-c-&-c-A-G-A-c-u-G-A-A-w-A m

DrosophiZa

pG-C-C-G-A-A-A-U-A-G-C-U-C-A-G-D-D 2 A pG-C-G-G-G-U-U-U-A-G-C-U-C-A-G-D-D

-G-G-G

-A-G-A-G-C-G-$-$-A-G-A-C-U-G-A-A-G-A

-G-G-G -G-G-G

-A-G-A-G-c-&J-c-A-G-A-~u-~-A-A-I;-A U * -A-G-A-G-C-G-$-$-A-G-A-C-U-G-A-A-G-A

-G-G-C

-A-G-A-G-C-~-JI-JI-A-G-A-~-U-S-A-A-);-A

m

6

t t

t

PHENYLALANINE

F12,FlS F16

2

N. crassa ( c y t )

F17 F18

F19

1,2

2

L

2

Bombyx mori

pG-C-C-G-A-A-A-U-A-G-C-U-C-A-G-D-D

B o v i n e Lens

pG-C-C-G-A-A-A-U-A-G-C-U-C-A-G-D-D

2

m

2

U

*

0

m

U

F20

E. qrac. ( c y t )

pG-C-C-G-A-C-U-U-A-G-C-U-C-A-G-D-D

-G-G-G

-A-G-A-G-C-G-$-$-A-G-A-&-U-$,-A-A-W-A

F21

R. mbmm

pG-C-C-C-G-G-G-U-A-G-C-U-C-A-G-C-D

-G-G-D

-A-G-A-G-C-A-C-G-U-G-A-C-U-G-A-A-A-A

F22

Spinach ( c p t )

pG-U-C-G-G-G-A-U-A-G-C-U-C-A-G-C-U-D

-G-G-D

-A-G-A-G-C-A-G-A-G-G-A-C-U-G-A-A-A-A

m

m

m

Y s s

I s o a c - Anticodon Stem

V a r i a b l e Loop

No. c e p t o r -

T+€ Stem

T$C Loop

T$C Stem

Aminoacyl Stem

394041424344454647: 1 2 3 4 5 6 7 8 910111213141516484950~152~354555657585960616263646~6667686g70717273747576 + 5 m

iM20

-C-C-C-A-G-G-A-$-C-G-A-A-A-C-C-U-G-G-C-U-C-U-G-A-U-A-C-C-A

-C-C-C-A-C-A-G-G-D

+

iM21

-C-$-C-A-A-A-A-G-U

22iM

-A-A-C-C-C-G-G-A-G-A-U

m

5

-C-A-G-A-G-G-A-$-C-G-A-A-A-C-C-U-C-U-C-U-C-U-G-C-U-A-C-C-A L

-C-G-G-U-A-*-C-C-A-A-U-C-U-A-C-C-U-C-C-C-G-C-U-A-C-C-A

>lETHIONINE 5M

-C-C-C-U-G-A-G-G-U

-C-G-G-G-G-G-T-$-C-G-A-U-C-C-C-C-U-C-C-G-C-C-G-C-U-A-C-C-A

+

?46

-G - C -C-G- A- A- A- G-D

h17

-G-G-C-G-G-C-A-G-U

Rbl

-U-C-C-G-G-A-G-G-U

-C-U-C-G-G-G-*-$-C-G-A-U-C-C-C-C-G-A-U-C-C-C-G-G-C-A-C-C-A

PI9

-C-IJ-C-A-A-A-A-&U

-C-A-C-A-G-G-A-*-C-G-A-A-A-C-C-U-C-U-C-U-C-U-G-C-U-A-C-C-A

hll0

-A-U-A-A-k-U-A-A

-U-G-A-A-G-C-T-III-C-A-A-U-U-C-C-U-U-C-U-A-C-A-A-G-U-A-C-C-A

+ll

-C-A-C-U-A-G-T-I-C-G-A-A-U-C-U-A-G-U-A-G-C-A-G-G-C-N-C-C-A -C-A-U-U-G-G-T-*-C-A-A-A-U-C-C-A-A-U-A-G-U-A-G-G-U-A-C-C-A

5

m

PHENYLALANINE

+

F12,FlS

-$-C-U-G-A-A-G-G-D

F16

-$-C-U-A-A-A-G-G-11

2

5

F17

-$-C-II-G-A-A-G-G-D

F18 F19

-$-C-U-A-A-4-G-G-D

1,2

+ *

+ + *

-I-C-U-A-A-A-G-G-D

F20

-$-C-U-A-A-A-G-C-U

F21

-$-C-A-C-G-C-U-G-U -u-c-c-u-c-G-u-c-u

m

-C-C-C-C-G-G-T-$-C-A-A-U-C-C-C-G-G-G-U-U-U-C-G-G-C-A-C-C-A

m

5

-C-G-U-G-U-G-T-*-C-G-A-U-C-C-A-C-A-C-A-A-A-C-C-G-C-A-C-C-A

* *

t m

5

* m

-C-C-C-U-G-G-T-*-C-G-A-U-C-C-C-G-G-G-U-U-U-C-G-G-C-A-C-C-A -~-~-C-U-G-G-T-~-C-G-A-U-C-C-C-G-G-G-U-U-U-C-G-G-C-A-C-C-A

+ +

F22

m

-C-G-C-G-U-G-T-$-C-G-A-U-C-C-A-C-G-C-U-C-A-C-C-G-C-A-C-C-A

+ T I

m

-C-C-C-U-G-G-T-JI-C-G-A-U-C-C-C-G-G-G-A-G-$-C-G-G-C-A-C-C-A -C-G-G-U-G-G-T-$-C-G-A-C-U-C-C-G-C-C-C-C-C-G-G-G-C-A-C-C-A -C-A-C-C-A-G-T-$-C-A-A-A-U-C-U-G-G-U-U-C-C-lJ-G-G-C-A-C-C-A

IsoNo. -

acceptor

Organism

Aminoacyl Stem

D Stem

D Loop

Anticodon Stem Anttcodon Loop

D Stem

91011121314151617:1 2181920:1 2 3 22 24 26 28 30 32 34 36 38 21 23 25 27 29 31 33 35 37 PROLINE

pG-G-C-#-C-G-U-U-G-G-U-C-$-A-G

-G-G-G-D

* * m -A-U-G-A-U-U-C-U-C-G-C-N-U-N-G-G-G-$

3P

Mouse,growing cells Phage T5

pC-U-C-C-G-A-U-U-A-G-C-U-C-A-A-U-U

-G-G-C-D

-A-G-A-G-U-A-C-A-C-C-G-U-U-U-G-G-G-G

4P

H. volcanii

pG-G-G-C-C-G-G-U-G-G-G-G-$I-A-N-C-U-U

-G-G-U

-A-U-C-C-U-U-C-G-G-C-C-U-U-C-G-G-G-$

P5

Spinach (chl)

PA-G-G-G-A-U-G-U-A-G-C-G-C-A-G-C-U-U

-G-G-D

-A-G-C-G-C-$-$I-U-U-G-U-$-U-N-G-G-N-$

s11

S. pombe (sup)

pG-U-C-A-C-U-A-U-G-U-C-C-G-A-G-D

-G-G-D-D

-A-A-G-G-A-G-$-U-A-G-A-C-U-U-C-A-A-A

s12

Beef Heart (mto)

pG-A-A-A-A-A-G-U-A-U-G

C-A-A-G-A-A-C-U-G-C-U-A-A

s12

Bovine (mto)

pG-A-A-A-A-A-G-T-A-T-G

C-A-A-G-A-A-C-T-G-C-T-A-A

513

Human (mto)

pG-A-G-A-A-A-G-C-U-C-A

C-A-A-G-A-A-C-U-G-C-U-A-A

514

S. cerevisiae (Sup)

pG-G-C-A-C-U-A-U-G-G-C-C-G-A-G-D

-G-G-D-D

R. volcanii

PG-U-U-G-C-G-G-U-A-G-C-C-A-A-N-C-C-U

4-G-C-C-C

P2,PZ'

1,2

2

m

4 3

m

A

m A

*

i

SERINE K

K

0

0

3

A *

-A-A-G-G-C-G-A-C-A-G-A-N-U-U-U-A-A-A U

t

t

i

-A-A-G-G-C-G-C-U-G-G-G-U-U-G-C-U-N-A 0 B

15s

1

165

2

H. volcanii

pG-C-C-G-A-G-G-U-A-G-C-C-9-A-N-C-C-C

-G-G-C-C

S17

1

Yeast (mto)

pG-G-A-A-A-A-U-U-A-A-C-U-A-D-A

-G-G-D

-A-A-A-G-U-G-A-U-U-A-$-$I-U-G-C-U-A-A

518

2

Yeast (mto)

pG-G-A-U-G-G-U-U-G-A-C-U-G-A-G-D

4-G-D-U-U

ol¶ -A-A-A-C-U-G-$-G-A-U-A-$-U-U-GA-G-C

T5

Spinach (chl)

pG-C-C-C-C-U-U-U-A-A-C-U-C-A-G-U

4-G-D

-A-G-A-G-U-A-A-C-G-C-C-A-U-G-G-U-A-A

T6

N. crassu (mto)

pG-C-C-U-G-G-U-U-A-G-C-A-U-A-A-A

-A-G-D

-A-A-U-G-C-A-A-U-U-G-$-U-U-U-G-U-A-A

-A-A-G-G-C-G-G-U-A-G-A-U-U-C-G-A-A-A 1

m

THREONINE T t

Isoac- Anticodon Stem

No. ceptor -

V a r i a b l e Loop

39404~424344454647:1 2 3 4 5 6 7 8 9

10

12

11

T$C Stem

T$C Loop

T$C Stem

Aminoacyl Stem

14 1 6 4 9 51 5 3 5 5 5 7 59 6 1 6 3 6 5 67 69 7 1 73 75 13 1 5 48 50 52 54 56 5 8 60 62 64 66 68 70 72 74 76

PROLINE P2,PZ' 1,2 4 - C - G - A - G - A - G - G - D

+

m

5 5

-C-C-C-G-G-G-$-$-C-A-A-A-U-C-C-C-G-G-A-C-G-A-A-G-C-C-C-C-C-A

+

3P

-C-G-G-U-G-G-G-G-U

-U-G-A-A-G-G-T-~-C-G-A-G-U-C-C-U-U-C-A-U-U-G-G-A-G-A-C-C-A

4P

-G-G-C-C-G-U-A-A

-C-C-U-C-A-G-$-$-C-G-A-A-A-U-C-U-G-A-G-C-C-G-G-C-C-C-A-C-C-A

A 5

+

P5

m a

-A-C-A-A-A-A-U-G-U

-C-A-C-G-G-6-T-$-C-A-A-A-U-C-C-U-G-U-C-A-U-C-C-C-U-A-C-C-A

s11

-$-C-U-A-A-U-G-G-G-C-U-U-U-G-C-C-C-G

-C-G-C-A-G-G-T-~-C-A-A-A-U-C-C-U-G-C-U-G-G-U-G-A-C-G-C-C-A

s12

-U-U-C-U-A-U-G-C-U

-C-C-C-A-U-A-U-C-U-A-A-U-A-U-A-U-G-G-C-U-U-U-U-U-C-G-C-C-A

s12

-T-T-C-T-A-T-G-C-T

-C-C-C-A-T-A-T-C-T-A-A-T-A-T-A-T-G-G-C-T-T-T-T-T-C-G

S13

-C-U-C-A-U-G-C-C

-C-C-C-A-U-G-U-C-U-A-A-C-A-C-A-U-G-G-C-U-U-U-C-U-C-A-C-C-A

S14

-$-C-U-G-U-U-G-G-G-C-U-C-U-G-C-C-C-G

5 -C-G-C-U-G-G-T-$-C-A-A-A-U-C-C-U-G-C-U-G-G-U-G-U-C-G-C-C-A 5 5

SERINE

to

GI W

15s

1

m

5

* *

-C-U-C-A-G-U-G-G-C-G-U-C-A-A-G-C-C

-C-C-G-G-G-G

-$-C-G-A-A-U-C-C-C-C-G-C-C-G-C-A-A-C-G-C-C-A

-$-C-G-A-A-U-C-U-C-A-C-C-C-U-C-G-G-C-G-C-C-A

2

16s

2

-$-C-U-A-C-U-G-U-C-C-A-U-U-C-G-GA-C-A

5 -C-G-U-G-A-G

S17

1

-G-U-A-A-U-U-G-A-A-U-U-G-U-A-A-A-U-U-C-U

-U-A-U-G-A-G-T-JI-C-G-A-A-U-C-U-C-A-U-A-U-U-U-U-C-C-G-C-C-A

S18

2

-$-A-U-C-A-U-U-A-G-U-C-U-U-U-A-U-U-G-G-C-U-A

-C-G-U-A-G-G-T-$-C-A-A-A-U-C-C-U-A-C-A-U-C-A-U-C-C-G-C-C-A

THREONINE

+

T5

-G-G-C-G-U-A-A-G-D

-C-A-U-C-G-G-T-JI-C-A-A-A-U-C-C-G-A-U-A-A-G-G-G-G-C-U-C-C-A

T6

-$-C-A-A-U-A-G-A

-A-G-C-A-A-G-T-G-C-G-A-U-A-C-U-U-G-C-A-C-U-G-G-G-C-U-C-C-A

IsoNo. -

acceptor

D Stem

Aminoacyl Stem

Organism

D Loop

A n t i codon Stem Anticodon Loop

D Stem

91011121314151617:1 2181920:1 2 3

21

22

23

24

25

26

27

28

29

30

31

32

33

34

35

36

37

38

TRY PTOPHAK

w9

N. crassu

w10

*

*

*

*

PA-A-G-A-G-U-A-U-A-G-U-U-U-A-A-U

-G-G-D

-A-A-A-A-C-A-G-A-A-A-G-C-U-N-C-A-N-C

Yeast (mto)

PA-A-G-G-A-U-A-U-A-6-U-U-U-A-A-D

-G-G-D

w11

Spinach (chl)

pG-C-G-C-U-C-U-U-A-G-U-U-C-A-G-U-U-C

-G-G-D

-A-A-A-A-C-A-G-U-U-G-A-$-U-N-C-A-N-A 2

12w

E. coli (S"+7)

PA-G-G-G-G-C-G-U-A-G-U-U-C-A-A-D-D

5. pombe

pC-U-C-C-U-G-A-U-G-G-U-G-JI-A-G-D-D

B. s u b t i l i s

8Y Y9

(mto)

S

-A-G-A-A-C-G-$-G-G-G-$-C-U-C-C-A-A-A

*

-G-G-D

-A-G-A-G-C-A-C-C-G-G-U-C-U-C-U-A-A-A

-G-G-D-D

-A-U-C-A-C-A-C-C-C-G-G-C-U-G-$-A-A-A

pG-G-A-G-G-6-G-U-A-G-C-G-A-A-G-U

4-G-C-U-A

H. w l c a n i i

pC-C-G-C-U-C-U-U-A-G-C-U-C-A-N-C-C-U

-G-G-C

-A-A-C-G-C-G-G-C-G-G-A-C-U-Q-U-A-A-A A -A-G-A-G-C-A-G-C-C-G-A-C-U-G-U-A-G-A

Yeast (into)

PG-G-A-G-G-G-A-U-U-U-U-C-A-A-U-G-U-D

-G-G-D-A-G

-U-U-G-G-A-G-$-U-G-A-G-C-U-G-U-A-A-A

* S

TYROSINE Y6 M

6 0

7Y

1,2

m

S

i

m

U

m

*

VALINE

1

Rat L i v e r

pG-U-U-U-C-C-G-U-A-G-U-G-JI-A-G-D-G

-G-D-D

-A-U-C-A-C-G-$-U-C-G-C-C-U-N-A-C-A-C

14V

1

B. s u b t i l i s

pG-G-A-G-G-A-U-U-A-G-C-U-C-A-G-C-tI

4-G-G

-A-G-A-G-C-A-U-C-U-G-C-C-U-U-A-C-A-A

V15

Spinach ( c h l )

PA-G-G-G-C-U-A-U-A-G-C-U-C-A-G-U-U-A

4-G-D

-A-G-A-G-C-A-C-C-U-C-G-U-U-U-A-C-A-C

V16

N. crassa (mto)

pG-A-G-A-G-A-U-U-A-G-C-U-C-A-G-U-U

-G-G-D

V17 18V

4

m

5

2

V13

*

a

6

-A-G-A-G-C-A-A-C-C-G-U-U-U-U-A-C-A-C

lhwsophita

pG-U-U-U-C-C-G-U-G-G-U-G-+-A-G-C-G

-G-D-U

-A-U-C-A-C-A-+-C-U-G-C-C-U-I-A-C-A-C

8. votcanii

pG-G-G-U-U-G-G-U-G-G-U-C-$-A-G-U-C-U

-G-G-U-U

-A-U-G-A-C-A-C-C-U-C-C-U-U-G-A-C-A-U

5

Isaac- Anticodon Stem

NO. ceptor -

Variable Loop

TJlC Stem

T$C Loop

TJlC Stem

Aminoacyl Stem

3 9 4 0 4 ~ 4 2 4 3 4 4 4 ~ 4 6 4 71: 2 3 4 5 6 7 8 91011121314151648495051525354555657585g6061626364656667686g707172737~7576 TRYPTOPHAN

w9

-C-U-U-U-A-A-A-U

-U-C-U-U-A-G-T-JI-C-G-A-G-U-C-U-A-A-G-U-A-C-U-C-U-U-G-C-C-A

w10

-$-C-A-A-U-C-A-U

-U-A-G-G-A-G-T-JI-C-G-A-A-A-U-C-U-C-U-U-U-A-U-C-C-U-U-G-C-C-A

w11

-A-C-C-C-G-A-U-G-N

-C-G-U-A-G-G-T-$-C-A-A-G-U-C-C-U-A-C-A-G-A-G-A-C-G-U-G-C-C-A

+

12w

-A-A-C-C-G-G-G-U-G-U

-U-G-G-G-A-G-T-$-C-G-A-G-U-C-U-C-U-C-C-G-C-C-C-C-U-G-C-C-A

TYROSINE

+

Y6

m

5

-C-C-G-G-U-U-G-G-U

-C-G-C-U-A-G-T-JI-C-G-A-U-U-C-U-G-G-C-U-C-A-G-G-A-G-A-C-C-A

-$-C-C-6-C-U-C-C-C-U-C-A-G-G-U-U

-C-C-G-C-A-G-T-JI-C-6-A-A-U-C-U-G-C-C-C-C-C-C-U-C-C-A-C-C-A

8Y

-$-C-G-G-C-U-U-G-U

-C-C-C-C-C-G-JI-JI-C-G-A-A-U-C-G-G-G-G-A-G-A-A-G-C-G-G-A-C-C-A

Y9

-U-G-A-C-U-U-A-G-G-U-C-U-U

-C-A-U-A-G-G-T-JI-C-A-A-U-U-C-C-U-A-U-U-C-C-C-U-U-C-A-C-C-A

t9

5

7Y

1,2

5

m

2

VALINE

+ V13

1

-G-C-G-A-A-A-G-G-D

14V

1

-C-C-A-G-A-G-G-G-U

m

5 5

-C-C-C-C-G-G-U-JI-C-G-A-A-A-C-C-G-G-G-G-C-G-G-A-A-A-C-A-C-C

+ -C-G-A-G-C-G-G-T-JI-C-G-A-G-C-C-C-G-U-C-A-U-C-C-U-C-C-A-C-C-A

+

v15

-C-G-A-G-A-A-A-G-N

-C-U-A-C-G-G-T-~-C-G-A-G-U-C-C-G-U-A-U-A-G-C-C-C-U-A-C-C-A

V16

-A-C-G-G-A-A-G-G-U

-U-G-G-G-U-G-T-JI-C-G-A-A-U-C-A-C-C-C-A-U-U-U-C-U-C-A-C-C-A

+

m -C-C-C-C-G-G-T-$-C-G-A-U-C-C-C-G-G-G-C-G-G-A-A-A-C-A-C-C-A

V17

lev

4

-G-C-A-G-A-A-G-G-C -G-G-A-G-G-A-G-G-C

5

m a

2

-C-G-G-C-A-G-J1-$-C-G-A-A-U-C-U-G-C-C-C-C-A-A-C-C-C-A-C-C-A

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ADDENDUM: Additional tRNA Sequences References and Footnotes for Additional Sequence List'

ASPARACINE N3. Walker 256 mammary carcinosarcoma: B. A. Roe, A. F. Stankiewicz, H. L. Rizi, C. Weisz, M. N. DiLauro, D. Pike, C. Y. Chen, and E. Y. Chen, NARes 6,573 (1979). This sequence differs from that of a typical mammalian tRNAAsnonly in the "wobble" position of the anticodon, i.e., the cancer sequence contains a G instead of Q in position 34 of the tRNA.

CYSTEINE C3. (g) Saccharomyces cerevisiae (mitochondrial): J. L. Bos, K. A. Osinga, (3. Van der Horst, and P. Borst, NARes 6, 3255 (1979). This sequence was deduced from gene analysis of the mitochondrial DNA of tRNACys,hence modified components could not be determined.

GLUTAMIC ACID E8. Rat liver (1):J. C. Chan, J. A. Yang, M. J. Dunn, P. F. Agris, and T. Wong, NARes 10, 4605 (1982). E9 (g). Euglena gracilis chloroplast: M. J. Hollingsworth and R. B. Hallick, JBC 257, 12795 (1982). This sequence was deduced from gene analysis of the chloroplast DNA of tRNAG'";hence, modified components could not be determined.

GLYCINE (318 (g). Euglena gracilis chloroplast: M. J. Hollingsworth and R. B. Hallick,JBC 257, 12795 (1982). This sequence was deduced from gene analysis of the chloroplast DNA of tRNAG'y;hence, modified components could not be determined.

HISTIDINE H7 (g). Euglena gracilis chloroplast M. J. Hollingsworth and R. B. Hallick, JBC 257, 12795 (1982). This sequence was deduced from gene analysis ofthe chloroplast DNA of tRNAHlS;hence, modified components could not be determined.

ISOLEUCINE 17. Spinach (chloroplast-1): M. A. Francis and B. S. Dudock,JBC 257, 11195 (1982).

LYSINE K11. Rat liver (mitochondrial): E. Randerath, H. Aganval, and K. Randerath, BBRC 103, 739 (1981). U34 is probably 5-methoxycarbonylmethyluridine. 12K. Bacillus subtilis (1,3):B. S. Vold, D. E. Keith, Jr., M. Buck, J. A. McCloskey, and H. Pang, NARes 10,3125 (1982). The sequence of isoacceptor 3 is given

* Note: g in parentheses indicates the sequence is derived from its gene structure. 243

244

RAM P. SINGHAL

et al.

here. Isoacceptor 1 differs from isoacceptor 3 in position 30, in not having a modified C; in position 37, it contains a mixture of PA and ms2PA. K13. Drosophila melanogaster tRNAy: D. L. Gibbs, I. C. Gillam, and G. M. Tener, NARes 10, 6393 (1982). U34 is probably 5-methylcarboxymethy1-2thiouridine, and A37 a mixture of PA and its modified derivatives and a component reacting with cyanogen bromide. K14 (g). Rat (1,2,3): T. Sekiya, R. Nishizawa, K. Matsuda, Y. Taya, and S. Nishimura, NARes 10,6411 (1982). The sequence was deduced from gene analysis of the rat DNA of tRNALy'; hence, modified components could not be determined. The sequence of the three tRNALysspecies is the same.

INITIATOR-METHIONINE 23iM. Halococcus morrhuae: Y. Kuchino, M. Ihara, Y. Yabusaki, and S. Nishimura, Nature 298, 684 (1982). N57 is probably mlI. 24iM. Sulfolobus acidocaldarius: Y. Kuchino, M. Ihara, Y. Yabusaki, and S. Nishimura, Nature 298,684 (1982). N26 and N57 are probably m%Gmand m'I, respectively. 25iM. Thermoplasma acidophilum: Y. Kuchino, M. Ihara, Y. Yabusaki, and S. Nishimura, Nature 298, 684 (1982). N32 consists of a mixture of C and Cm. N57 is probably I. 26iM. Streptomyces griseus: Y. Kuchino, I. Yamamoto, and S. Nishimura, NARes 10, 6671 (1982).

METHIONINE 8M. Thermoplasma acidophilum: M. W. Kilpatrick and R. T. Walker, Zbl. Bakt. H y g . , 1. Abt. Orig. C3, 79 (1982). This is a corrected version of a sequence published earlier, code 8M. (The authors claim that C32 is modified to Cm.) M11 (g). Euglena gracilis chloroplast: M. J. Hollingsworth and R. B. Hallick,JBC 257, 12795 (1982). The sequence was deduced from gene analysis of the chloroplast DNA of tRNAMet;hence, modified components could not be determined.

PHENYLALANINE F23. Blue green algae, Agmenellum quadruplicatum: L. I. Hecker, W. E. Barnett, F. K. Lin, T. D. Furr, J. E. Heckman, U. L. RajBhandary, and S. H. Chang, NARes 10, 6433 (1982). U12 is an unidentified modified uridine base. F24. Mouse liver: Y. Kuchino, E. Borek, D. Grunberger, J. F. Mushinski, and S. Nishimura, NARes 10,6421 (1982). F25. Ehrlich ascites tumor and neuroblastoma: Y. Kuchino, E. Borek, D. Grunberger, J. F. Mushinski, and S. Nishimura, NARes 10, 6421 (1982). This sequence differs from that of a typical mammalian tRNAPhein positions 32, 34, and 37 of the anticodon: the tumor tRNAPhecontains a C instead of Cm in position 32, G instead of Gm in position 34, and undermodified wye in position 37. In position 37, Ehrlich ascites tumor tRNAPhecontains an undermodified wye, but the neuroblastoma mostly mlG.

PROLINE P6. Torulopsis utilis: K. Ogawa, M. Kawakami, Y. Shimizu, and S. Takemura, J . Biochem. 91, 1241 (1982).

STRUCTURE OF TRANSFER

RNAS

245

P7 (g). Rat (1,2,3): T. Sekiya, R. Nishizawa, K. Matsuda, Y. Taya, and S. Nishimura, NARes 10, 6411 (1982). The sequence was deduced from gene analysis of the rat DNA of tRNAPro;hence, modified components could not be determined. The sequence of tRNAP differs from tRNAi;; in position 34 of the anticodon: tRNAp sequence contains a C instead of an A. In position 50, tRNAP differs from tRNA,P'"in having a U instead of a C.

TRYPTOPHAN 8W. Escherichia coli, a temperature-sensitive mutation due to a lesion in the gene of tRNAT? S. P. Eisenberg, L. Soll, and M. Yarus,JBC 254,5562 (1979).The mutant varies from the wild type in only one base change: G7 + A7. W13 (g). Euglena gracilis chloroplast: M. J. Hollingsworth and R. B. Hallick,JBC 257, 12795 (1982). The sequence was deduced from the gene analysis of the chloroplast DNA of tRNATq; hence, modified components could not be determined.

TYROSINE Y10 (g). Euglena gracilis chloroplast: M. J. Hollingsworth and R. B. Hallick,JBC 257, 12795 (1982). The sequence was deduced from the gene analysis of the chloroplast DNA of tRNATyr;hence, modified components could not be determined.

No.

Isoacceptor

Organism

A m i n o a c y l Stem

D Stem

D Loop

1 2 3 4 5 6 7 8 91011121314151617:192D:1

’

A n t i codon Stem A n t i c o d o n Loo

D Stem

3~2425262728293D~,3233343~363,~8

ASPARAGINE N3

Mammary c a r c i n o sarcoma

CYSTEINE

n

a

t

4-G-D-U-

-A-G-C-G-C-G-$-$-C-G-G-C-U-G-U-U-A-.I

(mto)

pG-G-A-G-A-U-G-U-U-G-U-U-U-U-A-A-

-G-G-U-U-

-A-A-A-C-U-A-U-U-A-G-A-U-U-G-C-A-A-k

Rat l i v e r

~u-c-c-c-A-c-A-u-G-G-u-c-*-A-G-c-

-G-G-D-D-

-A-G-G-A-U-U-C-C-U-G-G-,~-U-~-U.C-A-C

E . gracizis (Cpt) pG-C-C-C-C-C-A-U-C-G-U-C-U-A-G-A-

-G-G-C-C-U-

-A-G-G-A-C-A-U-C-U-C-C-C-U-U-U-C-A-C

E. g m c i l i s (Cpt) pG-C-G-G-G-U-A-U-A-G-C-U-C-A-G-U-U-

-G-G-U-

-A-G-A-G-C-G-U-G-G-U-C-C-U-U-C-C-A-A

S. cerevisiae

c3

m 2

pG-U-C-U-C-U-G-U-G-G-C-G-C-A-A-D-C-

GLUTAMIC ACID E8

1

E9 (91

m

GLYS INE

GI8 (g) N Ip

m

HISTIDINE H7 ( 9 )

ISOLEUCINE

I7

1

LYSINE R a t l i v e r (mto)

K11

12K K13

3 5

K14 ( 9 ) 1,2,3

B. s u b t i l i s

2 * -A-G-C-G-(I-U-A-A-C-C-U-U-U-U-A-A

m 2 pC-A-U-U-G-C-G-A-A-G-C-U-U-A-ti-

* pG-A-G-C-C-A-U-U-A-G-C-U-C-A-G-U-D-

2

-G-G-D-

-A-G-A-G-C-A-U-C-U-ti-A-C-U-U-U-U-2-A

*

Drosophi l a

pG-C-C-C-G-G-A-U-A-G-C-U-C-A-G-D-C-

-G-G-D-

-A-G-A-G-C-A-*-JI-G-G-A-C-U-U-U-U-A-A

Rat

pG-C-C-C-G-G-C-U-A-G-C-U-C-A-G-U-C

-G-G-U-

-A-G-A-G-C-A-U-G-A-G-A-C-U-C-U-U-A-A

-G-G-A-G-

-A-u-U-C-C-~-G-C-G-G-G-C-U-C-A-U-A-A

-G-G-A-G-U-

-A-U-C-C-C-G-C-A-G-G-G-C-U-C-A-U-A-A

-G-G-A-

-A-A-U-C-C-G-A-U-G-G-G-C-U-C-A-U-A-A

INITIATOR-METHIONINE 23iM

H. morrhuae

24iM

S.

?5iM

T. acidophilwn

.~A-G-c-G-G-G-A-u-G-G-G-A-*-A-G-c-c-A-

2 acidocatdarius pA-G-C-G-G-C-G-U-N-G-G-G-A-A-C-U-G-

PA-G-C-G-G-G-G-U-G-G-G-G-*-A-G-U-C-A-

2

m

T

*

m

*

NO.

Isoac- Anticodon ceptor Stem

T$€

Loop

Stem

L

~ ~ ~ ' 4 1 ~ ~ 4 3 1~ 2~ 34 45 5~ ~ 4 7 :

IVC Stem

Aminoacyl Stem

13141516 4 8 4 9 5 0 5 1 5 2 s ~ 4 5 5 5 6 5 7 5 8 5 g ~ ~ 6 1 6 2 6 5 6 4 6 5 6 6 6 7 6 8 6 g 7 0 7 1 7 2 7 ~ 4 7 5 7 ~

ASPARAGINE +

u3

m

-C-C-C-A-A-A-G-G-D

-U-G-G-U-G-G-N-*-C-G-A-G-C-C-C-A-C-C-C-A-G-G-G-A-C-G-C-C-A

-U-C-U-A-C-U-U-A-U

-U-A-A-G-A-G-U-U-C-G-A-U-U-C-U-C-U-U-C-A-U-C-U-C-U-U-C-C-A

CYSTEINE

c3

GLUTAMIC ACID

E8

1

E9 ( 9 )

5 5

-C-C-A-G-G-C-G-G

-C-C-C-G-G-G-T-*-C-G-A-C-U-C-C-C-G-G-U-G-U-G-G-G-A-A-C-C-A

4-C-A-G-G-C-A-A-

-C-G-G-G-G-A-U-U-C-G-A-A-U-U-C-C-C-C-U-G-G-G-G-G-U-A-C-C-A

-G-U-C-C-A-A-U-G-U-

-U-G-C-G-U-G-U-U-C-G-A-A-U-C-A-C-G-U-U-A-C-C-C-G-C-U-C-C-A

-U-C-C-U-U-C-A-U-U-

-C-G-C-G-G-G-U-U-C-G-A-U-C-C-C-C-G-U-C-A-U-U-C-A-C-C-C-C-A

-$-+-G-G-C-G-A-A-U-U

-C-G-U-A-G-G-T-*-C-A-A-U-U-C-C-U-A-C-U-G-G-A-U-G-C-A-C-C-.4

-G-U-U-A-A-A-G-U-U

-A-

m

GLYCINE

G18 ( 9 ) HISTIDINE

H7 ( 9 ) ISOLEUCINE

I7

1

LYSINf Klf 12K

3

K13

5

K14(g) 1,

+

-G-A-C-A-C-A-A-C-A-A-A-U-C-U-C-

-C-A-C-A-A-U-C-A-C-C-A

-$-C-A-G-A-G-G-G-U

-C-G-A-A-G-C-T-*-C-G-A-G-U-C-C-U-U-C-A-U-G-G-C-U-C-A-C-C-A

-+-C-C-A-A-G-G-G-0-

-C-C-A-G-G-G-T-$-C-A-A-G-U-C-C-C-U-G-U-U-C-G-G-G-C-G-C-C-A m

-U-C-U-C-A-G-G-G-U-

-C-G-U-G-G-G-U-U-C-G-A-G-C-C-C-C-A-C-G-U-U-G-G-G-C-G-C-C-A

5

m

2.3 INITIATOR-METHIONINE 23iM

-C-C-C-G-C-A-G-A-U

-C-A-G-U-A-C-$-+-C-~-A-A-U-C-U-A-C-U-U-C-C-C-G-C-U-A-C-C-A

24iN

-C-C-C-U-G-A-G-G-U

-C-C-C-U-G-G-U-U-C-~-~-A-U-C-C-A-G-G-C-G-C-C-G-C-U-A-C-C-A m m

25iM

-C-C-C-G-U-A-G-A-U

-C-G-A-U-G-G-+-$-C-~-~-A-U-C-C-A-U-C-C-C-C-C-G-C-U-A-C-C-A

5

Iso-

Anti codon D Stem Stem Anticodon Loo 192 0 : l 2 3 ~ 2 4 2 5 2 6 2 7 2 8 2 9 3 0 3 1 3 2 3 3 3 4 3 5 3 6 3 7 3 ~

Organism

Aminoacyl Stem D Stem 1 2 3 4 5 6 7 8 9 11 13 1415161 7 : l 2

Streptomyces griseus

pC-G-C-G-G-G-G-U-G-G-A-G-C-A-G-C-U-C- -G-G-D-

E . gmciZis(Cpt)

PG-G-C-U-C-A-G-U-A-G-C-U-C-A-G-A-

F23

Blue green algae

pG-C-C-A-G-G-A-U-A-G-C-U-C-A-G-U-0-

F24

Mouse l i v e r

pG-C-C-G-A-A-A-U-A-G-C-U-C-A-G-D-D-

No. -

acceptor

26iM

!??

-A-G-C-U-C-G-C-U-G-G-G-C-U-C-A-U-A-A

ME'IHIONINE

(g)

-G-G-A-U-

-A-G-A-G-C-A-G-G-G-G-A-U-U-C-A-U-A-A

PHENYLALANINE b

2

m

4-G-Dm

4-G-G-

2

kors

pc-C-C-G-A-A-A-U-A-G-C-U-C-Z-G-D-D- 4 - G - G -

T. u t i l i s

~G-GC-C-G-C-G-U-F-G-U-C-~-A-G-D--G-G-D-

-A-U-G-A-U-A-C-U-C-G-C-U-U-N-G-G-G-$

Rat

pG-G-C-U-C-G-U-U-G-G-U-C-U-A-G-G-

4-G-U-

-A-U-G-A-U-U-C-U-C-G-C-U-U-C-G-G-U

8W

E. coZi

PA-G-G-G-G-C-A-U-A-G-U-U-C-A-A-D-D-

-G-G-D-

-A-G-A-G-C-A-C-C-G-G-U-C-U-C-C-A-A-A

W13 (g)

E . g r a c i l i s (Cpt)

pG-C-G-C-U-U-U-U-A-G-U-U-C-A-A-U-U-

-G-G-U-

-A-G-A-A-C-G-U-A-G-G-U-C-U-C-C-A-A-A

E . g r a c i l i s (Cpt)

pG-A-G-U-U-G-U-U-G-C-C-C-G-A-G-U-

-G-G-U-U-A-

-A-U-G-G-G-G-G-C-G-G-A-U-U-G-U-A-A-A

F25 PROLINE P6 p7 (9)

1

m

TRYPTOPHAN S

TYROS I NE

y10 (g)

Isoac- Anticodon Stem

No. ceptor -

Variable Loop

T$C Stem

T$C Loop

T$C Stem

hinoacyl Stem

39404~424344454647:1 2 3 4 S 6 7 8 910111213141S164849S0s1S2s3545~56s7S8~g60616263646s6667686g70717273747576 26iM

m

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

-C-G-C-A-G-G-U-(I-C-A-A-A-U-C-C-U-G-U-C-C-C-C-G-C-U-A-C-C-A

4-C-C-C-U-U-G-G-

-C-A-C-A-G-G-U-U-C-A-A-A-U-C-U-U-G-U-C-U-G-A-G-C-C-A-C-C-A

MElH ION INE M11 ( 8 )

PHENYLALANINE

F23

-C-G-G-C-G-G-T-$-C-A-A-U-U-C-C-G-C-C-U-C-C-C-G-G-C-A-C-C-A

+

F24

-(I-C-U-A-A-A-G-G-U-

F25

5

-~-~-C-U-C-G-T-~-C-G-~-U-C-C-C-~-~-~-U-U-U-C-~-~-C-A-C-C-A

+

5 m -C-C-C-U-G-G-T-$-C-G-A-U-C-C-C-G-G-G-U-U-U-C-G-G-C-A-C-C-A

+

5 5 -C-C-A-G-G-G-T-$-C-A-X-~-~-~-~-~-~-~-~-U-C-G-G-C-C-C-C-C-A

-$-C-U-A-A-A-G-G-U-

PROLINE

P6 P7 (8)

-G-$-G-A-G-U-G-G-D

1

4-C-G-A-G-A-G-G-U-

-C-C-C-G-G-G-U-U-C-A-A-A-U-C-C-C-G-G-A-C-G-A-G-C-C-C-C-C-A

TRYPTOPHAN 8W

-A-C-C-G-G-G-U-G-U

-U-G-G-G-A-G-T-$-C-G-A-G-U-C-U-C-U-C-C-G-C-C-C-C-U-G-C-C-A

W13 ( 8 )

-A-C-C-U-G-A-U-G-U-

-A-G-U-A-G-G-U-U-C-G-A-A-U-C-C-U-A-C-A-G-A-G-C-G-C-G-C-C-A

-U-C-C-G-C-A-G-U-U-C-A-U-C-U-U-U-

-C-G-C-U-G-G-U-U-C-G-A-A-U-C-C-A-G-C-A-C-G-A-C-U-C-A-C-C-A

TYROSINE Y 1 0 (g)

This Page Intentionally Left Blank

ADDENDUM: References to 32 Additional Sequences

Several important articles on the structure-function of the 5 S RNA have recently appeared, and 32 more sequences of this type of RNA have been published since the review article was written. A survey of these papers indicates publications on a structural model of the 5 S RNA (E.coli) based on intramolecular crosslinking evidence (I), a comparison of generalized models for eukaryotic 5 S RNAs ( 2 ) , and generalized structures of the 5 S RNA based on sequences from a wide range of the organisms ( 3 ) .Other recent publications deal with the study of the reaction sites between the 5 S RNA (E.coli) and the proteins (L18and L25)(4)and with the study of the changes in yeast 5 S RNA structure brought about by various nucleases, to provide evidence for the minimal secondary structure (model) containing five helical regions (4a). Evidence for the existance of two native conformations for E . coli 5 S RNA (5)as well as for rat liver 5 S RNA (6)has been published by two different groups. In order to establish phylogenic positions of various organisms in the evolutionary scheme, a large number of 5 S RNA structures have been analyzed in the past several months. For example, the 5 S RNAs of such diverse organisms as photosynthetic and nonphotosynthetic bacteria (7), pathogens (8), an aphid (9), protozoa ( l o ) ,molluscs (I]), sponges (12),silkworms (13),water molds ( 1 4 ) ,algae (15,16), rotifers (17),nematodes (17),jellyfish (18),a legume (19),fungi (2,20),and a fern ( 2 1 ) have been studied. Presently, the reader is referred to these publications (see the list of organisms and references below). We plan to update this work in the near future. Paracoccus denitrijcans (nonphotosynthetic, purple non-sulfur bacteria) (7) Prochloron species (a plastid, photosynthetic prokaryote) (7) Spiroplasma species BC 3 (a honeybee pathogen) (8) Mycoplasma mycoides sp. capri PG3 (a pathogen) (8) Acyrthosiphon magnoliae (an elder aphid) (9) Paramecium (ciliated protozoa-ciliophora of Class 11) (10) Tetrahymena (ciliated protozoa of Class 11) (10) Blepharisma (ciliated protozoa of Class 111) (10) Helix pomatia (mollusc, the snail) ( 1 1 ) Arion mfus (mollusc, the snail) ( 1 1 ) Mytilus edulis (mollusc, the mussel) ( 1 3 ) Halichondria panicea (a sponge) (12) Hymeniacidon sanguinea (a sponge) (12) Haliclona oculata (a sponge) (12) Philosomia Cynthia ricini (silkworm, posterior gland) (13) 25 1

ADDENDUM

Saprolegnia ferar (an oomycete water mold) (14) Pythium hydrosperum (an oomycete water mold) (14) Blastocladiella simplex (a chytrid water mold) (14) Phlyctochytrium irregulare (a chytrid water mold) (14) Porphyra yezoensis (a red alga) (15) Scenedesmus obliquus (a green alga, cytoplasmic fraction) (16) Brachionus plicatilis (a rotifer) (17) Rhabditis tokai (a nematode) (17) Caenorhabditis elegans (a nematode) (17 ) Spirocodon saltatrix (a hydrozoan jellyfish) (18) Nemopsis dofleini (a hydrozoan jellyfish) (18) Aurelia aurita (a scyphozoan jellyfish) (18) Chrysoora quinquecirrha (a scyphozoan jellyfish) (18) Lupinus Zuteus (a yellow lupin-legume) (19) Phycomyces blakesleeanus (a lower fungus) (20) Thennomyces lanuginosus (a thermophilic fungus) ( 2 ) Dryopteris acuminata, chloroplasts (a fern) (21)

REFERENCES 1. 2. 3. 4. 4a. 5. 6.

J. Hancock and R. Wagner, NARes 10, 1257 (1982). A. G. Wildeman and R. N. Nazar, JBC 257, 11395 (1982). N. Delihas and J. Andersen, NARes 10, 7323 (1982). M. Speek and A. Lind, NARes 10,947 (1982). R. A. Garrett and S. 0. Olesen, Bchem. 21,4823 (1982). M. J. Kime and P. B. Moore, NARes 10,4973 (1982). I. Toots, R. Misselwitz, S. Bohm, H. Welfle, R. Villems, and M. Saarma, NARes 10,

3381 (1982). 7. R. M. MacKay, D. Salgado, L. Bonen, E. Stackebrandt, and W. F. Doolittle, NARes 10, 2963 (1982).

8. R. T. Walker, E. T. J. Chelton, M. W. Kilkpatrick, M. J. Rogers, and J. Simmons, NARes 10,6363 (1982). 9. Y. Kawata and H. Ishikawa, NARes 10, 1833 (1982). 10. T. Kumazaki, H. Hori, S. Osawa, T. Mita, and T.Higashinakagawa, NARes 10,4409 (1982). 11. B.-L. Fang, R. DeBaere, A. Vandenberghe, and R. De NARes 10,4679 (1982). 12. E. Dams, A. Vandenberghe, and R. DeWachter, NARes 10,5297 (1982). 13. G. Xian-Rong, K. Nicoghosian, and R. J. Cedergren, NARes 10,5711 (1982). 14. W. F. Walker and W. F. Doolittle, NARes 10, 5717 (1982). 15. F. Takaiwa, M. Kusuda, N. Saga, and M. Sugiura, NARes 10,6037 (1982). 16. G. A. Green, J. M. McCoy, and D. S. Jones, NARes 10,6389 (1982). 17. T. Kumazaki, H. Hori, S. Osawa, H. Ishii, and K. Suzuki, NARes 10 (1982). 18. H. Hori, T. Ohama, T. Kumazaki, and S. Osawa, NARes 10,7405 (1982). 19. J. A. Rafalski, M. Wiewiorowski, and D. Soll, NARes 10, 7635 (1982). 20. J. Anderson, W. Andresini, and N. Delihas,JBC 257,9114 (1982). 21. F. Takaiwa and M. Sugiura, NARes 10,5369 (1982).

Index

A

Chloroplasts, sequences of 5 S RNA, references, 202 Codon(s), misreading, isoacceptor tRNAs and, 94-96 Cross-linking, of rRNA and protein, 26 analysis of products, 30-37 methods and reagents, 27-30

Aging, translational fidelity and, 84-85 Alanine tRNAs, sequences for, 228,229 Amino acid(s) sequence, of ribosomal protein, S1,

109-111 starvation, translational fidelity and,

82-83

D

Antibiotics error-producing, translational speed and accuracy and, 94 translational fidelity and, 83 Archaebacteria, sequences of 5 S RNA, references for, 202 Arginine tRNAs, sequences for, 228, 229 Asparagine tRNAs, sequences for, 228,

DNA, synthesis, yeast cell cycle and, mitochondrial, 159-160 nuclear, 153- 156 plasmid, 156-158 DNA, transcription, function of ribosomal protein S1 in, 130

229

E

Aspartate tRNAs, sequences for, 228,

229 Elongation process, inhibitors, translational speed and accuracy and, 92-

B Bacteriocins, translational fidelity and,

84 C Cations, polyvalent, accuracy of in uitro translation systems and, 89-90 Cell cycle, methods for analysis, 146-

149 Cell density, yeast cell cycle and, 149-

150 Cell differentiation, change of ratio of queuine to guanine in tRNA during,

66-67 Cell division, control, yeast cell cycle and, 167-169 Cell mass, yeast cell cycle and, 149-150 Cell volume, yeast cell cycle and, 149-

150

93 Energy, regeneration, accuracy of in uitTO translation systems and, 91 Enzymic activities, yeast cell cycle and, 162-167 Escherichia coli,rRNA derivation of structures for, 8-12 protein cross-linking of, 26-37 three-dimensional packing of, 37-41 Eukaryotes sequences of 5 S RNA, references for, 200-202 structure of 5 S RNA base-paired regions, 190-191 homology in primary structures, 185 invariant regions, 185-190

G Gene, for ribosomal protein S1, 120-121 Glutamate tRNAs, sequences for, 230,

Cell walls, yeast cell cycle and, 150-152

231 253

INDEX

Glutamine tRNAs, sequences for, 230, 23 1 Glycine tRNAs, sequences for, 230,231 Guanine, change in ratio to queuine in tRNA during cell differentiation, 6667 Guanosine triphosphatase, ribosomal, translational speed and accuracy and, 93 H

Hinge region, of ribosomal protein S1, 119-120 Histidine tRNAs, sequences for, 230, 23 1 Homologous stretches, repeating, of ribosomal protein S 1 and, 111-1 13 I

Inhibitors, of ribosomal protein S1 function, 133-134 Isoleucine tRNAs, sequences for, 232, 233 L Leucine tRNAs, sequences for, 232, 233-234,235 Lysine tRNAs, sequences for, 234, 235 M

Membranes, yeast cell cycle and, 150152 Messengers natural, ribosomal protein S1 function and, 132-133 synthetic, ribosomal protein S1 function and, 132 Metabolic activities, yeast cell cycle and, 152-153 Methionine tRNAs, sequences for, 236, 237 Methionine-initiator tRNAs, sequences for, 234,235-236,237 Mitochondria DNA synthesis in, yeast cell cycle and, 159-160

sequences of 5 S RNA, references for, 202 Molecular mass, of ribosomal protein s l , 106 Mutant form, of ribosomal protein S1, 114- 115 N

Neoplastic cells, queuine in, 78-79 Nucleases, reactivity of prokaryotic 5 S RNA with, 195 Nucleic acid interaction with ribosomal protein S 1, 123-124 question of second binding site, 125-126 strong binding site, 124-125 Nucleic acid-binding fragment, of ribosomal protein S1, 118-119 Nucleus, DNA synthesis in, yeast cell cycle and, 153-156

0 Oligonucleotides, complementary, binding with prokaryotic 5 S RNA, 195 P

pH, accuracy of in oitro translation systems and, 91 PhenylaIanine tRNAs, sequences for, 236,237 Plasmid(s), DNA synthesis in, yeast cell cycle and, 156-158 Polyamines, translational fidelity and, 83-84 Prokaryotes conformation of 5 s RNA, 191-193 binding with complementary oligonucleotides, 195 exchangeable protons, 195 physical measurements, 196 reactivity for chemicals, 193-194 reactivity with nucleases, 195 reconstitution of ribosomes from, 196 sequences of 5 S RNA, references for, 198-200

INDEX

structure of 5 S RNAs base-paired regions, 183-185 homology in primary structures, 178 invariant segments, 178-183 Proline tRNAs, sequences for, 238, 239 Protein rRNA interaction binding sites on rRNA, 24-26 cross-linking in E. coli, 26-37 yeast cell cycle and, 162-167 Protein synthesis, ribosomal protein S1 and, 130 inhibitors of function, 133-134 natural messengers and, 132-133 synthetic messengers and, 132 Proteolysis, limited, of ribosomal protein

S1, 116-117 Protons, exchangeable, of prokaryotic 5 S RNA, 195

Q

Qp replicase, interaction with ribosomal protein s l , 126-127, 128 inactive 30 S subunits and, 128 rRNA and, 127 Qp replication, function of ribosomal protein S1 in, 129-130 Queuine, change in ratio to guanine in tRNA during cell differentiation,

66-67 Queuosine absence in tRNA of tumor cells, 63-66 analogs, possible applications of, 70-71 biological function in tRNA, 67-70, 79 biosynthesis in tRNA in animal cells, 59-60 tRNA-guanine transglycosylase, 57-59 in uiuo experiments, 56-57 distribution in animals and plants and isolation of derivatives, 55-56 history of discovery of, 50-54 skeleton, biosynthesis of, 62-63

R Radius of gyration, of ribosomal protein

S1, 106-107 Readthrough, translation and, 87-88 Ribosomal protein S1, 102-104

discernable structural and functional domains, 113-114 evidence for hinge region, 119-120 mutant form M l S l , 114-115 nucleic acid-binding fragment, 118-

119 ribosome-binding fragment, 117-118 SH groups, 119 stable fragment by limited proteolysis, 116-117 functions of, 128-129 in protein synthesis, 130-134 in Qp replication and DNA synthesis, 129-130 gene for, 120-121 general properties, 105 interactions of with nucleic acid, 123-126 within ribosome and Qp replicase, 126-128 isolation procedure, 104-105 location within ribosome, 121-123 mode of action of, 134-137 physical constants, 106-107 probable shape, 107-108 structure amino-acid sequence, 109-11 1 four repeating homologous stretches,

111-1 13 secondary elements, 113 Ribosomal RNA protein interactions binding sites on rRNA, 24-26 cross-linking in E. coli, 26-37 secondary structure, 5-8 derivation of structures for E. coli rRNA, 8-12 evidence for alternative conformations, 21-24 structures proposed for rRNA from other sources, 12-21 sequences of, 2-5 three-dimensional packing in E. coli electron microscopy and, 39-41 intra-RNA cross-linking, 37-39 Ribosome(s) GTPase activity, translational speed and accuracy and, 93 location of ribosomal protein S1 within, 121-123

256

INDEX

mutant, message decoding and, 86 reconstitution, with prokaryotic 5 S RNA, 196 role in message decoding, 85-86 Ribosome-binding fragment, of ribosomal protein S1, 117-118 RNA, 5 S conformation of prokaryotic, 191-193 binding with complementary oligonucleotides, 195 exchangeable protons, 195 physical measurements, 196 reactivity for chemicals, 193-194 reactivity with nucleases, 195 reconstitution of ribosomes from, 196 eukaryotic base-paired regions, 190-191 homology in primary structures, 185 invariant segments, 185-190 prokaryotic base-paired regions, 183-185 homology in primary structure, 178 invariant segments of structure, 178183 secondary structure of, 8-9 sequences archaebacterial, 202 chloroplasts, 202 eukaryotes, 200-202 mitochondria, 202 notes, 198 prokaryotes, 198-200 references and footnotes for sequence list, 198-202 RNA, 16 S and 23 S, secondary structure of, 9-12 RNA, synthesis, yeast cell cycle and, 160-162

s Septa, yeast cell cycle and, 150-152 Serine, tRNAs, sequences for, 238,239 Soluble factors, accuracy of in oitro translation systems and, 91-92 Sulfhydryl groups, of ribosomal protein s1,119

T

Temperature accuracy of in oitro translation systems and, 90 change, translational speed and accuracy and, 93-94 Threonine tRNAs, sequences for, 238, 239 Transfer RNA abbreviations in, 216-219 accuracy of in uitro translation systems and, 90 biological function of queuosine in, 67-70 biosynthesis of queuosine in, 56-60 change in ratio of queuine to guanine in during cell differentiation, 6667 code numbers, 215-216 isoaccepting, codon misreading by, 94-96 lacking queuosine, presence in tumor cells, 63-66 organization, 211-215 references for, 215 sequences, 228-241 references and footnotes for, 219227 Transfer RNA-guanine transglycosylase metal ion requirement, 76 properties of, 60-62 queuosine biosynthesis and, 57-59 size and subunits, 75-76 substrate specificity, 76-78 Translation accuracy in uitro, 88 energy regeneration and, 91 effect of pH, 91 polyvalent cations and, 89-90 soluble factors, 91-92 temperature, 90 tRNA and, 90 agents affecting fidelity in uiuo amino acid starvation, 82-83 antibiotics, 83 bacteriocins, 84 biological aging, 84-85 polyamines, 83-84

257

INDEX

message decoding and readthrough translation, 87-88 ribosomal mutants, 86 role of ppGpp, 86-87 role of ribosomes, 85-86 relationship between speed and accuracy error-producing antibiotics, 94 general, 92 inhibitors of elongation process, 92-

93 ribosomal GTPase activity, 93 summary, 94 temperature change, 93-94 Tryptophan tRNAs, sequences for, 240,

24 1 Tumor cells, presence of tRNA lacking queuosine in, 63-66, 78-79 Tyrosine tRNAs, sequences for, 240, 241

V Valine tRNAs, sequences for, 240,241

Y Yeast cell cycle control of cell division and, 167-169 patterns of growth cell walls, septa and membranes,

150- 152 DNA synthesis, 153-160 metabolic activities, 152-153 proteins and enzymic activities,

162-167 RNA synthesis, 160-162 volume, cell mass and density, 149-

150

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Contents of Previous Volumes Volume 1 "Primer" in DNA Polymerase Reactions-F. /. B o l l u r n M . S. S m e l l i e The Biosynthesis of Ribonucleic Acid in Animal Systems-R. The Role of DNA in RNA Synthesis-Jerard H u r w i t z a n d J. T. A u g u s t Polynucleotide Phospharylase-M. G r u n b e r g - M a n a g o Messenger Ribonucleic Acid-Fritz Lipmann The Recent Excitement in the Coding Problem-F. H. C. C r i c k Some Thoughts on the Double-Stranded Model of Deoxyribonucleic Acid-Aaron Bendich a n d H e r b e r t S. Rosenkranz Murmur, R . R o w n d , atid C. L. Denaturation and Renaturation of Deoxyribonucleic Acid-J.

Schildkruut Some Problems Concerning the Macromolecular Structure of Ribonucleic Acids-A. The Structure of DNA as Determined by X-Ray Scattering Techniques-Vittoria Molecular Mechanisms of Radiation E f f e c t e A . Wacker Volume 2 Nucleic Acids and Information Transfer-Liebe

F.

S. Spirin

Luzzati

C a v a l i e r i a n d B a r b a r a H. Rosenberg

Nuclear Ribonucleic Acid-Henry Harris Plant Virus Nucleic Acids-Roy Markhum R. Lehnznn The Nuclearer of Escherkbb cofi-1. Specificity of Chemical Mutagenesis-David R . Krieg Column Chromatography of Oligonucleotides and Polynucleotides-Matthys Mechanism of Action and Application of Azapyrimidines-/. Skoda The Function of the Pyrimidine Base in the Ribonuclease Reaction-Herbert Preparation, Fractionation, and Properties of sRNA-G. L. B r o w n

Staehelin

Witzel

Volume 3 Isolation and Fractionation of Nucleic Acids-K. S. Kirby Cellular Sites of RNA Synthesis-Duoid M . Prescott Ribonucleases in Taka-Diastase: Properties, Chemical Nature, and Applications-Fuji0 Egami, K e n j i Takahashi, and Tsuneko U c h i d a Chemical Effects of Ionizing Radiations on Nucleic Acids and Related Compounds-Joseph /. Weiss The Regulation of RNA Synthesis in Bacteria-Frederick C. N e i d h a r d t Actinamycin and Nucleic Acid Function-E. R e i c h a n d 1. H. G o l d b e r g Nisman and/. Pelmont De Novo Protein in Synthesis in Vitro-B. Mandel Free Nucleotides in Animal Tissues-P. Volume 4 Fluorinated Pyrimidines-Charles H e i d e l b e r g e r Genetic Recombination in Bacteriophage-E. Volkin M . Keir DNA Polymerases from Mammalian Cells-H. The Evolution of Base Sequences in Polynucleotides-B. J. M c C a r t h y Osawa Biosynthesis of Ribosomes in Bacterial Cells-Syozo 5-Hydroxymethylpyrimidines and Their Derivatives-?'. L. V. Ulbright

259

260

CONTENTS OF PREVIOUS VOLUMES

Amino Acid Esters of RNA, Nucleotides, and Related Compounds-H. Feldmann Uptake of DNA by living Cells-L. Ledoux

G.

Zachau a n d

H.

Volume 5 Introduction to the Biochemistry of 4-Arabinosyl Nucleosides-Seymour Cohen Effects of Some Chemical Mutagens and Carcinogens on Nucleic Acids-P. D.L a w l e y Nucleic Acids in Chloroplasts and Metabolic DNA-Tatsuichi I w a m u r a Enzymatic Alteration of Macromolecular Structure-P. R. Srinivasan a n d E r n e s t Borek R. Tata Hormones and the Synthesis and Utilization of Ribonucleic Acids-]. J. Fox, K y o i c h i A. Watanabe, a n d Alexander B l o c h Nucleoside Antibiotics-Jack A. Thomas, J r . Recombination of DNA Molecules-Charles Appendix I. Recombination of a Pool of DNA Fragments with Complementary Single-Chain Ends-G. S. Watson, W. K . Smith, a n d Charles A, Thomas, J r . Appendix II. Proof that Sequences of A, C, G, and T Can Be Assembled to Produce Chains of Ultimate Length, Avoiding Repetitions Everywhere-A. S. Fraenkel a n d J . G i l l i s The Chemistry of Pseudauridine-Robert Warner Chambers The Biochemistry of Pseudouridine-Eugene Goldwasseer and Robert L. Heinrikson

s.

Volume 6 Nucleic Acids and Mutability-Stephen Zamenhof Miura Specificity in the Structure of Transfer RNA-Kin-ichiro M. Michelson, J . Massoulii, and W. Guschbauer Synthetic Polynucleotides-A. The DNA of Chloroplasts, Mitochondria, and Centrioles-S. Granick a n d A h a r o n G i b o r Behavior, Neural Function, and RNA-H. H y d k n The Nucleolus and the Synthesis of Ribosomes-Robert P. Perry P. Georgieo The Nature ond Biosyntheris of Nuclear Ribanucleic Acids-G. Replication of Phage RNA-Charles Weissmann a n d Seuero Ochoa Volume 7 Autoradiographic Studies on DNA Replication in Normal and leukemic Human ChromosamesFelice Gauosto Proteins of the Cell Nucleus-Lubomir S. H n i l i c a The Present Status of the Genetic Code-Carl R. Woese Chantrenne, A. Burny, a n d G. M a r The Search for the Messenger RNA of Hemoglobin-H. baix Ribonucleic Acids and Information Transfer in Animal Cells-A. A. Hadjiolou Transfer of Genetic Information during Embryogenesis-Martin Nemer Enzymatic Reduction of Ribonucleotides-Agne Larsson a n d Peter Reichard The Mutagenic Action of Hydroxylamine-1. H. P h i l l i p s a n d D.M. Brown Mammalian Nucleolytie Enzymes and Their Localization-David Shugar a n d H a l i n a Sierak o ws k a Volume 8 Nucleic Acids-The First Hundred Years-J. N . Dnvidson Nucleic Acids and Protamine in Salmon Testes-Gordon H. Dixon a n d M i c h a e l Smith Experimental Approaches to the Determination of the Nucleotide Sequences of large OligonuW. H o l l e y cleotides and Small Nucleic Acids-Robert Alterations of DNA Base Composition in Bacteria--6. F. Cause Chemistry of Guanine and I t s Biologically Significant Derivatives-Robert Shapiro

26 1

CONTENTS OF PREVIOUS VOLUMES

Bacteriophage 4x174 and Related Viruses-Robert L. Sinsheimer W. Rushizky a n d The Preparation and Characterization of Large Oligonucleotides-George Herbert A. Sober Purine N-Oxides and Cancer-George Bosworth B r o w n The Photochemistry, Photobiology, and Repair of Polynucleotides-R. B. Setlow What Really Is DNA? Remarks on the Changing Aspects of a Scientific Concept-Erwin Chargafl Recent Nucleic Acid Research in China-Z'ien-Hsi Cheng a n d Roy H. D o i

Volume 9 The Role of Conformation in Chemical Mutagenesis-B. Singer a n d H. Fruenkel-Conrat Polarographic Techniques in Nucleic Acid Research4. PaleEek RNA Polymerase and the Control of RNA Synthesis-John P. Richardson Radiation-Induced Alterations in the Structure of Deoxyribonucleic Acid and Their Biological Consequences-D. T. K a n a z i r Tsi Yang a n d Tatsuya Optical Rotatory Dispersion and Circular Dichroism of Nucleic Acids-Jen Sameji mu The Specificity of Molecular Hybridization in Relation to Studies on Higher Organisms-P. M . B. Walker Quantum-Mechanical Investigations of the Electronic Structure of Nucleic Acids and Their Constituents-Bernard P u l l m a n a n d Alberte P u l l m a n K . Kochetkoz; a n d E. I. Budowsky The Chemical Modification of Nucleic Acids-N.

Volume 10 Induced Activation of Amino Acid Activating Enzymes by Amino Acids and tRNA-Ahn H. M e h l e r Transfer RNA and Cell Differentiation-Noboru Sueoka a n d Tamiko Kano-Sueoka N@-(A'-Isopentenyl)adenosine: Chemical Reactions, Biosynthesis, Metabolism, and Significance to the Structure and Function of tRNA-Ross H. H a l l Nucleotide Biosynthesis from Preformed Purines in Mammalian Cells: Regulatory Mechanisms and Biological Significance-A. W. M u r r a y , D a p h n e C. E l l i o t t , a n d M . R . Atkinson Ribosome Specificity of Protein Synthesis in Vitro-Orio Ciferri a n d B r u n o Parisi Synthetic Nucleotide-peptides-Zoe A. Shabarova The Crystal Structures of Purines, Pyrimidines and Their Intermolecular Complexes-Donald Voet a n d Alexander Rich

Volume 1 1 The Induction of Interferon by Natural and Synthetic Polynucleatides-Clarence Colby, Jr. Ribonucleic Acid Maturation in Animal Cells-R. H. B u r d o n Liporibonucleaprotein as an Integral Part of Animal Cell Membranes-V. S . Shapot a n d S. Ya. Davidoua Uptake of Nonviral Nucleic Acids by Mammalian Cetls-Pushpa M. Bhargava and G. Shanmugam The Relaxed Control Phenomenon-Ann M . Ryan a n d E r n e s t Borek 1. Douern Molecular Aspects of Genetic Recombination-Cedric E . Kennel/ Principles and Practices of Nucleic Acid Hybridization-David Recent Studies Concerning the Coding Mechanism--Thomas H . Jukes a n d L i l a G a t l i n The Ribosomal RNA Cistrons-M. L. Birnstiel, M . Chipchase, a n d J. Speirs Three-Dimensional Structure of tRNA-Friedrich Cramer Becker a n d Jerard Hurwitz Current Thoughts on the Replication of DNA-Andrew Reaction of Aminoacyl-tRNA Synthetases with Heterologous tRNA's-K. Bruce Jacobson On the Recognition of tRNA by Its Aminaacyl-tRNA Ligase-Robert Chambers

w.

262

CONTENTS OF PREVIOUS VOLUMES

Volume 12 Ultraviolet Photochemistry as o Probe of Polyribonucleotide Conformation-A.

1.L o m a n t

and

Jacques R . Fresco Some Recent Developments in DNA Enzymology-Mehran Goulian Minor Components in Transfer RNA: Their Characterization, Location, and Function-susumu Nishimura B. toftfield The Mechanism of Aminoacylation of Transfer RNA-Robert K . F. Bautz Regulation of RNA Synthesis-Ekkehard The Poly(dA-dT) of Crab-M. Laskowski, Sr. The Chemical Synthesis and the Biochemical Properties of Peptidyl-tRNA-Yehuda L a p i d o t a n d N a t h a n de Groot Volume 13 Reactions of Nucleic Acids and Nucleoproteins with Formaldehyde-M. Ya. F e l d m a n P. Deutscher Synthesis and Functions of the -C-C-A Terminus of Transfer RNA-Murray Mammalian RNA Polymerases-Samson T. Jacob Poly(adenosine diphosphate ribose)-Z’akashi Sugimura The Stereochemistry of Actinomycin Binding to DNA and Its Implications in Molecular BiologyH e n r y M. Sobell H. Richmond Resistance Factors and Their Ecological Importance to Bacteria and to Man-M. Lysogenic Induction-Ernest Borek a n d Ann R y a n Recognition in Nucleic Acids and the Anticodon Families-Jacques Ninio Translation and Transcription of the Tryptophan Operon-Fumio lmamoto lymphoid Cell RNA’s and Immunity-A. Arthur C o t t k e b Volume 14 DNA Modification and Restriction-Werner A r b e r Mechanism of Bacterial Transformation and Transfection-Nihd K . N o t a n i andJane K . Setlow DNA Polymerares II and 111 of Ercherichia c o l i - M a l c o h L. Gefter The Primary Structure of DNA-Kenneth M u r r a y a n d Robert W. Old RNA-Directed DNA Polymerase- Properties and Functions in Oncogenic RNA Viruses and CellsM a u r i c e Green a n d G r a y F. G e r a r d Volume 15 Information Transfer in Cells Infected by RNA Tumor Viruses and Extension to Human NeoplasiaD. Gillespie, W. C. Saxinger, a n d R . C. G a l l o Mammalian DNA Polymemses-F. J. Bollurn Eukaryotic RNA Polymerares and the Factors That Control Them-E. B . Elswas, A. Ganguly, a n d D. D a s Structural ond Energetic Consequences of Noncamplementary Bare Oppositions in Nucleic Acid Helices-A. J. L o m a n t a n d Jacques R. Fresco The Chemical Effects of Nucleic Acid Alkylation and Their Relation to Mutagenesis and Carcinogenesis-E. Singer Effects of the Antibiotics Netropsin and Distomycin A on the Structure and Function of Nucleic Acids-Christoph Zimmer Volume 16 Initiation of Enzymic Synthesis of Deoxyribonucleic Acid by Ribonucleic Acid Primers-Erwin Chargaff Transcription and Processing of Transfer RNA Precursors-John D. Smith

263

CONTENTS OF PREVIOUS VOLUMES

Bisulfite Modification of Nucleic Acids and Their Constituents-Hikoya Hayatsu The Mechanism of the Mutagenic Action of Hydroxylamines-E. I . Budowsky Diethyl Pyrocarbonate in Nucleic Acid Research-L. Ehrenberg, 1. Fedorcsrik, a n d

F. Sol-

YmosY Volume 17 The Enzymic Mechanism of Guanosine 5', 3'-Polyphosphate Synthesis-Fritz

L i p m a n n a n d Jose

SY

Effects of Polyamines on the Structure and Reactivity of tRNA-Ted T.Sakai a n d Seymour S. Cohen Information Transfer and Sperm Uptake by Mammalian Somatic Cells-Aaron Bendich, E l l e n Borenfreund, Steven S. Witkins, D e l i a Beju, a n d P a u l J . H i g g i n s Studies on the Ribosome and Its Components-Pnina Spitnik-Elson a n d D a v i d Elson Classical and Postclassical Modes of Regulation of the Synthesis of Degradative Bacterial Enzymes-Boris Magasanik Characteristics and Significance of the Polyadenylate Sequence in Mammalian Messenger RNAGeorge B r a w e r m a n Polyadenylate Polymerases-Mary Edmonds a n d M a r y Ann Winters Kim Three-Dimensional Structure of Transfer RNA-sung-Hou Insights into Protein Biosynthesis and Ribosome Function through Inhibitors-Sidney Pestka Interaction with Nucleic Acids of Carcinogenic and Mutagenic N-Nitroso Compounds-W. L i j i n s k y Biochemistry and Physiology of Bacterial Ribonuclease-Alok K . D a t t a a n d S a l i l K . N i y o g i Volume 18 The Ribosome of Ercherichia coli-R.

Brimacombe,

K . H. Nierhaus, R . A.

Garrett and

H. G.

Wittmann Structure and Function of 5 S and 5.8 S RNA-Volker A. E r d m a n n High-Resolution Nuclear Magnetic Resonance Investigations of the Structure of tRNA in SolutionD a v i d R. Kearns Premelting Changes in DNA Conformation-E. PaleEek Quantum-Mechanical Studies on the Conformation of Nucleic Acids and Their ConstituentsB e r n a r d Pullman a n d A n i l Saran

Volume 19: Symposium on mRNA: The Relation of Structure a n d Function I. The 5'-Terminal Sequence ("Cap") of mRNAs Caps in Eukaryotic mRNAs: Mechanism of Formation of Reovirus mRNA 5'-Terminal m'GpppGmC-Y. F u r u i c h i , S. Muthukrishnan, J . Tomasz a n d A. J . Shatkin M. Rottman, R o n a l d C . D e Nucleotide Methylation Patterns in Eukaryotic mRNA-Fritz srosiers a n d K a r e n F r i d e r i c i Busch, Structural and Functional Studies on the "5'-Cap": A Survey Method of mRNA-Harris F r i e d r i c h Hirsch, Kaushal Kumar Gupta, M a n c h a n a h a l l i Rao, W i l l i a m Spohn a n d B e n j a m i n C. Wu Modification of the 5'-Terminals of mRNAs by Viral and Cellular Enzymes-Bernard Moss, Scott A. M a r t i n , M a r c i a J . Ensinger, Robert F. Boone a n d Cha-Mer Wei Blocked and Unblocked 5' Termini in Vesicular Stomatitis Virus Product RNA in Vitro: Their Possible Role in mRNA Biosynthesis-Richard J . Colonno, Gordon A b r a h a m a n d A m i y a K. Banerjee The Genome of Poliovirus Is an Exceptional Eukaryotic mRNA-Yuan Fon Lee,Akio N o m o t o a n d E c k a r d Wimmer

264

CONTENTS OF PREVIOUS VOLUMES

It. Sequences and Conformations of mRNAs Transcribed Oligonucledide Sequences in Hela Cell hnRNA and

Hiroshi Nakazato, E. L. Korwek'and S. Venkatesan Polyadenylylation of Stored mRNA in Cotton Seed Germination-Barry

mRNA-Mary

Edmonds,

Harris a n d Leon Dure

Ill mRNAs Containing and Lacking Poly(A) Function as Seporate a n d Distinct Classes during Embryonic

Development-Martin Nemer a n d Saul Surrey Sequence Analysis of Eukaryotic mRNA-N.]. Proudfoot, C. C. Cheng a n d G. G. Brownlee T h e Structure and Function of Protamine mRNA from Developing Trout Testis-P. L. Dauies, G . H.

Dixon, L. N. Ferrier, L. Gedamu a n d K. Iatrou The Primary Structure of Regions of SV40 DNA Encoding the Ends of mRNA-Kiranur

N. Subramanian, Prabhat K . Ghoshi, Ravi Dhar, Bayar Thimmappaya, Sayeeda B. Zain, Julian Pan a n d Sherman M. Weissman Nucleotide Sequence Analysis of Coding and Noncoding Regions of H u m a n P-Globin mRNACharles A. Marotta, Bernard G . Forget, Michael CohenlSolal a n d Sherman M. Weissman Determination of Globin mRNA Sequences and Their Insertion into Bacterial Plasmids-Winston Salser, Jeff Browne, P a t Clarke, Howard Heindell, Russell Higuchi, Gary Paddock, John Roberts, Gary Studnicka a n d Paul Zakar The Chromosomal Arrangement of Coding Sequences in a Family of Repeated Genes-G. M. Rubin, D. J. Finnegan a n d D. S. Hogness Mutation Rates in Globin Genes: The Genetic Load and Haldane's Dilemma-Winston Salser a n d Judith Strommer Isaacson Heterogeneity of t h e 3' Portion of Sequences Reluted to Immunoglobulin K-Chain mRNA-ursula

Storb Globin mRNA-]ohn N. Vournakis, Marcia S . Flashner, MaryAnn Katopes, Gary A. Kitos, Nikos C. Vamvakopoulos, Matthew S. Sell a n d Regina M. Wurst Molecular Weight Distribution of RNA Fractionated on Aqueous and 70% Formamide Sucrose Gradients-Helga Boedtker a n d Hans Lehrach 111. Processing of mRNAs Bacteriophages T7 and T 3 as Model Systems for RNA Synthesis and Processing-].]. Dunn, c. W. Anderson,]. F. Atkins, D. C. Bartelt a n d W. C. Crockett The Relationship between hnRNA and mRNA-Robert P. Perry, Enzo Bard, B. David Hames, D a w n E. Kelley a n d Ueli Schibler A Comparison of Nuclear and Cytoplasmic Viral RNAs Synthesized Early in Productive Infection with Adenavirus 2-Heschel J . Raskas a n d Elizabeth A. Craig Biogenesis of Silk Fibroin mRNA: An Example of Very Rapid Processing?-Paul M. Lizardi Structural Studies on Intact a n d Deadenylylated Rabbit

Visualization of the Silk Fibroin Transcription Unit and Nascent Silk Fibroin Molecules on Polyribosomes of Bombyx mori-Steven L. McKnight, Nelda L. Sullivan a n d Oscar L. Miller,]r. Production and Fate of Balbiani Ring Products-B. Daneholt, S. T. Case, J. Hyde, L. Nelson

a n d L.Wieslander

Nuclear Ribanucleoprotein Complexes-Alan Kinniburgh, Peter B. Billings, Thomas]. Quinlan a n d Terence E. Martin IV. C h r o m a t i n Structure and Template Activity The Structure of Specific Genes in Chromatin-Richard Axel Distribution of hnRNA and mRNA Sequences in

J.

T h e Structure of DNA in Native Chromatin as Determined by Ethidium Bromide Binding-].

Paoletti, B. B. Magee a n d P. T. Magee Skeletons and RNA Messages-Ronald Robert Lenk a n d Sheldon Penman

Cellular

Herman, G a y Zieue, Jeffrey Williams,

265

CONTENTS OF PREVIOUS VOLUMES

The Mechanism of Steroid-Hormone Regulation of Tronscription of Specific Eukoryotic Genes-Bert W. O’Malley a n d Anthony R. Means Nonhirtone Chromosomal Proteins and Histone Gene Transcription-Gary Stein, Janet Stein, L e w i s Kleinsmith, W i l l i a m Park, Robert Jansing and Judith Thomson Selective Tronscription of DNA Mediated by Nonhistone Proteins-Tung Y. Wang, N i n a C . Kost r a b a a n d Ruth S. N e w m a n V. Control of Translation Structure and Function of the RNAs of Brome Mosaic Virus-Paul Kaesberg Effect of 5’-Terminal Structures on the Binding of Ribopolymers to Eukaryotic Ribosomes-s. Muthukrishnan, Y. Furuichi, G. W. B o t h a n d A. J. Shatkin Tronslotionol Control in Embryonic Muscle-Stuart M . H e y w o o d and D o r i s S. Kennedy Protein and mRNA Synthesis in Cultured Muscle Cells-R. G. Whalen, M . E. B u c k i n g h a m a n d

F. Gros VI. Summary mRNA Structure and Function-James

E. D a r n e l l

Volume 20 Correlotion of Biological Activities with Structural Features of Tronsfer RNA-B. F. C . Clark Bleomycin, on Antibiotic That Removes Thymine from Double-Stronded DNA-lverner E. G. MCll e r a n d Rudolf K . Z a h n Mammolian Nucleolytic Enrymes-Ha/ina Sierakowska a n d D a v i d Shugar C. Waters a n d B e t h C . Mullin Transfer RNA in RNA Tumor Viruses-Larry Integration versus Degradation of Exogenous DNA in Plants: An Open Question4‘aul F. Lurquin Initiation Mechanisms of Protein Synthesis-Marianne Grunberg-Manago a n d FrunGois Gros

Volume 21 lnformosomes ond Their Protein Components: The Present State of Knowledge-A. A. Preobrazhensky a n d A. S . S p i r i n Energetics of the Ribosome-A. S. S p i r i n Mechanisms in Polypeptide Chain Elongation on Ribosomes-Engin Bermek Synthetic Oligodeoxynucleotides for Anolysis of DNA Structure and Function-Ray Wu, Chander P. Bahl, a n d Saran A. N a r a n g The Tronsfer RNAs of Eukaryotic Orgonclles-W. E d g a r Barnett, S . D. Schwartzbach, a n d L. 1. Hecker Regulation of the Biosynthesis of Aminoacid:tRNA Ligoses ond of tRNA-Susan D. M o r g a n and D i e t e r Sol1

Volume 22 The -C-C-A End of tRNA ond Its Role in Protein Biosynthesis-Mathias Sprinzl a n d F r i e d r i c h Cramer The Mechanism of Action of Antitumor Platinum Compounds-J. J. Roberts a n d A. J. Thornson DNA Glycosyloses, Endonuclearer for Apurinic/Apyrimidinic Sites, and Bore Excision-RepoirThomas L i n d a h l Naturolly Occurring Nucleoside and Nucleotide Antibiotics-Robert J. Suhadolnik Genetically Controlled Variation in the Shapes of Enzymes-George Johnson E. Transcription Units for mRNA Production in Eukaryotic Cells and Their DNA Viruses-James Darnell, Jr.

266

CONTENTS OF PREVIOUS VOLUMES

Volume 23 The Peptidyltransferase Center of Ribosomes4lexander A. Krayeusky and Marina K. Kukhanoua Patterns of Nucleic Acid Synthesis i n Physarum polycoph~lu~Ceoflrey Turnock Biochemical Effects of the Modification of Nucleic Acids by Certain Polycyclic Aromatic Carcinogens-Dezider Grunberger a n d 1. Bernard Weinstein Participation of Modified Nucleosides in Translation and Transcripti0n-B. Singer and M. Kruger The Accuracy of Translation-Michael Yams Structure, Function, and Evolution of Transfer RNAs (with Appondix Giving Complete Sequences of 178 tRNAs)-Ram P. Singhal and Pamela A. M. Fallis Volume 24 Structure of Transcribing Chromatin-Diane Mathis, Pierre Oudet, and Pierre Chambon Ligond-Induced Conformational Changes in Ribonucleic Acids-Hans Canter Gassen Replicative DNA Polymerares and Mechanisms at a Replication Fork-Robert K. Fujimura and Shishir K. Das Antibodies Specific for Modified Nucleosides: An lmmunochemical Approach for the Isolation and Characterization of Nucleic Acids-Theodore W. Munns and M. Kathryn Liszewski DNA Structure and Gene Replication-R. D. Wells, T. C. Goodman, W. Hillen, G. T. Horn, R. D. Klein,]. E. Lurson, U. R. Milller, S. K. Neuendorf, N. Panayotatos, and S. M. Stirdiuant Volume 25 Splicing of Viral mRNAs-Yosef Aloni DNA Mathylation and Its Possible Biological Roles-Aharon Razin and Joseph Friedman Mechanisms of DNA Replication and Mutagenasis in Ultraviolet-Irradiated Bacteria and Mammalian Cells-]mnifer D. Hall and Dauid W. Mount The Regulation of Initiation of Mammalian Protein Synthesis-Rosemary Jagus, W. French Anderson, and Brian Safer Structure, Replication, and Transcription of the SV40 Genome-Coku; C. Das and Salil K . Niyogi Volume 26: Symposium on DNA: Multiprotein Interactions Introduction: DNA- Multiprotoin Interactions in Transcription, Roplication, and Repair-R. K. Fujimura Replicative DNA Polymerase and Its Complex: Summary-David Kern Enzyme Studies of 9x174 DNA Replication-Ken-ichi Arai, Naoko AraiJoseph Shlomai,/oan Kobwi, Laurien Polder, Robert Low, Ulrich Habscher, LeRoy B d s c h , and Arthur Kornberg The DNA Replication Origin (ari) of Eschrichia coli: Structuro and Function of tho ori-Containing DNA Fragment-Yukinori Hirota, Masao Yamada, Akiko Hishimura, Atsuhiro Oka, Kazunori Sugimoto, Kiyozo Asada, and Mitsuru Takanami Roplicotion of h o a r Duplox DNA in Vitm with Bacteriophage T5 DNA PalymorarcR. K . Fujimura, S . K . Das, D. P. Allison, and B. C. Roop Mechanisms of Catalysis of Human DNA Polymerares a and p-Dauid Korn, Paul A. Fisher, and Teresa S.-F. Wang Structural and Functional Properties of Calf Thymus DNA Polymerase 8-Marietta Y. W. Tsang Lee, Cheng-Keat Tan, Kathleen M. Downey, and Antero G. So Mochanirms of Transcription: Summary-R. K. Fujimura

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CONTENTS OF PREVIOUS VOLUMES

Regulatory Circuits of Bacteriophage Lambda-S. L. Adhya, S. Garges, a n d D . F. Ward Chromatin Transcription and Replication: Summary-Ronald L. Seale Site of Histone Assembly-Ronald L. Seale Chromatin Replication i n Tetmhymem pyriformis-A. T. Annunziato a n d C . L. F. Woodcock Role of Chromatin Structure, Histone Acetylation, and the Primary Sequence of DNA in the Exprersion of SV40 and Polyoma in Normal or Teratocarcinoma Cells-G. Moyne, M . Kutinka, S. Saragosti, A . Chestier, a n d M . Yaniv Control of Transcription in Eukaryotes: Summary-William J . Rutter Repair Replication Schemes i n Bacteria and Human Cells-Philip C. Hunawalt, Priscilla K . Cooper, a n d Charles Allen Smith Recent Developments in the Enzymology of Excision Repair of DNA-Errol C. Friedberg, C w r i e T. M. Anderson, Thomas Bonura, Richard Cone, E r i c H. Raduny, a n d Richard J . Reynolds Multiprotein Interaction i n Strand Cleavage of DNA Damaged by UV and Chemicals-Erling Seeberg In Vitm Packaging of Damaged Bacteriophage 17 DNA-warren E . Masker, Nancy B. Kuemmerle, a n d L o r i A. Dodson The Inducible Repair of Alkylated DNA-John Cairns, Peter Robins, Barbara Sedgwick, a n d Phillipa Talmud Functions Induced by Damaged DNA: Summary-Euelyn M. Witkin Inducible Error-Prone Repair and Induction of Prophage Lambda in Escherichia coli-Raymond Deuoret DNA and Nucleoride Triphosphate Binding Properties of recA Protein from Eschwichia coli-K. McEntee, G. M . Weinstock, a n d 1. R. Lehman Molecular Mechanismfor the Induction of “SOS” Functions-Michio Oishi, Robert M. Irbe, a n d Lee M. E. M o r i n Induction and Enhanced Reactivation of Mammalian Viruses by Light-Larry E. Bockstahler Comparative Induction Studies-Ernest C. Pollard, D. J . Fluke, a n d Den0 Kazanis Concluding Remarks-Ernest C. Pollard

Volume 27 Poly(adenosine diphosphate ribose)-Paul Mandel, Hideo Okazaki, a n d Claude Niedergang The Rogulatory Function of Poly(A) and Adjacent 3’ Soquencer in Translated RNA-vriel Littauer a n d Hennona S w e q tRNA-like Structures in the Genomer of RNA Viruses-Anne-Lise Haenni, Sadhna foshi, a n d Francois Chapeville Mechanism of Interferon Action: Progress toward Its UndentandingGanes Sen RNA-Helix-Dertabilizing Proteins-John 0.Thomas a n d Wlodzimierz Szer Nucleotide Cyclaser-Laurence S . Bradham a n d Wai Yiu Cheung Cyclic Nucleotide Control of Protein Kinawr-R. K . S h a n a

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