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TATA binding protein (TBP) plays a central role in transcription complex assembly and is regulated by a variety of transcription factors, including Mot1. Mot1 is an essential protein in Saccharomyces cerevisiae that exerts both negative and positive effects on transcription via interactions with TBP. It contains two conserved regions important for TBP interactions, another conserved region that hydrolyzes ATP to remove TBP from DNA, and a fourth conserved region with unknown function. Whether these regions contribute equally to transcriptional regulation genome-wide is unknown. Here, we employ a transient-replacement assay using deletion derivatives in the conserved regions of Mot1 to investigate their contributions to gene regulation throughout the S. cerevisiae genome. These four regions of Mot1 are essential for growth and are generally required for all Mot1-regulated genes. Loss of the ATPase region, but not other conserved regions, caused TBP to redistribute away from a subset of Mot1-inhibited genes, leading to decreased expression of those genes. A corresponding increase in TBP occupancy and expression occurred at another set of genes that are normally Mot1 independent. The data suggest that Mot1 uses ATP hydrolysis to redistribute accessible TBP away from intrinsically preferred sites to other sites of intrinsically low preference.
From Saccharomyces cerevisiae to human, the TATA binding protein (TBP) provides an indispensable role in nearly all RNA polymerase I, II, and III transcription events (29). TBP is the central component of a complex regulatory network governing transcription complex assembly (33). Consequently, TBP is subjected to an extraordinary level of regulation by numerous transcription factors, one of which is Mot1 (6). Mot1 is a conserved Snf2/Swi2-related ATPase (21) that regulates the dynamics of TBP-promoter interactions by removing TBP from DNA using the energy of ATP hydrolysis (6, 45, 47). The first 800 amino-terminal residues of Mot1 are both necessary and sufficient for TBP binding (2, 7). The Snf2-related ATPase domain resides within the last 600 carboxy-terminal residues (7). Genome-wide expression studies using temperature-sensitive mot1 alleles indicate that Mot1 regulates between 3 and 15% of the yeast genome, some negatively and others positively (4, 17, 24).
In vitro biochemical experiments have largely defined mechanisms by which Mot1 regulates TBP-DNA interactions. Mot1 can bind and stabilize TBP-DNA interactions, but in the presence of ATP, it dissociates TBP from DNA and, to some extent, Mot1 from TBP, thereby recycling both (2, 7, 12, 26). This reaction is important for two reasons. First, dynamic assembly and disassembly of the transcription machinery impart precise control over gene expression. Therefore, regulated recruitment of TBP to promoters must be accompanied by regulated removal of TBP, which Mot1 is well suited to do. In this context, Mot1 is a negative regulator.
Since TBP binds to the minor groove of DNA, which has limited sequence specificity, TBP has relative high affinity for nonspecific DNA (14). If bound inappropriately, this might lead to aberrant or nonproductive assembly of the transcription machinery. Biochemical experiments have demonstrated that Mot1 can remove nonspecifically bound TBP (41), perhaps acting as a chaperone allowing TBP to rebind in a productive mode. For example, at the URA1 gene, Mot1 can promote transcription by removing a nonproductive TBP bound in the reverse orientation (46). In this context, Mot1 operates as a positive regulator.
The mechanism by which Mot1 acts on TBP is well defined biochemically, and this provides a basis for interpreting less defined in vivo experiments. Because Mot1 is essential for growth in S. cerevisiae (19), in vivo functional analysis of important regions of Mot1 is not straightforward in that loss of function is lethal. Thus, an investigation into the genome-wide functions of essential proteins like Mot1 is hampered by the practical limitation that mutations that eliminate function cause cell death. Temperature-sensitive mutations might alleviate this problem to some extent, but they are difficult to target to specific regions of the protein and often vary in severity. To circumvent this limitation, we utilized a transient-replacement strategy (34) to investigate the contributions of conserved Mot1 domains to transcription and TBP recruitment genome-wide. Our study revealed that Mot1-regulated transcription is dependent on nearly all conserved regions of Mot1. Strikingly, transcriptional dependence for a subset of genes is specifically altered when the Mot1 ATPase domain is deleted. Genome-wide location analysis of TBP in a strain that lacks the Mot1 ATPase region corroborates the expression-profiling experiments, suggesting a direct effect on TBP. Furthermore, coimmunoprecipitation of TBP and the Mot1 ATPase deletion mutant demonstrates that the two directly interact. These findings reveal that Mot1-regulated genes are generally regulated by all parts of Mot1 and that a TBP-binding portion of Mot1 can alter the selectivity of TBP for promoters.
pCALF-T(PGK) (36) was converted to pCALF-FHT-T(PGK) 2.2 by inserting a 66-bp HIS-TEV oligonucleotide into the NdeI site. pUG6-FHT-P (4,170 bp) was made by PCR amplifying 259 bp containing the FHT (Flu-His-TEV) sequence from the pCALFHT-T(PGK) 2.2 plasmid. The PCR product was digested with SalI, and 161 bp was ligated into the SalI-digested pUG6 plasmid (4,009 bp) so that the orientation was FHT-loxP-kanMX-loxP.
A list of the yeast strains used in this study is provided in Table 1. S. cerevisiae strain BY4743 (9) (Invitrogen) was used as the parental strain. Initially, the strain was transformed with pSH47 (URA3) (25) encoding galactose-inducible Cre recombinase. Oligonucleotides (70-mer) were used to PCR amplify 1,991 bp of pFA6a-His3MX6-PGAL1 containing the HIS3 gene and GAL1 promoter (38). The PCR product was transformed into BY4743 using a high-efficiency lithium acetate method (25) to replace 550 bp of the endogenous MOT1 promoter with the GAL1 promoter, creating strain yLAC1. HIS+ homologous-recombination transformants were selected on complete synthetic medium lacking histidine and uracil (CSM-HIS-URA medium) and verified by colony PCR.
Regions of MOT1 were deleted by replacing coding sequences with an FHT tag. The FHT tag encodes three hemagglutinin (HA) (Flu) repeats, a decahistidine (H), sequence, and the TEV protease sequence (T). The kanamycin resistance region of pUG6-FHT-p was PCR amplified with 68-mer oligonucleotides with 50-bp homology to distinct regions of MOT1. The PCR products were transformed into yLAC1 and selected on CSM-HIS-URA (dextrose) plates containing 500 mg/ml G418 (Invitrogen). The kanamycin resistance cassette flanked by loxP sites was removed by induction of Cre recombinase with 2% galactose for 4 h, leaving the FHT tag coding sequence upstream of the mutation in MOT1. Kanamycin-sensitive colonies were identified by replica plating them on media containing and lacking G418. Additionally, mutations were verified by colony PCR with primers specific to each mutation.
Kanamycin-sensitive FHT-Mot1 strains (Table 1) were plated on CSM-HIS plus 5-fluoroorotic acid (5-FOA) to select cells that had lost pSH47 and verified by replica plating on CSM-HIS and CSM-HIS-URA. The strains were then transformed with pMR13 (MOT1 wild type [WT] and URA3), and transformants were selected on CSM-HIS-URA medium. The strains were plated on presporulation medium (1% yeast extract, 2% peptone, and 10% dextrose) for 2 days at 30°C. Cells were cultured in sporulation medium (0.3% potassium acetate, 0.02% raffinose) for 3 days at 30°C. Two hundred microliters of the culture was pelleted; resuspended in 1.2 M sorbitol, 10 mM Tris, pH 7.4; and treated with 20 units of (1 mg/ml) zymolyase (MP Biomedicals) at room temperature for 20 min. Tetrads were dissected according to standard yeast techniques on YPD (yeast-peptone-dextrose) plates. Spores were replica plated onto CSM-HIS-URA medium to select for the HIS3 gene (and therefore the GAL1 promoter). Mating types of the mot1 strains were confirmed with MATa and MATa sex tester strains. MATa leu− HIS+ LYS+ tetrads were selected. The strains were then transformed with pMOT221 (mot1-42 LEU2) or pAV20 (MOT1 WT LEU2) and selected on CSM-LEU medium. Cells that lost pMR13 (MOT1 WT URA3) were selected by plating them on CSM-LEU plus 5-FOA.
For colony PCR, 1× 25 mM MgCl2 buffer (Gene Choice), 2.5 U Taq polymerase (Gene Choice), 0.0002 U Pfu polymerase (Stratagene), 0.4 mM deoxynucleoside triphosphates (dNTPs), and 0.2 mM each primer were used per 50-ml reaction for 32 cycles.
MATa haploid FHT-Mot1 mutant strains carrying pMOT221 (mot1-42 LEU2) were grown at 25°C in YPR (yeast-peptone-3% raffinose) to mid-log phase. Cells (A600 = 0.5) were removed, and 2.5 μl of 10-fold serial dilutions was spotted on three sets of YPD (2% dextrose) and YPG (2% galactose) plates and incubated at 25°C, 30°C, or 37°C. Photographs were taken after 48 h.
Microarrays were performed essentially as described previously (13, 32). Briefly, cultures were grown to an A600 of ~0.6, induced with 2% galactose in YPR medium for 60 min at 25°C, and shifted to 37°C for 45 min to inactivate the temperature-sensitive copy of Mot1 encoded by the mot1-42 allele. Cells were harvested by centrifugation at room temperature, washed in RNase-free DEPC (diethyl pyrocarbonate)-treated double-distilled H2O, and frozen in liquid nitrogen.
Total RNA was isolated as described previously (32), and poly(A) tail mRNA was purified using oligo(dT) cellulose (Ambion) according to the manufacturer's instructions. Reverse transcription, labeling with fluorescent dyes (Cy3 and Cy5; Amersham), hybridization, and scanning were all performed as described previously (13, 32). Four micrograms of mRNA was used for hybridizations instead of the conventional 2 μg. Slides were treated with Dye Saver2 (Genisphere) according to the manufacturer's instructions to preserve signal intensity. Data sets were mode normalized by using R software (Bioconductor) to mode-center replicates (dye swaps) (data are available at http://atlas.bx.psu.edu/).
Genes were filtered by several criteria to minimize false positives. (i) Genes were eliminated if their signals on the array were greater than 25% saturated. (ii) The mean foreground signal minus the median background signal had to be greater than the standard deviation of background signal. (iii) Quality data were needed from both replicates of the dye swap. (iv) The directional change of the mutant's signal (relative to the reference) had to be equivalent in the replicates. False-discovery rates (FDRs) were determined using a modified version of a method described previously (35). The false-discovery rate is reported as a percentage of the number of expression changes above and below a given threshold (±0.59; log2 scale) in the homotypic control (yjdi420) compared to each FHT-Mot1 mutant expression experiment (yjdi410 to -416). The FDRs are 4.3%, 2.0%, 2.4%, 3.0%, and 2.9% for TBP1, TBP2, UK1, ATP1, and Null, respectively.
K-means clustering was performed using Cluster (22) on 515 genes that contained data in 80% of the experiments and that had a change of at least 1.5-fold (log2 ratio = 0.59) in one of the mutants. K was chosen to equal 6 clusters (K = 6). Clusters 5 and 6 were merged because they were visually indistinguishable. Clustering information was visualized using Treeview (22).
ChIP-chip was performed essentially as described previously (48), with minor changes noted below. Briefly, cultures were grown to an A600 of ~0.6, induced with 2% galactose in YPR medium for 60 min at 25°C, and shifted to 37°C for 45 min to inactivate the temperature-sensitive copy of Mot1 encoded by the mot1-42 allele. The cells were then fixed by adding formaldehyde to a final concentration of 1% for 2 h at 25°C (instead of a typical 15-min cross-link time) and quenched for 5 min with glycine. The cultures were diluted 2-fold with the same volume of temperature-adjusted distilled water just prior to addition of formaldehyde to achieve a medium temperature of 25°C. The harvested cells were lysed with glass beads, and the chromatin pellet was washed and sonicated. Sheared chromatin was immunoprecipitated with IgG-Sepharose. This ChIP-enriched DNA was amplified by ligation-mediated PCR (LMPCR) as described elsewhere (27), and 100- to 250-bp LMPCR-amplified fragments were gel purified according to the manufacturer's protocol (Qiagen) and subsequently hybridized to low-density tiled spotted microarrays containing >21,000 oligonucleotide probes as described previously (48). Briefly, each yeast gene is interrogated with a set of at least 3 oligonucleotide probes, which survey the relative occupancy levels for a given transcription factor at the −250 and −60 sites relative to the translational start site and the downstream portion of the open reading frame (ORF). Data were filtered and analyzed as previously described (49) (data are available at http://atlas.bx.psu.edu/).
Briefly, FHT-Mot1 mutant strains were grown as described above for ChIP-chip, except the cells were not cross-linked with formaldehyde. The harvested cells were then pelleted and flash frozen in liquid nitrogen. The cells were lysed with glass beads in NP-S buffer (10 mM Tris-Cl, pH 7.5, 0.5 mM Spermidine, 0.075% Igepal (Sigma), 50 mM NaCl, 5 mM MgCl2, 1 mM CaCl2). Chromatin pellets were washed in NP-S buffer, the chromatin was micrococcal nuclease digested (15 units) in a volume of 300 μl for 20 min, and then the chromatin was solubilized by washing the spun pellet with FA lysis buffer (50 mM HEPES, pH 7.5, 150 mM NaCl, 1% Triton X-100, 0.1% Na deoxycholate, 2 mM EDTA). This native nuclear extract was immunoprecipitated with anti-TBP rabbit polyclonal antibody serum, and the eluate was assayed by Western blot analysis for interacting FHT-Mot1 mutants.
For Western blotting, FHT-Mot1 mutant coimmunoprecipitation eluates were electrophoresed in 6% SDS-PAGE and transferred to a polyvinylidene difluoride (PVDF) membrane (Pall Gelman Laboratory) in Western transfer buffer for 60 min at 1.5 A. FHT-Mot1 mutants were detected with 1:3,300 anti-HA (HA.11; Babco) and 1:5,000 anti-mouse-horseradish peroxidase (HRP) antibodies (Amersham) and exposed to Hyperfilm (Amersham) with enhanced chemiluminescence (ECL) (Amersham).
The relationships to the top and bottom 10th percentiles of the expression and ChIP-chip data were calculated in Excel with the data downloaded from the referenced laboratory or journal's websites. The percent rank of the distribution was calculated with the PERCENTRANK function. Next, the numbers of genes that appear in the top 10% (>0.9 in PERCENTRANK) or the bottom 10% (<0.1 in PERCENTRANK) and also appear in each cluster were calculated. The CHITEST function of Excel was then used to calculate P values from the observed and expected values.
Mot1 is demarcated by several regions that are highly conserved from yeast to human (12, 16). To test their in vivo importance, we targeted four conserved regions for deletion, creating mutants named TBP1, TBP2, UK1, and ATP1 (Fig. 1A to C). The names reflect the associated functional regions (e.g., the TBP1 deletion removes one of two TBP interaction domains, UK1 is unknown, and ATP1 removes the ATPase domain). Deletion of each region of MOT1 was achieved by homologous recombination using a PCR-amplified cassette containing kanMX flanked by loxP Cre recombination sites (34). The cassette also contained coding sequences that allowed the deleted region to be replaced by an FHT epitope tag that encodes a triple-HA tag, a decahistidine tag, and a TEV protease cleavage site. After selection for recombinants on G418 plates and subsequent excision of kanMX with the Cre recombinase, the deleted region was replaced with the FHT tag and a single loxP site, both of which maintained an open reading frame through the replaced region. Cell viability was maintained in a resident mot1-42 temperature-sensitive allele, which allowed subsequent temperature inactivation of the mot1-42 allele at 37°C (Fig. 2A). The location of each deletion mutation was verified by PCR across the deletion borders, with the appropriate-size products detected (not shown). Expression of the deletion mutants was placed under the control of the GAL1 promoter. Immunoblot analysis demonstrated the presence of an appropriate-size band that reacted with anti-HA antibodies and was present only after the addition of galactose to the cells (Fig. 2B). The untagged Mot1 (WT) and null controls were not detected because both lack the FHT tag and thus are not recognized by the HA antibody. All mutants except UK1 were expressed at levels roughly equivalent to that of the wild-type Mot1 containing an FHT tag at the N terminus (WT1). UK1 was expressed at about 50% of the WT1 level. Importantly, the expression levels of the mutants were not diminished after inactivation of the mot1-42 allele at 37°C for 45 min.
Haploid strains carrying a galactose-inducible chromosomal copy of one of the mot1 deletion mutants and a plasmid-borne temperature-sensitive mot1-42 (16) were tested for the ability of the mot1 domain deletion mutants to support viability. The mot1-42 supporting cell viability was inactivated at 37°C and replaced with the domain deletion mutants by adding galactose to the medium. At 25°C and 30°C, mot1-42 remained functional, allowing cell growth, as expected. In the presence or absence of the galactose-induced domain deletion mutants (Fig. 3, 25°C and 30°C), viability was unaffected, indicating that the domain deletion mutants did not have a dominant-negative effect on cell growth. At 37°C, wild-type Mot1 (WT and WT1) and all mutants failed to support viability in dextrose medium, where these proteins are not expressed, verifying the temperature-sensitive nature of the mot1-42 allele (16). As positive controls, both the untagged (WT) and tagged (WT1) inducible Mot1 supported growth in galactose medium at 37°C. However, none of the Mot1 domain deletion mutants support growth, indicating that each of the four conserved domains is essential in S. cerevisiae. These findings are consistent with related studies presented elsewhere (1, 7, 16).
Inasmuch as Mot1 regulates genes both negatively and positively, we tested whether the four conserved regions of Mot1 make distinct gene-specific contributions to gene expression on a genome-wide scale. Expression of the Mot1 mutants was induced with galactose for 60 min, and then the resident functional mot1-42 allele was shut down by an abrupt temperature shift from 25°C to 37°C for 45 min (Fig. 2A). During this time, we expect heat shock-regulated genes to change in expression within 15 min of the temperature shift and then return to near-normal expression levels by 45 min (11, 23).
To place any changes in gene expression into the appropriate context, each expression profile conducted on a mutant was also conducted in parallel using a galactose-inducible untagged wild-type MOT1 allele (WT; yjdi420) (Table 1). Thus, if a mutant is as functional as wild-type Mot1, then no changes in gene expression are expected. Changes in gene expression were mode centered, meaning that the most frequent binned ratio (mutant/WT) corresponded to no change in gene expression. This centering is valid, since most genes are not appreciably regulated by Mot1 (4, 17, 24). Log2-transformed changes in gene expression are presented as a cluster plot (Fig. 4A), where red and green denote increased and decreased expression, respectively. Black denotes no change. Each row corresponds to a protein-coding gene, and each column corresponds to the expression profile for a Mot1 mutant. The 515 genes that met a specified cutoff (Fig. 4A) for changes in expression in at least one set of experiments are shown. The data were clustered by K-means (22) into five clusters, representing the maximum number of visually nonredundant clusters. The homotypic control expression profile, reflecting comparisons between two independent biological replicates of galactose-induced wild-type Mot1, produced no appreciable changes in expression (Fig. 4A, WT), as expected.
The homotypic control also provided a measure of intrinsic variability in the data. The mean log2 ratio was 0.045 (1.03- ± 0.12-fold [standard deviation] change). To ensure that the FHT tag was not perturbing expression, an FHT-tagged wild-type Mot1 control (Fig. 4A, WT1) was tested. Changes in gene expression were very modest compared to the untagged reference (log2 average = 0.033, or 1.02-fold ± 0.12-fold change), indicating that the FHT tag had little or no effect on expression genome-wide. In particular, the standard deviation in gene expression was not significantly different than that observed with the homotypic control.
Changes in gene expression of the galactose-induced null mutant (empty cassette in the mot1-42 strain) at 37°C provided a measure of the maximal level of expected change in expression and correlated well with changes in gene expression reported for the mot1-1 or mot1-14 allele (4, 17) (Fig. 4B). Thus, this transient-replacement system appears to provide an adequate reflection of Mot1 dependency.
From these expression-profiling experiments, we find genes that are negatively regulated by Mot1 (clusters 1 to 3 in Fig. 4A) and those that are positively regulated by Mot1 (cluster 5) each require the TBP1, TBP2, and UK1 conserved domains of Mot1 in that changes in gene expression were similar to that of the null mutant (Fig. 4A, compare columns 3 to 6). Thus, in general, Mot1 uses the same conserved domains to positively and negatively regulate transcription. Genes in cluster 1 are characterized as being stress induced, TATA containing, SAGA dominated, and negatively regulated by a wide range of TBP and chromatin regulators (Table 2). Cluster 2 genes have the same characteristics as those of cluster 1, except that they are not stress induced and are not inhibited by histones. Cluster 3 appears to be a mixture of cluster 1 and 2 genes. Genes in cluster 5 are characterized as being stress repressed, TATA-less, and TFIID dominated. In addition, the ribosomal protein genes dominate this group. The positive contribution of Mot1 to cluster 5 expression is in line with reports proposing that Mot1 positively regulates transcription by redistributing TBP throughout the genome (15, 41) and/or by dismantling transcriptionally inactive TBP (18, 46). Taken together, Mot1 typically regulates 6% of all TFIID-dominated genes in a positive manner, whereas 28% of all SAGA-dominated genes tend to be negatively controlled by Mot1.
The Mot1 ATP1 mutant, corresponding to a deletion of the ATPase domain, had mixed behavior. At most analyzed genes, reflected by clusters 2 and 5 in Fig. 4A, negative and positive regulation by Mot1 required the ATPase domain as much as it required the other conserved regions. Cluster 1 expression displayed less dependence on the ATPase domain of Mot1. The difference between clusters 1 and 2 may be subtle, such as a greater rate-limiting dependency on TBP binding versus ATP hydrolysis. Alternatively, since cluster 1 genes were generally induced and then partially repressed by the temperature regime used here, it is possible that while Mot1 binding is needed for this shutdown, the Mot1 ATPase activity may be partially dispensable. In other words, Mot1 binding in the absence of ATPase activity is sufficient to elicit some repression. This would be consistent with current models of Mot1 function in which simply binding to TBP would be sufficient to preclude binding to certain general transcription factors.
Cluster 4 genes appeared to be largely Mot1 independent, since expression of the null or any of the Mot1 domain deletions (except ATP1) had little effect on transcription (Fig. 4A, columns 3 to 6 in cluster 4). The ATP1 mutant caused an increase in transcription. Since the primary TBP binding regions of this protein are intact, conceivably the ATP1 mutant might promote DNA binding of TBP at these genes but is unable to dissociate TBP without the ATPase domain. In this case, the binding of the Mot1 ATP1 mutant to TBP would seem not to completely interfere with subsequent transcription complex assembly. A plausible rationale for this lies in our observation that cluster 4 genes tend to be repressed by the SSN6-TUP1 complex (P = 10−39) (Table 2). In this context, any enhancement of TBP binding by Mot1(ATP1), even in a weakened state, would provide some increase in expression. One implication is that stabilization of TBP binding at SSN6-TUP1-repressed genes circumvents to some extent SSN6-TUP1 repression.
Cluster 3 genes are generally inhibited by wild-type Mot1 but showed decreased expression when the ATPase domain was removed (Fig. 4A, column 2, cluster 3), paradoxically suggesting that the Mot1 ATPase region plays an apparently positive role at these genes while Mot1 as a whole plays a negative role. Two alternative explanations might account for the apparent paradox. First, as in cluster 4, the ATP1 mutant might stabilize TBP binding at the promoter of cluster 3 genes. However, this TBP-Mot1(ATP1) mutant complex seems to interfere with productive transcription complex assembly at these cluster 3 genes. Alternatively, the ATP1 mutant might promote nonspecific binding of TBP to the genome, which would reduce the amount of TBP that could be recruited to cluster 3 genes, resulting in decreased expression of cluster 3 genes. This would be consistent with the notion that Mot1 normally removes TBP from nontargeted regions of the genome (41). We explore these possibilities below.
Potential interpretations of the Mot1(ATP1) expression profile are predicated upon the Mot1(ATP1) mutant maintaining the ability to interact with TBP. Thus, to test whether Mot1(ATP1) directly interacts with TBP in vivo, we performed coimmunoprecipitation assays for TBP in the FHT-Mot1 mutant strains. Growth and Mot1 replacement were performed as in the expression studies. Importantly, to maintain native protein-protein interactions, the cells were not cross-linked throughout the coimmunoprecipitation procedure. TBP was immunoprecipitated from digested, soluble chromatin derived from Mot1 mutant cells. The ability of Mot1 mutants to interact with TBP on chromatin was revealed by immunoblot analysis (Fig. 5 A). The positive control, the tagged (WT1) inducible Mot1, showed an interaction with TBP (Fig. 5A, bottom). As in Fig. 2B, the untagged Mot1 (WT) and null controls were not detected, because both lack the FHT tag and thus are not recognized by the HA antibody. Among the Mot1 mutant derivatives, the Mot1(ATP1) mutant showed the strongest interaction in vivo with TBP, while the other mutants (TBP1, TBP2, and UK1) displayed either weak or no interaction. Therefore, the interaction between Mot1(ATP1) and TBP revealed by their coimmunoprecipitation supports one assertion of the interpretation that expression changes in clusters 3 and 4 may be a direct result of the Mot1(ATP1) mutant altering the DNA-binding status of TBP at these genes.
To further understand how the Mot1(ATP1) mutant might affect TBP recruitment to promoters, we used genome-wide location analysis (ChIP-chip) to monitor the changes in TBP, Mot1, and TFIID (Taf1 and Taf4 subunits) occupancy at every yeast gene in the Mot1(ATP1) mutant. Growth and Mot1 replacement were performed as in the expression studies. Each factor was immunoprecipitated from sheared soluble chromatin derived from formaldehyde cross-linked wild-type (WT1) and Mot1(ATP1) mutant cells. Bound DNA was differentially labeled and cohybridized to microarrays containing all intergenic regions. Median log2 changes in TBP and Mot1 occupancy for each of the clusters defined in Fig. 4A are plotted in Fig. 5B and compared to changes in gene expression for the Mot1(ATP1) and null mutants.
TBP and Mot1 occupancy changes at cluster 3 and 4 genes mirrored the expression output for these genes in the ATP1 mutant (a decrease at cluster 3 and an increase at cluster 4), suggesting that the expression change is a direct result of the Mot1(ATP1) mutant altering the DNA-binding status of TBP at these genes. The loss of TBP at cluster 3 genes suggests that when Mot1 lacks its ATPase domain, the corresponding loss in transcription is not due to stabilization of an inactive form of TBP at these promoters. Instead, the results are more consistent with the loss of TBP possibly being due to stabilization of TBP bound to other sites in the genome (e.g., cluster 4 genes) by the Mot1(ATP1) mutant. Indeed, TBP and Mot1 occupancy increased in the Mot1(ATP1) mutant at cluster 4 genes. Because cluster 3 genes normally have more TBP than cluster 4 genes (Fig. 5C), they have more to lose if TBP is distributed nonspecifically throughout the genome in the Mot1(ATP1) mutant.
One interpretation of the positive activity of Mot1 on expression at cluster 5 genes is that it removes an inactive form of TBP, allowing productive binding of TFIID. Accordingly a Mot1(ATP1) mutant might “lock down” TBP at cluster 5 genes, thereby preventing TFIID from binding. To test this hypothesis, we conducted ChIP-chip on the Taf1 and Taf4 subunits of TFIID, comparing its occupancy at cluster 5 genes in the wild type versus a Mot1(ATP1) mutant. Consistent with this hypothesis, Taf1 and Taf4 occupancy levels decreased at cluster 5 genes in the Mot1(ATP1) mutant, whereas TBP levels remained largely unchanged (Fig. 5D). A constant level of TBP is consistent with the hypothesis, in that one type of TBP (TAF free) replaces another type of TBP (TAF bound; TFIID).
The binding of TBP to DNA is generally considered to be a primary nucleating event in transcription complex assembly. TBP-DNA binding is therefore subjected to substantial positive and negative regulation. TBP not only binds to the TATA box located in promoters, it also binds to TATA-less promoter regions, and it binds to nonspecific DNA with fairly high affinity. Since nonspecific DNA binding by TBP can nevertheless nucleate transcription (14), robust mechanisms exist in vivo to prevent promiscuous assembly.
Mot1 plays an important role in removing TBP from inappropriate genomic sites (18, 41), which would free up TBP and/or the underlying DNA to engage in productive interactions (46). In this way, Mot1 would play a positive role at promoters that are rate limited either by the availability of TBP or by dissociation of an inactive TBP-promoter complex. Such an arrangement might predominate at ribosomal protein genes, where Mot1 plays a positive role (cluster 5 in Fig. 4A). Conceivably, a Mot1-inaccessible form of TBP (i.e., TFIID) binds to ribosomal promoters (12, 39). Nonproductive binding of a TFIID-independent form of TBP might antagonize TFIID recruitment or any other stage in transcription initiation. Removal of this nonproductive TBP would therefore positively impact transcription at these genes. Indeed, we find that stabilizing a TBP-Mot1 promoter interaction at Mot1-upregulated genes has the effect of displacing TFIID.
Stress-induced promoters often are highly transcribed even under nonstress conditions (11, 23). These promoters tend to rely more on a free form of TBP that does not involve TFIID. This form of TBP may be directed to the appropriate promoters via the SAGA complex, and since it is not TFIID, it may be more accessible to Mot1. Indeed, the abundance of accessible TBP at such promoters attracts Mot1, where it downregulates expression (cluster 1 in Fig. 4A). A moderately high level of expression is achieved through this balance of positive SAGA-TBP action (among other factors) and negative Mot1-TBP action (among other factors). Indeed, by using multidimensional chromatography coupled to mass spectrometry (5), a recent study found that Mot1 interacts with a variety of activators and transcriptional coregulators, such as Msn2, Hsf1, and RSC.
Since Mot1 has the ability to bind both TBP and DNA (44), it potentially can stabilize TBP-DNA interactions. However, this is not realized in general because Mot1 uses the energy of ATP hydrolysis to remove TBP from DNA. A form of Mot1 that lacks the ATPase domain provides a window into how potential stabilization of TBP-DNA interactions through Mot1 affects the distribution of TBP genome-wide. If the Mot1(ATP1) mutant were to increase TBP-DNA stability in an undirected way, then genomic loci that normally lack TBP should see an increase in TBP in the mutant. Those loci that normally have higher TBP levels should suffer a decrease in occupancy as TBP is sequestered at the vast number of nonspecific loci. Indeed we identified a set of genes (cluster 4) that had lower levels of TBP and whose levels of TBP, Mot1, and transcription increased in the Mot1(ATP1) mutant. We identified another set of genes (cluster 3) that had higher levels of TBP, which decreased in the Mot1(ATP1) mutant.
Taken together, our findings suggest that the four conserved regions of Mot1 are essential for viability and required for proper regulation of most Mot1-regulated genes. Loss of the ATPase domain, however, imparts some unexpected regulation on certain genes. Although only a fraction of all yeast genes are overtly regulated by Mot1, essentially all yeast genes require TBP and thus are potential targets for Mot1. Genes that are insensitive to loss of Mot1 likely reflect those that are relatively quiescent and thus lack TBP or those that involve TFIID, which is a Mot1-insensitive form of TBP. Allowing TBP to redistribute genome-wide in a potentially more nonspecific manner results in a net gain of TBP at TBP-deficient genes and a net loss at TBP-enriched genes. Thus, one consequence of Mot1 using ATP hydrolysis to remove TBP from DNA may be an increase in promoter selectivity.
We are grateful to David Auble for plasmids pMOT221 (mot1-42 LEU2), pMR13 (MOT1 WT URA3), and MOT1 WT (pAV20 LEU2). We thank Joe Reese for kindly providing the Taf1 and Taf4 antibodies.
This work was supported by National Institutes of Health grant GM059055.
Published ahead of print on 28 March 2011.