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Molecular and Cellular Biology, April 2007, p. 2886-2896, Vol. 27, No. 8
0270-7306/07/$08.00+0     doi:10.1128/MCB.00054-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Regulation of rRNA Synthesis by TATA-Binding Protein-Associated Factor Mot1{triangledown}

Arindam Dasgupta,1,{dagger} Rebekka O. Sprouse,1,{dagger} Sarah French,2,{dagger} Pavel Aprikian,3 Robert Hontz,1 Sarah A. Juedes,1 Jeffrey S. Smith,1 Ann L. Beyer,2 and David T. Auble1*

Department of Biochemistry and Molecular Genetics,1 Department of Microbiology, University of Virginia Health System, 1300 Jefferson Park Avenue, Charlottesville, Virginia 22908-0733,2 Department of Microbiology, University of Washington, 1959 NE Pacific St., Seattle, Washington 981953

Received 10 January 2007/ Accepted 1 February 2007


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ABSTRACT
 
Mot1 is an essential, conserved, TATA-binding protein (TBP)-associated factor in Saccharomyces cerevisiae with well-established roles in the global control of RNA polymerase II (Pol II) transcription. Previous results have suggested that Mot1 functions exclusively in Pol II transcription, but here we report a novel role for Mot1 in regulating transcription by RNA polymerase I (Pol I). In vivo, Mot1 is associated with the ribosomal DNA, and loss of Mot1 results in decreased rRNA synthesis. Consistent with a direct role for Mot1 in Pol I transcription, Mot1 also associates with the Pol I promoter in vitro in a reaction that depends on components of the Pol I general transcription machinery. Remarkably, in addition to Mot1's role in initiation, rRNA processing is delayed in mot1 cells. Taken together, these results support a model in which Mot1 affects the rate and efficiency of rRNA synthesis by both direct and indirect mechanisms, with resulting effects on transcription activation and the coupling of rRNA synthesis to processing.


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INTRODUCTION
 
rRNA synthesis is tightly coupled to cellular growth rate (36, 40, 62). rRNA accounts for the bulk of total RNA synthesis in exponentially growing cells but is dramatically down-regulated during nutrient limitation or stress. Budding yeast cells contain ~140 copies of the ribosomal DNA (rDNA) repeat, approximately half of which are transcriptionally active during exponential growth (11, 19). Recent evidence supports the hypothesis that the yeast regulatory response to the growth state is mediated by regulation of the number of active RNA polymerase I (Pol I) molecules rather than the number of active gene repeats (9, 19, 35). Therefore, a mechanistic understanding of how Pol I initiation is controlled is of fundamental importance for understanding how cells adjust their overall metabolic state in response to changes in extracellular nutrient levels.

In Saccharomyces cerevisiae, the Pol I preinitiation complex (PIC) assembles via cooperative interactions between TATA-binding protein (TBP), Rrn3, the multisubunit upstream activating factor (UAF), the core factor (CF), and Pol I itself (see references 3, 41, and 61 and references therein). UAF associates stably with the Pol I promoter and recruits CF and TBP, which provide a substrate for the Pol I-Rrn3 complex, which is the competent form of the polymerase (3, 7, 27, 28, 34, 35, 56). Association of Pol I-Rrn3 with the PIC also stabilizes the association of CF with the promoter, suggesting a model in which the CFs dissociate from the promoter following departure of Pol I and Rrn3 as a result of productive initiation. In this way, UAF has been proposed to facilitate multiple rounds of transcription characterized by cycles of association and dissociation of the basal machinery with the promoter (3).

Several different molecular mechanisms underlying Pol I transcriptional control have been described. The activity of the Pol I-associated factor Rrn3 is regulated by the cellular growth stage (9, 35). The activity of the mammalian homolog of Rrn3, TIF-1A, is also regulated by nutrient availability (33) and by stress (32). In mammalian cells, growth factor-mediated activation of rDNA transcription occurs by stimulation of transcriptional elongation rather than enhanced recruitment of Pol I (55). In yeast, the histone deacetylase Rpd3 participates in "closing" active rDNA repeats as cells enter stationary phase (48).

Despite the fact that TBP is required for transcription by RNA Pol I, RNA polymerase II (Pol II), and RNA polymerase III, TBP-associated factors typically have functions dedicated to just one transcription system (24, 29, 36, 41, 50). Mot1 is an essential TBP-associated factor in Saccharomyces cerevisiae that forms a complex with TBP that is distinct from other well-characterized TBP-containing complexes, such as TFIID (4, 16, 45). In vitro, Mot1 can displace TBP from TATA box-containing DNA in an ATP-dependent reaction, and extracts made from mot1 cells display higher levels of RNA Pol II-dependent transcriptional activity than extracts from wild-type cells (4). Genome-wide analyses have defined a broad role for Mot1 in control of Pol II-dependent genes, including repression of stress response-, diauxic shift-, and mating type-specific genes (1, 13, 23, 63). Consistent with Mot1's biochemical activity in vitro, TBP occupancy of Pol II promoters increases in mot1 cells (14). However, paradoxically, in wild-type cells, Mot1 occupancy of many promoters increases in proportion to promoter activity, suggesting that Mot1 may impose a limit on the extent of Pol II activation or perhaps Mot1 possesses an alternative biochemical activity that is distinct from TBP-DNA dissociation (1, 14, 23, 63). Here we expand the spectrum of Pol I transcriptional regulatory mechanisms by reporting a role for Mot1 in Pol I transcription and rRNA processing. The effect of Mot1 on Pol I transcription cannot be simply explained by Mot1's TBP-DNA dissociating activity, suggesting that Mot1 may have a novel function at the rDNA. Moreover, these results suggest that by coordinating the expression of both Pol I and Pol II genes, Mot1 plays a fundamental role in regulating the transcriptional response to the cellular growth state.


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MATERIALS AND METHODS
 
Yeast strains and growth conditions. Congenic wild-type and mot1 yeast strains used in this study have been described previously (12, 16). Haploid cells expressing chromosomal copies of FLAG-tagged Rrn5, Rrn7, Rrn9, or Rpa135 under endogenous promoter control were a generous gift from Beth Moorefield (Fred Hutchinson Cancer Research Center). These strains were mated to a haploid MOT1 shuffling strain (12), and haploid segregants were obtained that expressed the indicated FLAG-tagged construct in the MOT1 shuffling background. Hemagglutinin (HA)-tagged Utp8, Utp9, and Utp10 strains (17) were crossed to the same MOT1 shuffling strain, and haploid segregants were obtained that expressed the indicated HA-tagged Utp protein in the MOT1 shuffling background. For Northern blotting and chromatin immunoprecipitation (ChIP) experiments comparing wild-type cells to mot1-14 or mot1-42 cells, strains were grown in yeast extract-peptone-dextrose (YPD) or selective medium at 30°C to an optical density at 600 nm (OD600) of about 1.0. Cells were harvested for isolation of total RNA or were formaldehyde treated for ChIP experiments. For some Northern experiments, cells were rapidly shifted to 35°C by addition of an equal volume of prewarmed 40°C medium. A yeast strain expressing C-terminal Myc epitope-tagged Mot1 was obtained by transformation with a PCR-generated DNA cassette as described previously (30). The correct integration event at the MOT1 locus was confirmed by PCR and Western blotting.

Miller chromatin spreads of yeast cells. Starter cultures of yeast strains were grown at 30°C in YPD for 5 to 6 h before being diluted ~1:100 into YPD plus 1 M sorbitol and grown to mid-log phase (A600 = 0.3 to 0.5). One-milliliter samples were then transferred to prewarmed (30°C) tubes containing 5 mg zymolyase (20T; USBiological), rapidly mixed, and shaken at 30°C for 4 min. After this time, samples were transferred to a 1-ml microcentrifuge tube and centrifuged at maximum speed in a tabletop microcentrifuge for 10 s. The supernatant fluid was then withdrawn, the pellet was resuspended in 1 ml 0.025% Triton X-100, pH 9, and the resuspended cells were pipetted into an additional 3-ml volume of the Triton X-100 solution to ensure hypotonic lysis of the cells. All steps following the incubation with zymolyase were carried out as rapidly as possible. Cellular contents were then allowed to disperse in the Triton X-100 solution at room temperature for 20 to 60 min before a 1/10 volume of 0.1 M sucrose 10% formalin solution (pH 9) was added to the dispersed cell contents. Aliquots (70 µl) were centrifuged onto carbon-coated copper electron microscope (EM) grids at 10,000 rpm for 10 min in an HB-4 rotor (42). The grids were then stained with phosphotungstic acid and uranyl acetate and viewed in a JEOL 100 CX transmission electron microscope.

Analysis of polymerase density. Entire EM grids were scanned, and all rRNA genes visible were photographed. The number of RNA Pol I molecules (or nascent transcripts) associated with each gene that could be unambiguously traced from the 3' to the 5' end was determined.

Psoralen cross-linking assays. Psoralen cross-linking assays were performed as described previously (11, 48, 53). Stationary-phase cultures were inoculated into 50 ml fresh YPD to an OD600 of ~0.2 to 0.3 and shaken at 30°C for 4 h to an OD600 of ~1.0. Aliquots of ~1 x 108 cells were removed, washed in ice-cold 1x Tris-EDTA (TE), and frozen in liquid nitrogen. For cross-linking, cells were resuspended in 0.7 ml ice-cold 1x TE and placed in individual wells of a 24-well tissue culture plate; 35 µl of a 200-µg/ml solution of 4,5',8-trimethylpsoralen (Sigma) in ethanol were added to each well, and cells were irradiated on ice using a UV lamp (model B-100A; UV Products, Inc.) for 5 min at a distance of ~6 cm. This UV cross-linking process was repeated four more times. The cells were then lysed by addition of acid-washed glass beads and homogenization for 45 s at 4°C in a mini-bead beater, followed by treatment with 20 mg/ml proteinase K for 3 h at 50°C, phenol-chloroform extraction, and finally, ethanol precipitation. The pellet was resuspended in 1x TE. Five micrograms of DNA was digested with EcoRI for 5 h at 37°C and then resolved on a 1.3% Tris-borate-EDTA-agarose gel, transferred to a nylon membrane (Millipore Immobilon-NY+), and probed with a 32P-labeled XbaI rDNA fragment from pSB694 (8) that contains most of the 35S rDNA region.

RNA isolation and Northern blotting. Total RNA was isolated from cells by a hot acid-phenol extraction protocol (49). For Northern blots, 20 µg total RNA was resolved by electrophoresis on 1.2% formaldehyde-agarose gels and transferred to a Nytran membrane (Schleicher and Schuell). The Nytran membrane was hybridized with the 32P-labeled probe JS45 (48), which hybridizes to the 5' external transcribed spacer of the 35S rRNA. Hybridizations were performed for 90 min in Quikhyb solution (Stratagene) at 65°C. Blots were washed twice for 5 min each at room temperature with 2x SSC (1x SSC is 0.15 M NaCl plus 0.015 M sodium citrate)-0.1% sodium dodecyl sulfate (SDS) and twice more at 60°C and with 0.1x SSC-0.1% SDS. The ACT1 probe was derived by random priming using a PCR-generated portion of the ACT1 open reading frame and was used for Northern analysis as described previously (13).

In vivo RNA labeling and analysis. Wild-type and mot1 yeast cells were grown to an OD of ~0.3 to 0.4 in synthetic medium lacking methionine at 30°C. Cells were then resuspended in fresh prewarmed synthetic medium lacking methionine, incubated at 30°C for an additional 10 min, and then pulse-labeled with [methyl-3H]methionine (60 µCi/ml; 83 mCi/µmol) for 2 to 4 min at 30°C. A 1-ml aliquot of cells was removed at this point and frozen in liquid nitrogen ("time zero"). The label was then chased with the addition of unlabeled methionine (500 µg/ml). At various times, 1-ml aliquots were taken and cells were rapidly frozen in liquid nitrogen. The frozen cells were subsequently thawed and washed with ice-cold water, and total RNA was prepared by a hot acid-phenol extraction procedure (49). Labeled RNA was separated on 1.5% formaldehyde-agarose gels in 1x morpholinepropanesulfonic acid buffer. Subsequently, gels were submerged in the Amplify fluorographic reagent (Amersham) for 30 min and then dried on Whatman 3 MM filter paper using a gel drier at a low temperature. Dried gels were exposed to autoradiographic film for 2 to 14 days.

ChIP. ChIP was performed similarly to the procedure described previously (14). Cells were grown as described above and then treated with 1% formaldehyde for 15 min. Glycine was added to a final concentration of 125 mM, and the cultures were further incubated for 5 min. Cells were then washed once with cold TBS (20 mM Tris-HCl [pH 7.4], 150 mM NaCl) with 125 mM glycine, followed by a wash with TBS without glycine. Cells were frozen in liquid nitrogen and stored at –80°C for later analysis. Cell pellets were resuspended in 600 µl FA lysis buffer (50 mM HEPES-KOH [pH 7.5], 140 mM NaCl, 1 mM EDTA, 0.1% sodium deoxycholate, 1% Triton X-100) containing Roche "complete" protease inhibitor cocktail. The resuspended cell suspension was then mixed with an equal volume of acid-washed glass beads (425 to 600 µm), and cells were disrupted at 4°C using a FastPrep FP120 device (Bio Savant). Cell lysates were then sonicated to yield an average DNA fragment size of ~500 bp, and the sonicated material was clarified by centrifugation at 14,000 rpm for 30 min in a microcentrifuge. For ChIP analysis, chromatin protein was measured by a Bio-Rad protein assay using bovine serum albumin as the standard, and equal amounts of protein (1 to 2 mg) were immunoprecipitated overnight with 6 µl of TBP or TFIIB rabbit polyclonal antiserum (14) or with 5 µg of 9E10 anti-Myc monoclonal antibody. ChIP for Pol II was performed using 6 µl of RNA Pol II monoclonal antibody 8WG16 (58). The reactions were then incubated with 60 µl of protein A-Sepharose beads equilibrated in FA lysis buffer. ChIP for HA- and FLAG-tagged proteins was performed using 40 µl antibody-coupled beads (HA beads from Roche and FLAG beads from Sigma) in FA lysis buffer. The bead-bound immune complexes were recovered by centrifugation and washed twice each with 1.0 ml of FA lysis buffer, 1.0 ml of FA lysis buffer with a high salt concentration (50 mM HEPES-KOH [pH 7.5], 500 mM NaCl, 1 mM EDTA, 0.1% sodium deoxycholate, 1% Triton X-100), 1.0 ml LiCl wash buffer (10 mM Tris-HCl [pH 8.0], 250 mM LiCl, 0.5% NP-40, 0.5% sodium deoxycholate, 1 mM EDTA), and TE (10 mM Tris [pH 8.0], 1 mM EDTA). The immunoprecipitated material was eluted twice with 50 mM Tris [pH 8.0], 1% SDS, and 10 mM EDTA. The eluted material was incubated at 65°C overnight and purified using a PCR purification kit (QIAGEN). Quantitative PCR was performed using 1/100 to 1/50 of the material recovered after the immunoprecipitation or 1/500 to 1/10,000 of the input DNA. In all cases, titrations were performed to ensure that the yield of the PCR product was linearly related to the amount of added template. The primers for NTS2, 25S, and 18S were derived from regions as depicted in Fig. 5. The primers used to detect association with the 5' external transcribed spacer were identical to those used previously (20). The sequences of other primers are available on request. To account for the difference in gene copy number of the rDNA (~140 copies) and Pol II-dependent (single copy) genes, ChIP at the rDNA was quantified by performing 22 cycles of PCR, which pilot experiments showed was at the approximate midpoint of the linear response range for the assay. ChIP of Pol II promoters was quantified by performing 27 cycles of PCR, the midpoint of the linear response region for single-copy genomic sequences under our conditions. For Mot1 ChIP experiments, the relative intensities were normalized to band intensities of INO1, a Mot1-regulated gene (13). For TBP, TFIIB, and Pol II ChIP experiments, relative intensities were normalized to the band intensities of ACT1.


Figure 5
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  		FIG. 5. Mot1 is localized to rDNA. (A) Schematic representation of the rDNA in S. cerevisiae showing the initiation site and direction of 35S transcription by Pol I (arrow). Relative positions of NTS2, 18S, and 25S DNA amplified in ChIP experiments are shown by black bars. (B) ChIP analysis of Mot1-Myc binding to the indicated chromosomal loci. Relative ChIP signal obtained from three experiments is shown, normalized to the level of Mot1 associated with the Mot1-repressed INO1 promoter (1.0). The GAL1 promoter is a negative control. Note that in this and other ChIP experiments, PCR of rDNA was performed for 22 cycles whereas PCR of single-copy Pol II promoters (INO1 and GAL1 in this case) was performed for 27 cycles; these represent the midpoints of the linear response range of the ChIP assay. (C) TBP ChIP was performed using chromatin from wild-type (WT), mot1-14, or mot1-42 cells. TBP occupancy of the indicated loci is shown relative to that of the ACT1 promoter in wild-type cells (1.0). The data represent the mean values ± standard deviation determined from two experiments. (D) ChIP was performed for TFIIB as for panel C. (E) ChIP was performed as for panel C to determine the relative Pol II large subunit association with the indicated chromosomal regions. Each result represents the mean ± standard deviation obtained from two experiments using independently prepared chromatin samples.

In vitro analysis. In vitro transcription was performed as described previously (3) using a wild-type rDNA promoter fragment immobilized on magnetic beads and whole-cell extracts from wild-type or mot1-1 cells (16). In brief, extracts were prepared in parallel from identical numbers of cells. Equal volumes of extract (indicated in the legend to Fig. 9A) were incubated with promoter DNA for 45 min at room temperature. After extensive washing, transcription was initiated by addition of all four nucleoside triphosphates and allowed to proceed for ~20 min. Pol I-specific transcripts were then detected by primer extension (3). The results shown in Fig. 9B were obtained using whole-cell extracts prepared from yeast strains derived from W303-1a as described previously (3). Following incubation of extract with bead-bound Pol I template, the unbound material was removed by extensive washing (3), the beads were boiled in SDS sample buffer, and Mot1 bound to the Pol I promoter was detected by Western blotting (5). Cells with complete open reading frame deletions in the essential genes RRN3, RRN5, RRN7, and RPA190 were maintained by transformation with the multicopy plasmid pNOY103, which provides for rRNA synthesis under control of the Pol II-driven GAL7 promoter (39). A strain harboring a temperature-sensitive allele of TBP (51) was a gift of Steve Hahn.


Figure 9
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  		FIG. 9. In vitro recruitment of Mot1 to the Pol I promoter depends on CF and RNA polymerase I. (A) In vitro transcription analysis using whole-cell extracts from wild-type (WT) or mot1-1 cells was performed using identical volumes of extract (0.5, 1, 2, or 4 µl, indicated by the ramps) obtained from identical numbers of cells prepared in parallel (see Materials and Methods and reference 3). Note the decrease in Pol I transcript levels directed by extracts from mot1-1 cells compared to those for wild-type cells. (B) Whole-cell extracts from the indicated strains were incubated for 1 h at room temperature with a Pol I promoter fragment immobilized on magnetic beads. After extensive washing, the beads were boiled in SDS sample buffer and promoter-bound Mot1 was detected by Western blotting ("bead-bound" samples). The relative level of Mot1 in the starting extracts is shown in the lower panel (WCE [whole-cell extract]). Note that Mot1 binding to the Pol I promoter was readily detected using extract from wild-type cells and that this interaction was strongly dependent on Pol I and Rrn7 and markedly reduced in the absence of Rrn3.


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RESULTS
 
Elevated levels of 35S RNA in mot1 cells. Inactivation of MOT1 results in derepression of stress response, mating type, and diauxic shift genes (13). We wondered if mot1 cells possess other features of cells entering stationary phase. Since Pol I promoter activity is sensitive to the cellular growth stage (62), we examined by Northern blotting the levels of the primary rRNA transcript, 35S RNA, in wild-type and mot1 cells. As shown in Fig. 1, 35S RNA levels were elevated about twofold in mot1 cells compared to levels in wild-type cells. This effect was observed in each of two different temperature-sensitive mot1 strains following a shift for 45 min to 35°C. Elevated 35S RNA levels were also observed in mot1-14 cells grown at 30°C, a temperature at which these cells grow slowly (12). Since the increase in 35S RNA was seen in mot1-14 cells at 30°C, this effect is not a result of a response of the Pol I machinery to heat stress (43). Additionally, the Mot1 protein is virtually undetectable in extracts from mot1-14 cells (12), indicating that these effects on transcript levels are unlikely to be an underestimation resulting from partial inactivity of Mot1.


Figure 1
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  		FIG. 1. Elevated 35S RNA in mot1 cells. Wild-type (WT) and mot1 cells were grown in rich medium at 30°C to an OD600 of ~1.0. Cells were then harvested or heat shocked for 45 min at 35°C prior to harvest. Twenty micrograms of total RNA from each strain was resolved by electrophoresis, transferred to a nylon membrane, and probed with a radiolabeled 35S-specific probe. For normalization of the 35S RNA band intensity, the blot was stripped and reprobed for ACT1 message. The graph shows the normalized, relative 35S RNA level determined from three independent experiments ± standard deviation.

rRNA synthesis rate is diminished in mot1 cells. The elevated level of 35S RNA in mot1 cells suggests that Mot1 inhibits rRNA transcription in normal cells. Alternatively, the accumulation of the 35S species could be due to a function for Mot1 in 35S RNA processing or effects of Mot1 on both transcription and processing. To directly assess the relative rates of rRNA synthesis and processing in wild-type and mot1 cells, pulse-chase experiments were performed. Cells were grown in methionine-free medium and then incubated with [methyl-3H] methionine for 2 min, followed by the addition of an excess of unlabeled methionine (see Materials and Methods). Cells were harvested at various time points following labeling, and rRNA species were analyzed by gel electrophoresis and fluorography. As shown in Fig. 2, unprocessed 35S RNA was barely detectable in wild-type cells but multiple processed and partially processed forms of rRNA were readily detected, as is typical for control yeast strains (e.g., see reference 59). After 10 min of chase, nearly all of the labeled RNA was processed to mature 18S and 25S species. In striking contrast, ~5-fold less rRNA was synthesized by mot1-14 cells during the 2-min labeling period, and the predominant rRNA species detectable at time zero was the fully unprocessed 35S RNA. Despite the processing defect evident following pulse-labeling, 35S RNA was almost quantitatively processed to the mature 18S and 25S forms by 10 min. Since mot1-14 cells grow slowly, and rRNA synthesis occurs in proportion to the growth rate, it is not clear to what extent the reduced rRNA synthesis in mot1-14 cells is a consequence of a reduced growth rate versus a defect in Pol I transcription caused directly by a defect in Mot1. Nonetheless, these results indicate that both rRNA synthesis and processing are defective in mot1 cells.


Figure 2
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  		FIG. 2. Reduced rRNA synthesis and processing in mot1 cells. Wild-type and mot1-14 cells were grown at 30°C in synthetic medium lacking methionine to an OD600 of ~0.3 to 0.4. Cells were then pulse-labeled with [methyl-3H]methionine for 2 min (time zero) and then incubated with unlabeled methionine for the times indicated above the lanes prior to harvest. Total RNA was fractionated on a formaldehyde agarose gel, and radiolabeled rRNA species were detected by autoradiography as described in Materials and Methods.

Reduced polymerase density and defective processing in mot1 cells. To better determine how a defect in Mot1 affects synthesis of rRNA, the Miller chromatin spreading method was used to visualize individual nucleolar genes from two pairs of mot1 mutant and congenic wild-type strains. The wild-type control strains had polymerase density distributions similar to those seen in other wild-type strains, which typically display a wide range of polymerase densities with an average of ~50 transcripts/gene (19, 48). Wild-type strain AY51 (13) had a polymerase density of 56 (n = 55), similar to the density of the wild-type strain JD194 (16), which had a polymerase density of 50 (n = 27). The average number of polymerases/gene for mot1-14 (strain AY86) (13) was 48 (n = 75), while that for mot1-1 (strain JD215b) (16) was 30 (n = 130). The polymerase density for genes from mot1-14 cells was significantly lower than the density for the wild-type control cells, with an associated P value of just under 0.05. The reduced polymerase density on genes from mot1-1 cells versus those from congenic control cells has an associated P value that is well below 0.05, indicating that this difference is highly significant. As can be seen in the plot in Fig. 3A, all strains analyzed had a wide distribution in polymerase density, but their modal, or most common, Pol I density values were similar for both mot1 strains (20 to 25 pols/gene) and were less than the modal value for the wild-type strain (~50 polymerases/gene).


Figure 3
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  		FIG. 3. rRNA genes from mot1 cells display fewer transcripts per gene on average and slowed rRNA processing on those transcripts. (A) Distribution of polymerase density on genes from mot1-14 and mot1-1 strains compared to that for the wild-type control strain. The number of polymerases per gene was determined for 55 rRNA genes from wild-type cells, 75 genes from mot1-14 cells, and 130 genes for mot1-1 cells. (B and C) Electron micrographs of representative rRNA genes from the wild-type (B) or mot1-14 (C) strain, with 55 and 37 transcripts, respectively. Below each gene is an interpretive tracing of the rDNA (dotted line) and RNA transcripts (solid lines). The wild-type gene (B) displays typical cotranscriptional rRNA processing events seen on yeast rRNA genes (43), including formation of large 5'-terminal particles, which encompass the pre-small-subunit RNA, followed by cleavage of these SSU processomes from nascent transcripts (arrows). The mot1-14 rRNA gene (C) displays transcripts that do not acquire SSU processome components and are not cleaved while nascent (arrows), characteristic of slowed or delayed rRNA processing.

The observation of reduced polymerase density is consistent with the pulse-labeling results of Fig. 2, which suggests that transcription of rRNA is impaired in mot1 cells. rRNA processing commences on nascent rRNA transcripts and can be visualized by EM (43). EM analysis showed that in addition to a defect in transcription, rRNA processing was consistently delayed or slowed in mot1 strains relative to that in the wild type. This can be seen on the representative rRNA genes in Fig. 3B and C. Specifically, normal rRNA processing events, involving compaction of pre-18S rRNA into the small ribosomal subunit (SSU) processome and its subsequent cleavage from the nascent transcript, were seen along the wild-type rRNA gene as expected (Fig. 3B) but not in rRNA genes from mot1-14 cells (Fig. 3C) or mot1-1 cells (discussed below). Rather, the rRNA transcripts in the latter cells reached the end of the gene prior to substantial packaging and without RNA cleavage. This could represent a specific kinetic defect in rRNA processing or an unlinking of transcription and rRNA processing as a result of reduced Pol I transcription (see Discussion). Importantly, these differences in Pol I density and rRNA processing were observed in each of two different mot1 strains, indicating that these effects are not dependent on the mot1 allele or the strain background. Psoralen cross-linking analysis (Fig. 4) demonstrated that the proportion of active rRNA genes was the same in wild-type and mot1 cells. Consistent with the EM analysis, this observation suggests that Mot1 affects the utilization of individual transcriptionally competent rRNA genes rather than the number of active templates.


Figure 4
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  		FIG. 4. The number of active rDNA repeats is the same in wild-type and mot1 cells. Psoralen cross-linking of the ribosomal DNA slows migration of the actively transcribed repeats. Wild-type (WT), mot1-14, and mot1-42 cells were harvested in log phase and then psoralen and UV treated as described in Materials and Methods. Genomic DNA preps were then digested with EcoRI, resolved on a 1.3% agarose-Tris-borate-EDTA gel, transferred to a membrane, and probed with a 35S rDNA fragment. The actively transcribed repeats (open) and the inactive repeats (closed) are indicated on the blot; the percentages of open repeats are indicated in the bar graph.

Localization of Mot1 to rDNA and role in PIC and SSU processome assembly. An effect of mot1 on Pol I transcription could result indirectly from an effect of Mot1 on transcription of a Pol I factor(s), a direct function for Mot1 in Pol I transcription, or some combination of the two possibilities. As shown in Fig. 5A and B, by ChIP, Mot1 was readily detected on each of three regions of the rDNA, including the Pol I promoter (NTS2). The midpoint of the linear response range for Mot1 ChIP to the rDNA occurred after 22 cycles of PCR, 5 fewer than the number of cycles required to reach the midpoint of the linear range for Mot1 ChIP to INO1, a well-characterized Mot1-repressed gene (14). Since the rDNA is present in multiple copies, a direct comparison to the level of Mot1 found on single-copy Pol II promoters was not possible. However, the tight correlation of Mot1 localization with sites of its action at Pol II promoters (1, 13, 23) suggests that the robust Mot1 ChIP signal obtained at the rDNA is due to a direct functional role for Mot1 in rRNA synthesis. As expected, TBP was also localized to NTS2, but the TBP ChIP signal was not affected by mutation of Mot1 (Fig. 5C). These results suggest that the role for Mot1 in Pol I transcription cannot be ascribed to Mot1-catalyzed TBP displacement from NTS2.

The indistinguishable TBP occupancy of NTS2 in wild-type and mot1 cells appeared to be at odds with the differences in the rRNA transcription rate in these strains. Since Mot1 has a role in control of Pol II transcription initiation and start-site selection (44), one possibility was that Pol II transcription complexes assemble spuriously on the rDNA in mot1 cells. To examine this possibility, rDNA was analyzed by ChIP for occupancy by Pol II, as well as the Pol II-specific general transcription factor TFIIB. As shown in Fig. 5D and E, levels of TFIIB and Pol II on the rDNA were markedly less than the levels of these factors present on the ACT1 promoter. While it is not possible to obtain an absolute measure of occupancy at the rDNA versus ACT1 due to differences in gene copy number, under these conditions the NTS2 TBP ChIP signal was similar to TBP ChIP to ACT1 (Fig. 5C), whereas the extent of TFIIB and Pol II ChIP to rDNA was only a fraction of that at ACT1 (Fig. 5D and E). Since the ratios of TFIIB to TBP and Pol II to TBP at the rDNA were markedly less than these ratios at ACT1, we conclude that formation of Pol II PICs was disfavored on rDNA in both wild-type and mot1 cells compared to the case with ACT1. Collectively, these results suggest that Mot1 might have a direct function in Pol I transcription that is distinct from its role in Pol II transcription.

To further explore the possible mechanism of Mot1 action at the rDNA, we next measured occupancy of NTS2 by three Pol I general transcription factors (UAF subunits Rrn5 and Rrn9 and CF subunit Rrn7), as well as the Pol I subunit Rpa135. FLAG-tagged Rrn5, -7, and -9 and Rpa135 strains grew indistinguishably from wild-type cells (Fig. 6A), indicating that the tagged proteins are fully functional. With the exception of a slight decrease in the growth rate of FLAG-Rrn5 mot1-42 cells compared to that of the untagged mot1-42 strain, there were no synthetic growth defects observed in mot1-42 cells harboring these FLAG-tagged proteins (Fig. 6A). Surprisingly, however, Western blot analysis of yeast whole-cell extracts demonstrated that the levels of all of these factors were reduced in mot1-42 cells compared to wild-type cells (Fig. 6B). Nonetheless, association of Rrn5, Rrn7, Rrn9, and Rpa135 with NTS2 was somewhat greater in mot1-42 cells than in wild-type cells (Fig. 6C), suggesting that in wild-type cells, Mot1 imposes a limit on the extent to which these factors associate with the rDNA.


Figure 6
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  		FIG. 6. Recruitment of UAFs and the CF complex is enhanced in mot1 cells. (A) Serial dilution spot assays of strains expressing the indicated FLAG-tagged proteins in WT and mot1-42 cells. The YPD (rich medium) plates were incubated at the indicated temperatures for 2 days prior to photography. (B) Western blot showing protein levels in WT or mot1-42 cells. Protein levels (with the exception of Rrn7, which is unaffected) are decreased in mot1 cells. (C) Quantification of ChIP to NTS2 (region of DNA amplification shown in Fig. 5). Recruitment of both UAFs and the CF complex are modestly increased in mot1 cells. Note that the significant decrease in protein levels does not affect the ability for these proteins to be recruited for transcription.

The results in Fig. 2 and 3 suggest an inefficiency and delay in the assembly of the 35S RNA processing machinery at the rDNA locus. To further investigate this apparent defect, strains were constructed in which the SSU processome component Utp8, Utp9, or Utp10 (17) was tagged with the HA epitope in either the wild-type or mot1-42 background. These Utp proteins associate with both the promoter-proximal region of rDNA and the 5' end of 35S pre-rRNA and are involved in coupling rRNA transcription and processing (20). The HA-Utp8 and HA-Utp9 strains grew similarly to the untagged parent wild-type and mot1-42 strains, indicating that the tagged Utp8 and Utp9 proteins were fully functional (Fig. 7A). In contrast, the HA-Utp10 strain grew somewhat more slowly than isogenic wild-type cells and displayed a synthetic sick phenotype when the tagged gene was combined with the mot1-42 allele (Fig. 7A). Western blot analysis of yeast whole-cell extracts showed that the levels of Utp8, Utp9, and Utp10 were reduced in mot1-42 cells compared to those in wild-type cells (Fig. 7B). Despite their reduced levels, the rDNA association of Utp8 and Utp9 was not detectably affected by mutation of MOT1 (Fig. 7C), suggesting that the processing defect in mot1-42 cells is not simply a result of reduced chromatin association as a consequence of a reduced overall protein level. A similar result was obtained for HA-Utp10 ChIP, although the ChIP results were associated with a larger error than for HA-Utp8 and HA-Utp9. The synthetic growth defect in HA-Utp10 mot1-42 cells is consistent with a functional interaction between Utp10 and Mot1, but since the tagged Utp10 protein appears to not be fully functional in vivo, perhaps the impaired activity of HA-Utp10 and the sickness of the strains accounts for the variability in the ChIP results for this factor.


Figure 7
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  		FIG. 7. Recruitment of SSU processome components to chromatin is not impaired in mot1 cells. (A) Spot assay showing growth of 10-fold serial dilutions with HA-tagged Utps in WT and mot1-42 strains. HA-Utp10 cells show a severe synthetic defect with mot1-42. (B) Western blot detecting Utp levels in WT and mot1-42 strains. Mot1 affects the expression level of all three Utps tested. The asterisk indicates the position of Utp10, which is expressed at low levels even in WT cells. (C) Quantification of Utp ChIP results. Graph shows the mean value ± standard deviation obtained using two independently prepared batches of chromatin. Note that the extent of Utp association with chromatin is not dependent on the overall protein level in the whole-cell extracts.

Efficient rRNA synthesis and processing require Mot1 ATPase activity. Mot1's ATPase activity drives displacement of TBP from DNA and is required for its essential function in vivo (4). Inasmuch as TBP ChIP levels at NTS2 were unaffected in mot1 cells, it was important to determine if ATPase activity was important for the function of Mot1 in rRNA synthesis. To address this question, mot1-1 cells were transformed with plasmid-borne copies of wild-type MOT1, mot1-505, or vector alone. mot1-505 harbors multiple mutations in the Walker B motif of the ATP binding pocket, destroying ATPase activity (54). Importantly, mot1-505 is expressed at wild-type levels in vivo (54). As shown in Fig. 8A, mot1-1 cells expressing mot1-505 had elevated levels of unprocessed 35S RNA similar to those with vector alone, whereas addition of wild-type MOT1 restored the normal low level of 35S RNA. EM analysis demonstrated that rDNA loci in mot1-505 cells had polymerase densities that looked very similar to those for mot1-1 cells, and transcripts produced in these cells lacked the terminal knobs indicative of efficient 35S RNA processing (Fig. 8B to D). Thus, Mot1's ATPase activity is required for its function in Pol I transcription.


Figure 8
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  		FIG. 8. Mot1 ATPase activity is required for proper transcription of 35S RNA. (A) Results of Northern analysis showing relative 35S RNA levels in each of three strains. mot1-1 cells were transformed with vector plasmid or plasmids carrying the wild-type (WT) MOT1 or mot1-505 (DEAD box mutant) genes, and Northern analysis was performed as for Fig. 1. Relative 35S RNA levels were normalized to ACT1 message levels in each strain. The graph shows the average ± standard deviation obtained from two independent experiments. (B to D) Electron micrographs of rDNA from mot1-1 cells transformed with a wild-type MOT1 (B), mot1-505 (C), or vector (D) plasmid. Note that the lower number of transcripts and defects in cotranscriptional processing seen with the vector-carrying strain were recovered only by coexpression of wild-type MOT1 but not mot1-505, which is defective for Mot1 ATPase activity.

Recruitment of Mot1 to the Pol I promoter in vitro. Previous results indicated that Mot1 is not required for Pol I transcription in vitro (4). In light of the defects in rRNA synthesis in mot1 cells described above, a possible role for Mot1 in Pol I transcription in vitro was reexamined. As shown in Fig. 9A, consistent with previous results, whole-cell extracts from mot1-1 cells supported Pol I transcription. Since Mot1 protein is not detectable in extracts from mot1-1 cells (not shown), this result indicates that Mot1 is not essential for Pol I transcription in vitro. Titration of whole-cell extracts from wild-type and mot1-1 cells revealed that mot1-1 extracts had a reduced capacity for Pol I transcription compared to extracts from wild-type cells (Fig. 9A), but this quantitative difference was likely due to the combined effects of depletion of Mot1, as well as reduced levels of Pol I enzyme and a subset of Pol I general transcription factors in the extract (Fig. 6B).

Consequently, a more direct approach was employed to determine if Mot1 is recruited to the Pol I promoter via association with TBP and/or other components of the Pol I general transcription machinery. The association of Mot1 with the Pol I promoter was analyzed in vitro using an immobilized template assay (3). Whole-cell extracts were incubated with bead-bound Pol I template to allow transcription complexes to form. Unbound factors were then removed by washing the beads, and the template-associated Mot1 was analyzed by Western blotting. As shown in Fig. 9B (lane 1), using extracts from wild-type cells, Mot1 association with the Pol I promoter was readily detected. To determine if Pol I or Pol I-specific general transcription factors were involved in recruitment of Mot1 to the Pol I promoter, we took advantage of a yeast strain in which rRNA synthesis is supported by the Pol II-dependent GAL7 promoter (39). In this strain, Pol II bypasses the requirement for the Pol I apparatus, permitting loss of genes encoding Pol I transcription factors that are otherwise essential. Levels of Mot1 in extracts from rrn3{Delta}, rrn7{Delta}, and rpa190{Delta} cells were comparable to the level in extracts from wild-type cells (Fig. 9B). Interestingly, recruitment of Mot1 to the Pol I promoter in vitro was strongly dependent on Rrn7, a component of CF, as well as Pol I itself, as demonstrated by the loss of Mot1 binding to the promoter in extracts missing the Pol I-specific subunit Rpa190 (Fig. 9B, lanes 4 and 5). In contrast, association of Mot1 with the Pol I promoter was less dependent on the Pol I-associated factor Rrn3. Mot1 recruitment was also reduced using extracts depleted of the UAF subunit Rrn5, but there was less Mot1 present in the rrn5{Delta} extract, making the assessment of Rrn5's role in Mot1 recruitment unclear. Overall, these results support a model in which Mot1 associates with the Pol I promoter via cooperative interactions among Pol I-specific factors that stabilize formation of the Pol I PIC.


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DISCUSSION
 
In this study, we provide evidence for a new role for Mot1 in rRNA synthesis. The conclusion that Mot1 participates in rRNA synthesis is supported by Northern blotting, EM analysis, and pulse-labeling experiments, each of which provides complementary insight into the function of Mot1 at the rDNA. In contrast to the case with other factors dedicated to Pol I transcription and rRNA processing, rather than eliminating rRNA synthesis entirely, loss of Mot1 resulted in a kinetic defect in production of mature rRNA. The fact that rRNA production did proceed, albeit inefficiently, in mot1 cells might explain why Mot1 has not been uncovered in previous screens for Pol I transcription and processing factors. Since rRNA production is regulated in proportion to the cellular growth rate (62), the defect in rRNA expression might also explain, at least in part, why mot1 cells grow slowly. The lower growth rate of mot1 cells is consistent with slower production of ribosomes. Importantly, a role for Mot1 in rRNA expression was supported by analysis of different mot1 alleles and in different strain backgrounds, pointing to a fundamental role for Mot1 in this process in yeast cells. A stimulatory role for Mot1 in rRNA synthesis fits well with the previously described role for Mot1 in control of stress response, mating, sporulation, and diauxic shift genes (13). Thus, in cells growing in rich medium, Mot1 functions to promote the synthesis of rRNA and repress the expression of genes important for survival during nutrient limitation and stress. A previous study (10) demonstrated that the abundance of mature rRNA species was similar in wild-type and mot1 cells. This observation is consistent with our own observations (not shown) indicating that the levels of stable, mature rRNA species are unaffected in mot1 cells. In vitro transcription data reported here (Fig. 9) are also consistent with published results demonstrating that Mot1 is not essential for Pol I transcription (4). Instead, experiments performed by titrating the whole-cell extracts suggest that in vitro, Pol I transcription activity is modestly decreased in extracts from mot1 cells.

Two general classes of models can explain the role for Mot1 in rRNA production. The first possibility is that decreased rRNA synthesis in mot1 cells is an indirect effect of a Pol II-specific (or perhaps a TBP-specific) effect of Mot1. Our previous global expression analysis (13) did not support a role for Mot1 in expression of genes involved in rRNA synthesis. However, while this paper was under revision, we obtained new microarray data using more-sensitive analysis that reveals a role for Mot1 as a weak activator of several UTP genes (R. O. Sprouse and D. T. Auble, unpublished observations). Preliminary analysis indicates that Mot1 appears to stimulate UTP gene expression by about twofold. These modest stimulatory effects cannot account for the substantial changes in some rRNA synthesis factors that were detected in whole-cell extracts (Fig. 6 and 7), but they do caution against ruling out indirect effects of Mot1 mutation on rRNA production. Mot1 might also control the expression of some novel rRNA transcription or processing gene or regulate the abundance or activity of a bona fide Pol I factor in a novel way. Alternatively, in principle, an indirect effect of mot1 on rRNA synthesis might result from a global defect in the specificity of Pol II PIC formation in mot1 cells. Genetic analysis identified a role for MOT1 in Pol II start site selection (31), and evidence supports a role for Mot1 in redistributing TBP among chromatin sites on a global scale (10, 37). Therefore, inactivation of Mot1 might lead to inappropriate assembly of stable TBP-containing complexes on the rDNA. Such complexes could potentially interfere with transcribing Pol I molecules by forming roadblocks to elongation, for example. This hypothesis is not supported by EM analysis, however: blocks to elongation would be predicted to stall elongating Pol I molecules, thereby allowing them to accumulate on transcribed regions. In fact, the opposite situation was observed, in which the density of elongating Pol I enzymes was reduced in mot1 cells compared to that in wild-type cells. Furthermore, while ChIP analysis did demonstrate significant TBP occupancy of 18S and 25S transcribed sequences (Fig. 5C), impaired Mot1 function did not lead to an increase in TBP occupancy as was seen at Mot1-controlled Pol II promoters (14). Additionally, TBP binding to Pol I-transcribed regions did not efficiently support loading of TFIIB or Pol II.

Regardless of the possible indirect contributions of a defect in Mot1 to rRNA synthesis, the results presented here provide several lines of evidence in support of a second model in which Mot1 participates directly in rRNA synthesis. Most noteworthily, data in Fig. 5B demonstrate robust association of Mot1 with rDNA in vivo. Geisberg et al. (23) reported that Mot1 occupies the rDNA, but at quite low levels. The explanation for the quantitative difference in Mot1 occupancy observed here compared to the findings of Geisberg and Struhl is not obvious but could be due to differences in experimental detail. As a general comment, since Mot1 catalyzes TBP displacement, we suggest that since Mot1 is transiently associated with chromatin, the overall ChIP signal may not reflect in all cases the extent of its contribution to regulation at sites where it operates. Thus, even a low Mot1 ChIP signal could be physiologically important if the signal is misleadingly low due to the high catalytic efficiency of the enzyme at that locus. On the other hand, Mot1 does not appear to simply function as a TBP-DNA-dissociating enzyme at NTS2, because inactivation of Mot1 led to no apparent change in TBP occupancy. This lack of change in NTS2 TBP occupancy in mot1 cells is in contrast to the increase in TBP occupancy observed at Pol II promoters, such as INO1 (Fig. 2) (14). These results suggest that Mot1 functions differently in Pol I transcription than in Pol II transcription.

The results presented here are the first to directly implicate Mot1 in rRNA synthesis, but our retrospective examination of high-throughput biochemical and genetic results suggests an astonishing number of previously unanticipated interactions between Mot1 and the rRNA synthesis and processing machinery. In fact, despite extensive studies on the role of Mot1 in regulation of Pol II transcription, isolation of native, Mot1-containing complexes has uncovered more potential interactions between Mot1 and components of the rRNA synthesis machinery than between Mot1 and Pol II factors. Of note, a physical interaction was reported between Mot1 and the Pol I subunit Rpa135 (21), consistent with our conclusion that Mot1 functions in Pol I initiation. Interactions have also been reported between Mot1 and both Net1 and Reb1, as well as between Mot1 and histone H4 (21, 22). As a subunit of the RENT complex, Net1 is localized to rDNA and is required for Sir2-mediated rDNA silencing and nucleolar integrity (57). Interestingly, Net1 also stimulates Pol I transcription in vitro and in vivo, an effect attributable to the interaction between Net1 and Pol I (52). Reb1 is involved in the expression of both Pol I and Pol II genes (2, 7, 18, 25, 46, 47), and the significance of the physical interaction between Mot1 and Reb1 is supported by the overlap in the synthetic lethal profiles of these two genes (15, 38). In addition to being a general structural component of chromatin, histones H3 and H4 are subunits of UAF (26) and H3 depletion results in a marked inhibition of Pol I transcription (59). Since Mot1, Reb1, and H4 are localized to NTS2, the interactions of Mot1 with Reb1 and H4 could reflect Mot1's function in Pol I transcription initiation.

High-throughput purification also suggests a functional link between Mot1 and rRNA processing and ribosome biogenesis. Mot1 has been purified in association with a number of ribosomal proteins, including Rpl4, Rpl5, Rpp0, Rps3, Rps4, and Rps5 (21, 22). rRNA processing occurs cotranscriptionally (20, 43) and requires a large ribonucleoprotein complex called the SSU processome (17). Remarkably, the SSU processome contains five small ribosomal subunit proteins, one of which is the Mot1-interacting protein Rps4 (6). It is difficult to conclude whether the rRNA processing defect in mot1 cells is due to a direct or indirect effect of Mot1, but previous studies have linked rDNA binding proteins, rRNA transcription, and rRNA processing (20, 59). Since fully processed ribosomal RNAs do accumulate in mot1 cells, but inefficiently, whereas the amount of Utp8, -9 and -10 recruited to the rDNA is not significantly affected as judged by ChIP, the rRNA processing defect in mot1 cells might indicate a role for Mot1 in stabilizing SSU processomes on nascent Pol I transcripts. The modest but significant increase in Utp8 ChIP to rDNA in mot1 cells might also result from a kinetic defect in mature SSU processome assembly, suggesting a possible role for Mot1 in transferring a subset of Utps (the t-Utps, of which Utp8, -9 and -10 are members) from rDNA to the 5' end of nascent rRNA (20).

It has been proposed that the Pol I PIC undergoes dynamic cycles of assembly and disassembly at the promoter (3). The localization of Mot1 to NTS2 in vivo (Fig. 5B), the cooperative assembly of Mot1 with the Pol I promoter in vitro (Fig. 9), and the requirement for Mot1's ATPase activity (Fig. 8) suggest that Mot1 might contribute to the assembly or disassembly of the Pol I PIC. Such an effect on PIC dynamics might explain why the loss of Mot1 affects the efficiency of rRNA synthesis rather than being an essential component of the PIC. An effect of Mot1 on PIC dynamics is also consistent with ChIP results showing somewhat elevated levels of Rrn proteins, Rpa135, and Utp8 on rDNA in mot1-42 cells. If Mot1 functions to regulate PIC dynamics, the results are consistent with the possibility that normal PIC dynamics are involved in coupling rRNA transcription and processing, such that in mot1 cells there is a delay in normal rRNA processing until after transcription. The EM results showing unprocessed 5' ends in mot1 cells are consistent with results in Fig. 1 showing higher amounts of the 5' external transcribed spacer, which is typically very short lived and processed cotranscriptionally (60). The EM results are also consistent with the RNA labeling experiment (Fig. 2) showing a longer-lived 35S precursor. An alternative possibility is that mot1 cells manifest a specific kinetic defect in rRNA processing because Mot1 has a novel, direct function in processing regulation or dictating its kinetics.


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ACKNOWLEDGMENTS
 
This work was supported by NIH grants GM55763 to D.T.A. and GM63952 to A.L.B. This work was also supported by American Heart Association grant AHA-0555490U to J.S.S.

Special thanks go to Loan Vu for help and advice with the pulse-labeling experiments, to Susan Baserga and Steve Hahn for yeast strains, and to Beth Moorefield for providing yeast strains prior to publication. We are also grateful to members of the Auble and Beyer labs for discussions and comments on the manuscript.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biochemistry and Molecular Genetics, University of Virginia Health System, 1300 Jefferson Park Avenue, Charlottesville, Virginia 22908-0733. Phone: (434) 243-2629. Fax: (434) 924-5069. E-mail: dta4n{at}virginia.edu Back

{triangledown} Published ahead of print on 12 February 2007. Back

{dagger} These authors contributed equally. Back


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Molecular and Cellular Biology, April 2007, p. 2886-2896, Vol. 27, No. 8
0270-7306/07/$08.00+0     doi:10.1128/MCB.00054-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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