Structure And Synthesis Of Ribosomal Rna

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Copyright I970. All rights reserved

STRUCTURE AND ~ SYNTHESIS OF RIBOSOMALRNA CIUSEPPE

ATTARDI

AND

F~CESCO

AMALDI

Annu. Rev. Biochem. 1970.39:183-226. Downloaded from arjournals.annualreviews.org by 192.244.210.205 on 10/25/05. For personal use only.

Division o/Biology, California Institute of TechnologyPasadena,California. Centro degll Addl Nuddcl del C.N.R., lst~tuto di F~slologia Generale Universitgt di Roma,Italy CONTENTS

INTRODUCTION ..................................................... HIGH MOLECULARWEIGHT RIBOSOMAL RNA .................. SIZE ................................................................. ~. .................................................. PRI~AR"C$~ucxum Base composition, major component~ ................................. Boze composition, minor compon¢nts .................................. Partial sequences .................................................... SECONDARY AND TERTIARY

STRUCTURE ...................................

192

SYNTheSIS........................................................... Bacterial sy~tem~ .................................................... Eukaryotic system~ .................................................. 5S RIBOSOMAL RNA................................................ SIz~ ................................................................. S’r~uc~n~ ........................................................... Primary struaure ................................................... ~evondar~ stmvt~re ................................................. SYNTHESIS

195 195 200 209 209 209 209 210 210

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

LOCATION, REDUNDANCY, AND VARIABILITY RNAGENES........................................................ CONCLUDING REMARKS ...........................................

183 185 185 186 186 188 190

OF RIBOSOMAL 213 218

INTRODUCTION The great advances made in the last decade in.the analysis of the mechanism of protein synthesis have led to the elucidation of the general features of this process (1). It is now clear that further progress in this area will depend to a great extent on the development of our knowledge concerning the molecular organization of ribosomes. For this reason, studies on the structure and assembly of the RNAand protein components of ribosomes are of particular importance, Seen in another context, the synthesis and pro* The authors thank Dr. R. Monier for sending a manuscript prior to publication. Research cited in this review which was carried out in the authors’ laboratories was supported by a grant from the National Institutes of Health (GM11726) and by a grant from the Italian National Research Council (C.N.R.). 183

720

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184

ATTARDI & AMALDI TABLE 1. Sedimentation coefficient of ribosomes and sedimentation eoeffleient and molecular weight of high molecular weight ribosomal RNAfrom some representative organisms Ribosomes

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Organism

High molecular

Sedimentation Sedimencoefficient tation (S value) coefficient Larger (S value) Smaller species species

Prokaryotic celts Bacteria Escherichia coli Bacillus cereus Streptomycesgriseus Blue-green algae Anaboena Nostoc Oscillatoria Eukaryoti6 cells Algae Chlordla Chlamydomonas Protozoa Acanlhamoebacaslellonll Euglena gracilis Tetrahymena pyriformis Paramecium Fungi Diclyoslelium Aspergillus Neurospora crassa Saccharomycescere~isiae Higher plants Pea

s69.1 a68.6

23.50

s16.3

°22.5

°15.8

s71.6

weight rRNA Molecular weight 4) (daltons X 10

References

Larger species

Smaller species

i,l1.07 k1.10

i,|0.55 k0.56

1.11

0.56

8 9, 10

1.07 1,07 1.07

0.55 0.56 0.56

9, 10 10 10

$, 6. 7

a83

t25.1

f17.6

1.28 1.30

0.69 0.69

I0 11~ 12, 10

b86 a78 a84.5

g24 f25.4 h25.5

g20 f17 h17.7

1.53 1.30 1.30 1.31

0.89 0.85 0.69 0.69

I0 13, 10 14~ 15, 10 16, I0

77o ~ 81.3 76.90 a81.5

25f

17f

f25 h24.6

f17.4 h16.2

1.30

0.73

1.30

0.72

a80

!24.7

16.5f~

b80

t25

16t J

1.27-1.31

0.70-0.71

h28. S 29.40 t28

h18. O °18.8 /18

Bean Animals a80 Ascaris lumbricoides Drosophila melanogaster Arbacia Xenopuslaevis (liver, ovary) Chick (liver)

17 9, 10 18 9, 19 20, 21, 10 22, 10

1.40 1.40 1.54 1.58

0.73 0.68 0.69 0.70

23 24, 10 25j 10 10 10

The sedimentation coefficients of ribosomes were determined either in the analytical ultracentrifuge with schlieren optics (corrected for infinite dilution) (a), or U V optics (d), or by sucrose gradient trifugation in the preparative ultracentxifuge with rat liver ribosomes (80S) (b), E. col i rib osomes (70S) (e), as a standard. The sedimentation runs of rRNAwere carried out at a salt concentration between 0.01 Mand 0.1 M, in general in the absence of Mgions. The sedimentation coefficients were determined either in the analytical ultracentrifuge with ~chlieren optics (corrected for infinite dilution) (h), or UVoptics or by sucrose gradient centrifugation in the preparative ultracentrifuge with E. coli rRNAspecies (16-16.7S, 23S) (f), or rat liver rRNAspecies (18S, 29S) (g), as standards. The molecular were determined by measurements of: sedimentation and viscosity (i), sedimentation equilibrium (j), light scattering, sedimentation and viscosity (k), intrinsic viscosity 0), or electrophoretic mobility with appropriate standards (all others).

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AND SYNTHESIS

OF RIBOSOMAL

RNA

185

TABLE 1. (Continued) Ribosomes

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Organism

Sedimentation coefficient (S value)

Sedimentation coefficient (S value)

Molecular weight (daltons X 10~

References

Larger species

Smaller species

Larger species

Smaller species

e29.8 f29.5 e30.0

e16.8 f17.8 e17.4

1.72 1.75 1.641 |1.9

0.70 0.70 0.671 0.71|

26, 27, 10 28, 18, 10 29 30, 31

a67 b67

f22.4 f23

f16.7 f16

1.07-1.11

0.56

11, 12 22, 10

°73.2

f23 h22.4

f16 h17.8

dRabbit (reticulocyte~ 78 a83 Rat 0ive~ Mouse(Jensensarcoma) ad Man(HeLa cells) Organdies of t~ukaryoti~~ells Chloroplasts Chlamydomonas Higher plants Mitoehondria Neurost)ora¢rassa Saccharomyce$ cerevisiae

High molecular weight rRNA

18, 32 19

cessing of ribosomal RNAs, both in bacteria and eukaryotic cells, represent a model for the study of transcription and its regulation and for that of posttranscriptional modifications of RNA. This article reviews the experimental and conceptual developments which have led to our present understanding of the structure and synthesis of ribosomal RNA(rRNA). Particular emphasis has been placed on the evolutionary aspects of the structure of rRNA. Ribosomes from all organisms contain as intrinsic constituents three RNAspecies (2) : two species of high molecular weight, one pertaining the large subunlt and the other to the small subunit, and one species of low molecular weight, known as 55 RNAfor its sedimentation coefficient, which is associated with the large ribosomal subunit. In some animal cells another low molecular weight RNAcomponent, 75 RNA, has been isolated from the larger high molecular weight rRNA species by processes which disrupt hydrogen bonding (3, 4). This component apparently results from a physiological splitting of the 285 RNAwhich occurs early in the maturation process of the larger subribosomal particle. HIGH

MOLECULAR

WEIGHT

RIBOSOMAL

RNA

A large amount of experimental evidence has pointed to the existence in living cells of two classes of ribosomes: one, with a sedimentation coefficient of about 705, which is present in bacteria and in organelles (chloroplasts and mitochondria) of eukaryotic cells, and the other, with a sedimentation coefficient of about 805, which is found in eukaryotlc cells (Table 1 ). The high molecular weight rRNA in bacterlal-type 705 ribosomes has been uniformly found to consist of a larger component with a sedimentation

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186

ATTARDI & AMALDI

constant of about 238 and a molecular weight of about 1.1 X 10" daltons, and a smaller component ~vith a sedimentation constant of about 16S and a molecular weight of about 0.55 X I0" daltons (Table 1). By contrast, the 80S ribosomes of eukaryoti¢ cells have been shown to vary in the size of their rRNAcomponents in relationship to the evolutionary position of the organism. Thus, the 80S ribosomes of lower eukaryotie cells and plant cells contain a 25S RNAspecies with a molecular weight of about 1.3 X 10., and a 16 to 18S RNAspecies with a molecular weight of about 0.7 X 10~ [the occurrence of a small rRNAspecies of a somewhat larger size has been reported in some protozoa (Amoeba, Euglena) (10), but the siguificanee of these ob;ervations is not clear] ; on the other hand, ribosomes from animal cells, which contain a 17-18S minor rRNA component, vary in their major rRNAspecies, which apparently increases in size from the lower to the higher animal forms (from a mol wt 1.4 X 10" in Arbacia and Drosophila to a mol wt 1.7-1.9 X 106 in mammals) (Table 1). PRIMARYSTRUCTURE Base composition, major components.--The lack of correlation between the base composition of rRNAand that of DNAin various organisms was implied in the first base composition analyses performed on total RNA (which consists mainly of rRNA) from different bacteria (33). This of correlation is not surprising since we know now that rRNAis coded by a very small fraction of the genome (see below). Table 2 summarizes the base composition data for the two purified high molecular weight rRNAcomponents from various representative organisms. An analysis (34) of Table 2 and Figure 1 indicates that the percentage GC varies considerably among different organisms for the major rRNA component and, to a lesser extent, also for the minor one. More significantly, the base composition o~ the two species tends to vary in parallel. Furthermore, an evolutionary trend in this variation is apparent. In fact, in lower eukaryotic and plant cells the GCcontent is in general lower than in animal species, and within the latter there is a tendency for the percentage of GCto increase when going from the lower to the more evolved forms. It also appears from Table 2 that from lower eukaryotic and plant cells to animal cells there is a tendency to a reduction in the proportion of bases which are noncomplementary. Both types of observations mentioned hbove suggest that the evolution of rRNAhas proceeded towards an increase in secondary structure, in agreement with other findings (see below). Mitochondrial rRNAof Neurospora, which is similar in sedimentation properties to bacterial rRNA(18), has a base composition not only different from that of the cytoplasmic rRNAof the same cells but also from that of any bacterial-type rRNA (18, 38). Only unfractionated rRNA from higher plant chloroplasts has been analyzed, and it appears to be rather similar in base composition to the cytoplasmic rRNAfrom the same source (55).

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187

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188

~ 6c

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-6 4O

GC% in larger high M.WrRNA FIG. 1. Relationship in GCcontent betweenthe larger and the smaller high molecular weight ribosomal RNA. + E. coli (7), Aerobacter aerot~enes (53), B. subtilis (35), Pseudomonas aerug,nose (36), Proteus vulgaris (53). Saccharomyces cerevisiae (37), Neurospora crasse (38), Coprlnus micaceus (39), Psalliota campestrls (39). O Potato (40), pea (40), cauliflower (39), cabbage (39), parsnip (39), celery (39), [] Mitochondri~. ~rom Ne~rospora crassa (38), Saccharomyces cerevlsiae (37). 0 Ascari.~ lumbrlcoides (23). Drosophila melanogaster (2 4). <~ Arbacia pun tulata (25). Xenopu¢ laevls (41). [] Duck (42 ). A Rabbit (43 ), rat (43) (102), hamster (54), pig (54), man

Base composition, minor components.--The high molecular weight rRNAcontains, besides the four commonnucleotides, a small amount of unusual nucleotides. The presence of pseudourldylle acid has been demonstrated in rRNA extracted from various sources (44, 56-60), and it has been clearly shown not to be due to contamination by tRNA (44, 58). In coli 23S RNApseudouridylic acid has been found in the proportion of 0.150.30 moles percent, and in 16S RNAin the proportion of 0.06-0.12 moles percent (58, 59). The amount of pseudourldylic acid appears to be higher rRNAfrom animal and plant cells than in/~. coli rRNA(1.2-1.8 mole percent) (44, 54, 57) ; in HeLacells, in contrast E. col i, the pseudouridylic acid content, in mole percent, of the smaller rRNAcomponent is higher (by about 40%) than that of the larger rRNAcomponent (44). Methylated nucleotides have been found in high molecular weight rRNA from various organisms (54, 58, 61-68). In E. coli 23S RNAthe methylated

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STRUCTURE AND SYNTHESIS

OF RIBOSOMAL RNA

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nucleotides represent about 0.6 mole percent; in 16S RNA,about 0.95 mole percent (61). In animal and plant cells the reported level of methylation somewhat higher (1.2-1.7 mole percent) (63, 64, 66, 68). As E. col i, the specific methylation o~ the smaller rRNAcomponent (18S RNA)in animal cells has been found to be higher than that of the larger component [by 30 to 50% in HeLa cells (63, 66, 69) and 15 to 20% in L cells (68) and Krebs II ascites tumor cells (65)]. Methylatlon of a ribonucleotide can occur either on the base or on the ribose moiety (2’-OH group) : the latter results in the resistance of the internucleotide linkage to alkali or ribonuclease digestion, due to interference with the formation of 2’,3’-cyclic phosphate intermediates (70). In E. coli rRNA more than 80% of the methyl groups are present in bases (61). contrast, in L cells and HeLa cells between 80 and 90% of the methyl groups have been reported to occur in the ribose moieties (63, 66, 68) : however, evidence indicating that losses of unstable methylated bases during the manipulations may increase the apparent relative proportion of methylated ribose has been recently presented for HeLa cell rRNA(67) ; in this material 50 to 60%of the methyl groups have in fact been found to be associated with the bases when a milder treatment was employed. The above-mentioned higher proportion of methyl groups in the smaller rRNAcomponent relative to the larger one appears in general to concern both the methylated bases and the methylated riboses. The study of the methylated bases in rRNAhas been seriously complicated by the lability of manyof these compounds which results in their destruction or conversion to other methylated bases (67). The identified methylated bases found in E. coli high molecular weight rRNAinclude derivatives of guanine (N*-methylguanine, 1-methylguanine, 7-methylguanine), adenine (N6-methyladenine, N6-dimethyladenine, 2-methyladenine) and cytosine (5-methylcytoslne, N*-methylcytoslne, and probably a dimethyl- derivative of cytosine) (61). In both components of high molecular weight 4rRNAfrom HeLa cells the main methylated base has been found to be N methylcytosine, which is present in the average in the proportion of 7 residues (out of about 40 methylated nucleotides) per 18S molecule and 13 residues (out of about 70 methylated nucleotides) per 28S molecule (67). Other methylated bases reported to be present in 18S and 28S rRNA from HeLa cells are derivatives of adenine (N6-methyladenine, N6-dimethyladenine, 1methyladenine), cytosine (3-methylcytosine) and uracil (3-methyluracil); methylated derivatives of guanine (1-methylguanine, N2-methylguanine, dimethylguanine, 7-methylguanine) have also been found in 18S RNAand in trace amounts in 28S RNA(67). Reported analyses of the methylated bases in high molecular weight rRNAof other eukaryotic cells [rat liver (54), L cells (68), S. cerevis~ae (71)] have revealed only some of the residues detected in I-IeLa cells, particular N6-methyladenine and N6-dimethyladenine. The failure to detect in these materials the N*-methylcytosine may be due to the extreme lability

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of this base (67). Note that the exclusive presence of N~-dimethyladenine in the smaller high molecular weight rRNA component has been reported for E. coli (61), L cells (68), and by several authors for HeLa cells (63, 69). At variance with these observations is the above-mentioned report of the presence of this methylated base also in 28S RNAfrom HeLa cells (67) ; the reason for this discrepancy is not clear. Partial sequences.--The analysis of nucleotide sequences in high molecular weight rRNAhas been carried out to the present time only at the level of the partial sequences, either terminal or internal, isolated after enzymatic or alkali digestion. The 5’-terminal sequences of rRNA molecules were studied in various organisms (72, 73) by using a method based on enzymatic phosphorylation with radioactive orthophosphate of the Y-terminal hydroxyl group (after removal of the terminal phosphate by alkaline phosphatase), followed by hydrolysis with pancreatic or T1 RNase and identification of the radioactive fragments. By this procedure, the main 5’-terminal fragment in E. coli 16S RNA was found to be pApApApUpGp- and in 23S RNA pGpGpUp-. In three Bacillus species (B. cereus, B. subtilis, and B. stearothermophilu~) 16S and 23S RNAterminated at the 5’-end respectively with pUpXpXpXpXpGp- and pUpXpXpXpG- (where X is one of the four cleosldes). Sarcina lutea, on the other hand, had pUpXpXpXpGp-as 5’terminal sequence in 16S RNAand pApApGpPyp- in 23S RNA. These observations indicate that related bacterial species may have identical 5’terminal sequences, whereas these differ in unrelated species. Fewer data are available on the 3’-terminal sequences of bacterial rRNA. The predominant 3’-terminal sequences of E. coli 16S and 23S RNA were found to be -pGpCpA and -pPpU respectively (74): these observations were made ~vith cells grown on glucose-salts or succinate-salts media. Later investigations by the same authors (75) suggested that when the cells are grown in richer media the terminal sequence -pPypU becomes the predominant 3’-terminal sequence also in 16S RNA.The significance of these observations with respect to the problem of heterogeneity of rRNAis uncertain (see below). The above-mentioned observations concerning the difference in 5’terminal and 3’-terminal sequences between 16S and 23S RNAof bacterial species agree with the structural data discussed above and below and with the DNA-RNA hybridization data (see below) in arguing against the possibility that the major rRNAspecies is a dimer of the minor species. Concerning the terminal sequences of the high molecular weight rRNA in eukaryotic cells the available data are more fragmentary. In Saccaromyces cerevlslae, with the method of enzymatic phosphorylation, pUpXpXpXpXpGp-and pUpUpGp- were detected as 5’-terminal sequences in 16S and 25S RNArespectively (73). In L cells pup- has been reported the main 5’-end group of 18S RNAand pCp- as the main 5’-end group of 28S RNA(76). In the same cells the main 3’-terminal nucleosides were

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OF RIBOSOMAL RNA

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found to be A in 185 RNAand U in 28S RNA(76). Similar results have been reported for the rRNAfrom rabbit reticuloeytes, where the 3’ ends have been found to be -pPypA in 18S RNAand -pPypU and -pGpU in 28S RNA(77) In one plant (cauliflower) A and U have been reported as 3’terminal nucleosides of both 18S and 28S RNA(78). Partial sequence analyses of high molecular weight rRNAfrom various sources have been carried out by digesting the RNAwith pancreatic or T1 RNase and separating the resulting oligonucleotides by paper or column chromatography. By the use of this method, 16S and 23S RNAfrom E. coli were shown to be significantly different from each other in the frequency of the partial sequences released by enzyme digestion (79). No differences were found among samples of total RNAprepared from various E. coli strains or from closely related bacterial species (80) ; on the contrary, base sequence differences were detected among rRNAsamples prepared from species with very different GC content in their DNA(80). In Pseudomonas aert,ginosa and E. coli an influence of the conditions of growth on the partial sequence distribution in 16S RNAand, to a lesser extent, in 23S RNA, has been reported (81). An analysis of the sequences released by T1 digestion which contain methylated nncleotldes was performed on 23S and 16S RNAfrom E. coll (61). A marked difference in the methylatlon pattern between the two rRNA components was found. An interesting observation is that in 23S RNAsubstantially all methylated sequences are repeated twice in the molecule ; this could mean that the 23S RNAmolecule consists of two similar or identical portions (see below). Implicit in these results is the conclusion that the methylated nucleotides are not distributed at random.. This conclusion has been confirmed, as concerns the 2~-O-methylated nucleotides, by an analysis of the distribution of alkali-resistant dinucleotides (59, 82). It interesting that 6 out of the 10 pseudouridylic acid residues of the 23S RNA are located in the mcthylated sequences released by T1 RNase (representing less than 4%of the molecule), a finding which suggests a strong topographical correlation between methylation and location of pseudouridyllc acid (61). A partial sequence analysis after pancreatic RNase digestion of 28S and 18S RNAfrom HeLa cells (44) showed that these components are very different from each other in the frequency of mono-, di-, and trinueleotides. Only a rough agreement was found between the observed frequency of partial sequences and that expected from base composition under the assumption of a random distribution of the bases. The same analysis was carried out for 18S and 28S RNAprepared from different human tissues (45): significant differences in the frequency of partial sequences were detected among the 18S RNApreparations, and respectively 28S RNApreparations, from HeLa cells and the various human tissues examined. An analysis of longer oligonueleotides (hepta-, octa-, and nonanucleotides) released hy pancreatic RNase digestion of IqeLa cell 28S and

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RNArevealed very marked differences in their relative frequency in the two rRNA components (83). As in the case of E. coli rRNA,the distribution of pseudouridylic acid in HeLa cell 28S and 18S RNAwas found to be nonrandom with respect to that of uridylie acid (44). Similarly, the distribution of 2’-O-methylated nueleotides in these rRNAspecies appears to be nonrandom, as revealed by the observation that the frequencies of alkali resistant dlnueleotldes do not reflect the rRNA base composition (84). A similar observation has been made for wheat germ rRNA(57, 64). SECONDARY

AND

TERTIARY

STRUCTURE

A considerable amount of experimental evidence indicates that rRNA molecules both in bacteria and eukaryotic cells are constituted by continuous polynucleotide chains, which are folded and internally base paired in various degrees (secondary structure) depending upon the environmental conditions (85-88). The experimental results obtained by the analysis of the properties of UVabsorption, optical rotatory dispersion, X-ray diffraction, viscosity, and infrared spectroscopy have led to the development of a plausible model for the secondary structure of the high molecular weight rRNAwhich has been discussed in detail (89). Under conditions of moderate ionic strength and at room temperature a large fraction (up to 60-75%) of the nucleotide residues of the long rRNA molecule, both from bacterial and eukaryotic sources, appears to be involved in intramolecular base pairing (85, 90-101). The flexible single-stranded RNAchain can fold upon itself so that two of its sections run in antiparallel directions and, if the nucleotide sequences are complementary, a short double-stranded helical region can be formed. It has been pointed out that double-stranded configurations involving adjacent regions of the RNA molecule are much more probable than those involving distant regions (91). The size of the helices has been estimated to range from 4 to 17 base pairs, with some unpaired nucleotide residues (at least three) at the loop region (95). Moreover, some additional nucleotide residues can loop out of the double helix if the corresponding nucleotide sequences are not perfectly complementary; up to one-fifth of the nucleotides in the helical regions shorter than six base pairs and one-third of the nucleotides in longer helices can loop out without destroying their stability (91). If one considers that molecule of high molecular weight rRNAconsists of 1500 to 5000 nucleotide residues, the number of double-stranded regions in each molecule can be expected to be of the order of 100. Upon heating an rRNA solution or lowering its ionic strength, the secondary structure, i.e. the helical regions of the rRNAmolecules, melts out (90, 94, 95). Because of their heterogeneity in stability (due to differences in size, base composition and sequences, regularity of base pairing, relationship to neighbor regions, etc.), the helical regions in a single molecule do not melt out abruptly all together at a critical temperature or ionic

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strength. The ~rst helices to melt will be the most labile, while those relatively more stable will melt only at higher temperatures (lower ionic strength); the melting of a single rl~NA molecule as a whole thus takes place over a relatively wide temperature (ionic strength) interval. Any heterogeneity of the rRNAmolecules within a population may contribute to this broad melting behavior. Physical consequences of the melting of the secondary structure of the rRNAare (a) a hyperchromic effect (90, 94, 95, 102-104) and a decrease optical rotatory dispersion to almost zero (90, 94, 98), due to breaking hydrogen bonds between paired bases; and (b) a manyfold increase in specific viscosity and a fall in sedimentation velocity, due to the transition from a compact globular conformation to an unfolded configuration (94, 103). When the temperature or the ionic strength of a melted rRNAsolution are brought back to normal values, there is a reversion of the above-mentioned physical changes, indicating that the original (or a different) secondary structure has been reacqulred. From what has been said it is clear that the rRNAmolecules do not have a fixed structure in solution, but they can acquire a different structure according to the prevailing conditions. The molecules are in a dynamic equilibrium, always trying, by "lateral mobility" (91) of the various regions, to find the most stable conformation for the particular conditions. Concerning the tertiary structure of rRNA, i.e. the mutual arrangement of the helical regions within each rRNAmolecule, the positive UVdichroism of solutions of rRNAmolecules oriented by an electrical field (with their long axes presumably parallel to the direction of the field) indicates some regularity in the orientation of the helices. Interpretation of the data thus obtained has provided a model for the tertiary structure of the high molecular weight rRlq’A (g9). At very low ionic s~reugth, when the electrostatic repulsion is high, the rRNAmolecule, as discussed above, is completely melted out or almost so. As the ionic strength is increased to moderate values, there is an organization of the moleculc in short double-helical regions which will be mainly oriented perpendicularly to the long axis of the molecule. The two possible variants of the model assume (a) that the helices radiate in all directions from the long axis of the molecule, and (b) that the helices are stacked up to form a rod-shaped structure. The forces involved in this orientation of the helices are mainly the electrostatic repulsion, still high enough at moderate ionic strength, and/or metallic bridges and hydrogen bonds. A solution of oriented RNAmolecules in this configuration will have a positive UVdichroism due to the predominant orientation of the base planes parallel to the long axis of the molecules. A further increase in ionic strength will reduce the electrostatic repulsion to such an extent that the helices will be no longer held in the oriented configuration and the molecule will take the structure of a compact coil. All possible intermediate configurations can be acquired by the RNAmolecule at intermediate ionic strengths.

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What has been said above refers to purified rRNAin aqueous solutions. It is important to know, of course, what is the secondary and tertiary structure of rRNAwhen it is associated with proteins in the ribosomal subunits. Studies of hypoehromism, optical rotatory dispersion, and X-ray diffraction carried out on whole ribosomes have indicated that the rRNA structure ~vithin the ribosome is very similar to that of free rRNAin solutions of moderate ionic strength and at room temperature (98, 105-111 ). An orientation of double helices of the rRNAmolecules in ribosomes has been demonstrated by X-ray diffraction analysis of ribosomes of various origins (E. coll, Drosophila, rat, and rabbit) : the patterns observed are consistent with a model according to which part of the rRNAis in form of four or five parallel double helices 45 to 50 A apart (112). The secondary and tertiary structure of the high molecular weight rRNAdescribed above makes the rRNAmolecule nonrandomly accessible to degradative enzymes. Mild treatments with nueleases break the molecule first at the most sensitive sites, probably the unpaired regions exposed on the outside of the coil (87, 113-116). This nonrandom degradation can occur during rRNAextraction if not enough care is used to avoid RNase action (87). As a result of enzymatic attack at sensitive sites, the continuity of the molecule is broken, though the coll can be still held together by secondary bonds (hidden breaks); only upon heating do the fragments come apart (116, 117). The nonrandom degradation of rRNA under conditions of mild RNase digestion, followed by analysis of the resistant fragments by polyacrylamide gel electrophoresis, sedimentation, or melting tests, has been utilized to study the helical regions of rRIXlA (94, 99), or to detect homologies and differences between rRNAsamples from various sources (118-120). In the latter connection, note that whereas differences between two l~lq’As in the electrophoretic or sedimentation pattern of the RlXlase resistant fragments very likely reflect sequence differences, even extensive variations in primary structure are compatible with a similar secondary and tertiary structure which is responsible for the pattern of sensitivity to mild l~lX~ase digestion (120). By the method of limited RN’ase digestion evidence has been obtained which suggests that the secondary structure of rRNAhas differentiated appreciably in the course of evolution, as indicated by a general trend towards an iucrease in resistance to enzymatic attack (119, 120) : this is in agreement with the above-mentioned increase in GC content and reduction in the proportion of noncomplementary bases. However, from a comparison of the differences in RNA-DNAhybridization behavior among rRNAs from various mammalian and microbial species with the differences in the electrophoretlc pattern of the l~lXlase-reslstant fragments of the digested rRN’As, it would appear that the secondary or tertiary structure of rRNA has evolved more slowly than the nucleotide sequence (120). analysis by the method of limited RNase digestion of purified rRIXlA components from various human tissues and HeLa cells revealed no significant

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differences between them in the sedimentation pattern of the resistant fragments (45). The hydrolysis of the rRNAmolecules at a few sensitive sites under controlled conditions of enzymatic digestion will presumably be useful for the study of the nucleotide sequence of high molecular weight rRNA, as it has been for the analysis of the sequences of 4S (121) and 5S RNA(122124).

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SYNTHESIS

The process leading to the formation of mature high molecular weight rRNAboth in prokaryotic and eukaryotic cells involves a transcription step and posttranscriptional events, i.e. secondary modifications of the nascent polynucleotide chains. These secondary modifications may already start on growing chains attached to the DNAtemplate and continue on the completed chains, in parallel with their acquisition of a definitive secondary and tertiary structure and their association with ribosomal proteins. In bacteria, each species of high molecular weight rRNAappears to be synthesized and processed as an independent unit (and possibly, in the case of 23S RNA, as half-size precursors), and the maturation process involves mainly, if not exclusively, methylation of bases and ribose moieties and conversion of uridylic acid residues to pseudouridylic acid (and possibly the dimerization of 23S RNAfrom its half-size precursors). In animal cells, on the contrary, yeast cells and presumably, all other eukaryotic cells, the two species of high molecular weight rRNAderive from a unique precursor molecule, which contains the sequences of one molecule of the larger species, one molecule of the smaller species and, in addition, a nonrlbosomal portion destined to be discarded : here, therefore, the maturation process seems more complex, and includes, besides secondary modifications of nucleotides, a specific splitting of the long precursors to give final products of the appropriate size. Below, in discussing the series of events leading to the formation of mature rRNAin bacteria and animal cells, the E. coll system and, respectively, the HeLa cell system, which have been the most extensively studied in this connection, will be generally used as models. Bacterial systems.--The first systematic investigation of the synthesis of rRNAand of its association with proteins to form ribosomal subunits in E. coli was carried out by the Carnegie Group of Washington (125-129). After exposure of exponentially growing cells for different times to 14C-uracil, radioactivity was found by these workers, first in an l~lgA fraction sedimenting between 8 and 20S and peaking at 14S (eosome), then in ribonucleoprotein particles with sedimentation constants of about 30S and 43S (neosomes), and finally in mature 30S and SOS ribosomal subunits. A kinetic analysis of the flow of newly synthesized IRNAand of the apposition to RNAof newly synthesized protein led the authors to postulate that the

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eosome fraction contains precursors of both 16S and 23S RNA. Unfinished RNAchains from this pool, after completion and addition of some protein components, would become 30S neosomes, and these, after further apposition of protein, mature 30S subunits; likewise, other eosome chains, after growth to completion and addition of protein, would give rise to 43S neosomes (in part directly, in part through a 30S neosomestage), and these, further apposition of protein, would become mature 50S ribosomal subunits. Later investigations showed that a substantial portion of the eosome fraction is represented by unstable messenger RNA(mRNA)(130-132), which complicates the interpretation of this kinetic analysis. However, the main principles which emerged from these investigations, i.e. the existence of delay points, represented by pools of precursors, in the .flow of newly synthesized RNA,and the stepwise addition of protein have been confirmed by other groups of investigators. Osawa and collaborators, using shift-up cultures of E. coli, in which mRNAsynthesis is markedly reduced as compared to that of rRNA (81, 133), observed that the radioactivity after a aH-uridine pulse first appeared in 18S and 22S components, then in three classes of particles sedimenting at about 26S, 30S, and 40S, and finally in 30S subunits (distinguishable from the 30S pulse-labeled particles by the lower density in CsCI after formaldehyde fixation) and in 50S subunits (2, 134). A considerable amount of particles with sedimentation constants of 22S, 26S, 30S, and 405, indistinguishable in the nature of the rRNAcomponent and in protein composition from the above-described pulse-labeled particles, was found also in cell extracts from E. coli grown in the presence of low concentrations of chloramphenlcol (CAP) (0.6-1.5/zg/ml) (2). 22S and 26S part icles cont ained RNAsedimenting as 17S; in the tow CAP particles this had a level of methylatlon corresponding to 15-20% of that of mature 16S RNAfound in 30S subunits (2, 133). The 30S and 40S nascent and low CAP particles contained RNAsedimenting at about 23S, and with approximately 60% as many methyl groups as mature 23S RNAfound in the 50S subunits (2, 133). The protein content increased progressively, going from the 22S to the 26S particles to the 30S subunits, and from the 30S to the 40S particles to the 50S subunits (2, 135). These results suggested a sequence of events leading from the nascent rRNAto the 30S subunit via two intermediates, the 22S and 26S, and to the 50S subunits via the intermediates 30S and 40S

(2, 135). Pulse-labeled 26S particles containing "16S" RNAand 32S and 43S particles containing 23S RNAhave also been described in cultures of fragile E. coli cells, which can be lysed by very gentle procedures, thus allowing the separation of the mRNAassociated with polyribosomes from rRNAprecursors (136). These particles behaved as concerns their kinetics of labeling precursors of 30S and, respectively, 50S subunits. The 26S, 32S, and 43S precursors are presumably equivalent respectively, to the 26S, 30S, and 40S precursors detected in shift-up cultures. Similarly, the 26S and 30-32S particles probably correspond to the 30S neosomes, and the 40-43S particles to

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the 43S neosomes of the Carnegie Group. The radioactive 18S and 22S components detected after a very brief pulse in shift-up cultures were not observed in the fragile E. coli cultures. Instead, a spectrum of heterogeneous components sedimenting slower than 26S was observed (136). The labeled RNAextracted from these components sedimented slower than 16S RNA and competed in RNA-DNAhybridization experiments with purified 16S and 23S RNA.Several lines of evidence indicated that in the lysate these RNAchains sedimented faster because they were associated with proteins; these resembled ribosomal proteins in their chromatographic behavior on carboxymethylcellulose columns. These observations suggest that ribosomal proteins may become associated with rRNA chains while these are still growing on the DNAtemplate (136), thus excluding the presence of complete "free" rRNAin the cytoplasm of E. coli, at variance with earlier reports (134). However, the interpretation of these results is complicated the demonstrated tendency of free RNAto form complexes with proteins present in cell extracts (137, 138). This phenomenon may account for the discrepancy in the results reported on the sedimentation properties of the smaller precursors of ribosomal particles. The question whether the conversion from the precursor to the mature forms of 16S and 23S rRNAinvolves only addition of methyl groups and, possibly, conversion of uridylic to pseudouridylic acid, or whether, on the contrary, there is also a change in the polynucleotide size, is not yet settled. It is generally believed that the slightly higher sedimentation constant of the nascent 16S RNAand the lower affinity for methylated albumin of both nascent rRNAcomponents is related to a difference in secondary structure (139, 140), possibly due to undermethylation of the precursor forms. Changes in the secondary structure of polynucleotides have indeed been found to occur as a result of methylation (141). However, a dit~erenee electrophoretie mobility which has been detected between precursor and mature forms of rRNA, and which persists after heating and fast cooling, has been interpreted to indicate a somewhat larger size of the nascent species as compared to the mature ones [about 25% for the 16S RNAand 5-10% for the 23S RNA(142)]. The completion of methylation of the nascent rRNA appears to occur late during the conversion of the 26S and 40S nascent ribosomal particles to the 30S and 50S subunits respectively, or even after completion of the subunits (2, 135). It is possible that this late methylation indispensable for the attachment of the last ribosomal protein to the 26S and 40S precursor particles. The time required to synthesize a 16S RNAchain, as estimated by different methods in E. coli, has been found to be about 30 sec for a doubling time of 45 rain (143) and 120 see for a doubling time of 120 rain (136). B. subtilis growing with a doubling time of 60 mln, the time reported is 18 sec (144). These times correspond to rates of movement of RNApolymerase ranging from 13 to 90 nucleotides per second, which is about the same range reported for the rate of growth of various mRNAspecies (145-147). It is not yet clear to what extent the differences in the reported rate of

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rRNAchain growth are related to differences in growth rate of bacteria. It can be calculated that in cells growing in glucose-salts medium with a doubling time of 45 rain the synthesis of the complement of 16S RNA(about 20,000 molecules per genome) (148) from the known number of 16S cistrons per genome (5 to 6) (149, 150) would require approximately 16S RNAmolecules to be made per minute per gene copy: for a rate of growth of the RNAchain of about 55 nucleotides per second (147), this would imply that about 30 molecules of RNApolymerase must be working in tandem on each cistron at the same time. Considering the size of the RNApolymerase [about 100 A in average dimension (151)] and the length of a 16S RNAcistron (about 6000 A), it thus appears that about half of the length of each gent is covered by RNApolymerase molecules under the conditions considered here. In very fast growing cultures it is therefore likely that the rRNA genes are saturated with RNApolymerase molecules (148). This may suggest that the region of the DNAduplex which corresponds to these genes may be permanently "melted." In an exponentially growing culture an equal number of 16S and 23S RNAchains are synthesized per unit time. Since the 23S RNAmolecule is twice the size of the 16S molecule, if the same number of gent copies exists in a cell for both rRNAspecies, as suggested by the available evidence, either the rate of movement of RNApolymerase on a 23S cistron is twice as high as that on a 16S cistron or, more likely, there are twice as manymolecules of RNApolymerase working at the same time on a 23S cistron as compared to a 16S cistron, i.e. there is a constant number of enzyme molecules per unit length of rDNA.That a further constraint exists is suggested by the observation that in E. coli fragile cultures about 2 rain were required to synthesize an entire chain of either 16S or 23S RNA(136). This would imply that, if the rate of RIgA polymerase movement is equal for the two rRNAspecies, the enzyme molecules involved in the synthesis of 23S RNA must produce chains of the same size as 16S RNA, with each 23S molecule being then formed by covalent linkage of two 16S chains: both of these half-size precursors probably would have to be different in sequence from the 16S RNAbecause of the evidence discussed earlier concerning the difference in 3’ and Y-terminal sequences (72, 74) and in the partial sequence distribution after ribonuclease digestion between 16S and 23S RNA (79), and because of the behavior of the two speeies in hybridization competition experiments (see below). Evidence supporting the idea that 23S arises from the joining of two half-size molecules has come from the observation that after a short sI-I-uracil pulse (30 see), when all nascent chains should be labeled to about the same extent, the radioactivity in the sedimentation pattern of the RNAextracted from E. coli fragile cultures is distributed mainly in the region 4S to 16S, with relatively little between the and 23S peaks, and after longer pulses accumulates in finished 16S and 23S RNAchains (136). It is also interesting that the methylation pattern of coli 23S RNAsuggests that this species is made o{ two identical or similar halves.

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Various conditions which inhibit protein synthesis in bacteria without immediately affecting RNAsynthesis, like addition of chloramphenicol (152, 153), puromycin (153-156), streptomycin (157), chlortetracycline (158), deprivation of an essential amino acid in a culture of a RCr~ mutant of E. coli (159-162), removal of + f rom a culture of a K +- dependent mut ant (163), lead to the accumulation of particles with sedimentation coefficients about 18S and 25S. Similar particles accumulate in early phases of Mgion replenishment following a prolonged starvation for these ions (164). Particularly well investigated have been the particles induced by chloramphenlcol (at: a concentration of more than 3/zg/ml) (CAP particles). The 18S particles contain 17S RNA(139, 165-167), with a content of methyl groups eqnivalent to 15 to 20% of that of mature 16S RNA(58, 168, 169) ; the 25S CAP particles contain 23S RNA(139, 165-167) with a level of methylation equivalent to 60% of that of mature 23S RNA(58, 168, 169). Thus, from the point of view of the sedimentation properties and the level of methylation the RNAof CAPparticles [and likewise that of the RCre1 (58, 140, 170), streptomycin (58, 157), chlortetracycline particles (158), etc.] is lar to the RNAof nascent precursor particles. Also the content of pseudouridine in rRNAof 18S and 25S CAP particles is lower than that of mature rRNA(58). The CAPparticles (and the particles of similar nature) an average protein content of about 25%, with some indication of heterogeneity (152, 153) : this protein content is considerably lower than that of the 30S and 50S ribosomal subunits (about 38%) and also slightly lower than that of nascent particles (2). After removal of the block of protein synthesis, the RNAof the CAPparticles (and similarly the RNAof the other defective particles) becomes fully methylated and acquires its final sedimentati’.on characteristics (140), while it is incorporated into mature ribosomal particles (161-163, 171, 172). The reader is referred to an earlier review (2) for further details concerning the nature and fate of the CAPand equ.ivalent particles. An accumulation of 43S ribosomal precursor particles has been observed in E. coli grown in the presence of p-fluorophenylalanine (173) and in mutant strains of E. coli (174, 175). E. coli cells grown in the presence of 5fluorouracil (5-FU) accumulate 28S and 32S particles (134, 165, 176, 177), which contain respectively "16S" and 23S RNA(167) : in these there is a partial replacement (up to 70%) of uracil by the analogue, resulting in changes in their physical properties. Upon removal of 5-FU, the abnormal rRNAis not incorporated into normal subunits, but is degraded (177, 178), which suggests that the modification brought in the structure of rRNAby the analogue, rather than the presence of abnormal ribosomal proteins due to 5FU-induced miscoding, prevents its proper utilization. In agreement with this interpretation is the observation that 32S particles accumulate and no 50~¢’, subunits are formed in the presence of the analogue even under conditions where normal proteins, accumulated during a preceding phosphorus starvation step, are available (179). The idea that nascent rRNAmay function as mRNAfor the synthesis of

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ribosomal proteins, originally suggested by the observation of a preferential protein synthesis (occurring even in the presence of actinomycin D) and the labeling patterns after a radioactive amino acid pulse in a methioninerequiring RCrel mutant of E. coli recovering from methionine starvation [see Osawa (2)], has received support from several observations. Thus, extracts of the mutant where there has been accumulation of nascent rRNA as a result of the methlonlne deprivation, a considerable amount of this appears to be associated with 70S ribosomes to form "heavy 70S" and polysome-like material, and these complexes have amino acid-incorporating activity in vitro (180). Furthermore, nascent rRNAextensively purified from CAPor RCre~ particles can act as messenger in an in vitro protein-synthesizing system (181) :.the bulk of the product formed in vitro by this cell-’free system has electrophoretic mobilities similar to those of ribosomal proteins and distinct from those of the products of mRNA-directed synthesis. Mature rRNAhas no amino acid incorporation-stimulating activity in these in vitro systems (182-184) ; however, it becomes a template for amino acid incorporation, especially in the presence of neomycin, if its secondary structure is destroyed by heating (185). It is possible that the messenger activity of nascent rRNAis related to its relatively low methylation level as compared to that of mature rRNA, resulting in a more loose secondary structure. Some negative reports on the existence of messenger activity of nascent rRNAhave appeared (186, 187) ; however, the apparently heavy contamination of the preparations by mRNA,which could obscure the activity due to nascent rRNAin these experiments, makes these observations inconclusive. The idea that translation Of nascent rRNA, as is the case for mRNA,plays an indispensable role in its removal from the template, thus creating a feedback control by protein synthesis on rRNAsynthesis, has been suggested by Stent (188). The amount of information contained in the two rRNAclasses together, assuming that each is homogeneous, would be su~cient to code for only about 20% of the different species of ribosomal proteins present in the 30S and 50S subunits (189). Therefore, if nascent rRNAshould function the exclusive source o~ mRNA for ribosomal proteins~ at least five different species for each rRNAclass would be necessary. If so, the gene redundancy for rRNA(see below) would serve an indispensable informational role. Eukaryotic systems.--The nucleolar location of the synthesis of rRNA in animal cells, originally suspected on the basis of cytochemical (190-192) and autoradiographic observations (193-198), was first corroborated by the demonstration of the effect of UVmicroirradiation of the nucleolus on the synthesis of cytoplasmic RNA[which is predominantly rt~NA (199)] and by the recognition of similarity in base composition between nucleolar and cytoplasmic RNAextracted from microdissected subcellular fractions (200203). Stronger evidence was provided by the observation that treatment with low doses of actinomycin D blocks selectively the synthesis of nucleolar RNA, as detected by autoradiography, and the synthesis of the

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4SS rRNA precursor (see below) (204, 20S). In agreement with nucleolar origin of rRNAwere also elcctronmicroscopic observations concerning the presence in the nucleolus of ribonuclease-sensitive granules about 150 /~ in diameter, similar to the large ribosomal subunits (206214), and electronmicroscopic-autoradiographic evidence that the delay in the labeling of these granules after a pulse with an rRNA precursor corresponded well to the time course of processing of the newly synthesized 45S RNAto mature 28S RNA(210, 211, 213) (see below). Nucleoli were also shown to contain the enzymatic apparatus for the synthesis and methylation of RNA(215-218). More recently, a nucleolus-associated RNA polymerase with different properties from the nucleoplasmic enzyme has been isolated from the nuclei of rat liver and sea urchin embryos (219). The introduction of techniques for fractionating nuclei into nucleoli and nucleoplasm (220, 221) has made it possible to obtain unequivocal biochemical evidence that 45S synthesis occurs exclusively in nucleoli and that all 45S and 32S rRNAprecursors are located in these organelles. Direct evidence associating the genes for rRNAwith the nucleolar organizer was first provided by the demonstration that mutants of lae~;s lacking the capacity to form nucleoli do not synthesize rl~NA (222). Further evidence has come from the observation of a proportionality between the amount of rRNA genes, as detected by RNArDNA hybridization, and dosage of nucleolar organizers in various strains of Dro~ol>hila melanogaster (223, 224) and in wild-type Xenopus embryos and in Xenopus individuals homozygous and heterozygous for the anucleolate mutation (225, 226). An enrichment of rRNA genes in a crude nucleolar fraction from HeLa cells (227) and rat liver (228) has been observed. More recently, the electronmicroscopic visualization of the genes for rRNAprecursors in the extrachromosomal nucleoli of amphibian oocytes has been reported (229) (see below). The reader is referred to a review by Perry (230) for further information concerning the role of the nucleolus in the synthesis of rRNA and assembly of ribosomes. The biochemical analysis of the synthesis of rRNAstarted with the discovery, made in HeLaand L cells, that a large-size molecule, with a sedimentation constant of about 45S, is the precursor of mature rRNA(204, 205, 2~1-2~7). This component can be shown as a peak emerging over a background of heterogeneous RNAin the sedimentation pattern of total RNApreparations from cells subjected to ~H-uridine or ~C-uridine pulses as short as 3-5 mln (236); with increasing pulse length, the 45S peak becomes more prominent, and after about 25 min radioactivity appears in a 32S component and in 18S RNA; after a still longer pulse 28S RNA comes labeled, as shown by the broadening o{ the 32S profile to overlap the 28S UVabsorbance peak (220, 2~7). The use of actinomycin D, after a short pulse with an RNAprecursor, to block further RNAsynthesis, causes the disappearance of the 45S component accompanied by appearance of label in the 32S and 18S components and later in 28S RNA(232). Furthermore, the

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base composition of 45S and 32S components labeled with s2P-orthophosphate resembles that of rRNA(232). These early observations led to the view that the 45S RNAis destined to give rise by cleavage to 18S and 32S RNA, the latter being then converted to 28S RNA.In agreement with the idea that 45S RNAis an rRNAprecursor were various observations, which indicated that under a variety of conditions in which there is a decreased synthesis of this component,as in anueleolate mutants of Xenopus(222), in the early stages of embryogenesis(238-240), or as a result of selective inhibition with low doses of actlnomycin D (204, 205), there is also a lack of synthesis of rRNA. The developmentof techniques for fractionating cells into cytoplasmic, nucleoplasmie, and nucleolar fractions (220, 221) has made it possible follow the topography of the above-described conversions of 45S RNA. Thus, the 18S RNAarising from cleavage of 45S RNAis rapidly transferred to the cytoplasm(about 25 rain after the beginning of incorporation of radioactive precursor), whereas the 32S RNAremains in the nucleolus and is transformed into 28S RNA,which is then found in the nueleoplasm and later (about 60 rain after the beginning of labeling) in the cytoplasm (220, 237). Furthermore, 50S and 30S ribonueleoprotein particles identical to the cytoplasmic subribosomal particles as concerns their RNAcomponent and behaving kinetieally as their precursors have been found in the nueleoplasm(241). The nuclear pool of 50S particles is less than one-sixth that the larger cytoplasmic ribosomal subunits (60S), whereas the nuclear pool of 30S particles is less than 1%of the pool of 45S cytoplasmicsubunits. The 50S particles remainin the nucleus for about 20 rain, whereasthe 30S particles remain only a few minutes. It is not knownwhether these nuclear precursors of cytoplasmic subribosomalparticles are present in the nueleoplasm in vivo or whether they are extracted from the nucleoll by the high ionic strength-deoxyribonuelease treatment used in the fractionation procedure. The latter alternative is suggested by the above-mentioned eleetronmieroseopie evidence of the existence of particles of the size of the larger ribosomalsubunits in the nueleolus (206-214). Simultaneously or immediately following their synthesis the 45S molecules undergochemical modifications through the activity of methylating enzymes (69, 236). Almost all the methylation of rRNAoccurs at this stage. The pattern of ribose methylation of the 45S RNAcorresponds to that of an equimolar mixture of 28S and 18S 1RNA(84) ; likewise, the pattern 2’-O-methylation of the 32S RNAis similar to that of 28S RNA(84). The yield of methylated bases of 32S RNAappears also to be equivalent to that of 28S RNA.By contrast, the amount of methylated bases in 45S RNAis significantly lower (to the extent of about two methyl groups per molecule) than that expected for the sum of the two rRNAcomponents (84). agreement with this finding is the observation that the 45S RNAlacks the 6-dimethyladenine residue present in 18S RNA(69). Evidence for a secondary methylation of the 45S RNAoccurring in the nueleolus at the time of the precursor cleavage and leading to the formation of dimethyladenine in

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18S RNAhas been presented (242). The amount of pseudouridylic acid 45S RNA(88) suggests that the conversion of uridylic to pseudouridylic acid in rRNAoccurs exclusively or in its majority at the stage of the 45S precursor, possibly while the polynucleotide chain is still being synthesized. The observation that methyl-labeled methionine selectively labels among the high molecular weight RNAspecies the rRNA precursors has made it possible to use this labeling to follow the conversion of the precursors to mature rRNAby a chase experiment (236). The results obtained have confirmed the time sequence of events and the location of the intermediates found in previous experiments utilizing continuous uridine incorporation. By using pulse labeling with ~4C-methylmcthionine, it has been estimated that the synthesis of the 45S molecule takes place in 2.3 rain if all methylation occurs near the growing point, or in a longer time if some methylation takes place on complete 45S molecules (236). Thus, the rate of chain growth of this RNAspecies in animal cells appears to be similar to that of rRNAin E. coli. Recent electronmicroscopic evidence obtained on rRNA precursor sites in DNAfrom amphibian oocytes (229) supports the idea that numerous rRNA precursor molecules are being synthesized simultaneously on each site as in bacteria. By utilizing isolated nucleoli as a source of rRNA precursors from HeLa cells it has been possible to obtain RNApreparations consisting almost exclusively of these components (220, 221). The UVabsorbance sedimentation profile in sucrose gradient of nucleolar RNAhas revealed two main components corresponding to 45S and 32S RNA; in the better resolved patterns a 28S RNAcomponent is recognizable as a shoulder or a partially resolved peak (220, 221), and there is an indication of UVabsorbing material between 45S and 32S RNA(243). The introduction of polyacrylamide gel electrophoresis for the analysis of nucleolar RNA,besides allowing a better resolution of 28S RNAand the recognition of a small amount of 18S nucleolar RNA, has revealed the presence of small peaks corresponding to RNAspecies with tentatively assigned sedimentation constants of 41S, 36S, 24S, and 20S (243, 244). The analysis of the kinetics of labeling of nucleolar RNAduring short pulses of 14C-methylmethionine and the results of pulse-chase experiments utilizing actlnomycin D have indicated that and 36S RNAbecome labeled after 45S RNAand, in the presence of actinomyein D, decay after the disappearance of the precursor, concomitantly with the transfer of radioactivity to the 32S peak (243). Nueleolar 28S RNAin these experiments is the first 28S species to be labeled; furthermore, 20S becomes labeled before 18S. 41S RNAhas about the same number of methyl groups as 45S RNA, and 24S and 20S about the same number of methyl groups as 18S RNA(244). These results have been interpreted indicate that 41S and possibly 36S RNAare precursors of 32S RNA, and 24S and 20S RNAare precursors of 18S RNA(see below). Poliovirus infection interferes with the normal processing of nucleolar RNAand leads to accumulation of 41S, 28S, 20S, and 18S RNA(243). The sedimentation properties of 45S RNAand its electrophoretic mobil-

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ity in polyaerylamide gel indicated that each molecule of this species is longer than the sum of one 28S and one 18S molecule; likewise, the sedimentation and eleetrophoretie behavior of 32S RNAsuggested that this molecule is somewhat larger than 28S RNA(243). The relationship between 45S and 32S rRNA precursors and the mature rRNA species was clarified when it was shown that 40 to 50% of the 45S molecule and 20 to 30% of the 32S molecule are represented by sequences of nonribosomal type, whleh are not conserved in the conversion of the precursors to mature rRNA. Evidence for this nonconservative process has come from various types of observations : 1. Molecular weight determinations by sedimentation equilibrium have indicated that the molecular weight of 45S RNA(~4.4 × 10~ daltons) exceeds by about 106 daltons the sum of the molecular ~veights of one 32S molecule (~2.4 × 10n daltons) and one 18S molecule (~0.7 × 106 daltons); likewise, the molecular weight of 32S RNAis larger than that o~ 28S RNA (~1.9 × 10~ daltons) (31). 2. Accurate base composition analyses of purified rRNAprecursors and mature rl~NA components have revealed dlserepaneies between 45S t~NA and the weighted average of 28S and 18S RNA, and between 32S and 28S RNA(88, 245, 246). Furthermore, oligonueleotide mapping after pancreatic ribonuclease digestion has sho~vn that an "average" 45S molecule contains at most the sequences of one 28S molecule and one 18S molecule (88, 246), this implying that about 50% of its length is not conserved; the same type of analysis has indicated that about 30% of the 32S RNAmolecule is represented by nonrlbosomal sequences (88, 246). 3. The specific methylation (i.e. the ratio of l~C-methyl to pyrimidines or total bases) in 45S RNAhas been found to be about one-half that expected for a weighted average of 28S and 18S RNA(243, 244) ; similarly, the specific methylation of 32S RNAis 20 to 30% lower than that of 28S RNA (243, 244). On the basis of the evidence suggesting that almost all methylation occurs at the 45S RNAstage (69, 236), and that the methyl groups 45S RNAare conserved in the processing to 28S and 18S RNA(84), the above-mentioned data indicate the loss of about one-half of the 45S precursor and of 20 to 30% of the 32S intermediate in the maturation process. Similar results have been obtained by analyzing the content in various rRNAspecies of alkali-resistant dinucleotides, which provide a measure of the 2’-O-methylation of the ribose moieties (66). 4. RNA-DNA hybridization e~periments have indicated that the ~raction of DNAcomplementary to 18S, 28S, and 45S RNAis proportional to the molecular weight of these species (247). Furthermore, 45S RNAcompeted with 28S and 18S RNAfor sites in DNAto the extent expected if 35% and 13%, respectively, of the precursor molecule was involved; similarly, 32S RNAcompeted with 28S RNA to the extent expected if about 70% of the molecule was involved (247). These observations have provided unequivocal evidence for the noncon-

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STRUCTURE AND SYNTHESIS

OF RIBOSOMAL RNA

205

servative processing of the 45S RNAprecursor and of the 32S intermediate. Furthermore, they support the view, originally proposed on the basis of kinetic arguments, that the 45S RNAis a precursor of both 28S and 18S RNA.This view has also been corroborated by the observation that the 2’O-methylation pattern of 45S RNAcorresponds to that of an equimolar mixture of 28S and 18S RNA(84). As concerns the distribution of 28S and 18S polynucleotide stretches among the 45S molecules, RNA-DNA hybridization experiments have indicated that the 28S and 18S genes in Xenopus and Drosophila alternate along the ribosomal DNA(rDNA) and are interspersed with stretches of DNAof higher GC content, in such a way that a large {ractlon of single-stranded rDNA chains of an average molecular weight of 3 × 10’ daltons, i.e. considerably shorter than the 45S RNAprecursor, contain sequences homologous to both 28S and 18S RNA(41, 248, 249). These results have, therefore, provided conclusive evidence that the 28S and 18S sequences are contained in the same precursor molecule. The nonribosomal portion of both the 45S precursor and the 32S intermediate is characterized by very high GC content (about 77%) and low A content (7--8%) (88, 245). The analysis of the relative methylation level and pattern in the ribosomal RNAprecursors and mature ribosomal RNA suggests that the nonribosomal portion has no or few methyl groups (84, 244). On the contrary, it contains pseudouridyllc acid, although in only about one-third the proportion as found in the ribosomal portion of the precursor (88). The oligonucleotide pattern after RNase digestion of the nonribosomal portion of 32S RNAis very similar to that of the nonribosomal portion of the other half ot the 45S molecule (88): this observation, together with the very high GCcontent of these stretches and the capacity of 32S RNA to compete with 45S RNA in RNA-DNAhybridization to a greater extent than expected {tom the molecular weight ratio, suggests that the nonribosomal portion of the 45S precursor may contain repetitive sequences (247). This would speak against a messenger role for this nonribosomal portion, and would rather favor the view that it may have a transient structural role--that of creating, through formation of an appropriate secondary structure, the precise configuration which allows the proper interactions between the RNAprecursor and modifying enzymes or specific cleaving enzymes or structural ribosomal proteins (88). The possibility of this nonribosomal portion of the precursor functioning as mRNAfor ribosomal proteins seems also to be excluded by its base composition, which is drastically different (especially in the very low A content) from that expected for proteins with amino acid compositions such as those found in ribosomal proteins of different (including mammalian) sources (250, 251). Whatsoever the role of these nonribosomal stretches may be, they are ultimately destined to be rapidly destroyed, because there is no evidence that they accumulate in the cell. As concerns the minor nucleolar RNAspecies, from the methylation level and the estimated molecular weight it would appear that the 41S RNA’

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derives from the 45S RNAby the loss of a nonribosomal stretch of about 106 daltons. The 24S and 20S components appear to be precursors of 18S RNAcontaining a nonribosomal stretch of about 7.5 × 105 and 3 × 105 daltons, respectively (244) ; less certain is the significance of the 36S component (244). Further experiments are needed to establish conclusively the relationship of these minor species with the 32S, 28S, and 18S RNAcomponents, and the significance of their presence as related to a normal or abnormal processing of 45S RNA.These experiments should also clarify the precise anatomy of the 45S molecule, that is the relative arrangement of the 28S and 18S stretches and of the nonribosomal stretch(es). After the 45S precursor molecule has been synthesized or, possibly, while it is being assembled, it becomes associated with protein. Two types of nascent ribonucleoprotein particles have been isolated from nucleoli of HeLa cells, with sedimentation coefficients of about 80S and SSS in the presence of EDTA(252). The 80S particles contain 45S RNAand some 32S RNA, the 55S particles 32S RNA, a small amount of 28S RNAand 5S RNA; furthermore, the proteins of the 55S particles show in polyacrylamide gel electrophoresis a pattern strikingly similar to that of the 50S cytoplasmic particles, and behave in pulse-chase experiments as their precursors. The absence of 7S RNA(see below) shows that the 55S nucleolar nascent ribonucleoprotein particles are distinct from the SOSnucleoplasmic particles and are presumably their immediate precursors ; the 80S particles, on the other hand, are precursors of the 55S nascent particles. Similar nucleolar ribonucleoprotein particles containing 45S or 32S RNAhave been described also in L cells (253-255) and in amphibian oocytes (256). Isolated nncleoli are capable of carrying out some of the initial steps in the processing of 45S RNA(257, 258). In particular, by using nucleoli isolated from cells prelabeled with nridine and methionine, the formation in vitro of 41S and 32S RNAspecies having the same methylafion level as the in vivo counterparts and of an unmethylated fragment with a sedimentation constant of about 26S, has been described (257). This represents a promising approach for the biochemical dissection of the maturation process of 45S RNA. . Considerable evidence indicates that 5S RNAdoes not derive from 45S RNAand is not even synthesized in the nucleoli (see below). On the contrary, the 7S RNA,a small molecule about 150 nucleotides long, which is not covalently linked to 28S RN’A and which can be released from it by treatments which disrupt hydrogen bonding (urea, heat, dimethylsulfoxide), arises during the conversion of 32S RNAto 28S RNA(3, 4) ; its presence can be already recognized in the 28S RNAextracted from the nucleoplasmic 50S particles (3), but not in the nucleolar 55S particles (252). The existence of large precursors of rRNAis not exclusive to mammalian cells. Evidence has been presented for the occurrence in amphibian oocytes of a 40S rRNA precursor equivalent to the 45S RNAof mammalian cells, and of a 30S and a 20S species which are intermediate precursors to

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STRUCTURE AND SYNTHESIS

OF RIBOSOMAL RNA

207

28S and 18S RNA,respectively (256, 259). Likewise, a 38S precursor and 30S and 20S intermediates have been isolated from nucleoli of cells from insects (Chironomu~ tentans) (260), and a 45S precursor and 40S and 35S intermediates have been identified in Saccharomyces carlsbergensis (261). Recently, it has been possible to observe in the electronmicroscope the genes coding for the 40S rRNAprecursors in the extrachromosomal nucleoll of amphibian oocytes (229). Visualization of the individual genes in the DNase-sensitive fiber of the nucleolar core has been made possible by the existence of a halo of RNase- and protease-sensitive fibrils emanating from each gene and regularly increasing in length from one end of the unit to the other: these fibrils presumably represent growing rRNAprecursor molecules coated with protein. From the number of fibriIs per gene (about I00) and from the dimensions of the RNApolymerase molecule (151) it was estimated that about one-third of each gene was covered with polymerase molecules. Individual genes were separated by stretches of DNAthat apparently were not transcribed at the time of synthesis of the rRNAprecursor. Base analogues and inhlbltors have been extensively applied to the study of the synthesis and processing of rRNAprecursors. An analogue of guanine [8-azaguanine (205, 234)] and two analogues of adenosine [toyocamycln (262), tubercidin (262)], which are incorporated into the polynucleotlde chain, do not affect the synthesis of 45S RNA;they interrupt, however, its processing at various stages, which indicates that the presence of proper bases is essential for the normal cleavage and secondary modifications of the precursor to occur. Another analogue of adenosine (cordicepin) causes premature termination of 45S RNA(263): the prematurely terminated molecules do not give rise to 32S RNA,but produce appreciable amounts of 18S RNA,suggesting that the 18S portion of the 45S precursor is synthesized first. Thloacetamlde also blocks some step in the maturation process, leading to accumulation of 45S and 32S RNAin the nucleoli (264, 265). Actinomyc{n, which blocks RNAsynthesis, does not affect the conversion of 45S RNAsynthesized prior to drug treatment into 28S and 18S RNA; however, the transfer of the mature species, especially of 28S RNA,to the cytoplasm is greatly reduced (266, 267). Inhibition of protein synthesis by cycloheximide reduces the rate of 45S RNAsynthesis in HeLa cells (267, 268), slowing down its processing at the same time so that the RNAcontent of the nucleolus remains relatively constant for at least 1 hr (268). 32S RNAcontinues to be formed and converted to 28S RNA, and the latter to be exported to the cytoplasm for several h0i~rs, though at a greatly reduced rate (267, 268) ; by contrast, no, or very little, 18S RNAappears either in the nucleoll, nucleoplasm or cytoplasm, which suggests a rapid degradation of this species after its formation (268). Similar effects of eyclohexlmide have been described in L cells (269). Also in the presence of puromycln, HeLa and L cells, 45S RNAcontinues to be synthesized and methylated and to undergo the initial processing to 32S RNAand 18S RNA; no, or very

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little, mature rRNA, however, emerges from the nucleolus, which suggests that 18S and 32S (or more probably 28S) are rapidly degraded (267, 270). The pattern of ribose and base methylatlon of the rRNAprecursors synthesized in the presence of puromycin in L cells appears to be substantially identical to that of the rRNAsynthesized under normal conditions, except for the substantially lower content of 6-dimethyladenine (271) : the latter observation confirms the result, suggesting that this methylated base is formed late in the maturation process (69, 242). That the 45S RNAmade in the presence of puromycin is normal is indicated by its ability to be converted to mature rRNA once the drug is removed (270). The results obtained with cycloheximide indicate that cessation of protein synthesis cannot account by itself for the complete block of the transfer of mature rRN’Ato the cytoplasm in the presence of puromycin. Some evidence that prematurely terminated polypeptldes resulting from puromycln action may be involved here has been presented (270). Under conditions of methlonine starvation, undermethylated 45S is formed and this gives rise by cleavage to undermethylated 32S RN’A; no or very little mature rRNAappears in the cytoplasm or in the nuclear subribosomal particles, this finding pointing again to a rapid degradation of 18S RNAand to a block in the processing of 32S RNA (272). Upon restoration of methionine, the undermethylated 45~ RNAcompletes its methylation and, at least in part, undergoes normal processing (272). The results discussed above suggest that complete methylation of the 45S RNAis not essential for the initial steps of maturation leading to 32S RNA; however, it is required for the completion of the process. That the block in the rRNAmaturation in these experiments is not due to the lack of a growth-essential amino acid, independently of its role as a methyl donor, is suggested by the analysis of the effects of deprivation of valine and other essential amino acids (273). In fact, under these conditions, although there is a considerable decrease in the rate of synthesis of 45S RNAand a grossly parallel decrease in the rate of its conversion to 32S RNA,the appearance of the two mature rRNAspecies in the cytoplasm continues for many hours at a reduced rate: here, presumably, the necessary supply of the missing amino acid derives from protein turnover. The observations discussed above on the effects of base analogues, or inhibitors of protein synthesis or methionine starvation on the maturation of the rRNAhave clearly indicated that whenever rRNAprecursors or mature rRNAcannot be utilized for the assembly of normal ribosomal particles because of the presence of abnormal bases or of deficient methylation or of the lack of essential proteins, they are rapidly degraded. Besides this rcgulation by degradation of nonutilizable products, animal cells also must have a way of regulating the rate of synthesis and processing of the 45S precursors. That some proteins may play a role in this control is suggested by the observations made on the effects of inhibition of protein synthesis by cyclohexlmide (268) and of valine starvation (273).

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AN.D SYNTHESIS

OF RIBOSOMAL

RNA

209

5S BIBOSOMAL ttNA

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The major ribosomal subunits from all bacterial (274-278), animal (277-284), plant (285, 286), and lower eukaryotic cells (277, 287) thus far examined, contain a low molecular weight rRNA component with a sedimentation coefficient of about 5S. In the two cell types where its complete sequence has been determined, E. coli (122, 123) and KB cells (a human line) (124), the 5S RNAmolecule has been shown to be of identical length, 120 nucleotide residues. TABLE3. Nucleotide composition of 5S ribosomal RNAfrom some representative organisms Moles

GC%

Organism Escherichiacoli Saccharomyces cerevisiae Blastocladiella emerson~ Pea Rat (liver) Man(Kb cells) Man(HeLa cells)

C

A

U

G

30.0 23.8 24.6 25.0 27.1 27.5 28.7

19.2 24.1 19.2 21.2 18.1 18.4 18.8

16.7 a 24.7 b23.7 22.0 21.6 21.6 22.9

34.1 27.4 32.5 31.8 33.2 32.5 29.7

64.1 51.2 57.1 56.8 60.3 60.0 58.4

Reference

122 287 289 290 281 124 292

Includes 1.3%pseudouridylic acid. Includes ,-~0.5% pseudouridylic acid. STRUCTURE

Primary structure.--The base composition of 5S RNA, although determined only for a limited number of organisms, does not appear to be correlated with that of the high molecular weight rRNA(Table 3). This composition has been found to be of high GC type in 5S RNAfrom E. coli (288, 289), B. emersonii (289), plant (290), and mammalian cells [rat (281), mouse (291), and man (124, 292)] ; only in 5S RNAfrom yeast (287) somewhat different base composition has been reported, with the four bases in rather similar proportions. A property of 5S RNAwhich distinguishes it from the high molecular weight rRNAis the apparently complete absence of methylated nucleotides and, in most eases examined, also of pseudourldylic acid (276, 279-281, 288-292). In E. coli 5S RNAvarious sequences are repeated twice in the molecule, and the whole molecule can be divided into two halves which display a considerable similarity (122, 123). This has been interpreted to reflect the origin of the present 5S RNAgene by duplication of a smaller gene. Alternatively, one can think that a certain degree of symmetry is required for the

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function of the molecule (two binding sites ?). The two E. coli 5S RNAterminal sequences complement each other and are probably involved in base pairing; in the molecule there are also other regions with complementary sequences which can base pair. The 5"-terminal nueleotide is uridylic acid, the 3’-terminal nueleoside, nridine. It is interesting that two forms of 5S RNA,differing by one nueleotide in only one position, have been found in an E. coli strain in about equal amounts; in another strain, two alternative nueleotides have been found in another position in the molecule (122, 123). KB cell 5S RNA,which has a base composition very similar to that of E. coli 5S RNA,has a different nucleotide sequence (124), with only a limited amount of homology to the E. coli counterpart. Like the latter, it has complementary terminal sequences, probably involved in base pairing; other regions of the molecule are also complementary, but they are not located in the same. positions as those in E. coli 5S RNA. In KB cell 5S RNAthere are also some sequences repeated twice, but these are close to one another in the molecule and not in the two halves, as in E. coll 5S RNA.In KB cell 5S RNApGp- has been detected at the 5r-end, although only in half-molar yield; at the 3’-end, two alternate sequences, -pCpUpU and -pCpUpUpU, have been found, each in nearly half-molar amount. A partial sequence analysis of t-IeLa cell 5S RNAhas given results which are in general agreement with those reported for KB cell 5S RNA (292). However, among the pancreatic RNase digestion products of HeLa 5S RNAa trinucleotide (ApGpCp) was found which was absent in the 5S RNAdigest. This finding and the fact that several of the larger oligonucleotides released by RNase both from HeLa and KB cell 5S RNAwere reproducibly found in a much lower than expected.molar yield suggest the occurrence of alternate 5S sequences in human cells. Another interesting finding in I-IeLa 5S RNAis the presence at the 5’ terminus, in addition to pGp-, of ppGp- and pppGp-, which represent the major portion of the 5’ end (292). This is the first instance of the presence of a nucleoside di- or triphosphate at the 5~ terminus of a naturally occurring RNA,other than viral. Secondary structure.--On the basis of the sequences which were particularly resistant to RNases, a model in the form of a ringlike structure containing three short helical regions, therefore with substantially less basepairing than tRNA, has been proposed for E. coli 5S RNA(122, 123). However, analysis of the conformation of this RNAspecies in aqueous solution by several optical techniques has indicated a considerable degree of helical structure, with 60 to 80%of the nucleotide residues being involved in base pairing (293-295). On the basis of these measurements and of theoretical considerations, several models have been suggested for the secondary structure of both E. coli and KB cell 5S RNA, which resemble the cloverleaf model of tRNA (293-296).

RNA-DNA hybridization

experiments

have shown that in B. subtilis

the

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STRUCTURE AND SYNTHESIS

OF RIBOSOMAL RNA

211

cistrons for 5S, 16S, and 23S RNAare located in two regions of the chromosome: 60 to 80% of the rRNAclstrons, as judged from their time of replication in germinating spores, are interspersed with one another in a proximal region with respect to the replication origin (297, 298), and the rest is located in the last quarter of the chromosome(297). It is not known, however, whether the 5S RNAcistrons are transcribed as independent units with respect to the 16S and 23S RNA. From their relative amount in the cell and their identical labeling kinetics it has been estimated that an equal number of 5S, 16S, and 23S RNAmolecules are synthesized in E. coli in a given period of time (299). The observation that there are twice as many loci for each of the two high molecular weight rRNAcomponents as for 5S RNAin B. subtilis chromosome (297) would speak against the existence a commonprecursor including the sequences of one 16S or 23S RNAmolecule and one 5S RNAmolecule. The occurrence of a precursor of 5S RNA in B. subtilis has, however, been suggested by the kinetics of incorporation of radioisotopes into 5S RNAand by the fact that some 5S RNA~[ormation can occur in the apparent absence of transcription (300). The existence or pool of 5S RNAor a precursor thereof in E. coli is also in agreement with the lag of appearance of labeled 5S RNArelative to labeled 23S RNAin newly formed 50S ribosomal subunits (301). During inhibition of protein synthesis (by chloramphenicol or puromycin or by amino acid starvation of an auxotrophic RCre1 strain), several 5S RNA-like low molecular weight RNA’s accumulate in the E. coli high-speed supernatant fraction (302). These RNAspecies have the same nucleotide sequence as 5S RNAbut have one or a few more nucleotides (either pU or pUpUor pUpUpUor pApUpUpU) at the p end. T he s ame l abeled s pecies h ave b een i solated f rom cells subjected to a short ~2P-orthophosphate pulse (302). They are presumably, therefore, precursors of 5S RNA, either independently transcribed or derived by sequential nucleolytic attack from the longer sequence terminating with pApUpUpU at the 5" end. The fact that they accumulate in cells which are not able to form complete ribosomes suggests that the clipping of the precursors to the final 120 nucleotide length probably occurs after their incorporation into the nascent ribosomal particles. The 40-435 precursors of the 50S ribosomal subunits contain little 5S RNA(135, 301) ; however, evidence suggests that these particles may contain 5S RNAin a loosely bound form which is easily lost during the purification, especially at low Mgion concentrations (302). The 43S-associated 5S RNAis identical in sequence to mature 5S RNA. These results suggest that the 5S RNAprecursor is inserted into the ribosomal structure at or close to the 43S stage (303), and that shortly afterwards it is reduced to the length of mature 5S RNAwhile its binding to the subunlt becomes stabilized by the addition of the last ribosomal proteins. As concerns the synthesis of 5S RNAin eukaryotlc cells, the earlier suggestion that this RNAspecies derives from the 45S precursor of the high molecular weight rRNA(230) has not been corroborated by the available evidence. Thus, it has been observed that anucleolate mutants of Xeno-

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& AMALDI

pus laevis, thoughlacking almostcompletely thegenesfor thehighmolecular weight rRNA, contain a normal complement of DNAcomplementary to 5S RNA(304); furthermore, the amplification of the genes for 18S and 28S RNA, which has been shown to take place during oogenesis in amphibians (305, 306), is not accompanied by an increase in the number genes for 5S RNA(305). To the same conclusion has led the observation that the DNAcomplementary to 5S I~NA in Xenopus has a different buoyant density from the DNAwhich contains the genes for 18S and 28S RNA (304). In HeLa cells, the bulk of the genes for 5S RNAhas been found in chromosomes different from those which carry the genes for 18S and 28S RNA(307). In agreement with a separate synthesis of 5S RNAand 45S RNAare other observations also made in HeLa cells. In these cells, the newly synthesized 5S RNAappears in the cytoplasm after the newly formed 285 RNA: this suggests the occurrence of a nuclear pool of 5S l~lklA amounting to 20 to 30%of the total cellular 5S RNA,i.e. the existence of a 20 to 30% molar excess of 5S RNA relative to 28S RNA (283). The above-mentioned existence of ppGp and pppGp at the 5’-end of the major part of the 5S molecules in HeLa cells (292) would restrict any possibility of derivation of 5S RNAfrom the 45S rRNA precursor to one copy per molecule, in correspondence with the 5"-terminal segment: this result, being incompatible with the large excess of 5S sites over 45S sites in HeLa cell DNA(308), brings further evidence against the existence of any relationship between 45S RNAand 5S RNA. The presence of 5"-di- and triphosphate groups in 5S molecules also indicates that 5S RNAin HeLa cells, and presumably other animal cells, does not derive from a precursor containing an extra sequence at its 5’ end as in E. coli; furthermore, it implies that transcription of 5S RNAfrom the multiple 5S cistrons on the DNAmust occur in the form of discrete units, and not as polycistronie RNAchains secondarily cut into 5S size. Although the 5S cistrons have a different location from the 18S and 28S ¢istrons in the eukaryotic genome, the synthesis of these three rRNAspecies is coordinated. This is most strikingly exemplified by the absence of synthesis of 5S RNAin anucleolate Xenopus embryos which do have 5S RNAgenes (304), and by the coordinately accelerated synthesis o{ all three RNAspecies in amphibian oocytes, where no amplification of the 5S genes is detectable (305). It is possible that the higher degree of redundancy of genetic information for 5S RNA, as compared to that for 18S and 28S RNA(see below), reflects the need for extra genes during oogenesis, so that a transcriptional amplification may match the transient gene dosage amplification for the high molecular weight rRNAcomponents (305). On the other hand, the nonequivalence of gene dosage for RNAand the high molecular weight rRNA components and the different location in the genome could be related to some other function of the 5S species besides that of a structural RNAcomponent of ribosomes.

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LOCATION,

OF RIBOSOMAL RNA

REDUNDANCY, AND VARIABILITY BIBOSOMAL BNA GENES

213

OF

The use of the technique of RNA-DNA hybridization has made it possible to identify and titrate the rRNAgenes both in bacteria and in eukaryotic cells. The following information has emerged from this type of investigation. 1. Distinct genes exist for the two high molecular weight rRNA components, both in the prokaryotic and the eukaryotic genome (52, 224, 227, 247, 249, 309-312), in agreement with the differences in nucleotide composition and base sequence detected between the two RNAspecies. Whenever highly purified rRNAcomponents have been used, the hybridization assay has, in TABLE.4. Numberof DNAsites for ribosomal RNAcomponents and 48 RNA in various organisms (per haploid genome)as estimated by RNA-DNA hybridization

Organism

High molecular weight: rRNA

Escherichia¢oli Bacillus sublilis Saccharomycescere~islae Neuros#ora ¢rassa cyt: Pea cyt Tobacco Drosophilamelanogaster Xenopuslae~is Chick HeLa cells

5-6 9-10 140 125 4500 3450 130 450 100 280

5S rRNA 4S tRNA

11 4-5

50 42 320-400

>25,000

860 1150

2000

1260

Size of haploid genome (daltons)

References

D 2.8X10 °3.9 XlO ~0 1.25 XI0 m 2.4X10 xs 3)<10

149, 150, 313, 311, 314 297,315 312 52, 316 310, 316 3X101~ 316, 317 1.2)<1011 224, 318 TM 304 1.8Xl0 tt X10 224 7.2 ts 3.1 )
general, proven to be able to discriminate clearly between the DNAsites for the two species. The partial cross-hybridization observed between 16 and 23S E. coll rRNA(136, 1S0) may reflect the evolutionary relationship between the genes for the two RNAspecies. 2. Multiple copies of rRNAgenes occur both in bacterial and in eukaryotic cells. The redundancy of information, which is the same for the two rRNAspecies, appears to be ~5-10 copies per genome in the bacteria examined (149, 150, 297, 298, 313), ~100 to several hundred copies per haploid genome in lower eukaryotie cells (52, 312) and animal cells (224-228, 247, 304), and of one order of magnitude greater in higher plants (310, 317) (Table 4). In the eukaryotie cells there appears to be no direct correlation between size of the haploid genome and number of ribosomal genes. It is interesting that the ratio of 4S RNAsites to rRNAsites is fairly constant in the genomes of various organisms, the reported values varying between 2.5 and 10 (308). The relative constancy in the comparative amount genes complementary to transfer RNA(tRNA) and rl~NA in the whole evolutionary scale may reflect the quantitative requirement for sites of tran-

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Annual Reviews www.annualreviews.org/aronline ATTARDI & AMALDI 214 scription to keep the proportion of these two componentsof the protein-synthesizing machineryinvariant for maximum efficiency of protein synthesis. As concerns 5S RNAgenes, the available information concerning their amountis still too scarce to allow any generalization. However,it wouldappear that, whereas in bacteria the redundancy of information for 5S RNA is similar to that for high molecular weight rRNA,in animal cells it is one or several orders of magnitudehigher (304, 307, 308). The reason for this difference is not known,although, as mentionedearlier, it mayreflect the need for a permanentreservoir of genes to be called upon during oogenesis to match the selective replication of the genes for high molecular weight rRNA. 3. RibosomalRN~I genes are clustered both in the prokaryotic and the eukaryotic 9enome.By using sequentially replicated segments of the genome of synchronized E. coli cells or germinating B. subtilis spores, it has been possible to localize by RNA-DNA hybridization the genes for rRNAin these species. Thus, in B. subtili,, as mentionedearlier, 60 to 80%of the genes for 5S, 16S, and 23S rRNA,as well as of the genes for tRNA,have been found interspersed with one another in the region of the chromosome close to the origin of replleatlon (297, 298), wherethe streptomycin resistance locus is, while the rest is localized in the last quarter of the chromosome. The observation that the same fragments of single-stranded DNA with an average molecular weight of 1.9 X 10~ which hybridize with 23S RNAalso hybridize with 16S RNAindicates that the cistrons for the two rRNAgenes are situated very close and possibly adjacent to one another (319). In E. coli also, two separate regions of the chromosomeappear to be complementary to 16S and 23S rRNAas well as to tRNA, and one of these regions is very close to the streptomycin resistance locus (320). Additional information on the location of rRNAgenes in E. coli has comefrom the analysis of the pattern of synthesis of rRNAin synchronized cultures. In fact, although the segments of the genomecontaining the rRNAgenes in this organism appear to be continuously transcribed, an enhancementof the transcription activity of the two DNAregions homologous to rRNAhas been observed at about the time these regions are replicated (320). Earlier observations had indicated the occurrence of two bursts of synthesis of rRNAat two different stages of the chromosome replication cycle in two E. coli strains with different chromosomalpolarity (321): a correlation these findings with the genetic mapindicated that one of these bursts in both strains corresponded to the streptomycin locus. It is likely that this enhancementof rRNAsynthesis at the time of rDNAreplication reflects the exposition of single-stranded DNAor some other physical property of replicating DNA. In eukaryotic cells, as was mentioned earlier, RNA-DNA hybridization studies utilizing DNAfrom mutants of Drosophila (223, 224) and Xenopus (222, 225, 226) or DNAfrom isolated nucleoli (227, 228) or fractionated chromosomesof HeLacells (322) have pointed to the nucleolar organizer

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(i.e. the segment of DNAassociated with the nucleolus) as the site rRNAgenes. The available evidence indicates that these genes are closely clustered in the nucleolar organizer (41,223, 225, 229), although they do not appear to be adjacent to one another (229). A selective replication of the chromosomal nucleolar organizer occurs at early stages of oogenesis in amphibia and results in the formation of several hundred, up to 1000, extrachromosomal nucleoli per nucleus (305, 306, 323, 324). These are comparable to somatic nucleoli in ultrastructure and are active in producing rRNAthroughout lampbrush chromosome stages and yolk deposition (203, 239, 306, 325-327). The extrachromosomal copies of the nucleolar organizer become nonfunctional and are discarded into the cytoplasm at the first meiotic reduction division, and upon fertilization and cleavage are diluted out or degraded (305). The selective replication of rDNAleading to the production of extrachromosomal nu¢leoli in the amphibian oocytes presumably reflects the need for extra templates to support the very high rate of synthesis of rRNAat early stages of oogenesis. In oocytes from animal species other than amphibia, where there is no formation of extrachromsomal nucleoli but there is instead a conspicuous enlargement of the single nucleolus (239) and of the nucleolar associated chromatin [leading to the formation of the "Giardina body" in insects (328, 329)], a similar replication of the rRNAgenes takes place; this is indicated by RNA-DNA hybridization tests involving DNAfrom oocytes of worms (305), clams (305), and insects (328, 329). The selective amplification of high molecular weight rRBIA genes which occurs during oogenesis seems to be a unique phenomenon, at least in its proportions. An examination of different cell types of the same organism which differ greatly in the rate of rRNAsynthesis has revealed no significant differences in their amount of rDNA(224, 304, 330). Regulation of rate of transcription appears therefore to be the general mechanism by which production of rRNAis adjusted to the individual needs in different cell types or stages of development. Besides a regulation of rRNAgene activity related to the functional state of the cell, a dosage regulation has been revealed by the observation that Drosophila melcmo~Taster individuals with one, two, or three doses of the rDNAsegment have the same rRNAcontent (331), and, similarly, by the finding that the rate of synthesis of rRNA about equal in wild-type Xenopus lc, evis containing two nucleolar organizers and in individuals heterozygous for the anucleolate mutation (222). The use of mutants of Drosophi[c~ containing deletion of the nucleolar organizer (bobbed strains) appears to be a promising approach to gain insight into the regulation of expression of rRNAgenes (332). Two main explanations, not mutually exclusive, can be entertained for the redundancy of rRNAgenes. The first is that the multiplicity of rRNA genes merely satisfies a quantitative template requirement to maintain the rate of rRNAsynthesis needed for cell growth or differentiation. The second is that the rRNAgenes are not all equivalent, but rather belong to

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functionally different sets which become selectively activated in different cell types or stages of development. Wementioned the evidence that the multiple rRNAgenes in E. coli must all be simultaneously transcribed by many RNApolymerase molecules in order to produce the normal cell complement of rRNA. Similar calculations in HeLa cells, based on an average rRN’A content per cell of 24 pg (227), an estimate of about 1100 45S genes per cell (247), a minimumtime of 2.3 rain for the synthesis of one 45S molecule (236), and a maximumnumber of 100"RNApolymerase molecules operating simultaneously on each gene, as in the active extrachromosomal nucleoli of amphibian oocytes (229), indicate that at least 100 genes would have to continuously transcribed at maximumrate during the 24 hr doubling time to produce the average cell complement of rRNA. Therefore, the redundancy of genetic information for rRNA, at least in part, is used to satisfy a quantitative requirement both in bacteria and eukaryotic cells. The observation that the deletion of somewhat more than half of rDNAproduces in Drosophila metanogaster when homozygous, the bobbed phenotype (224, 332, 333), characterized by slow development, short bristles, low viability--and fecundity--a phenotype which would be expected to result from a general defect of the protein-synthesizing machinery would also speak in favor of a quantitative requirement for a redundant rRNAgene complement, rather than support the idea of a functional differentiation of rRNAgenes. Also the observation that the nucleolar organizer locus in Zea (334) and in Chironomus (335) can be broken by X-ray irradiation into fragments each retaining a nucleolar organizing activity sufficient to sustain normal development is in keeping with a quantitative role of rRNAgene redundancy. Strictly related to the question discussed above is the problem of rRNA gene heterogeneity: are the rRNA genes, and as a consequence the rRNA molecules in the same cell type, all identical ? And, if not, what is the degree and the physlologlcal significance of this heterogeneity? The rapid initial rate with which rl~lgA hybridizes with rDNA(41), the apparently regular and complete hydrogen bonding of rRNAmolecules to rDNA sites in these hybrids (247), and the renaturation kinetics of rDNA(336) suggest indeed a close similarity in sequence, if not an identity of rRNAgenes. Information concerning the possible sequence heterogeneity of rRNAgenes can be derived from an analysis of their transcription products. If the rRNAgenes in the same genome are not identical, two extreme possibilities, which correspond to the above discussed interpretations of their redundancy, have to be considered: (1) There is a functional differentiation of rRNAgenes, i.e. the pattern of expression of the various genes is different under different conditions; in this case, it should be possible, for example, to detect some differences between rRNAsamples extracted from bacteria grown in different media or from various tissues or developmental stages of the same higher organism. (2) The various rRNA genes are expressed in a coordinate fashion, though at a different rate, under all condi-

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tions; in such a case all rRNAsamples prepared from the same organism will be equally heterogeneous. The first possibility has been ~videly tested. Some differences in base composition (337) and partial sequences (81) have been described in molecular weight rRNA extracted from E. coli and Pseudomonas aeruginosa grown in poor or rich media; however, rRNA preparations from spores and vegetative B. subtilis have been reported to have the same base composition and hybridization properties (35, 338). In higher organisms, the cases where well-purified high molecular weight rRNAspecies have been analyzed, no difference between the rRNAs from different tissues or developmental stages has been found by using a variety of techniques: sedimentation analysis, mcasurement of optical properties, base composition and partial sequence analysis, DNA-RNA hybridization, and limited RNase digestion (23, 25, 43, 45, 119, 222, 339-346). As for the second possibility, the available evidence suggests that if heterogeneity exists in an RNApopulation, this is limited. Both in bacteria and in eukaryotic cells each high molecular weight rRNAcomponent, when prepared and analyzed under conditions which exclude degradation or aggregation, has been found to be homogeneous from the sedimentation and chromatographic point of view. Some heterogeneity has been found for the terminal sequences of high molecular weight rRNAof various sources (75, 77, 78), but in these cases the heterogeneity is restricted to two or very few types of molecules, and it seems difficult to correlate such a low degree of heterogeneity with the high redundancy of the rRNAgenes. Some evidence for heterogeneity has been obtained for L cell 18S RNA,in which some of the O’-methylated and base-methylated nucleotides are apparently present in only a fraction of the molecules (1/2 to 1/6) (68). However, since the methylation takes place after the rRNAchain has been synthesized, this heterogeneity might not involve the rRNAgenes. Homogeneity in primary structure of the high molecular weight rRNAis suggested in E. coli by the molar yield of unity or multiple thereof of methylated sequences (61) and in rabbit reticulocytes by the near-to-unity molar yield of rRNAfragments obtained by mild digestion with T1 RNase (116). The sequence studies on 5S RNAhave provided more incisive information concerning the heterogeneity of this low molecular weight rRNAspecies. In each of two strains of E. coli two forms have been found (with one in common), in about equal amounts, which differ from each other in sequence in one position, with some indications of smaller amounts (10 to 20%) of other forms of 5S RNAwith one or two base changes (122, 123). These forms presumably correspond to the various 5S genes that RNADNAhybridization experiments have revealed in E. coli (313). It is surprising, however, that only such a low degree of variability exists between these forms. Even more surprising is the fact that it has been possible to determine a main sequence also in 5S RNAfrom animal source, in partleular, KB cells (124) (a cell line of human origin) which contain presumably

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many thousands of genesfor this RNA species.The deviations from unity of themolaryieldsof thelargeoligonucleotides released by pancreatic or Tl ribonuclease digestion of K]3 5S RNA suggest, however,the presence of smaller amounts of variant forms of this RNAspecies, differing from the main sequence at various sites in the molecule. The presence in substantial amount of one or several alternate forms of 5S RNAdiffering presumably in one or more bases from the main sequence is indeed indicated by the fingerprint analysis after pancreatic RNase digestion of 5S RNAfrom HeLa cells (292). On the other hand, it is interesting that no differences have been found between the T1 and pancreatic RNase fingerprints of 5S RNAfrom two mouse cell lines and that of KB 5S RNA(347). The low degree of sequence variability of 5S RNAobserved within a given bacterial species and, in animal cells, within the same species and among different species of mammalspresumably reflects the great restrictions imposed upon evolutionary base sequence changes by the structural requirements for 5S RNAfunction. It is possible that the scqucnce variability of the 5S RNAgenes is greater than detected in the "functional" 5S RNA molecules isolated from the ribosomes, either because of inactivity of some of the genes or of destruction of the too aberrant transcription products. Considerations similar to those made above concerning 5S RNAapply to the apparently low degree of variability of the high molecular weight rRNA species. As to the mechanismwhich may operate in keeping low the variability of the highly redundant rRNAgenes in eukaryotic cells, one may think of a mechanism like that proposed by Callan (348, 349), involving one "master" gene and multiple "slave" copies, which are matched against the master and corrected for mistakes due to mutation or recombination once per life cycle of the organism or even per cell division. Alternatively, one can postulate a selective replication of a master gene in gametes with destruction of the old replicas. Experimental evidence speaks, on the contrary, against a selective replication of a master gene at each cell division (350). CONCLUDING IIEMAtlKS The extraordinary pace at which new knowledge concerning the structure and synthesis of rRNAboth in bacteria and eukaryotic cells has accumulated in recent years has certainly become apparent to the reader of this review. One can anticipate that within a reasonably short time the application of rapid methods for sequencing RNA(351) will lead to the determination of the complete sequence(s) of the high molecular weight rRNA components, at least from bacterial sources. This analysis will also make it possible to critical|y probe at the level of primary structure the question of the possible heterogeneity of rRNApopulations within the same cell and in different ceils of the same species. The mechanism by which the large families of repeated rRNAgenes in the eukaryotic genome are prevented from diverging in evolution is of a great theoretleal interest; possibly, this mechanismwill prove to be of gen-

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eral importance in the maintenance of reiterated gene families. Direct experiments should be able to test whether a system involving a "master" gene and multiple "slave" copies which are periodically put in register with the master, as postulated by Callan (348, 349), operates here, or whether gametes represent a "bottleneck," where replication of a new family of rRNA genes proceeds from one master gene with destruction of the old replicas. The availability of purified rDNA genes both from bacteria (319) and eukaryotic cells (225, 304) will soon allow the investigation of in vitro synthesis of rRNA and of the factors which regulate it. In eukaryotic cells, further biochemical dissection of the processing of rRNA precursors and of the assembly of ribosomal precursor particles will greatly depend on in vitro studies with isolated nucleoli and with subnucleolar fractions. Regulation of rRNA synthesis both in oogenesis and in somatic cell growth and differentiation represents another area of great interest for ture investigations. The selective amplification of rDNA which occurs early in oogenesis in amphibia and other organisms may represent just an example of a widespread phenomenon of gene dosage regulation operating in cell differentiation, and therefore the analysis of its mechanism may lead to ings o~ broad significance. It can be foreseen that the study o~ rRNA synthesis will continue to be very rewarding for the understanding of the organization and expression of the bacterial and eukaryotic gcnomc. LITER/~TURE CITED 15. Kumar, A., Biochim. Biophys. ,~cta, 186, 326 (1969) 16. Reisner, A. H., Rowe, J., Macindoe, H. M., J. Mol. Biol., 82, 587 (1968) 17. Ceeearini, C., Maggio, R., Biochim. Biophys. Acta, 166, 134 (1968) 18. Kfintzel, H., Noll, H., Nature, 215, 1340 (1967) 19. Rogers, P. ~., Preston, B. N., Titehener, E. B., Linnane, A. W., Biochem. Biophys. Res. Commun., 27, 405 (1967) 20. Ts’o, P. O. P., Bonnet, J., Vinograd, J., B~ocMm.Bioph~s. ,4cta, 80~ 570 (1958) 21. Click, R. ~., Tint, ]3. L., J. ~ol. B~oI., 25, 111 (1967) 22. Stutz, E., Noll, H., Proc. Natl. ,4cad. Sci. U.S., 57~ 774 (1967) 23. Grummt, F., Bielka, H., B~och~m. Biophys. Acta, 161, 253 (1968) 24. Hastings, J. R. B., Kirby, K. S., Biochem. f., 100, 532 (1966) 25. Slater, D. W., Spiegelman, S., Biophys. I., 6, 385 (1966) 26. Ts’o, P. O. P., Vinograd, ~’., Biochim. Biophys. Acta, 49, 113 (1961)

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Annu. Rev. Biochem. 1970.39:183-226. Downloaded from arjournals.annualreviews.org by 192.244.210.205 on 10/25/05. For personal use only.