Are upstream activating factor (UAF) and upstream binding factor (UBF) the same thing?

Are upstream activating factor (UAF) and upstream binding factor (UBF) the same thing?

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During ribosome pre-40S and pre-60S synthesis, many sources state the importance of UAF or UBF in initiation complex of ribosome DNA transcription. None of the sources I've seen mentions the other when mentioning UAF or UBF, so I want to know if they are actually the same thing.

Another source of confusion is that Wikipedia has article for both:

Thanks in advance.

Differential roles of phosphorylation in the formation of transcriptional active RNA polymerase I

Regulation of rDNA transcription depends on the formation and dissociation of a functional complex between RNA polymerase I (pol I) and transcription initiation factor Rrn3p. We analyzed whether phosphorylation is involved in this molecular switch. Rrn3p is a phosphoprotein that is predominantly phosphorylated in vivo when it is not bound to pol I. In vitro, Rrn3p is able both to associate with pol I and to enter the transcription cycle in its nonphosphorylated form. By contrast, phosphorylation of pol I is required to form a stable pol I-Rrn3p complex for efficient transcription initiation. Furthermore, association of pol I with Rrn3p correlates with a change in the phosphorylation state of pol I in vivo. We suggest that phosphorylation at specific sites of pol I is a prerequisite for proper transcription initiation and that phosphorylation/dephosphorylation of pol I is one possibility to modulate cellular rDNA transcription activity.

Precise regulation of rRNA synthesis according to environmental conditions plays a key role in ribosome biosynthesis, which is tightly controlled through all prokaryotic and eukaryotic organisms (1, 2). The level of rRNA production correlates with the activity of the RNA polymerase I (pol I)-containing transcription machinery that generates the precursor transcript of the mature 18S, 25S, and 5.8S rRNA. Transcription efficiency depends on the formation of active initiation complexes on the rDNA promoter, which requires several transcription initiation factors and the recruitment of initiation competent pol I. In the yeast Saccharomyces cerevisiae, the first pol I-dependent initiation factor identified was the TATA binding protein TBP (3, 4), which could be isolated within a stable complex of 240 kDa (5). TBP contacts polypeptides of two multiprotein complexes, the core factor (CF refs. 6, 7) and the upstream activating factor (UAF refs. 6 and 8). One crucial role of TBP in pol I transcription is to recruit the pol I machinery to the promoter in a manner completely dependent on UAF (6, 8, 9). CF is composed of three stably associated proteins encoded by the genes RRN6, RRN7, and RRN11 (7, 10, 11) whereas UAF is a multisubunit complex consisting of at least five different proteins: Rrn5p, Rrn7p, Rrn10p (12), and the histones H3 and H4 (13). It has been proposed that binding of UAF upstream to the promoter is necessary to efficiently recruit CF and, finally, the initiation active form of pol I to the start site of rRNA synthesis (8). Initiation-competent pol I is stably associated with the pol I-specific initiation factor Rrn3p (14, 15). By binding the C-terminal part of Rrn6p and the pol I-specific subunit A43, Rrn3p connects CF to pol I, which results in recruitment of pol I to the promoter and in the formation of an active initiation complex (16). Recently, it was suggested that pol I-Rrn3p, together with CF, cycles on and off the promoter, whereas UAF remains bound (17). One possibility to regulate transcription initiation is the formation and dissociation of the pol I-Rrn3p complex, which represents a molecular switch to regulate rRNA synthesis. This idea was supported by the following previously published data (15). First, stable association of pol I with Rrn3p in solution is a necessary prerequisite to initiate transcription second, only a minor proportion of both Rrn3p and pol I were found in a stable complex with each other in whole cell extracts, whereas most of both pol I and Rrn3p were uncomplexed and were found to be inactive in transcription initiation third, the pol I-Rrn3p complex dissociated during one round of transcription in vitro and lost its capacity for subsequent reinitiation and fourth, stationary cells, which are silent in rDNA transcription, show a strong reduction of pol I-Rrn3p complexes however, they contain substantial levels of both free Rrn3p and pol I (15). Because both pol I and Rrn3p exist in parallel as uncomplexed and as stably associated molecules, it is conceivable that different posttranslational modifications have to exist in either one or both components to trigger initiation activity.

Recently, the human homolog of yRrn3p has been identified. It is able to replace yeast Rrn3p in vivo (18) and was shown to be identical with TIF-IA, the factor mediating growth-dependent control of mammalian ribosomal RNA synthesis (19). In accordance with its yeast counterpart, human Rrn3p was found to be essential to recruit initiation-competent pol I to the promoter, thereby linking human pol I to the promoter selectivity factor SL1 (20). Similar to the situation in yeast, only a minor proportion of human pol I complexes were found in stable association with hRrn3p to result in an initiation-competent enzyme (20). These findings suggest that the reversible interaction between Rrn3p and pol I by posttranslational modifications might reflect a conserved basal mechanism to regulate rDNA transcription in eukaryotes.

Phosphorylation is a common posttranslational modification that is involved in many regulatory pathways. Because previous results demonstrated that the pol I subunits A190, A43, A34.5, A23, and A19 are phosphorylated in vivo (21, 22), we asked whether phosphorylation of pol I is a possible target that could modulate the interaction between pol I and Rrn3p. In deed, association of pol I with Rrn3p is paralleled by a change in the pol I phosphorylation pattern, suggesting a transition in the phosphorylation state of pol I to reach initiation competence. Dephosphorylation of pol I in vitro reduced initiation activity of yeast pol I, resulted in destabilization of the preformed pol I-Rrn3p complex, and inhibited complex formation with bacterially expressed Rrn3p. By contrast, initiation competence of Rrn3p did not require its phosphorylation. From these results, we propose that phosphorylation of pol I is one possibility to modulate pol I-Rrn3p interaction and, thus, to control activity of the transcription process.

Structural organization of the rRNA transcription unit

In higher vertebrates, a standard rDNA transcription unit encodes the precursor to 18S, 28S, and 5.8S rRNAs. Each unit also contains important sequence elements that regulate pre-rRNA transcription, such as the rDNA promoter, enhancers, spacer promoters, an origin of replication, transcription terminators, and a replication fork barrier that prevents replication forks from colliding with transcribing RNA polymerase I during S phase. The tandem arrangement of multiple rDNA genes may have been useful to increase gene dosage and to maintain the well-recognized rRNA sequence homology. With the exception of closely related species, eukaryotic rDNA promoter sequences have diverged significantly. Consistent with this sequence disparity, rDNA transcription is generally specific to taxonomic orders, the promoter of one group not being recognized by the transcription machinery of others (for review, see Heix and Grummt 1995). With a few exceptions, rDNA promoters share a common modular organization, consisting of a start site proximal core promoter (CP) and an upstream control element (UCE). The stereospecific alignment of both sequence elements is crucial for efficient transcription initiation. Analysis of structural parameters of ribosomal gene promoters from human to lower plants revealed the conservation of specific structural features, rather than base sequence, that are fundamental for promoter function (Marilley and Pasero 1996 Marilley et al. 2002). Apparently, a structural code, in addition to primary sequence, directs specific DNA–protein interactions at the rDNA promoter and may play an important function in transcriptional control.


It has long been known that pol I transcription is localised to discrete sites called nucleoli these can be likened to ribosome factories, in which rRNA is synthesised by pol I in the fibrillar centres and then processed and assembled into ribosomes in the surrounding granular regions ( 3). This has always been regarded as a peculiarity of the pol I system, but a recent study has shown that pol II and pol III both also carry out tran­scription at discrete locations ( 4). Confocal and electron microscopy of HeLa cells revealed that pol III transcription occurs at

2000 sites within the nucleoplasm ( 4). Each site has a radius of

20 nm and contains, on average, five molecules of active pol III ( 4). Pol II is not found at these sites, but functions at its own spatially separate locations, of which there may be

Although transcripts made by pol I and pol II are thought to be processed close to the site of their synthesis, at least part of the tRNA processing pathway may occur in the nucleolus ( 5). Thus, fluorescent in situ hybridisation reveals unprocessed pre-tRNAs within the nucleoli of Saccharomyces cerevisiae ( 5). Furthermore, the RNA subunit of RNase P, a pol III product which helps cleave pre-tRNA, can be found in both the nucleolus and the nucleoplasm in yeast and mammals ( 5, 6). Processing of tRNA within the same compartment as ribosome synthesis may provide opportunies for coordinating production of distinct components of the translational apparatus.


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. ​ Fig.1, 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 35ଌ. Elevated 35S RNA levels were also observed in mot1-14 cells grown at 30ଌ, a temperature at which these cells grow slowly (12). Since the increase in 35S RNA was seen in mot1-14 cells at 30ଌ, 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.

Elevated 35S RNA in mot1 cells. Wild-type (WT) and mot1 cells were grown in rich medium at 30ଌ to an OD600 of 𢏁.0. Cells were then harvested or heat shocked for 45 min at 35ଌ 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- 3 H] 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. ​ Fig.2, 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, 𢏅-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.

Reduced rRNA synthesis and processing in mot1 cells. Wild-type and mot1-14 cells were grown at 30ଌ in synthetic medium lacking methionine to an OD600 of 𢏀.3 to 0.4. Cells were then pulse-labeled with [methyl- 3 H]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 � 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. ​ Fig.3A, 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 (� polymerases/gene).

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. ​ Fig.2, 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. ​ (Fig.3B) 3B ) but not in rRNA genes from mot1-14 cells (Fig. ​ (Fig.3C) 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. ​ (Fig.4) 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.

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. ​ (Fig.5C). 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. ​ (Fig.5C), 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. ​ (Fig.6A), 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. ​ (Fig.6A). 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. ​ (Fig.6B). 6B ). Nonetheless, association of Rrn5, Rrn7, Rrn9, and Rpa135 with NTS2 was somewhat greater in mot1-42 cells than in wild-type cells (Fig. ​ (Fig.6C), 6C ), suggesting that in wild-type cells, Mot1 imposes a limit on the extent to which these factors associate with the rDNA.

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. ​ Fig.5). 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. ​ Fig.2 2 and ​ and3 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. ​ (Fig.7A). 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. ​ (Fig.7A). 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. ​ (Fig.7B). 7B ). Despite their reduced levels, the rDNA association of Utp8 and Utp9 was not detectably affected by mutation of MOT1 (Fig. ​ (Fig.7C), 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.

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. ​ Fig.8A, 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.

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. ​ Fig.1. 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. ​ Fig.9A, 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. ​ (Fig.9A), 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. ​ (Fig.6B 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. ​ Fig.9B 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 rrn3Δ, rrn7Δ, and rpa190Δ cells were comparable to the level in extracts from wild-type cells (Fig. ​ (Fig.9B). 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. ​ (Fig.9B, 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 rrn5Δ 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.

Materials and methods

Strains and media

To generate JS490, JS493 and JS566, the RPD3, SIN3 or SIR2 open reading frames (ORFs), respectively, were deleted from JS311 and replaced with the kanMX4 marker by a one-step gene replacement method ( Smith et al., 1999 ). Miller spreads were performed on the S288C derivatives, JS772 and JS777. The genotype for each strain is described in Table II.

Strain Genotype
JS311 a a Smith et al. (1999) .
MATα his3Δ200 leu2Δ1 met15Δ0 trp1Δ63 ura3-167 RDN1::Ty1-MET15, mURA3-HIS3
JS490 a a Smith et al. (1999) .
JS311 made rpd3Δ::kanMX4
JS566 JS311 made sir2Δ::kanMX4
JS218 b b Smith and Boeke (1997) .
MATα his3Δ200 leu2Δ1 ura3-167 sir2Δ::HIS3 RDN1::Ty1- mURA3
JS772 c c Winzeler et al. (1999) .
MATa his3Δ0 leu2Δ0 ura3Δ0
JS777 JS772 made rpd3Δ::kanMX4

Psoralen cross-linking

Psoralen cross-linking assays were performed as described previously ( Dammann et al., 1993 Smith and Boeke, 1997 ). Stationary phase cultures were used to inoculate fresh 250 ml YPD cultures to an A600 of ∼0.2. Aliquots of ∼1 × 10 8 cells were harvested at the indicated time points. Washed cells were resuspended in 1.4 ml of ice-cold TE and 0.7 ml of this mixture was aliquoted into a 24-well tissue culture plate 40 μl of a 200 μg/ml solution of 4,5′,8-trimethylpsoralen (Sigma) were added to each well. The cells were then UV irradiated (UV lamp model B-100A Ultraviolet Products, Inc.) on ice for five doses of 5 min at a distance of 6 cm. Cells were spheroplasted with zymolyase, lysed, proteinase K treated, phenol–chloroform extracted and the resulting nucleic acid was ethanol precipitated. The pellet was resuspended in TE and normalized to a nucleic acid concentration of 2 mg/ml, and 4 μl were digested with EcoRI in a 30 μl reaction. Cleaved DNA was separated on a 1.3% agarose gel and transferred to a positively charged nylon membrane. rDNA-specific bands were detected by hybridization with a 3.6 kb XbaI–XbaI fragment containing most of the 35S region of yeast rDNA (Figure 1A).

RNA blot analysis

Yeast cultures were grown as described above for psoralen cross-linking. Cells were harvested (∼4 × 10 8 ) at the indicated time points and total RNA was isolated using the acid–phenol method ( Ausubel et al., 2000 ). Total RNA (15 μg) was separated on a 1.2% agarose–2% formaldehyde gel and transferred to a nylon filter. The filter was hybridized for 90 min with the 32 P-labeled probe JS45 (5′-TCGGGTCTCTCTGCTGCC GGAAATGCTCTCTGTTCA-3′), which hybridizes to the 5′ ETS of the 35S rRNA. Hybridization was performed in Quikhybe solution (Stratagene) at 65°C. Filters were washed twice (5 min) at room temperature with 2× SSC/0.1% SDS and then twice more at 60°C with 0.1× SSC/0.1% SDS. 35S rRNA bands were visualized by autoradiography. The ACT1 oligonucleotide probe was JB2351 (5′-GTC ACCGGCAAAACCGGCTTTACACATACCAGAACCGTTATCAAT AACCAAAGCAGCAAC-3′). Hybridization with this probe was the same as with JS45 except that the hybridization temperature was 60°C and the high temperature wash was at 55°C.

Transcriptional run-on assay

In vivo run-on assays were performed as described previously ( Elion and Warner, 1986 Cormack and Struhl, 1992 ), with several modifications. Cells were harvested (∼8 × 10 7 ) from log phase cultures (OD600 = 0.5) or cultures that were approaching stationary phase (OD600 = 5.6). Cells were washed twice with ice-cold TMN buffer (10 mM Tris–HCl pH 7.4, 100 mM NaCl, 5 mM MgCl2), resuspended in 1 ml of 0.5% sodium N-lauroyl sarcosine and incubated for 15 min on ice. Cells were then pelleted and resuspended in a 100 μl reaction mixture containing 50 mM Tris–HCl pH 7.9, 100 mM KCl, 5 mM MgCl2, 1 mM MnCl2, 2 mM DTT, 0.5 mM ATP, 0.25 mM CTP, 0.25 mM GTP, 10 mM phosphocreatine, 10 mg/ml creatine phosphokinase, 100 μCi [α- 32 P]UTP (800 Ci/mmol) and 25 μg/ml α-amanitin. This reaction was incubated at 25°C for 10 min, and then stopped by the addition of 1 ml of ice-cold TMN supplemented with 1 mM UTP. Total RNA was isolated and separated by RNA blotting as described above. Labeled RNA was detected by autoradiography and quantitated using a Molecular Dynamics Storm PhosphorImager.

Electron microscopy

Yeast was grown in YPD plus 1 M sorbitol to mid- (A600 = 0.4) or post- (A600 = 2.5) log phase. Miller chromatin spreads of yeast strains were prepared after brief digestion of the yeast cell wall with zymolyase followed by hypotonic lysis in 0.025% Triton X-100 pH 9 ( Hamkalo and Miller, 1973 Rattner et al., 1982 ). Volumes of 1 and 0.2 ml were harvested, and zymolyase concentrations of 5 and 25 mg/ml were used for mid- and post-log phase cells, respectively. Cell wall digestion was carried out at 30°C with shaking in the presence of actinomycin D (0.1 mg/ml). After 4 min, the digests were spun at maximum speed in an Eppendorf microcentrifuge for 6 s and the yeast pellet was resuspended in 1 ml of the lysis solution, which was then further diluted with an additional 3 ml of lysis solution. The samples were swirled gently for 20 min to allow for dispersal of cellular contents before being fixed with 0.4 ml of 0.1 M sucrose, 3.7% formalin and centrifuged onto carbon-coated EM grids. The grids were then stained with phosphotungstic acid and uranyl acetate and examined in a JEOL 100 CX microscope.

Western blotting

WCEs were separated on a 12% SDS–polyacrylamide gel and then transferred to PVDF membranes. Antibodies specific for acetylated H4 (penta H4), acetylated H3 (K9,14-acetyl) and H3 methylated on K4 were kindly provided by David Allis, and were incubated with the membrane at 1:5000 dilution in PBS-T (1× PBS, 0.05% Tween-20) containing 2% milk for 1 h at room temperature. All other antibodies were incubated at 1:10 000 dilutions for 2 h at room temperature. Membranes were washed with PBS-T (2% milk). Antibodies specific for histone H4 acetylated on lysine 5, 8, 12 or 16 were purchased from Serotec. The secondary antibody was a goat α-rabbit–HRP conjugate used at a dilution of 1:5000 (Amersham Biosciences). Post-translationally modified histones H3 and H4 were detected using ECL reagents from Amersham Biosciences.


YPD cultures (50 ml) of JS311 and JS490 were grown into log phase (A600 = 1) or early stationary phase (A600 = 7), then fixed for 1 h with 1% formaldehyde. Chromatin extraction and immunoprecipitations were performed as described previously ( Kuo and Allis, 1999 ). Cell pellets were resuspended in 400 μl of FA-lysis buffer (50 mM HEPES–KOH pH 7.5, 140 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% deoxycholic acid, 1 mM PMSF, 1 μg/ml leupeptin, 1 μg/ml pepstatin). Cells were disrupted with glass beads in a BioSpec Products Mini Beadbeater. The extract was sonicated to fragment the chromatin and then centrifuged at 14 000 r.p.m. for 20 min to clarify the WCE.

For the immunoprecipitations, 200 μl of FA-lysis buffer were combined with 40 μl of WCE and 1 μl of the histone antibodies. This mixture was incubated at 4°C overnight on a rotator. Bound chromatin was precipitated with protein A–Sepharose beads (Amersham Biosciences) for 2 h at 4°C. The beads were washed extensively and the immune complexes were eluted twice with 200 μl of 1% SDS/0.1 M NaHCO3 at room temperature. Following reversal of cross-linking, the recovered material was treated with RNase A and then proteinase K. The remaining DNA was phenol extracted, ethanol precipitated and resuspended in a final volume of 150 μl of TE. Each sample (3 μl) was used in 50 μl PCRs specific for the NTS1, NTS2, 5′-ETS or 25S regions of the rDNA, or the INO1, MATα and CUP1 genes. PCR products were separated on a 2% agarose–1× TBE gel, stained with ethidium bromide and images captured using an AlphaImager 2000 documentation and analysis system from Alpha Innotech. Antibodies were the same as those used for western blotting.


The nucleolus was identified by bright-field microscopy during the 1830s. [6] Little was known about the function of the nucleolus until 1964, when a study [7] of nucleoli by John Gurdon and Donald Brown in the African clawed frog Xenopus laevis generated increasing interest in the function and detailed structure of the nucleolus. They found that 25% of the frog eggs had no nucleolus and that such eggs were not capable of life. Half of the eggs had one nucleolus and 25% had two. They concluded that the nucleolus had a function necessary for life. In 1966 Max L. Birnstiel and collaborators showed via nucleic acid hybridization experiments that DNA within nucleoli code for ribosomal RNA. [8] [9]

Three major components of the nucleolus are recognized: the fibrillar center (FC), the dense fibrillar component (DFC), and the granular component (GC). [1] Transcription of the rDNA occurs in the FC. [10] The DFC contains the protein fibrillarin, [10] which is important in rRNA processing. The GC contains the protein nucleophosmin, [10] (B23 in the external image) which is also involved in ribosome biogenesis.

However, it has been proposed that this particular organization is only observed in higher eukaryotes and that it evolved from a bipartite organization with the transition from anamniotes to amniotes. Reflecting the substantial increase in the DNA intergenic region, an original fibrillar component would have separated into the FC and the DFC. [11]

Another structure identified within many nucleoli (particularly in plants) is a clear area in the center of the structure referred to as a nucleolar vacuole. [12] Nucleoli of various plant species have been shown to have very high concentrations of iron [13] in contrast to human and animal cell nucleoli.

The nucleolus ultrastructure can be seen through an electron microscope, while the organization and dynamics can be studied through fluorescent protein tagging and fluorescent recovery after photobleaching (FRAP). Antibodies against the PAF49 protein can also be used as a marker for the nucleolus in immunofluorescence experiments. [14]

Although usually only one or two nucleoli can be seen, a diploid human cell has ten nucleolus organizer regions (NORs) and could have more nucleoli. Most often multiple NORs participate in each nucleolus. [15]

In ribosome biogenesis, two of the three eukaryotic RNA polymerases (pol I and III) are required, and these function in a coordinated manner. In an initial stage, the rRNA genes are transcribed as a single unit within the nucleolus by RNA polymerase I. In order for this transcription to occur, several pol I-associated factors and DNA-specific trans-acting factors are required. In yeast, the most important are: UAF (upstream activating factor), TBP (TATA-box binding protein), and core binding factor (CBF)) which bind promoter elements and form the preinitiation complex (PIC), which is in turn recognized by RNA pol. In humans, a similar PIC is assembled with SL1, the promoter selectivity factor (composed of TBP and TBP-associated factors, or TAFs), transcription initiation factors, and UBF (upstream binding factor). RNA polymerase I transcribes most rRNA transcripts 28S, 18S, and 5.8S) but the 5S rRNA subunit (component of the 60S ribosomal subunit) is transcribed by RNA polymerase III. [16]

Transcription of rRNA yields a long precursor molecule (45S pre-rRNA) which still contains the ITS and ETS. Further processing is needed to generate the 18S RNA, 5.8S and 28S RNA molecules. In eukaryotes, the RNA-modifying enzymes are brought to their respective recognition sites by interaction with guide RNAs, which bind these specific sequences. These guide RNAs belong to the class of small nucleolar RNAs (snoRNAs) which are complexed with proteins and exist as small-nucleolar-ribonucleoproteins (snoRNPs). Once the rRNA subunits are processed, they are ready to be assembled into larger ribosomal subunits. However, an additional rRNA molecule, the 5S rRNA, is also necessary. In yeast, the 5S rDNA sequence is localized in the intergenic spacer and is transcribed in the nucleolus by RNA pol.

In higher eukaryotes and plants, the situation is more complex, for the 5S DNA sequence lies outside the Nucleolus Organiser Region (NOR) and is transcribed by RNA pol III in the nucleoplasm, after which it finds its way into the nucleolus to participate in the ribosome assembly. This assembly not only involves the rRNA, but ribosomal proteins as well. The genes encoding these r-proteins are transcribed by pol II in the nucleoplasm by a "conventional" pathway of protein synthesis (transcription, pre-mRNA processing, nuclear export of mature mRNA and translation on cytoplasmic ribosomes). The mature r-proteins are then imported into the nucleus and finally the nucleolus. Association and maturation of rRNA and r-proteins result in the formation of the 40S (small) and 60S (large) subunits of the complete ribosome. These are exported through the nuclear pore complexes to the cytoplasm, where they remain free or become associated with the endoplasmic reticulum, forming rough endoplasmic reticulum (RER). [17] [18]

In human endometrial cells, a network of nucleolar channels is sometimes formed. The origin and function of this network has not yet been clearly identified. [19]

In addition to its role in ribosomal biogenesis, the nucleolus is known to capture and immobilize proteins, a process known as nucleolar detention. Proteins that are detained in the nucleolus are unable to diffuse and to interact with their binding partners. Targets of this post-translational regulatory mechanism include VHL, PML, MDM2, POLD1, RelA, HAND1 and hTERT, among many others. It is now known that long noncoding RNAs originating from intergenic regions of the nucleolus are responsible for this phenomenon. [20]


Cell Culture

N1S1 cells or NISIC3 cells, which stably incorporate a FLAG tagged β subunit of RNA Pol I (15), were grown in RPMI supplemented with 5% horse serum and 1% fetal bovine serum. The preparation of S100 extracts from N1S1 cells has been previously described (6). Where indicated, S100 extracts were prepared from NISIC3 cells treated with 100 μg/ml cycloheximide (CHX) (Sigma, St. Louis, MO) for 1 h. S100 extracts were dialyzed against C-20 (20 mM HEPES, pH 7.9, 20% glycerol, 100 mM KCl, 5 mM MgCl2, 0.2 mM EDTA) in a Pierce (Rockford, IL) Slide-A-Lyzer mini dialysis unit overnight at 4ଌ before use to remove endogenous NTPs. Active, recombinant human FLAG tagged Rrn3 was expressed in Sf9 cells and purified as previously described (8).

DNA Templates

A 920-bp fragment of the rat 45S rDNA promoter was subcloned into the BamHI and EcoRI sites of pUC 19 and used to generate rDNA templates for the immobilized template assays. Wild-type template was amplified with one of two primer pairs using a common 5′ primer (5′-GCTCACTCATTAGGCACC CCAGG-3′), based on pUC 19 sequences upstream of the rDNA insert. The reverse/downstream primers were either 5′-GGAAAACCCTTCCAGTCG-3′ or 5′-GTGCAACTCGGGAGGCACACAG-3′, which generate products of 680 or 857 bp, respectively, containing 90 bp of pUC, and fragments of the rat rDNA gene extending from � to either +303 or +480, respectively. pUC 19 was used as a nonspecific DNA template in some experiments. pUC template was amplified using forward 5′-CAGGGGATAACGCA GG-3′ and reverse 5′-GACGCCGGGCAAGAGCA AC-3′ primers, which generate a 1300-bp product. Primers were purchased from Integrated DNA Technologies (Coralville, IA). PCR products were purified using a Qiagen (Valencia, CA) MinElute PCR purification kit. Following elution from the Qiagen spin columns, the DNA was phenol extracted, ethanol precipitated, and resuspended in 10 mM Tris, pH 8.0, 1 mM EDTA (TE).

Formation, Isolation, and Analysis of PIC Complexes

Five microliters of S100 (5𠄸 μg protein) was incubated with 60 ng PCR template (wild-type or pUC as indicated) for 20 min at room temperature in a total volume of 30 μl. Heparin or sarkosyl (Sigma) were added where indicated in a total volume of 20 μl and the incubation continued for 15 min at 30ଌ. Anti-FLAG M2 agarose beads (Sigma) were washed three times with C-20 and stored as a 50% slurry in the same buffer. Twenty microliters of the 50% slurry of anti-FLAG beads was added and the mixture was tumbled for 1 h at 4ଌ. The slurry was centrifuged at 2,000 rpm (360 × g) for 20 s and the supernatant removed. The beads were washed three times with 100 μl of C-20. The bead pellet was resuspended in 50 μl of distilled water followed by the addition of 5 μl of 1 mg/ml proteinase K (Fisher). The mixture was incubated at 65ଌ for 15 min. The beads were centrifuged and the supernatant removed to a fresh tube. Nucleic acids were purified by extraction with an equal volume of phenol/chloroform/isoamyl alcohol (25:24:1) and the aqueous phase was then extracted with 10 μl of chloroform. The DNA from the entire reaction was amplified using 2 ng of the forward and reverse primers described above with 50 μl of Promega (Madison, WI) PCR Master mix. PCR was performed in a GeneMate Genius (ISC BioExpress, Kays-ville, UT) under the following conditions: 94ଌ, 1 min (1 cycle) 95ଌ, 45 s, 55ଌ, 30 s, 72ଌ, 1 min (35 cycles) 72ଌ, 7 min (1 cycle). Twenty-five microliters of this reaction was further amplified in a second PCR reaction. The second reaction contained 24 ng of each primer and proceeded for 20 cycles. Control reactions demonstrated that these conditions ensure primer excess. Ten microliters from the second reaction was electrophoresed on a 1% agarose gel DNA was visualized by ethidium bromide staining. Gels were scanned on an AlphaEaseFC Imaging System (Alpha Innotech Corp., San Leandro CA). Real-time PCR (RT-PCR) was performed with a Roche Light Cycler (Roche Molecular Biochemicals, Mannheim, Germany). The forward primer was 5′-CCTGTCATGTTTATCCCTG-3′ and the reverse 5′-GGTGCAAGCCTCTTGGAACG-3′, which generates a 135-bp product. For RT-PCR experiments DNA and extract in a final volume of 30 μl were incubated for 10 min on ice, heparin was added at the concentrations indicated to a final volume of 50 μl, and the incubation continued for an additional 30 min at 30ଌ. Twenty microliters of the 50% slurry of anti-FLAG beads was added and the reaction tumbled at 4ଌ for 45 min. The supernatant was removed from the slurry, digested with proteinase K, and the DNA purified as described above. The first round of PCR was the same as for ethidium bromide stained gels. Two microliters of this reaction was further amplified in the second RT-PCR reaction using the Qiagen, QuantiTect SYBR Green PCR kit. PCR conditions were 95ଌ, 15 min (1 cycle) 95ଌ, 1 s, 55ଌ, 10 s, 72ଌ, 27 s (65 cycles).

In Vitro Transcription: Immobilized Template Assays

All templates were generated by PCR and immobilized on avidin-magnetic beads (Dynal Corp, Oslo, Norway) through a biotin incorporated in a common 5′ primer. The two templates were distinguished by the 3′ PCR primer resulting in transcription of either 480- or 303-nt transcripts. Binding of the biotinylated template to the magnetic beads was performed as recommended by the manufacturer. After 60μl of S100 was incubated with 260 ng of immobilized template for 45 min at room temperature, the beads were collected with the magnet. Beads were washed with C-20 and the incubations continued at 30ଌ followed in some experiments by a second wash with C-20. Recombinant Rrn3 was added to some incubation (300 ng) at the time points indicated. Transcription was initiated by the addition of NTPs ± [α- 32 P]UTP as indicated. The products of transcription were analyzed by denaturing urea/PAGE as previously described (44). In vitro transcription reactions using templates in solution were performed using 0.1 μg p5.1E/X linearized with EcoRI as template per reaction as previously described (44). The reactions were supplemented with heparin or sarkosyl as indicated.

Western Blot Analysis

SDS-PAGE and electroblot analysis were performed as described previously (15). Polyclonal rabbit antisera to mouse rpa43 were raised to recombinant rpa43 expressed in E. coli (Capralogics, Inc., Hardwick, MA). Monoclonal anti-FLAG antibody (M2) was purchased from Sigma Chemical Co. (St. Louis, MO) and was used as previously described (15).


SHU1 overexpression is toxic in uaf30Δ strains

To identify novel genes that genetically interact with SHU1, an SDL screen was conducted using an overexpressing SHU1 plasmid introduced into the yeast deletion library. The SHU1 gene was cloned under a copper-inducible promoter into a 2-μm vector. In the presence of increasing copper concentrations, SHU1 expression was induced and visualized by protein blot using a FLAG-tagged Shu1 (Supplemental Figure 1A). Overexpression of SHU1 or SHU1-FLAG does not cause any growth defects (Supplemental Figure 1B) and complements the MMS sensitivity of a shu1Δ strain (Supplemental Figure 1C), demonstrating that the SHU1 plasmids encode functional alleles. The untagged SHU1 plasmid was introduced separately into the ∼4800 viable haploid strains of the yeast deletion library, and nine disruptions failed to grow when SHU1 was overexpressed. One of these disruptions, uaf30Δ, was confirmed for its SDL interactions after direct transformation of the SHU1 plasmid into the deletion strain (Figure 1A and Supplemental Figure 1A). An SDL interaction is seen even without the addition of copper, suggesting that the basal level of Shu1 expression, likely due to small amounts of copper in the medium, is sufficient to cause this phenotype. This observation is consistent with the finding that the uninduced SHU1 plasmid suppresses the MMS sensitivity of a shu1Δ strain (Supplemental Figure 1C).

FIGURE 1: Overexpression of SHU1 causes an SDL interaction with deletion of UAF30, a gene that alters rDNA recombination in a FOB1-dependent manner. (A) Fivefold serial dilutions were plated onto selective media with increasing copper concentrations with the empty vector (pWJ1530) or the SHU1 overexpression plasmid (SHU1) transformed into WT or uaf30Δ. (B) Fluorescence microscopy of yeast cells containing Uaf30-YFP and Top1-CFP was analyzed for colocalization (Merge). (C) The frequency of rDNA recombinants (CAN R , ade2) was measured in WT, uaf30Δ, fob1Δ, and uaf30Δ fob1Δ yeast strains, and they were plotted with SD. Note that rDNA recombination frequencies vary in uaf30Δ cells, likely due to the destabilization of the rDNA array.

Uaf30 is a nucleolar protein important for regulating rDNA recombination

Uaf30 was originally copurified as a component of the UAF complex, which promotes transcription of the rDNA by RNA polymerase I and represses RNA polymerase II (Siddiqi et al., 2001 Hontz et al., 2008). Because UAF functions in the nucleolus at the rDNA, we analyzed cells expressing Uaf30–yellow fluorescent protein (YFP) and Top1–cyan fluorescent protein (CFP), a known protein that resides in the nucleolus (Edwards et al., 2000 Huh et al., 2003), for their localization. Figure 1B shows that Uaf30-YFP colocalizes with Top1-CFP in the nucleolus. These results confirm a previously published genome-wide study in which Uaf30 was localized to the nucleolus (Huh et al., 2003) and contradict another genome-wide report in which it was found to be cytoplasmic (Huang et al., 2003).

Because Uaf30 is nucleolar and the absence of other components of the UAF complex results in rDNA expansion, we examined the potential role of UAF30 in rDNA recombination using a marker loss assay. In this assay, the ADE2 and CAN1 genes were inserted into one of the 100–200 rDNA repeats, and recombination frequencies were calculated by measuring the simultaneous loss of both markers. Figure 1C shows that uaf30Δ cells exhibit increased rDNA recombination frequencies compared with the wild-type (WT) parental strain. Furthermore this increase is dependent upon FOB1, a gene important for rDNA replication fork stalling that leads to a low frequency of spontaneous DSBs in the array (Figure 1C) (Kobayashi et al., 1998, 2004 Burkhalter and Sogo, 2004).

Shu1 also localizes to the nucleolus and affects rDNA recombination

Because Uaf30 localizes to the cell nucleolus and uaf30Δ cells are sensitive to SHU1 overexpression, we asked whether Shu1 is also localized to the nucleolus. Previously we found that a doubly YFP-tagged Shu1 localizes to the nucleus, but we did not look carefully at its nucleolar localization (Shor et al., 2005). Here we show, upon further analysis, that Shu1 is also found in the nucleolus as it colocalizes with Nop1-CFP, a known nucleolar protein (Figure 2A).

FIGURE 2: Shu1 localizes to the nucleolus and affects rDNA recombination. (A) Yeast with Shu1-YFP-YFP and Nop1-CFP were analyzed by fluorescence microscopy for colocalization (Merge). (B) The frequency of rDNA recombinants (CAN R , ade2) was measured in WT, shu1Δ, uaf30Δ, and uaf30Δ shu1Δ yeast strains, and they were plotted with SD. (C) The amounts of rDNA in WT, shu1Δ, uaf30Δ, and uaf30Δ shu1Δ strains were quantitated by analyzing the amount of rDNA resulting from restriction digest of total DNA, revealing a 9.1-kb fragment as described in Materials and Methods. SD are plotted. (D) WT and uaf30Δ strains were transformed with the empty vector (pWJ1530) or the SHU1 overexpression plasmid (pWJ1530-SHU1). Strains were grown in the presence of 100 μM copper (CuSO4), and the frequency of rDNA recombinants (CAN R , ade2) was measured and the SE are plotted. Note that the recombination frequencies in uaf30Δ cells were different relative to (B), likely due to their growth in synthetic minimal medium, which was needed to maintain the plasmid.

Because the Shu complex was previously found to promote recombination through HR (Shor et al., 2005 Mankouri et al., 2007), we hypothesized that in its absence, recombination would be suppressed at the rDNA. Indeed, we found that shu1Δ largely suppresses the increased rDNA recombination of uaf30Δ cells (Figure 2B). In contrast, in the absence of Shu1 alone, rDNA recombination is neither increased nor decreased (Figure 2B and data not shown). These results suggest that Shu1 functions to promote HR processing of rDNA recombination intermediates created by uaf30Δ. Furthermore these findings are not specific to disruption of SHU1 because disruption of other SHU genes (i.e., SHU2 and CSM2) also lower the increased rDNA recombination frequency of uaf30Δ cells (Supplemental Figure 2). In contrast, deletion of PSY3 does not significantly lower uaf30Δ-increased rDNA recombination and is highly variable (Supplemental Figure 2). In addition, csm2Δ cells alone increase rDNA recombination relative to WT (Supplemental Figure 2). Indeed, different members of the Shu complex can have distinct functions in various processes because differences between Shu complex members have been reported for gross chromosome rearrangements (Huang et al., 2003) and Rad52 focus formation and gene conversion rates in S. pombe (Martin et al., 2006).

Disruption of SHU1 decreases the rDNA recombination levels observed in uaf30Δ cells. To determine whether SHU1 disruption suppresses rDNA instability in general, we analyzed the effect of deleting SHU1 on the increased rDNA recombination that is observed in top1 mutants (Christman et al., 1988 Gangloff et al., 1996). We found that this increased recombination frequency is not significantly altered in a shu1Δ top1Δ double mutant, supporting the notion that the genetic interaction between the Shu complex and Uaf30 is specific (Supplemental Figure 3).

In the absence of the UAF genes, cells expand their rDNA (Nogi et al., 1991 Keys et al., 1996). Because Uaf30 is a component of the UAF complex, we assumed that uaf30Δ strains would exhibit increased rDNA copy number, and indeed they do (Figure 2C). Because Shu1 largely suppresses the recombination frequency observed in uaf30Δ strains (Figure 2B), we hypothesized that its disruption would also suppress the increased size of the rDNA array observed in uaf30Δ. As predicted, shu1Δ uaf30Δ double mutants exhibit a reduced rDNA copy number compared with uaf30Δ (Figure 2C). However, shu1Δ shows a slight increase in rDNA copy number relative to WT (Figure 2C).

Finally, we asked whether overexpressing Shu1 would further increase rDNA recombination because the Shu complex is genetically involved in promoting recombination. Using the copper-inducible SHU1 plasmid characterized in Supplemental Figure 1, we found that the increased rDNA recombination observed in uaf30Δ cells is exacerbated by SHU1 overexpression (Figure 2D). Given that SHU1 overexpression is toxic in a uaf30Δ background, the recombination frequencies may actually be even higher than those observed if recombination events lead to death or growth arrest of these cells.

Shu1 functions in the same pathway as Srs2

Two of the Shu complex proteins, Shu1 and Psy3, are Rad51 paralogues. The Rad51 paralogues, along with Rad52, promote Rad51 filament formation. Importantly, in both budding and fission yeast, another member of the Shu complex, Shu2, physically interacts with the Srs2 helicase, which disrupts Rad51 filaments (Ito et al., 2001 Krejci et al., 2003 Veaute et al., 2003 Martin et al., 2006). Genetic and biochemical studies suggest that Srs2 functions as an anti-recombinase regulating Rad51-mediated strand exchange (Aboussekhra et al., 1989 Krejci et al., 2003 Veaute et al., 2003). Based upon these observations, we hypothesized that the Shu complex and Srs2 may be involved in the same pathway controlling rDNA hyperrecombination in uaf30Δ. Because Srs2 was previously shown to be enriched in the nucleolus (Torres-Rosell et al., 2007), we examined whether srs2Δ mutant cells, like shu1Δ, can also suppress increased rDNA recombination seen in the absence of UAF30. Disruption of SRS2 alone modestly increases the frequency of rDNA recombination (threefold over WT), consistent with its hyperrecombination phenotype observed at other loci. When combined with a uaf30Δ, the uaf30Δ srs2Δ double mutant shows similar recombination rates to uaf30Δ shu1Δ, indicating that both srs2Δ and shu1Δ suppress the uaf30Δ defect (Figures 3A and 2B). We also find that the uaf30Δ srs2Δ shu1Δ triple mutant exhibits the same level of suppression, suggesting that Srs2 and Shu1 function in the same pathway in response to uaf30Δ-induced DNA damage at the rDNA. Consistent with this view, we do not observe any synthetic growth defect in srs2Δ uaf30Δ double-mutant strains, unlike that reported in a genome-wide study (Pan et al., 2006).

FIGURE 3: Shu1 functions in the same pathway as Srs2 to suppress uaf30Δ rDNA recombination and alters Srs2 focus formation. (A) The frequency of rDNA recombination was measured in WT, shu1Δ, srs2Δ, shu1Δ srs2Δ, uaf30Δ, uaf30Δ srs2Δ, and uaf30Δ srs2Δ shu1Δ strains, and they were plotted with SD. Note the recombination frequency of the uaf30Δ shu1Δ strain was not conducted at the same time. (B) YFP-Srs2–expressing strains were analyzed for the percentage of spontaneous nuclear foci in WT, shu1Δ, uaf30Δ, and shu1Δ uaf30Δ cells. Images of Srs2 are shown with white arrowheads indicating foci. Each experiment was done in triplicate with a total of 400–500 cells analyzed. The graph shows the percentage of cells with foci along with the SE. (C) Cells expressing CFP-Rad51 were analyzed in WT and shu1Δ strains for the percentage of spontaneous nuclear foci. Each experiment was done in triplicate with a total of 150–200 cells analyzed with SE plotted. Note that the strains also contain a WT Rad51–complementing plasmid because CFP-Rad51 is not fully functional.

Shu1 affects Srs2 and Rad51 focus formation both spontaneously and at site-specific breaks

In vivo, Srs2 forms foci at sites of DNA replication and recombination, where it removes Rad51 nucleoprotein filaments (Burgess et al., 2009). Because Shu1 and Srs2 genetically interact, it is possible that Shu1 normally promotes recombination by inhibiting the anti-recombinase function of Srs2. To test this hypothesis, we analyzed whether the number of Srs2 foci change in shu1Δ, uaf30Δ, or shu1Δ uaf30Δ cells (Figure 3B). Interestingly, shu1Δ and shu1Δ uaf30Δ strains show an increased number of Srs2 foci (p ≤ 0.005 Figure 3B). In contrast, uaf30Δ strains, which exhibit increased rDNA recombination frequency, have fewer Srs2 foci relative to WT (p ≤ 0.05 Figures 1C and 3B). Because shu1Δ cells increase the number of spontaneous Srs2 foci, which may in turn increase Srs2 anti-recombinase activity, we analyzed whether the ability of Rad51 to form recombination foci is impaired by SHU1 disruption (Figure 3C). We find that the number of spontaneous Rad51 foci in a shu1Δ strain is decreased relative to WT (p ≤ 0.05 Figure 3C). Altogether these results are consistent with the notion that, in absence of Shu1, the activity of Srs2 is increased.

To test directly whether Shu1 regulates Srs2 recruitment to DSB sites, we took advantage of a system where an I-SceI endonuclease cut site was inserted into one rDNA repeat (Torres-Rosell et al., 2007) or outside the rDNA at the URA3 locus on chromosome V (Lisby et al., 2004) (Figure 4). In addition, a tandem array of Tet repressor–binding sites (224xtetO or 336xtetO, respectively) was positioned adjacent to each cut site. The localization of the cut site is revealed by expression of TetI fused to a monomeric red fluorescent protein (mRFP), which binds to TetO. Using this system, we analyzed the localization of fluorescently tagged Srs2 or Rad52 with respect to the DNA cut site after inducing a DSB in both WT and shu1Δ cells (Figure 4, A and B). Rad52, a central DNA repair protein, was used to monitor the efficiency of cutting of the DSBs (Lisby et al., 2004 Torres-Rosell et al., 2007). We find that Rad52 is recruited to either the rDNA break or a break in chromosome V, even in the absence of SHU1 (p ≤ 0.01). In contrast, Srs2 recruitment to either DSB increases significantly in a SHU1 disruption (Figure 4, A and B, p ≤ 0.025 and p ≤ 0.05, respectively). These results show that Shu1 normally functions to inhibit Srs2 recruitment to DSBs.

FIGURE 4: Shu1 inhibits Srs2 recruitment to DNA breaks. (A) An I-SceI cut site was integrated into the rDNA adjacent to a tandem array of Tet repressor–binding sites (224xtetO). Location of the rDNA break is revealed by expression of a TetI fused to mRFP. Rad52-CFP and YFP-Srs2 were monitored for their recruitment to rDNA breaks in WT and shu1Δ cells expressing a GAL-I-SceI plasmid. The results are quantitated in the graph with SE plotted. (B) An I-SceI cut site was integrated at the URA3 locus on chromosome V adjacent to a tandem array of the Tet repressor–binding sites (336xtetO). Rad52-CFP and YFP-Srs2 were monitored in WT and shu1Δ cells for recruitment to the cut site in strains expressing a GAL-I-SceI plasmid. The results are quantitated in the graph with SE plotted.

The simplest hypothesis to explain these observations is that the Shu complex normally inhibits Srs2, and in its absence, increased Srs2 activity alters the equilibrium to remove Rad51 filaments. An alternative explanation is that the Shu complex is directly involved in Rad51 filament formation (acting like a mediator), and in its absence, fewer filaments are formed. If the latter explanation were correct, then Rad51 focus formation would be reduced in a shu1Δ, as we observe in Figure 3C, but would also be reduced a shu1Δ srs2Δ double mutant because the presence or absence of Srs2 should not alter Rad51 filament formation. Indeed, when we disrupt the Rad51 filament mediator, RAD55, fewer Rad51 foci are seen in the absence of SRS2 (Supplemental Figure 4 p ≥ 0.025). Alternatively, if Shu1 inhibits Srs2 directly, then Rad51 focus formation would increase in a shu1Δ srs2Δ double mutant relative to WT, which is precisely what we found. We find that the shu1Δ srs2Δ double mutant exhibits as many Rad51 foci as the srs2Δ strain (Figure 5 p ≥ 0.05). Although we have not completely ruled out that the Shu complex plays some role in Rad51 filament formation, our results strongly suggest that a major role of the Shu complex is to inhibit Srs2.

FIGURE 5: Rad51 filament formation is not inhibited in shu1Δ srs2Δ cells. WT, shu1Δ, srs2Δ, and shu1Δ srs2Δ cells were analyzed for the percentage of spontaneous CFP-Rad51 foci. Each experiment was done in triplicate with a total of 200 cells analyzed with SE plotted. Note that the strains also contain a WT Rad51–complementing plasmid because CFP-Rad51 is not fully functional. This configuration likely results in fewer Rad51 foci observed in srs2Δ cells than we previously reported (Burgess et al., 2009).

Materials and methods

Plasmids, strains and media

Plasmids and yeast strains used in this study are listed in Tables I and II, respectively. Standard yeast genetic techniques and media were used ( Sherman et al., 1979 ).

Plasmid Description Vector Reference/donor
Yeast plasmids
pGP5 CEN, TRP1, RPA43 pRS314 this study
pGP5-4 CEN, TRP1, rpa43–4 pRS314 this study
pGP5-6 CEN, TRP1, rpa43–6 pRS314 this study
pGP5-14 CEN, TRP1, rpa43–14 pRS314 this study
pGP5-18 CEN, TRP1, rpa43–18 pRS314 this study
pGP5-24 CEN, TRP1, rpa43–24 pRS314 this study
pGP32 2μ, LEU2, GAL4[768–881]-RRN6[865–894] pACT2 this study
pGP40 2μ, LEU2, GAL4[768–881]RRN6[772–8894] pACT2 this study
pGP44 2μ, LEU2, GAL4[768–88]RRN6 pACT2 this study
pAS-RRN3 2μ, TPR1, GAL4[1-147]-RRN3 pASΔ J.Steffan
yCPA43 CEN, URA3, RPA43 yCP Thuriaux et al. (1995)
pSYCYES2 2μ, URA3, RRN3 pSYC Cadwell et al. (1997)
pL1 2μ, URA3, SPT15 pFL44 Cavallini et al. (1989)
pNOY102 2μ, URA3, RDN under control of the GAL7 promoter pC1/1 Nogi et al. (1991)
pSIRT mini 26S Ye30-Δ6 Musters et al. (1989)
Plasmids for expression in E.coli
pGP4 HA-HIS-RPA43 under control of the T7 promoter pRSET5d this study
pGP34 HA-HIS-RRN3 under control of the T7 promoter pRSET5d this study
pGP37 RRN6[772–894] under control of the T7 promoter pRSET5d this study
pGP45 A43 under control of the T7 promoter pACYC184-11b this study
pGP47 GST-RRN6[772–894] under control of the T7 promoter pGEX3X this study
Strain Description Reference
GPy9 MATa rpa43::LEU2 ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1/pGP5 this study
GPy11-4 MATa rpa43::LEU2 ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1/pGP5-4 this study
GPy11-6 MATa rpa43::LEU2 ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1/pGP5-6 this study
GPy11-14 MATa rpa43::LEU2 ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1/pGP5-14 this study
GPy11-18 MATa rpa43::LEU2 ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1/pGP5-18 his study
GPy11-24 MATa rpa43::LEU2 ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1/pGP5-24 this study
GPy21 MATα rpa43::LEU2 rrn3-8 ura3-52 trp1 leu2 his3Δ200 his7Δ2 ade/yCPA43 this study
GPy43 MATα rpa43::LEU2 rrn3-8 ura3-52 trp1 leu2 his3Δ200 his7Δ2 ade/pNOY200 this study
GPy44 MATα rpa43::LEU2 rrn3-8 ura3-52 trp1 leu2 his3Δ200 his7Δ2 ade/pNOY102 this study
GPy52 MATa ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1 rrn6-Δ1 this study
MATα ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1 RRN6
D101-I2 MATa rpa43::LEU2 ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1/yCPA43 Thuriaux et al. (1995)
D128 MATα rpa43::LEU2 ade2-101 ura3-52 lys2-801 trp1-Δ63 his3Δ200 leu2-Δ1/pNOY102 Thuriaux et al. (1995)
yCC95 MATα rrn3-8 ade5 ura3-52 trp1-289 his7-2 leu2-112 Cadwell et al. (1997)
Y190 MATa ade2-101 trp1-901 his3 leu2-3,112 gal4 gal80 ura3-52::GAL1-lacZ::URA3 lys2::GAL1-HIS3 cyh R Harper et al. (1993)

Random mutagenesis

rpa43 mutant alleles were obtained by mutagenic PCR (200 μM MnCl2) and gap repair as described ( Muhlrad et al., 1992 ).

Purification of the B600 and B2000 fractions

Purification of the B600 and B2000 fractions and the Pol I–Rrn3 and TBP-containing complexes was performed as previously described ( Milkereit et al., 1997 ).

In vitro transcription assays

Non-specific activity of RNA Pol I was assayed as described ( Buhler et al., 1974 ). Specific transcription assays were performed as described ( Milkereit et al., 1997 ) with 40 ng of plasmid pSIRT ( Musters et al., 1989 ). Transcripts were separated by electrophoresis on a 6% polyacrylamide gel containing 7 M urea in TBE and submitted to autoradiography.

Preparation of recombinant A43, Rrn3 and Rrn6 proteins

Recombinant Rrn3 and Rrn6 proteins expressed in E.coli. Transformed BL21(DE3) cells were grown at 24°C in LB medium supplemented with ampicillin (500 μg/ml), chloramphenicol (34 μg/ml) and sorbitol (1 M). Expression of the recombinant protein was induced at 0.4 OD600 with 250 μM isopropyl-β- D -thiogalactopyranoside (IPTG). Cells were shaken for 4 h at 24°C, collected and resuspended in breaking buffer [100 mM Tris–HCl pH 8, 20% glycerol, 400 mM KOAc, 1 mM EDTA, 5 mM Mg(OAc)2, 5 mM β-mercaptoethanol and 0.1% Tween 20] supplemented with protease inhibitors (Complete Boehringer). Cells were lysed by two cycles of freezing and thawing in liquid nitrogen. Lysozyme (20 μg/ml) and benzonase (100 U/ml) were added and the suspension was incubated for 1 h at 4°C. Crude extract was centrifuged in a Beckman 45-Ti rotor for 1 h at 40 000 r.p.m.

Recombinant A43 and Rrn3 co-expressed in E.coli. Cell culture and induction were performed as described above. Cells were resuspended in breaking buffer [50 mM Tris–HCl pH 8, 20% glycerol, 100 mM KOAc, 5 mM Mg(OAc)2 and 0.1% Tween 20] supplemented with protease inhibitors and were lysed with an Eaton press and the crude extract was centrifuged as above.

Partial purification of rRrn3. The S100 fraction from a 12 l culture was loaded on a Q-Sepharose High-Load 20 ml column equilibrated with Q buffer [20 mM Tris–HCl pH 8, 20% glycerol, 5 mM Mg(OAc)2 and 0.1% Tween 20] containing 400 mM KOAc. After washing, proteins were eluted with a linear gradient from 400 mM to 2 M KOAc in Q buffer. Rrn3-containing fractions, as determined by western blot analysis using monoclonal anti-HA antibodies (16B12 Babco), were pooled and supplemented with 5 mM imidazole. The pool was loaded onto a 1 ml Hi-Trap nickel column equilibrated with Ni buffer [20 mM Tris–HCl pH 8, 20% glycerol, 500 mM KOAc, 5 mM Mg(OAc)2 and 0.1% Tween 20] containing 5 mM imidazole. The column was developed with a linear gradient from 5 to 200 mM imidazole in Ni buffer Rrn3-containing fractions were pooled and dialysed against 20 mM Tris–HCl pH 8, 20% glycerol, 50 mM KOAc, 5 mM Mg(OAc)2, 5 mM β-mercaptoethanol and 0.1% Tween 20.

[ 35 S]Rrn6[C1] synthesis. [ 35 S]Rrn6[C1] was synthesized in a wheatgerm extract (Promega) supplemented with T7 RNA polymerase (Promega), [ 35 S]methionine (0.8 mCi/ml) and 1 μg of purified plasmid pGP37.

Affinity chromatography

Chromatography of [ 35 S]Rrn6[772–894] was on a column of Rrn3 immobilized to protein G–Sepharose beads (Pharmacia) coated with 45 μg of purified 12CA5 antibodies, equilibrated in breaking buffer with 1% (w/v) low fat milk. S100 containing or not HA-tagged Rrn3 (1 ml) was then passed five times through the beads. The beads were extensively washed with breaking buffer supplemented with 1% (w/v) low fat milk then with binding buffer [20 mM Tris–HCl pH 8, 20% glycerol, 50 mM KOAc, 5 mM Mg(OAc)2, 5 mM β-mercaptoethanol and 0.1% Tween 20]. An aliquot of 50 μl of the [ 35 S]Rrn6[772–894] mixture, diluted in 300 μl of binding buffer, was then loaded five times onto the same column. After extensive washing with binding buffer, proteins were eluted at 37°C with 1 ml of binding buffer containing 1 μg/μl purified HA-peptide. Proteins were subjected to SDS–PAGE and analysed by autoradiography on BiomaxMS films (Amersham).

Chromatography of rRrn3 was on a column of immobilized GST–Rrn6[772–894]. Glutathione–Sepharose beads were prepared as above and 150 μl of partially purified HA-His6-Rrn3 were loaded on the column. Proteins were eluted with 1 ml of binding buffer supplemented with 30 mM glutathione and analysed by western blotting using monoclonal anti-HA antibodies (16B12 Babco).

Two-hybrid interaction between Rrn3 and Rrn6

The two-hybrid screening was performed essentially as described ( Flores et al., 1999 ). Strain Y190 transformed with pAS-Rrn3 was transformed with a DNA genomic library ( Fromont-Racine et al., 1997 ). Transformants were selected on 50 and 75 mM 3AT-containing medium and tested for activation of the lacZ reporter gene.

Specimen preparation for electron microscopy and image processing

Formation of immune complexes and image analysis of the data were performed as described by Klinger et al. (1996) .


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