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Temperature-sensitive RNA polymerase III (rpc160-112 and rpc160-270) mutants were analyzed for the synthesis of tRNAs and rRNAs in vivo, using a double-isotopic-labeling technique in which cells are pulse-labeled with [33P]orthophosphate and coextracted with [3H]uracil-labeled wild-type cells. Individual RNA species were monitored by Northern blot hybridization or amplified by reverse transcription. These mutants impaired the synthesis of RNA polymerase III transcripts with little or no influence on mRNA synthesis but also largely turned off the formation of the 25S, 18S, and 5.8S mature rRNA species derived from the common 35S transcript produced by RNA polymerase I. In the rpc160-270 mutant, this parallel inhibition of tRNA and rRNA synthesis also occurred at the permissive temperature (25°C) and correlated with an accumulation of 20S pre-rRNA. In the rpc160-112 mutant, inhibition of rRNA synthesis and the accumulation of 20S pre-rRNA were found only at 37°C. The steady-state rRNA/tRNA ratio of these mutants reflected their tRNA and rRNA synthesis pattern: the rpc160-112 mutant had the threefold shortage in tRNA expected from its preferential defect in tRNA synthesis at 25°C, whereas rpc160-270 cells completely adjusted their rRNA/tRNA ratio down to a wild-type level, consistent with the tight coupling of tRNA and rRNA synthesis in vivo. Finally, an RNA polymerase I (rpa190-2) mutant grown at the permissive temperature had an enhanced level of pre-tRNA, suggesting the existence of a physiological coupling between rRNA synthesis and pre-tRNA processing.
The existence of three nuclear transcription systems is documented for all eukaryotes investigated so far. RNA polymerase I synthesizes the three largest rRNAs, RNA polymerase II produces mRNAs and many noncoding RNAs, and RNA polymerase III makes tRNAs and 5S rRNA, as well as a few small noncoding RNAs. Exceptions to this transcriptional specialization are rare and mostly concern noncoding RNA species that can be produced by either RNA polymerase II or III, depending on the phylum considered (reference 39 and references therein). Given its universality, the triplication of the transcriptional apparatus must provide a major selective advantage to the eukaryotic cell, probably by facilitating the separate control of mRNA, rRNA, and tRNA synthesis in response to changes in the environment or in the cell growth rate. On the other hand, RNA polymerases I and III deliver matching amounts of tRNAs and rRNAs to the protein synthesis machinery and may thus need to operate in a closely coordinated way (references 21, 38, and 39 and references therein). In fact, the extent to which the three nuclear RNA polymerases are coordinated relative to each other remains largely undetermined, although this is presumably a key aspect of the transcriptional regulation of growth.
Yeast (Saccharomyces cerevisiae) is a particularly convenient model organism to study transcription and its regulation. Its three nuclear RNA polymerases are biochemically and genetically well characterized and contain 14, 12, and 17 subunits. Five of the subunits are common to the three enzymes and two others are shared by RNA polymerases I and III, thus providing a potential target for common regulatory controls (references 3, 5, and 42 and references therein). These common subunits are structurally conserved among eukaryotes, and the corresponding polypeptides are interchangeable in vivo between budding yeast (S. cerevisiae), fission yeast (Schizosaccharomyces pombe), and humans (20, 27, 28, 29). Yeast is also the only eukaryote from which temperature-sensitive mutants are available for each of the the three transcription enzymes (12, 22, 40). These mutants provide a unique opportunity to assess the interdependency of the three RNA polymerases by using such mutants to separately turn off each RNA polymerase in vivo and to examine how this affects the physiological activity of the other two transcription systems.
Yeast strains are listed in Table Table1,1, and their growth patterns on YPD (1% yeast extract, 2% Bacto Peptone, and 2% glucose) are shown in Fig. Fig.1.1. In vivo labeling with 33Pi and [3H]uracil was done in low-phosphate medium (YPD*) and in casein hydrolysate medium lacking uracil (25, 28), respectively. Growth was monitored with a Hach Ratio/XR turbidimeter. One turbidimetry unit corresponds to about 2 × 108 haploid cells per ml (for the wild-type strain W303-1b) and to an absorbance of 7.2 at 600 nm. Wild-type cells grown on YPD and YPD* had a doubling time of 110 min at 30°C, but growth was slightly slower under the conditions of 3H labeling (doubling time, 130 min).
The rpc160-112 and rpc160-270 mutants are well-characterized RNA polymerase III mutants that inhibit transcription in vitro (9, 34). The rpb1-1 (strain RY260) mutant is a tight conditional mutant of RNA polymerase II that rapidly stops mRNA synthesis when shifted to 37°C (22). Our data (see Fig. Fig.2)2) indicate a slower decrease in mRNA accumulation than in the original report. The presence of an extragenic suppressor of rpb1-1 in our RY260 isolate was ruled out by appropriate genetic crosses, but the possibility of a mild intragenic suppressor cannot be excluded. The rpb1-1 mutant also rapidly inhibits rRNA and tRNA synthesis at 37°C (22) (data not shown). The rpa190-2 mutant is an RNA polymerase I mutant with a strong temperature-sensitive growth defect (40) (Fig. (Fig.1).1). Yet our in vivo labeling data show that its RNA polymerase I defect, already quite strong at 25°C, is not much increased at 37°C (see Fig. Fig.33).
A total of 250 μCi of 33Pi (at 10 μCi/μl) was added to 5-ml log-phase cultures that were further grown for 10 min before the addition of 20 ml of ice-cold water. The culture was spiked with a 100-μl aliquot of wild-type cells labeled with [3H]uracil (see below). Labeling data were expressed by measuring the ratio between 33Pi- and 3H-labeling signals. The external 3H control notably improves the quantification of the 33Pi pulse-labeling assay, since it bypasses experimental variations in the efficiency of RNA extraction or recovery, assuming that the cells behave similarly during the extraction and RNA purification procedures. It also minimizes experimental artifacts related to gel loading and electrophoresis. Cells were harvested by centrifugation, washed twice with ice-cold water, frozen in an ice-ethanol bath, and stored at −80°C. 33Pi uptake, measured by counting the radioactivity left in the culture supernatants, ranged between 30 and 60% of the exogenous 33Pi. The reference sample of [3H]uracil-labeled wild-type cells (strain OG27GF) (Table (Table1)1) was prepared by adding 5 mCi of [3H]uracil (at 1 mCi/ml) to a 100-ml culture of cells grown exponentially at 30°C, at an optical density of 0.24 at 600 nm. Cells were further grown for 1.5 h, leading to a 95% incorporation of [3H]uracil, harvested by centrifugation, washed twice with ice-cold water, and resuspended in 20 ml of water. They were dispatched in 100-μl samples frozen in ethanol-dry ice and stored at −80°C.
Cell cultures (5 ml) were pelleted by centrifugation, suspended in 0.5 ml of 50 mM sodium acetate buffer (pH 5.3) with 10 mM EDTA and 1% sodium dodecyl sulfate, and mixed with an equal volume of buffered phenol prewarmed at 65°C. Cells were broken in an Eaton press with a homemade device or by extraction with 200 μl of glass beads. Comparable yields of RNA were obtained by either method. For the glass bead extraction, cells went through five consecutive cycles of vigorous vortexing at room temperature (2 min) and transfer to a 65°C bath (1 min) before being frozen in liquid nitrogen and thawed at 65°C. The whole procedure was done twice. After centrifugation (15,000 × g) for 20 min at 4°C and extraction in 0.5 ml of phenol-dichloromethane-isoamyl alcohol (25:24:1), nucleic acids were precipitated in 2.5 volumes of ethanol and 0.2 volume of 10 M LiCl, rinsed with 70% ethanol, and dissolved in 50 μl of RNase-free water treated with diethyl pyrocarbonate. This was treated for 2 h at 37°C in the presence of alkaline phosphatase (5 U/μg of RNA). After phenol extraction and ethanol precipitation, RNA was dissolved in 50 μl of RNase-free water and stored at −20°C. RNA radioactivity was measured by precipitating 1 μg of RNA with 4 μg of carrier tRNA in 1 ml of 5% trichloroacetic acid. After 2 h on ice, the RNA precipitate was filtered through a 0.45-μm-pore-size membrane (Millipore) and counted in biodegradable counting scintillant (Amersham Pharmacia Biotech). A specific radioactivity of about 10−2 μCi/μg of RNA was obtained in all cases.
RNA (10 μg) was separated by electrophoresis on 6% polyacrylamide–8 M urea gels and autoradiographed with Kodak Biomax MR film. In the case of the two large rRNA species, 1 μg of RNA was loaded onto 1% agarose denaturing formaldehyde gels. The gels and their autoradiograms were superimposed to locate the major stable RNA species (tRNAs and 5S, 5.8S, 18S, and 25S rRNA). This was facilitated by using 33P rather than 32P labeling, due to the superior resolution of the autoradiographic signals. The corresponding gel positions were punched with an awl, generating 2.3-mm-diameter spots, as illustrated in Fig. Fig.3.3. This material was rehydrated for 30 min in 25 μl of water and left overnight at room temperature in 0.5 ml of NCS-II tissue solubilizer (Amersham Pharmacia Biotech). Samples were equilibrated for 1 week in the dark in 1 ml of BCS-NA scintillant, and 33Pi and 3H activity was counted. Spots located between the autoradiographic signals were collected and measured in the same way, providing a measure of the background level of radioactivity. The signal-to-noise ratio was always higher than 10-fold (and usually close to 100-fold) for all experimental data presented here.
RNA levels were determined by Northern blot analysis (except for PEP4 mRNA; see below), using standard conditions. Briefly, 10 μg of RNA was separated on a 1% agarose denaturing formaldehyde gel, blotted onto a nylon membrane, and hybridized overnight to radiolabeled oligonucleotide probes in 0.5 M sodium phosphate buffer (pH 7.2) with 10 mM EDTA and 7% sodium dodecyl sulfate. ACT1, CYH2, and NME1 RNAs were hybridized to 5′-TGAAGAAGATTGAGCAGCGGTTTG-3′, 5′-CATGTTAATTCTGTGGTGATGTTGAC-3′, and 5′-CGTCATAACTATGGTTTAG-3′ probes, and PEP4 mRNA was quantified by reverse transcription-PCR (RT-PCR) of RNA from mutant or wild-type cultures spiked with a small aliquot of wild-type cells (strain OG27GF), as described above for the in vivo double-labeling assay. OG27GF has a deletion of the PEP4 gene and harbors the GFP gene (Table (Table1).1). GFP mRNA, amplified from the 5′-GTAACAAGACTGGACCATCACC-3′ and 5′-GGTGAAGGTGATGCTACTTACGG-3′ primers, served as an RNA recovery marker of the PEP4 mRNA, which was amplified from the 5′-GACCGGTCCAACCCTTCTTGG-3′ and 5′-GGTTCCTTGGCTTGTTTCC-3′ primers. One microgram of total RNA was reverse transcribed for 1 h at 42°C with 100 pmol of appropriate oligonucleotide primers. The RT-PCR amplification signals were directly proportional to the amount of RNA, over a range of 0.1 to 10 μg. RT was stopped by adding 180 μl of water to the 20-μl reaction volume. Ten-microliter samples were amplified by PCR (15 cycles) in the presence of 25 μCi of [α-32P]dCTP, using 10 pmol of the corresponding oligonucleotide primers. A sample of 5 μl of each reaction product was loaded on a 6% polyacrylamide–8 M urea gel, dried, and analyzed with a Molecular Dynamics PhosphorImager.
Unlike the RNA polymerase II rpb1-1 mutant, RNA polymerase I (rpa190-2) and III (rpc160-112 and rpc160-270) mutants continue to grow for at least 6 h after the temperature shift (Fig. (Fig.1)1) and thus have little effect on the synthesis of essential mRNAs. This was confirmed by measuring the levels of individual RNA polymerase II transcripts such as the PEP4, ACT1, and CYH2 mRNAs and the RNA of RNase MRP encoded by NME1 (Fig. (Fig.2).2). Likewise, cells that are deprived of the largest subunit of RNA polymerase I (by controlling its transcription with the galactose-repressible GAL1 promoter) have little effect on the synthesis of several ribosomal proteins (41). Thus, the level of RNA polymerase II-dependent transcription in vivo is largely uncoupled from the activity of the other two transcription enzymes.
RNA polymerase III (rpc160-112 and rpc160-270) mutants distinctly impair tRNA synthesis at 25°C (consistent with the detectable growth defect at this temperature) and completely prevent it at 37°C, as shown by in vivo labeling data (Fig. (Fig.3).3). Furthermore, Northern hybridization (Fig. (Fig.4)4) shows that the rpc160-112 mutant strongly reduces the steady-state level of pre-tRNALeu3 relative to the mature tRNALeu3. They also reveal a marked depletion in the SCR1 RNA component of the signal recognition particle. This RNA is predicted to be an RNA polymerase III transcript because of the presence of a typical RNA polymerase III terminator at its 3′ end (10). Previous experiments based on [3H]uracil pulse-labeling (32) indicated that RNA polymerase III mutants hardly affect 5S rRNA synthesis in vivo, despite overwhelming evidence that the latter is made by RNA polymerase III in vitro (26, 35). The more quantitative double-label technique used here shows that rpc160-112 and rpc160-270 cells distinctly affect 5S rRNA synthesis, albeit less than tRNA synthesis, thus reconciling the in vitro and in vivo data (Fig. (Fig.33 and and5A).5A).
Beyond their effect on RNA polymerase III transcripts, rpc160-112 and rpc160-270 cells also strongly reduce the synthesis of the 5.8S, 18S, and 25S rRNAs, which derive from a common single transcript made by RNA polymerase I. In the case of the rpc160-112 mutant, a shift to the restrictive temperature (37°C) leads to a tight adjustment of the de novo synthesis of large rRNAs in response to the temperature-sensitive RNA polymerase III defect. As seen in Fig. Fig.5B,5B, the relative rates of tRNA and rRNA synthesis were down to the wild-type level within 3 h after the shift, i.e., well before growth arrest (Fig. (Fig.1).1). In the case of the rpc160-270 mutant, this pleiotropic effect on rRNA synthesis also occurs at 25°C.
Since a 10-min pulse with 33Pi is close to the time needed to process pre-rRNA in vivo (36), our labeling data do not distinguish between transcriptional and posttranscriptional effects on rRNA biogenesis. Previous work from this laboratory suggests that RNA polymerase III defects may impair pre-rRNA processing in vivo. Thus, the rpc160-112 mutant and another conditional (rpc160-41) mutant (12) have a mild effect on the maturation of 5.8S rRNA at the semipermissive temperature of 30°C (13). Moreover, yeast mutants specifically defective in the biogenesis of 5S rRNA accumulate the 27S pre-rRNA precursor of 25S rRNA, with no effect on 20S, the precursor of 18S rRNA (7) (Fig. (Fig.4).4). rpc160-112 and rpc160-270 cells had a symmetrical effect on rRNA processing and distinctly enhanced the level of 20S pre-rRNA. In the rpc160-112 mutant, this occurred only at 37°C, while the rpc160-270 mutant had this effect at both temperatures (Fig. (Fig.4).4). Hence, the effect of these two mutants on 18S rRNA synthesis (as measured by pulse-labeling) correlates with, and at least partly results from, an rRNA processing defect.
Our Northern hybridization data also show a good match between the steady-state levels of mature rRNAs and tRNAs and their rates of synthesis as predicted by in vivo labeling data (Fig. (Fig.5C).5C). The rpc160-112 mutant has an almost threefold deficit in tRNAs when grown at 25°C, consistent with its limited effect on rRNA synthesis under these conditions. rpc160-270 cells accumulate tRNAs and rRNAs in the same ratio as wild-type cells, as expected from their coordinated effects on tRNA and rRNA synthesis, even when grown at 25°C. The allele-specific difference consistently observed between the rpc160-112 and rpc160-270 mutants is puzzling, given that they were constructed in the same isogenic background and have similar growth patterns (Fig. (Fig.1).1). Other RNA polymerase III mutants behave like the rpc160-112 mutant in the sense that they have a deficit in tRNA at the permissive temperature (12, 32) (data not shown), and there may thus be something special about the rpc160-270 mutant. A cryptic suppressor mutant seems unlikely, given the perfectly regular 2:2 segregation of its temperature-sensitive growth defect in meiotic crosses (data not shown). We note, however, that the rpc160-112 mutant has a direct catalytic defect (9) with a rapid transcriptional arrest at 37°C (Fig. (Fig.3),3), whereas the elongational defect of the rpc160-270 mutant correlates with a high level of the cleaving RNase activity of RNA polymerase III (34) and a somewhat delayed transcriptional arrest in vivo.
Given that RNA polymerase III mutants affect the overall rate of rRNA synthesis and also interfere with pre-rRNA processing, we wondered if RNA polymerase I mutants might have a reciprocal effect on tRNAs. In vivo labeling data suggest that this may be the case (compare the rates of tRNA and rRNA synthesis in rpa190-2 cells in Fig. Fig.3)3) but are inconclusive, because the transcriptional defect of the rpa190-2 mutant, already quite strong at 25°C, is hardly aggravated at 37°C (a similar situation was observed for the rpa190-1 mutant [data not shown]). Yet, rpa190-1 and rpa190-2 cells are strongly temperature sensitive in terms of growth, suggesting that rRNA synthesis may be especially growth limiting at 37°C. Northern hybridization data, however, show that rpa190-2 cells grown at 25°C have a high level of pre-tRNALeu3 (Fig. (Fig.4)4) and thus interfere with pre-tRNA processing, perhaps in relation to the nucleolar localization of tRNA processing enzymes (1). We cannot rule out the possibility that there is, in addition, some effect on the transcriptional synthesis of tRNAs, as is indeed suggested by the partial drop in the pre-tRNA/tRNA ratio when rpa190-2 cells are shifted to 37°C. However, the high content of 7SL RNA (SCR1) found in rpa190-2 cells at both temperatures argues against a general and massive effect on RNA polymerase III-dependent transcription.
Yeasts are fast-growing cells that invest a substantial amount of metabolic energy in ribosome biogenesis and may therefore need to precisely adjust the synthesis of rRNAs, tRNAs, and ribosomal proteins as a function of the growth rate. This regulation is fairly well understood as far as ribosomal proteins are concerned (38), but comparatively little is known of the control of tRNA and rRNA synthesis. In particular, the extent to which yeast RNA polymerases I and III are coregulated relative to each other and to the transcriptional synthesis of ribosomal proteins is still a moot point. In fact, the main evidence for coordinated control of the transcriptional synthesis of rRNA, ribosomal protein mRNAs, and tRNAs is that blocking protein secretion inhibits these three processes in a way that requires protein kinase C (19). On the other hand, rRNA and tRNA synthesis can be uncoupled under physiological conditions, such as amino acid starvation (6, 23). Even under balanced growth conditions, the cellular levels of tRNAs and rRNAs are roughly but not strictly constant, since rRNAs are more strongly affected than tRNAs in slow-growing cells (15, 32, 37). Finally, conditional mutants of RNA polymerase I or III have no effect on the transcription of ribosomal protein genes by RNA polymerase II (reference 41 and this study), showing that there is no obligatory link between the transcriptional synthesis of rRNA and of ribosomal protein mRNAs.
We show here that RNA polymerase III mutants turn off the formation of the three large rRNA species (25S, 18S, and 5.8S) in parallel to the reduced rate of tRNA synthesis, thereby adapting the flux of newly synthesized rRNA to the low level of tRNA synthesis and keeping the rRNA/tRNA steady-state ratio at the wild-type level. An obvious concern is that this could somehow be the indirect result of a common dependency on growth rate. The fast response of rpc160-112 cells when shifted to 37°C argues against this interpretation, since they reach a low rate of rRNA synthesis within one doubling time, well before growth arrest is observed. This coordinated synthesis of tRNAs and rRNAs could partly involve transcriptional effects, as in secretion-defective cells (19), and may perhaps also reflect changes in RNA turnover. However, our data strongly suggest an additional effect on pre-RNA processing, as shown by the increase of 20S pre-rRNA observed in the rpc160-112 and rpc160-270 mutants. This is consistent with previous data showing that RNA polymerase III mutants grown at 30°C have minor but distinct effects on pre-rRNA processing (13). Conversely, we also observed that RNA polymerase I mutant cells accumulate a high level of pre-tRNALeu3 and thus probably interfere with tRNA processing.
The mechanism by which RNA polymerase III may control pre-rRNA processing is unknown. One possibility is that a hypothetical RNA polymerase III holoenzyme (5) may contain or contact nucleolar proteins participating in pre-rRNA processing. This could arguably account for the allele-specific differences in the rRNA processing defects of the rpc160-112 and rpc160-270 mutants at 25°C, as these mutants are thought to have a different effect on the conformation of the elongating RNA polymerase III complex (34). Alternatively, RNA polymerase III transcripts could directly participate in pre-rRNA processing. This is the case for U3 snRNA in plants (16) or RNase MRP RNA in mammals (43), but the yeast counterparts are made by RNA polymerase II (reference 14 and this work). RNase P RNA is another candidate, as it affects 5.8S rRNA maturation in vivo (4) and is an RNA polymerase III transcript in organisms ranging from yeasts (17) to humans (2). Moreover, its high dosage partly suppresses a mutant defective in the RNA polymerase III initiation factor TFIIIC (18). Finally, yeast 5S rRNA mutants interfere with pre-rRNA processing, providing another link to RNA polymerase III (7). Native 5S rRNA is short-lived (probably reflecting its lack of nucleotide modification) (33) unless it is complexed by yeast ribosomal protein L1 (8). It could therefore operate as a sensor, stimulating pre-rRNA processing in response to RNA polymerase III activity. Unlike RNA polymerase III mutants, however, 5S rRNA mutants mainly interfere with 25S rRNA maturation, with little effect on 18S rRNA (7) (Fig. (Fig.44).
In human cells, transcriptional controls over tRNA and rRNA synthesis are probably critical to the (de)regulation of differentiated cell growth upon viral infection or tumorigenesis, as shown by the inhibitory effect of the retinoblastoma and p53 tumor-suppressing factors on RNA polymerases I and III (reference 39 and references therein). Our observation that yeast cells adjust pre-rRNA processing as a function of RNA polymerase III activity extends the repertoire of homeostatic controls of ribosome synthesis (21, 38). It would be interesting to know if a similar situation exists in human cells. Moreover, U6 snRNA and the signal recognition particle RNA are made by RNA polymerase III in organisms ranging from yeasts to humans, thus relating RNA polymerase III activity to mRNA splicing and cotranslational protein secretion. Taken together, these data underscore the highly pleiotropic role of RNA polymerase III in modulating the main steps of RNA and protein synthesis.
We thank Jean Labarre and Jean-Marie Buhler for useful suggestions, Michel Werner and anonymous reviewers for improving the manuscript, and André Sentenac for his kind support.
J.-F.B. had a fellowship from the Fondation de la Recherche Médicale, F.N. held a European Marie Curie Fellowship, and O.G. was supported by the Institut de Formation Supérieure Biomédicale. This work was partly funded by the European Training and Mobility Program (grant FMRX-CT96-0064).