Synthesis and processing of high molecular weight RNA by nuclei isolated from embryos of Rana pipiens

J. Mol. Biol. (1972) 69, 1938

Synthesis and Processing of High Molecular Weight RNA by Nuclei Isolated from Embryos of Rana pipiens J. Doua~ks C&xroNt MTD PAUL H. JONES Department of Anatomy and Devekqnnental Biology Center Case We&em Reserve University Cleveland, Ohio 44106, U.B.A. (Received 25 November 1971, and in revised fom~ 1 May 1972) When synthesized under conditions optimal for maximal methylation, two of the major classes of high molecular-weight RNA produced by isolated nuclei were indistinguishable from RNA pm-i&d from cytoplasmic ribosomes. The degree of methylation was found to influence the sedimentation velocity of only the heavier major RNA fraction. Upon becoming maximally methylated, this RNA sedimented at 27 s rather than 26 s. When comparing maximally-methyl&ted nuclear RNA to cytoplaamic rRNA, it was found that on the basis of sedimentation coefficient, base composition and electrophoretic mobility, the 07 million dalton nuclear RNA was equivalent to 18 s cytoplasmic rRNA, the 1.6 million dalton nuclear RNA was equivalent to the 27 s moiety of 28 s cytoplasmic rRNA, and the 0.058 million dalton nuclear RNA was the same as the 7 s moiety of 28 s cytoplasmic rRNA. In isolated nuclei, the 27 s and 7 s RNA did not associate to form the 28 s duplex found in mature cytoplasmic ribosomes. In addition to these three fractions, which accumulated throughout the incubation period, several rapidly turning over, high molecular-weight, methylated RNA fractions were identified. The largest had a sedimentation coefficient (45 s) and nominal molecular weight (4.4 million daltons) to be expected of the primary transcription product of the entire rRNA gene. The results have been used to deduce processing pathways for cleavage of the primary precursor into the two definitive rRNA species and several excess rRNA fragments.

1. Introduction The two large ribosomsl RNA molecules in eukaryotic organisms are transcribed as parts of a single molecule, the primary ribosomal precursor (for review see Attardi & Amaldi, 1970). This molecule is cleaved by way of multistep pathways to yield eventually the definitive rRNA species and the excess rRNA fragments. The synthesis and processing of rRNA has been studied most extensively in HeLa cells (summarized by Weinberg & Penmrtn, 1970), where the primary transcription product of the rRNA gene has been shown to have a molecular weight of 4-l to 4.5 million dsltons (McConkey & Hopkins, 1969). In contrast, studies of rRNA synthesis and processing in whole cells of Amphibia have indicated that the primary transcription product has a molecular weight of only 2.4 to 2-6 million daltons (Loening, Jones & Birnstiel, 1969; Perry et al., 1970). Since the amphibian rRNA gene is large enough to code for about 4.5 million daltons of RNA (Birnstiel, Grunstein, Speirs & Hennig, 1969; Dawid, Brown & Reeder, t To whom correspondence should be sent. 19

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1970; Wensink & Brown, 1971), the finding of the 2.4 to 26 million dalton precursor has been taken to mean that the amphibian system, unlike the mammalian system, transcribes only about one-half of its rRNA gene. Results reported in this paper, however, show that nuclei isolated from embryos of Ram pipiens, as well as from post-metamorphic frog liver, in situ, produce several discrete fractions of RNA with nominal molecular weights greater than 2.6 million daltons ; indeed, a 4.4 millioqdalton fraction (45 s) has been observed routinely. This suggested that the amphibian system can transcribe its entire rRNA gene. However, the isolated nuclear system also appeared to synthesize an anomalous class of RNA. Instead of producing 28 s and 18 s rRNA, incubations of isolated nuclei accumulated 25 s and 18 s RNA. Although the electrophoretic mobility (molecular weight) and base composition of the 25 s nuclear RNA and 28 s cytoplasmic rRNA were the same, it has been necessary to explain this difference in sedimentation velocity to assure that the nuclear system did not synthesize aberrant rRNA molecules. The work reported here shows that the difference in sedimentation behavior resulted from (1) under-methyl&ion of the 25 s nuclear RNA, and (2) the molecular structure of the 28 s cytoplasmic rRNA duplex (27 s : 7 s = 28 s). When these factors are considered, RNA synthesis in isolated nuclei, at least during the first 40 minutes of incubation, produces [molecules that are indistinguishable from their whole-cell counterparts. Also, results from the isolated nuclesr system have provided additional information about processing of rRNA precursor molecules and indicate that the amphibian rRNA system (and perhaps the system of other poikilothermic organisms) is more similar to the mammalian system than had been thought previously.

2. Material and Methods (a) Isolation and incubation

of nuclei

Nuclei were isolated from feeding-stage embryos of Rana pipiena (stage 25; Shumway, 1942) by the method of Mittermayer, Breun & Rusch (1966), in a medium composed of 0.25 M-sucrose, O-1 M-KCl, 1.0 mM-MgCl, and 0.1 m-Tris, pH 6.8. The nuclei were incubated at 27% for the times indicated, in incubation medium consisting of 0.25 M-sucrose, 5 mMMgCls and 60 mM-Tris, pH 7.8 ; nucleoside triphosphates were included at concentrations of 6 PM-ATP and GTP and 70 PM-CTP and, except where otherwise stated, ‘70 PM 3H-labeled UTP (0.01 to 24 Ci/m-mole), which proved to be optimal. (b) Purification

and analysis of RNA

At the times indicated, nucleic acids were extracted from the nuclear incubations by the sodium lauryl sulfate/cold phenol method. After treatment with DNase and subsequent purif%cation, RNA was fractionated and analyzed by sucrose gradient centrifugation and acrylamide gel electrophoresis (Bishop, Claybrook & Spiegelman, 1967; Loening, 1969). The gels, 6 mm in diameter, contained 2% acrylamide and 1% agarose. After trimming about 1 cm from the top and bottom, the gels were 95 mm long. Before loading the sample, the gels were subjected to standard electrophoretic conditions for 30 min and allowed to stand overnight at room temperature in the running buffer: 36 mm-Tris, 30 mM-NaHsPG4, 1.0 m&r-EDTA, O*2o/o sodium lauryl sulfate, final pH 7.8. Electrophoresis of the sample, which also contained several marker RNA’s, was for 2.5 hr at 50 V (about 5 mA per gel). After scanning at 260 nm in a Gilford spectrophotometer to determine the position of absorbency markers, the gels were sliced into sections of 1 mm thickness and processed for counting in a 3-channel liquid scintillation spectrometer equipped with external standardization. To achieve maximal resolution of the several molecular-weight classes of RNA, less than 03 Aas0 unit of total RNA was loaded on to the gels. To obviate problems associated with 23 s and 16 s RNA (0.1 matching the absorbency tracings with gel slices, 32P,-labeled

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Aaeo and 11,000 cts/min of each) were added to the original RNA extract whenever possible. These served both as internsd and external markers and permitted absolute resolution of several nuclear RNA classes (1.2, 1.0, O-9, 0.6 and 0.4 kkd)t from bacterial RNA (1.08 and 0.66 kkd). This procedure also permitted estimation of over-all recovery of the 3H-labeled RNA made by isolated nuclei. Thus, the analysis of each class of RNA at each time period could be treated in a quantitative as well as a qualitative manner. Before their use for analysis of nuclear RNA, sample gels from each betch were tested for their ability to separate RNA’s with molecular weights in the 0.3 to 4.4 l&d range. In these tests, labeled RNA from E. coli (32Pi-labeled 23 s and 16 s) and HeLa cells (3Hlabeled, 46 s) were mixed with RNA from one other source and electrophoresed under standard conditions. The RNA’s used as markers are listed with their assumed molecular weights in Table 1. Although there are uncertainties concerning molecular weight as determined by acrylamide gel electrophoresis (see Loening, 1969), with most RNA’s the method TABLE

Assumed molecular

Source of RNA

1

weight of RNA used a.s markers for acrylumide-gel electrophoretic analyses

Nominal mol. wt in millions of daltons

E. coli Rst

1.08 (23 8) and 046 (16 s)

Chick Rana pipiena

1.60 (28 8) and 0.68 (18 s) 1.61 (28 s) and 0.68 (18 s)

Xenopus laevti

1.51 (28 s) end 0.68 (18 s)

Standley & Bock, 1965 Hamilton, 1967; Petermann & Pavlovec, 1966 Relative to markers of E. coli and rat Relative to markers of E. coli and rat Relative to markers of E. coli and rat;

4.4 (45 s) (3H labeled)

McConkey & Hopkins, 1969

1.72 (30 s) end 0.68 (18 s)

also see Loening HeLa Cells

et al., 1969

seems to be at least m reliable se the usual hydrodynamic methods. For present purposes we refer to each RNA class by its nominal molecular weight, which in some cases differs slightly from its apparent molecular weight (see Table 4), in l&d. Base-composition determinations were performed on alkaline hydrolysates of RNA by thin-layer chromatogmphy (Randerath & Randerath, 1964) or by high-voltage paper electrophoresis using 0.05 m-ammonium formate, pH 3.5, as the buffer (Markham & Smith, 1952). (c) Analysis

of 28 s cytoplmic

rRiQA

The 28 s rRNA was purified from liver ribosomes of well-fed adult Rana pip’ena by several cycles of centrifugation through sucrose gradients. The 28 s duplex (40 to 200 pg of RNA/ml.) was disrupted by heating at 40 to 45°C for 10 min in 0.01 M-NaHPO,, pH 7.6, and then cooling rapidly (cf. King & Gould, 1970). For analysis by sucrose gradient centrifug&ion or aorylemide gel electrophoresis, the sample (either intact 28 s or disrupted 28 s RNA) was transferred to the appropriate buffer without an intervening precipitation with ethanol. (d) Labeling of liver RNA, in situ To label liver RNA, 1 mCi of 32P,-labeled or 0.5 mCi of 14C-labeled uridine (0.5 Ci/m-mole) and 0.6 mCi of [C3H,]methionine (3.3 Ci/m-mole) were injected into the peritoneal cavity of recently metamorphosed Rana p@ens. The labeled RNA wss extracted from total liver homogenates and purified as described above. t Abbreviation

used: kkd, million daltons.

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3. Results Most of the newly synthesized RNA that accumulated in reaction mixtures containing isolated nuclei separated into two major high molecular-weight fraotions upon centrifugation through sucrose gradients (Fig. 1) or electrophoresis in acrylamide gels (a) I.00 -

28 s

Fraction from bottom (0 75ml)

FIQ. 1. Sucrose-gradient analyses of RNA synthesized by isolated nuolei after 40 min of inoubstion in the presence (a) or absence (b) of a methyl donor. (a) RNA labeled with [8HjU’l!P (24 Ci/m-mole; -@-a-) was centrifuged with 18 s and 28 s cytoplesmio rRNA ((-----), As,,). Most of the absorbency in the low molecular-weight region resulted from partimlly degraded DNA. (b) RNA labeled with [W]UMP (0.125 Ci/m-mole; (---@-a--) and CsH&adenosylmethionine (1.26 Ci/m-mole; -- 0 --- 0 --). The position of 28 8 absorbenoy marker, whioh was inoluded in the centrifugation tube, is indicated by the errow.

(Figs 3(a), 6 and 7). The base composition (Table 2) and relative proportion of RNA in these fractions, as well as the electrophoretic mobilities, were suggestive of oytoplasmic rRNA, yet only one of the major fractions, 18 s, sedimented coincidentally with RNA purified from cytoplasmic ribosomes. The other sedimented at about 25 s, considerably slower than 28 s RNA from mature cytoplasmic ribosomes (Fig. l(a)). Thus, although by most criteria the RNA made by isolated nuclei appeared to be mostly normal rRNA, the sedimentation of the heavier, major fraction implied that one species of the newly synthesized nuclear RNA was somewhat abnormal. However, it seemed probable that the abnormality concerned some secondary modifkation such as conformation rather than a major difference in molecular weight or primary structure, because the cytoplasmic 28 s and nuclear 26 s RNA migrated together on aorylamide gels and, within experimental error, had the same base composition.

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TAEZLE2 Base wmpo&o~~ of RNA synthesized by isolated nuclei compared to that of mature cytopluamic ribowmd RNA

I. Made by isolated

AMP

32.2 32.2 27.8 28.4 32.1

14.3 17.1 21.9 21.2 20.4

32.6 32.0 28.1 28.2 31.6 29.1 28.8 28.4 32-O

Mole y. GMP

UMP

G+C

36.1 33.0 30.2 28.1 23.6

17.4 17.7 20.1 22.3 23.9

68.6 06.2 68.0 66.6 66.7

14.6 16.6 21.4 20.9 21.1

36.7 34.0 31.0 29-l 24.0

17-2 17.4 19-6 21.8 23.3

68.2 66-O 69.1 67.3 66.6

18.7 19.2 20.4 20.3

32.8 32.8 29.3 23.7

19.4 20.2 21-9 24.0

61-9 61.6 67.3 66.7

nuoleit

A. without CH3 donor 46 s$ 26 s$ 1-6 kkd 78

18 s$ B. with CH3 donor 46 s$ 27 s$ 1.6 kkd 7s 18 s$ II. Cytoplasmic 28 s 27 s 7s 18 s

CMP

rRNA§

t RNA was labeled with a mixture of the four aH-labeled ribonuoleoside triphosphetes results were not correoted for isotope dilution by endogeneous nuoleoside pool. $ Not homogeneous fmotione; see Fig. 7. 0 Caloukbted from molar absorption coeffioient of eech nucleoside monophosphate.

(a) Methyl&on

and the

of nuclear RNA

The RNA made by isolated nuclei probably had been incompletely methylated, because the nuclear preparations contained no detectable S-adenosylmethionine and because methyl donor had not been added to the reaction mixtures. Moreover, the degree of methylation might greatly influence the hydrodynamic behavior of large RNA molecules without significantly altering their electrophoretic mobility (molecular weight) or base composition. For this reason, the effect of methylation on the sedimentation and electrophoretic behavior of RNA synthesized by isolated nuclei was examined. In these experiments we have assumed that the maximal level of C3H3 incorporation represented the natural degree of methylation of the RNA. Maximal methylation could be accomplished with a concentration of S-adenosylmethionine of about 2 x 10m6 M or of methionine of about 1 x 10e4 M (Fig. 2). However, the concentration of S-adenosylmethionine that gave maximal incorporation also produced a 5 to 10% inhibition of RNA synthesis as measured by UTP incorporation. The inhibition was probably related to the adenosine (unpublished results) because inhibition of RNA synthesis w&9 not seen with methionine at concentrations up to about 5 x IO-4 M. The two methyl donors were equally effective in methylation of the RNA, and asauming no dilution of the added methyl donor, we calculate that under optimal conditions, 1.1 to 1.3% of the nucleoaides in 27 s (formally 25 s) and 1.6 to 13% of those in 13 s

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2x10-e

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1X10-5

P. H.

JONES

1X10-4

1110-3

Concentration of CH3 donor (~1

FIG. 2. Effect of methyl donor concentration on methylation of RNA by isolated nuclei. Nuolei were inoubated for 30 min in standard incubation medium containing either S-adenosylmethionine (-•--•--) or methionine (-- 0 --- 0 --). The specific activity was adjusted so that eaoh incubation tube contained 10 $i or C3Hs donor/ml.

RNA became methyl&ted. The distribution of methyl groups among the four ribonucleoside monophosphates has not been determined, nor has the degree of methylation of mature cytoplasmic rRNA of Ranapipiens been measured. However, the values given here are within the range reported for rRNA of other eukaryotic organisms (see review by Attardi & Amaldi, 1970). Two other RNA classes on which base composition data are available, 45 s and 7 s, showed an average degree of methylation of 1.07% and O-6 to l-2%, respectively (Table 4). Because of the limited amount of material, we have found no reliable way to assess directly the degree to which RNA molecules made in the absence of added methyl donor were methylated. We assume, however, that such RNA was incompletely methyl&ted because the calculations given above indicate that the specific activity of added methyl donor was diluted very little, if at all, by an endogenous pool. Also, direct measurements showed that standard nuclear preparrations contained less than 3 x lo-l1 mole of S-adenosylmethionine per ml. This concentration would not be expected to yield maximal methyl&ion. The only apparent effect of maximal methyl&ion was on the sedimentation velocity of the heavier major fraction of newly synthesized RNA. When maximally methylated, this RNA sedimented at about 27 s, whereas when incompletely methylated it sedimented at about 25 s (Fig. 1(b) ; Table 3). Maximal methyl&ion produced no apparent effect on the sedimentation of other RNA classes (Fig. l), nor did it alter the number of bands or the mobility seen on ecrylamide gel electrophoresis (Figs 3(a) and 7). Also, maximal methyl&ion failed to alter significantly the over-all base composition of the major RNA classes (Table 2). (b) Properties of 28

s

RNA of cytoplasmic ribosomes

Although when maximcllly methylated the heavier major fraction of nuclear RNA sedimented two S-units faster, under no condition was this RNA found to sediment ooinaidentally with 28 s RNA from mature cytoplasmic ribosomes. For this reason, we have investigated the molecular integrity of 28 s cytoplasmic rRNA to determine if it, like the comparable rRNA class of other metazosns, was a molecular duplex mada

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TABLE 3 Comparison of sedimentation coejkients of RNA synthesized under differing conditions by isolated nuclei to those of RNA puri$ed from mature cytoplasmic ribosomest Observed Heavy I. II.

Cytoplasmic

rRNA

RNA made by isolated presence of:

S value Light

28

18

nuclei in

A. 200 p&r-Methionine (7) B. 10 q-S-Adenosylmethionine (3) C. Without a methyl donor (10)

26.2-21.2 26.2-21.2 23.8-25.2

17.8-18.2 17.8-18.2 17.8-18.2

t Sedimentation coefficients measured by the method of Martin & Ames (1961) using 23 s or 28 s and 18 s RNA as standard markers. Numbers of determinations shown in parentheses.

TABLE

4

Estimated molecular weight of discrete RNA classesfound in incubations of isdated nucleic Nominal mol. wt VW 4.4 3.8 3.1s 2.6 2.2 1.911 1.8

14/l 1.5 (27 8) 1.2 1.011 0.9 0.7 (18 s) 0.6 0.471 0.37 0.058 (7 s)

Renge WV 4.35-4.50 3-70-3.90 3*05-3.20 2.45-2.75 2.15-2.30 1.80-2.02 1.72-1.78 l-65-1.70 1.49-1.53 1.15-1.22 0.94-1.15 0.86-l-04 0.65-0.71 0.68-0.62 0.38-0.42 0.28-0.31 0.05-0.06

Average W4 4,40 3.83 3.13 2.58 2.20 1.93 1.74 1.63 1.51 1.19 1.04 0.91 0.76 0.59 0.41 0.30 0.058

Number of methyl groupst per molecule 143

50

35

1-2

t Twenty-two measurements were made on 2% gels, 4 on 2.2%, 2 on 2.4% and 3 on 2.8%; all these gels were 65 mm long. In addition, 4 measurements were made on 2% gels that were 95 mm long. $ The number of methyl groups per molecule was calculated from the mole C?H,/mole 14Cq labeled UMP, the y0 UMP and assumed moleculsr-weight of a given fraction. The values listed represent the average of 3 determinetions on 4 samples end were within +5% (except the v&e for 7 s which was & 10%). 5 Small amount of radioactivity relative to adjacent fractions; not seen after 20 min of incubation. II Small amount of radioactivity relative to that in adjacent fractions. lJ Usually resolved only on 95 mm gels.

J. D. CASTON AND P. H. JONES

26

(b)

Dlstonce from origin (mm)

FIQ. 3. Analysis of [l*C]UMP (-•--a---) and C3H3 (-acrylamide/l~O agarose gel. Number over eaoh band refers &B determined by the position of 31P-labeled 23 8 and 16 nuclei after 36 min of incorporation. (b) RNA from frog liver

0 --- 0 --)-labeled RNA on 2.0% to nominal moleoular-weight in kkd 6 markers. (a) RNA from isolated labeled & eittl for 3 hr.

up of one 27 s moiety in a non-covalent association with 8 7 s moiety (of. pene, Knight & Darnell, 1968; Ring & Gould, 1970). The results given in Figure 4 show that heating et relatively low temperature (45°C for 10 minutes) completely disrupted highly purified 28 s cytopltunnio rRNA into 27 s and 7 s fractions. Eleotrophoretic analysis indicated molecular weights of 1.51 and 0.058 kkd for purified 27 s and 7 s RNA, respectively. According to these molecular weights, separation of the 28 s duplex yielded roughly stoiohiometrio amounts of 27 s and 7 s RNA (Table 5). These results are in sgreement with the findings of King & Gould (1970), who have presented a more thorough treatment and discussion of the phenomenon using 30 s rRNA of rabbit reticulocytes. No difference in electrophoretic mobility (Fig. 4) or over-all base composition (Table 2) between 28 s and 27 s rRNA could be detected. This is not surprising, since the 7 s RNA contributed so little (s,pprox. 3.5%) to the total mass of the 28 s duplex. It should be noted that the isolated nuclear system synthesized both the 27 s and 7 s molecules (Fig. 3). However, conditions were evidently unfavorable in the nuclear preparations for formation of the 28 s duplex. (0) Heterogeneity of nuclear RNA In addition to the two major high-molecular-weight fractions, a small but significant amount of newly synthesized RNA sedimented in discrete bands at about 45 s, 41 s, 36 s and 30 s (Fig. 1). To establish the identity of the sedimentation classes with

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a)

I.5 1.2 0 g 09 06 O.?

;!7s

3s

i 0 B P

/ -

95mm

0

-

95mm

Fm. 4. Sucrose gradient and aci$8mide gel 81dysis of 28 s duplex of cytoplaemic ribosomes. (8) Sedimentetion of dierupted duplex (-m--O--, A& in relation to intact duplex (oII-labeled, (----)). (b) Electrophoresic of 7 s moiety of 28 s duplex on 7.2% acryhunide gel. (c) Electrophoreeie of intact duplex on 2.0% ecrylamide gel (96 mm long). (d) Electrophoresis of disrupted duplex; solid line indicated a Uilford spectrophotometer recorder setting of 0 to 2 A,,, and deehed line indicatee 8 setting of 0 to 0.6 k&c. TABLE 5 Relative colztributtin of 27 S and ?’ S to 28 S duplex Amount of 28 s (Aa,o) 31.0 17.5 7.8

Amount of 27 s Waao)

Amount of 7 s (&JO)

29439 1.13 17.06 0.64 7.62 0.30 0.058 kkd (7 8) x 100 = 3.51 l-6 kkd (27 e)

2Fs x 100 3.78 3.75 3,8B

respect to electrophoretic mobility, the individual sucrose-gradient fractions were electrophoresed in acrylamide gels. As shown in Figure 5, each sedimentation class resolved into more than one electrophoretic band. The mobility, and hence nominal molecular-weight of RNA, in these bands corresponded exactly to that of bands routinely found upon electrophoresis of unfractionated, newly synthesized RNA (Figs 3 and 7, Table 4). When extracted from the gel slices and electrophoresed again, each band tested (4-4, 3.8, 2.6, l-2 and 0.7 kkd) migrated to its original position. Of the 16 fractions listed in Table 3, all except 3.1, l-9, l*O, O-6,0-4 and 0.3 kkd were

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I.76 I.51

k&L I.78

AL20

40

60

20

40

60

40

60

Distance from origin (mm)

FIG. 6. Acrylamide gel electrophoresis of RNA from sucrose-gradient f&&ions. 3H-bbeled RNA made by isolated nuclei w&s fractionated on sucrose gmdients sJld the individmd freations were electrophoresed as indicated with unlebeled rRNA from R. pip&u (Rp) end 1. wli (Ec); Azso (---) is plotted on the first and last frames. Number over each band refers to nomine molecular-weight in kkd. observed in every an&~&

of total RNA. Most of these fractions were probably present in all of the preparations but the 0.3 and 0.4 kkd fractions sometimes ran off 6Ei-mm gels. The other fractions may not have been clearly resolved on some gels because they were obscured by the relatively large amount of radioactivity in adjacent bands. Also, some of the fractions can be found only after characteristic periods of synthesis (Fig. 6; see section (d) below). The 1.2, 1.0, 0.9, O-7, O-6 and 0.4 kkd RNA could not have arisen from bacterial contamination, since 3aP-labeled 23 s (l-08 kkd) and 16 s (O-56 kkd) marker RNA never coincided on acrylamide gels with these 3H-labeled fractions (Fig. 7). Apart from the discrete high molecular-weight classes of RNA shown in Table 4, ribonucleoside phosphates were also incorporated into RNase-sensitive material, which formed an asymmetric zone in the heaviest regions of sucrose gradients and which trailed from the top of acrylamide gels into about the f&t centimetre. This material contained essentially no methyl label (Fig. 3(a)) and contributed less than 0.5% of the total radioactivity after 15 minutes of synthesis (Fig. 6). We assume that this RNA was equivalent to the “>5 kkd” fraction described in whole cells of XenoF l.aevis by Loening et al. (1969). This RNA has not been examined systematically but it appears to have been comprised of a very heterogeneous population of RNA molecules. The heterogeneity of RNA made by isolated nuclei was also seen in RNA made by frog liver, in situ. The results given in Figure 3(b) show that the only apparent difference in high molecular-weight RNA from the two sources concerned the relative amount of radioactivity present in the individual RNA fractions. Whether this indicates a real difference in the amount of a given RNA class, or whether it merely reflects differing kinetics with which the nuoleoside triphosphate pool of the two systems became saturated with label, has not been determined. However, the results

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I

-6 F: 0 .cx 4 E > 2 7.5 z 2 9 6

IO

20

30

40

50

60

IO

20

30

40

50 60

Distance from originhm)

FIG. 6. Time course in which labeled nucleoside phosphate appeared and accumulated in high molecular-weight fractions of RNA synthesized by isolated nuclei. At the times indicated, azPlabeled rRNA from E. co&i was mixed with 3H-labeled RNA extracted from about 3 x 10s nuclei (equivalent to 45 pg of DNA) and analyzed by electrophoresis as described in the text. Numbers over the several bands refer to molecular weight in kkd; the position of azP marker RNA is shown by (f). Recovery of 32P ranged from 93% at 20 min to 98% at 25 and 25 min.

definitely show that frog liver and isolated nuclei produced the same number of high molecular-weight fractions and that the individual fractions were equivalent at least in terms of their nominal molecular weight. These results provide additional evidence that during the 40-minute period of incubation the isolated nuclear system produced naturally occurring RNA molecules. (d) Time sequence of labeling All analyses of RNA samples taken at equivalent times during the linear period of synthesis gave essentially the same results as those shown in Figure 6. Before about four minutes of incorporation, no discrete class of radioactive RNA could be detected. Although the 2*5-minute samples contained a considerable amount of radioactive

J. D.:CASTON

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(a)

6-

10

20

30

40

50

60

IO

20

30

40

50

60

Distance from origin (mm)

FIQ. 7. Analysis of RNA purified from the same stsnderd nuclear incubation (without added CHa donor) Bfter different periods of synthesis. (a) After 10 min of continuous labeling; (b) after 20 min; (o) after 30 min; (d) after 20 min of continuous labeling followed by e IO-min chase with lOOO-fold excess of unlabeled UTP. Arrows indicate the position of 28 8 8nd 18 s rRNA of R. pipkna and 3aP.Labeled 23 s and 16 a rRNA whioh were interns1 markers. The gels were 2.0% eoryhbmide end conteined 1 o/o agerose.

RNA, most of the labeled molecules were localized in the first few millimetres of the gel. These comprise the “>5 kkd nuclear RNA.” The rest of the radioactivity seen at this time period formed a rather constant background throughout the gel. The first discrete fractions of RNA to become labeled to a degree sufficient for detection were observed in the four- (not shown) and five-minute samples. Although the trained eye can detect radioactivity in most of the discrete classes at this time, convincing bands of label were present only in the RNA with nominal molecular weights of l-2, 1.0 (not resolved well in these gels), O-9 and O-7 (18 S) kkd. After seven minutes of incorporation, most of the discrete RNA classes contained enough label to be unmistakably resolved from the background. The three classes not labeled at this time were O-3,0-4 and O-6 kkd, which became labeled between 16 and 17, and 17 and 18 minutes, respectively. Resolution of the 3.1 to 3.2 kkd fraction was lost after about 20 minutes of labeling. In only two fractions, l-5 (27 s) and O-7 (18 S) kkd, did label continue to accumulate through the entire period of RNA synthesis. The amount of radioactivity in each of the other classes eventually reached a plateau, but the length of time required to achieve maximal labeling differed according to individual fractions. Since molecules in these fractions were continuously turning over and being replaced with newlysynthesized molecules, we presume that the difference in time required to reach maximal labeling resulted from the original position of these molecules with respect to the 5’ and 3’ ends of the 4.4 kkd rRNA precursor from which they arose. Of special interest is the differential accumulation of label in O-7 and 15 kkd RNA with respect to time. Note that the 0.7~kkd fraction contained a sizable amount of label at time periods five and seven minutes when the 1.5-kkd fraction and the several precursor fractions contained very little radioactivity. Since these molecules represent

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stable process products of a single precursor moleoule, these results definitely show that the 0.7~kkd class arose from a position nearer the 3’ end of the primary rRNA precursor than did the l.Bkkd class, and that the two were separated by less than two minutes of labeling time (see Discussion). Beoause added substrates equilibrated instantaneously within the isolated nuclei (unpublished results), and because RNA synthesis was linear throughout the entire period of study, results presented in Figure 6 can be used to estimate the over-all rate at which RNA molecules were synthesized (initiated, polymerized and terminated). By restricting measurements to the primary rRNA precursor molecules, 4.4 kkd, rates of processing (turn-over) need not be considered, except to assume that the system was in a steady state. Although radioactivity was present in the 4.kkkd region from the earliest time period examined, because of the high background, reliable measurement of label in these molecules could not be made during the first ten minutes of synthesis. However, it appears that the measurements presented in Table 6 have accurately estimated the TAEZLE6 Time wurae in which kzbel accumulated in 4.4 kkd RNAf Time bm

Experiment

10 12 16 17 20 26 30

3700 6020 6400 7340 7620 7390 7440

1 (ots/min)

Experiment

2

3260 4740 6610 6310 6440 0430 6380

t Deta given in Fig. 6 were nonnelized by the amount of 3”P-lebeled internal marker reoovered from the gels et 1.08 and 0.66 kkd. The resultant ourves were idealized to estim8te background 8nd the emount of overlep between adjeoent bands. Maximal cts/min represent the ideelized area of each band oorrected for be&ground and overlsp.

accumulation of label in the 4.4.kkd fraction, because extrapolation of the linear portion of accumulation to zero time coincides with zero amount of label. Accordingly, the 4.4-kkd fraction became maximally labeled, and hence the individual molecules presumably contained label from the 5’ end all the way to the 3’ end, within about 17 minutes of synthesis. If labeled, incomplete molecules contributed little to these measurements, the over-all rate of initiation, polymerization and termination at 27°C was about 259,990 daltons of RNA per minute (about 13 nucleoside phosphates per second). This value agrees well with the rate of 12 to 15 nucleosides per second indicated for this system by kinetics of rifamycin inhibition (unpublished results). It is considerably lower than the apparent rate of 166 nucleosides per second calculated from experiments with HeLa cells (Greenberg & Penman, 1966) and about half the rate calculated for RNA chain growth in E. coli cultured at 29°C (Manor, Goodman & Stent, 1969). The value for isolated nuclei is in good agreement with the apparent rate of 12.5 nucleosides per seoond estimated for Euglenu gracilis growing at 25°C (Brown & Haselkorn, 1971).

32

J. D. CASTON

AND

P. H. JONES

(e) Stability of nuclear RNA fractions Because all of the discrete high molecular-weight RNA fractions were methylated (Figs 1 and 3), it seemed likely that they represented the primary rRNA precursor and its several processing fragments. This was further indicated by the observation that the amount of radioactivity in the 15 (27 s) and 0.7 (18 s) kkd RNA increased throughout the entire incubation period, whereas the level of radioactivity in the highest molecular-weight fractions reached plateau between 15 and 20 minutes (Figs 6 and 7). This suggested that the larger molecules were short lived and perhaps were being converted into definitive rRNA which accumulated, and into excess precursor fragments which were rapidly hydrolyzed into molecules of very low molecular weight. To test this possibility, RNA from pulse-chase experiments was analyzed on aorylamide gels to determine which of the high molecular-weight fractions were stable and which were short lived. Figure 7 compares the RNA pattern after 30 minutes of continuous labeling with that of a 20-minute pulse followed by a lo-minute chase (Fig. 7(d)) with a lOOO-fold excess of unlabeled UTP (final concentration about 7 mM). In addition to diluting the [3H]UTP, the increased concentration of UTP during the chase would be expected to inhibit total RNA synthesis by about 40% (unpublished results). Even so, it can be seen that the amount of label in the 15 (27 S) and 0.7 (18 s) kkd fractions of the two preparations (Fig. 7(c) and (d)) were about the same, whereas the amount of label in the other fractions decreased significantly or disappeared altogether during the chase. These results provide evidence that the 4.4 kkdfraction represented the primary precursor to rRNA and that the other short-lived fractions were intermediary precursors or excess rRNA process fragments.

4. Discussion (a)

Characteristics of nuclear RNA

The isolated nuclear system used in these experiments synthesized and accumulated sizable amounts of high molecular-weight RNA. Although the system was operating outside its normal cytoplasmic environment, it produced discrete RNA classes with gross properties similar to those of RNA produced by adult frog liver in situ (Fig. 3). It has not yet been possible to compare all the nuclear fractions to equivalent fractions from whole cells. However, the direct comparisons given below, as well as a preliminary comparison of the 5 s and 4 s fractions from the two sources (Jones, 1971; Caston & Jones, unpublished observations), indicate that the nuclear system produced naturally occurring classes of RNA. In comparing the 1.5 kkd RNA made by isolated nuclei to RNA from cytoplasmic ribosomes the duplex nature of the 28 s rRNA must be considered. The large cytoplasmic rRNA molecule of Ranu pipiens (Fig. 4), like that of rabbit reticulocytes (King & Could, 1970) and HeLa cells (Pene et al., 1968), was found to be a duplex composed of one 27 s and one 7 s molecule in non-covalent association. Comparison of these two molecules with the maximally methylated 1.5 and 0.058 kkd RNA made by isolated nuclei, showed that the RNA’s from the two different sources were indistinguishable on the basis of sedimentation velocity, eleotrophoretic mobility and over-all base composition. By these same criteria, the O-7 kkd RNA made by the isolated nuclei was likewise indistinguishable from the 18 s RNA of cytoplasmic ribosomes.

PROCESSING

OF rRNA

PRECURSOR

33

Only under conditions which gave incomplete methylation of newly synthesized RNA did the nuclear system appear to produce aberrant RNA molecules. Even then, the only discernible aberrancy concerned the sedimentation behavior of the 1.5 kkd RNA. When incompletely methylated this RNA sedimented at 25 S, whereas when maximally methylated it sedimented at 27 s. This increase in sedimentation coeacient implies a possible gain of about0.2 kkdmolecular weight (Kurland, 1960; Spirin, 1961). However, electrophoretic analyses, which were capable of resolving molecular weight differences of less than 0.1 kkd (Fig. 3 ; also see Loening, 1969), indicated that any gain in molecular weight produced by methylation was considerably less than this amount. Also, since the over-all base composit’ion of incompletely and maximally methylated 1.5 kkd RNA was the same, it seems unlikely that the levels of methylation achieved in these experiments altered the points at which the rRNA precursor molecules were cleaved. Indeed, maximal methylation of the RNA had no detectable effect on the synthesis or the accumulation of any of the discrete RNA classes. Whether this means that a level of methylation crucial for normal processing was achieved by an undetectable concentration of endogeneous methyl donor has not been determined. However, it seems probable that any molecular weight gained by the 1.5 kkd and other nuclear RNA as a result of maximal methylatian was restricted solely to the weight of the methyl groups added to the RNA and, possibly, a concomitant increase in bound water associated with them. This conclusion is strengthened by the fact that the characteristics (over-all base composition, sedimentation and electrophoretic properties) of the 4.4 and O-7 kkd nuclear RNA were unaffected upon becoming maximally methylated. Thus, by whatever means, the hydrodynamic properties of the 1.5 kkd RNA would appear to be very dependent on the degree to which the molecule is methylated. In addition to the direct comparisons given above, acrylamide gel analyses of RNA made by isolated nuclei gave results very similar to those of frog liver RNA, at least in terms of the number and molecular-weight distribution of the discrete high molecular-weight, methylated RNA fractions. This gross similarity of RNA from the cell-free and in vivo sources indicates that the isolated nuclear system produced natural RNA molecules and, in particular, that the termination mechanism was not altered (faulty) in the isolated nuclei. This is an important point, especially with regard to the RNA molecules in the 3-Oto 4.4 kkd (45 S) fractions. RNA molecules in this molecular-weight range have been described in several eukaryotic systems and have been implicated as part of the rRNA pathway (cf. Attardi & Amaldi, 1970). However, discrete 3.0 to 4.4 kkd fractions have not been observed in amphibian material previous to this study. Whether these fractions are produced only in certain types of amphibian material, or whether their presence is sometimes obscured by the high background produced by the “>5 kkd, heterogeneous, nuclear RNA” is uncertain. In our experience as well as in the experience of others, the 3.0 to 4.4 kkd fractions can not be demonstrated in all types of amphibian material. For example, in dissociated embryonic cells (cf. Landesman & Gross, 1969) and in cells under certain kinds of culture conditions (cf. Loening et al., 1969; Perry et al., 1970), the largest discrete RNA to be identified sedimented at 40 s and had a nominal molecular weight of about 2.6 kkd. The 2.6 kkd RNA previously reported for amphibian whole cells has generally been assumed to represent the primary transcription product of the rRNA genes. Because the amphibian rRNA gene appears to contain about 9 kkd of double-stranded DNA (Birnstiel et al., 1969; Dawid et al., 1970; Wensink & Brown, 1971), it has been further 3

34

J. D. CASTON AND P. H. JONES

assumed that only about one-half of the amphibian gene is transcribed. This assumption has been supported by electron micrographs of active rRNA genes (Miller & Beatty, 1969). However, detection of methylated RNA with nominal molecular weights in excess of 2.6 kkd and with gross properties similar to those of the mammalian rRNA precursor in frog liver and in isolated nuclei, forces a re-evaluation of previous interpretations of the amphibian rRNA system. In this respect, the 4.4 kkd RNA identified in these experiments is of special interest. Apart from its gross similarities to the mammalian 45 s rRNA precursor, the 4.4 kkd RNA is about the size to be expected if the entire rRNA gene were transcribed. Moreover, its over-all base composition and its level of methylation (Fig. 7, Table 4) are adequate to account for the composition of 27 s and 18 s rRNA. Furthermore, radioactivity in the 4.4 kkd RNA has been found to chase through other high moleoularweight, methylated RNA fractions into 27 s and 18 s rRNA. Thus, the labeling pattern indicates a precursor-product relationship. Because of these characteristics, we conclude that the 4.4 kkd RNA represents the primary rRNA precursor and that the 2.6 kkd RNA identified in these and in previous experiments represents but one of the intermediary precursors to RNA of cytoplasmic ribosomes. (b) Pathways for processing rRNA precursor Several schemes for processing the 4.4 kkd molecules can be deduced that will account for the methylated, high molecular-weight RNA seen in these experiments (cf. Table 3). The number of possible schemes can be reduced by assuming: (1) all 4.4 kkd molecules are identical ; (2) there is a precise number of specific cleavage sites ; (3) all of the process fragments arise by way of cleavage of larger molecules rather than by the joining together of smaller molecules (ligation); (4) all of the new 1.5 (27 s) and O-7 (18 s) kkd molecules are stable, e.g. 1.5 does not give rise to 1.2, 1.0 or 0.7 kkd molecules (see Fig. 6). A final requisite is that the processing schemes should be additive. That is, the sum of the molecular weights of the process fragments should equal the molecular weight of the precursor. Pathway A

4i:;; f?

Pathway C

4.4 kkd f-5 ;[06’[

-02

ito2

t -01 -0.j‘;9 I -02I.8 t 0.9 -03

-0,2 -0 I ‘i6 1 15 o-7 127s)

Pathway B

(IBSI

+7s! I284

j: Subsequlent processing OSinApa+hway

-01 -02+*

1

I.9 t

I.6 -01 t I.5 07 (27s) +75

(185)

f

FIG. 8. Pathways for processing rRNA precursor in isolated frog nuclei. Psthway A appears to be the dominant pathway used by isolated nuclei and accounts for all except one (3.1 kkd) of the discrete fractions listed in Table 4. Pathways B and C represent possible alternative pathways to account for the 3.1 to 3.2 kkd fraction. Direct precursors of 27 8 end 18 8 rRNA are shown in bold face type; excess rRNA fragments are given in italic type. The origin of the 7 s moiety of the 28 s duplex is uncertain.

PROCESSING

OF rRNA

PRECURSOR

::.?I

The proces&g schemes shown in Figure 8 were deduced from the nominal molecular weight of the fractions listed in Table 4 and from the labeling pattern illustrated in Figure 6. As discussed by Loening (1969) and by Weinberg 6 Penman (1970), there is some uncertainty about the accuracy of molecular-weight values as estimated by acrylamide gel electrophoresis. However, in most cases with RNA, such determinations appear to be at least as reliable as those estimated by the usual hydrodynamic methods. Even so, we have not attempted to assign actual molecular-weight values to these RNA classes. Instead, we refer to a nominal molecular weight, which we feel is adequate for deducing the major processing pathways. As with processing of rRNA in HeLa cells (cf. Weinberg & Penman, 1970), it has been necessary to postulate at least two pathways for processing rRNA in isolated nuclei to account for all of the fractions listed in Table 4. Pathway A can account for all but the 3.1 to 3.2 kkd RNA. This RNA was detected only during the first 15 to 20 minutes of synthesis in the isolated nuclei, but has been seen routinely in the RNA of adult frog liver. Thus, it appears to represent a naturally occurring class of RNA. That it was not observed in the nuclear system after 20 minutes of incorporation may mean that some component of the processing system became rate-limiting or that its particular processing system became disorganized in some way. The presence of 3.1 to 3.2 kkd RNA can be accounted for by pathways B and/or C. Pathway B, like pathway A, can account for all of the fractions but one, in this case the 3.8 kkd. Pathway C, however, can account for the 3.1 to 3.2 and the 3.8 kkd fractions, but not several other prominent fractions (2.2, 1.2 and O-9). The relative distribution of label amongst the several fractions indicates that pathways A and B operated in the isolated nuclei. One of the main features of these two pathways is that the 2.6 kkd (40 s) RNA would contain the 27 s but lack the 18 s rRNA region. In this regard, pathway C would be more consistent with the report that the 40 s RNA isolated from whole cells of Xenopus laevis contained both the 28 s and 18 s regions (Dawid et al., 1970). Although operation of any two of these pathways can account for each of the observed fractions, additional information is needed to establish unequivocally the precise manner in which rRNA is processed. Processing of the precursor molecules was very rapid in isolated nuclei. This is evident from the results given in Figure 6. The simultaneous appearance and subs+ quent accumulation of label in the 4.4 kkd and its large cleavage product (either the 3.8 or 3.1 to 3.2 kkd, depending on which pathway is used) must mean that the time between termination and processing is very short. Similar observations and conclusions have been reported for the rRNA system of Euglena gracilis, in viva (Brown & Haselkorn, 1971). Considering the short life-span of the precursor molecules and the fact that they normally fractionate in the leading edge of the heavily labeled >5 kkd nuclear RNA, it is not surprising that the precursor never appears to contain more label than some of its process products, notably those derived from its 3’ end. During the time that the precursor does contain more label than any of the process fragments, up to about 3 minutes of incorporation (Fig. 6), its radioactivity is eclipsed by that of the >5 kkd responfraction. These features of the amphibian system are, without doubt, partly sible for the difficulties encountered in demonstrating a discrete class of 45 s RNA in certain types of amphibian material. The time sequence in which individual molecular-weight classes became labeled (Fig. 6) can be used to establish the location of the 18 s and 27 s rRNA with respect to

36

J. D. CASTON

AND

P. H.

JONES

the 5’ and 3’ ends of the primary rRNA precursor. This is possible because the system was already engaged in RNA synthesis (i.e. it was pre-initiated and therefore contained a very large number of incomplete, growing precursor molecules) at the time radioactive substrate was added. Since labeled nucleoside triphosphates equilibrated within the nuclei almost instantaneously (within less than 15 seconds (unpublished observation)), the first appearance and subsequent accumulation of label in the stable process products must reflect the order in which these molecular regions were transcribed. This is true irrespective of how many different processing pathways operated, because at no time was there a significant hold-up of label in any of the putative intermediary precursors, which would have complicated interpretation of the labeling patterns. Thus, only the polarity of transcription, 5’-+ 3’ end of RNA and the over-all rate of synthesis, about O-259 kkd per minute need be considered. Accordingly, the 18 s was the first stable RNA to become labeled. It was detected almost simultaneously with its intermediary precursors, 1.2, 1.0 and 0.9 kkd, after four to five minutes of synthesis. This means that it originated from a position within about 1.2 kkd (0.259 kkd/minute x 5 minutes) of the 3’ end of the primary precursor. That the 18 s came from a position somewhat internal to the 3’ end of the precursor, had been inferred from the absence of label in the 0.7 kkd fraction before about four minutes of synthesis. If the 3’ end of the precursor contained 0.7 kkd of stable RNA, it should have been detected by 2.5 to 3 minutes of synthesis because it would have been labeled throughout most of its length. Since there was no label detected in any of the discrete, high molecular-weight fractions at this time, we conclude that most of the labeled RNA added at the growing 3’ end of the primary precursor during the first three minutes of synthesis was processed into unstable and/or low molecular-weight products. We further conclude that the 18 s RNA originated from a position about O-5 kkd internal to the 3’ end of the primary precursor. The position of the 27 s region can be approximated by similar reasoning. For example, after 7 minutes of synthesis, the primary precursor could have contained no more than 1.8 kkd of labeled RNA, all of which would be located within the 3’ portion. Since the 27 s RNA was already labeled at this time, its 3’ end must have originated from a position within something less than 1.8 kkd for the 3’ end of the precursor. (Processing required a small, but finite amount of time. Also, a certain amount of time was required to incorporate enough label into the 27 s region to allow for its subsequent detection.) Because the 18 s was located even nearer the 3’ end of the precursor, the 27 s region must have originated from a position internal to the 5’ side of the 18 s region. Whether the 27 s and 18 s regions are separated by a few thousand daltons of excess (spacer) RNA can not be deduced from present information. Transcription of rRNA gene -

-

-

Time in minutes from 3’end (termination) of complete rRNA precursor molecule 17 15 12 10 . !,,A %“A &+w-?Ls,,&-*.v>/ . .wb~“&b~ -?““%-mw +

27 s

7

5

2 :: t

18s

FIG. 9. Probable arrangement of the 27 s and 18 s rRNA regions within the rRNA gene of Rana pipkns. The position of the two rRNA regions with respect to the 3’ and 5’ ends of the gene has been inferred from the polarity of transcription, the over-all rate of RNA synthesis and the time sequence in which label appeared in the definitive rRNA molecules (see Fig. 6).

PROCESSING

OF rRNA

PRECURSOR

::‘i

As discussed earlier, the present results differ from previous reports on the rRNA system in amphibians. They differ ‘not only in the molecular size of the putative primary rRNA precursor, but also in the arrangement of the 27 s and 18 s regions within the rRNA genes. Results from experiments in whioh RNA polymerase purified from E. coli was used to transcribe DNA of rRNA genes purified from Xenopus Levis have been taken to indicate that the 18 s region of the transcription product was on the 5’ side of the 28 s region (Reeder t Brown, 1970). This would seem to imply that the order of the 27 s and 18 a regions within the genes of Xerwpus lumis and Ranu pipiens is reversed. Regardless of these apparent differences, it seems likely, even with the uncertainties inherent in our methodology, that the arrangement shown in Figure 9 represents a reasonable first approximation of the rRNA genes of Ranapipiens. The technical assistance of Mm Radka Vatev is gratefully acknowledged. This work was supported in part by U.S. Public Health Service grant HD00038 and a training grant to the Department of Anatomy. REFERENCES Attardi, G. & Amaldi, F. (1970). Ann. Rev. Biochem. 39, 183. Birnstiel, M. L., Grunstein, M., Speirs, J. & Hennig, W. (1969). Nature, 223, 1265. Bishop, D. H. L., Claybrook, J. R. & Spiegelman, S. (1967). J. Mol. Biol. 26, 373. Brown, R. D. & Haselkorn, R. (1971). J. Mol. Bill. 59, 491. Crippa, M. t Tocchini-Valentini, G. P. (1971). Proc. Nut. Acad. Sci., Wmh. 68, 2769. Dawid, I., Brown, D. D. & Reeder, R. H. (1970). J. Mol. BioZ. 51, 341. Greenberg, H. & Penman, S. (1966). J. Mol. BioZ. 21, 527. Hamilton, M. (1937). B&him. lriophys. Actu, 134, 472. Jones, P. H. (1971). Ph.D thesis. Case Western Reserve University. King, H. W. S. BEGould, H. (1970). J. Mol. BioZ. 51, 687. Kurland, C. G. (1960). J. Mol. BioZ. 2, 83. Landesman, R. & Grow, P. R. (1969). DeveZopmen&Z BioZ. 19, 244. Loening, U. E. (1969). Biochem. J. 113, 131. Loening, U. E., Jones, K. W. & Birnstiel, M. L. (1969). J. Mol. BioZ. 45, 353. Maden, B. E. H., Salim, M. & Summers, D. F. (1972). Nutwre New Biology, 237, 5. Manor, H., Goodman, D. & Stent, G. S. (1969). J. Mol. BioZ. 39, 1. Markham, R. & Smith, J. D. (1962). B&him. J. 52, 652. Martin, R. G. & Ames, B. N. (1961). J. BioZ. Chem. 236, 1372. McConkey, E. H. & Hopkins, J. W. (1969). J. Mol. BioZ. 39, 545. Miller, 0. L. & Beatty, B. R. (1969). Scielzce, 164, 955. Mittermayer, C., Braun, R. C%Rusch, H. P. (1966). B&him. biophys. Acta, 114, 536. Pene, J. J., Knight, E., Jr. & Darnell, J. E., Jr. (1968). J. Mol. BioZ. 33, 609. Perry, R. P., Cheng, T., Freed, J. J., Greenberg, J. R., Kelley, D. E. & Tartof, K. D. (1970). Proc. Nat. Acud. Sci., Wash. 65, 609. Petermann, M. L. & Pavlovec, A. (1968). Biochim. biophys. Acta, 114, 264. Randerath, E. & Randerath, K. (1964). J. Chrowu&graphy, 16, 126. Reeder, R. H. & Brown, D. D. (1970). J. Mol. BioZ. 51, 361. Shumway, W. (1942). Anat. Rec. 83, 309. Spirin, A. S. (1961). Biokhimiya, 26, 511. Standley, W. M. & Bock, R. M. (1965). Biochemistry, 4, 1302. Weinberg, R. A. & Penman, S. (1970). J. Mol. BioZ. 47, 169. Wensink, P. & Brown, D. D. (1971). J. Mol. BioZ. 60, 235. Note added in proof: A 4.4 kkd (45 s) RNA has been identified in oocytes of Xenopua &e&s and has been shown by competition-hybridization using purified rDNA to contain nucleoside sequences complementary both to 28 s and 18 s rRNA and to non-i-RNA (Crippa & Tocchini-Valentini, 1971). HeLa cell 41 s RNA has been shown to contain both

38

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28 s and 18 s sequence (Maden, Salim & Summers, 1972). Because of gross similarities between the amphibian and HeLa cell systems, this may mean that pathway C (Fig. 8) is the dominant processing pathway. However, it is not known whether the procedure uaed to enrich for 41 s RNA (i.e. poliovirus infection of HeLa cells) preferentially selected for products of pathway C.