Construction of conditional degron alleles of Reb1, Abf1 and the RSC ATPase Sth1

The mechanism by which Reb1:dT₇ induces formation of an NFR flanked by two nucleosomes carrying H2A.Z likely involves the recruitment of Reb1, although this was not tested directly in our previous study. We also hypothesized that Reb1 might recruit a chromatin remodeling enzyme to produce an NFR. A systematic study of protein interactions revealed that Reb1 physically associates with Rsc2, Rsc3 and Npl6, which are subunits of the essential chromatin remodeling complex RSC (Cairns et al., 1996; Cairns et al., 1999; Gavin et al., 2002). Since the Reb1-related factor, Abf1, has also been implicated in the formation of a nuclease-sensitive site (De Winde et al., 1993), we pursued its functional role.

As Reb1, Abf1, and the catalytic subunit of RSC (Sth1) are all essential proteins, we generated a series of conditional alleles Specifically, we used the temperature-sensitive degron system to engineer yeast strains in which we could control degradation of these proteins via the N-end rule pathway (Dohmen and Varshavsky, 2005). This pathway operates through the recognition of destabilizing N-terminal amino acids by the nonessential E3 ubiquitin ligase Ubr1. A N-terminal arginine is the strongest signal for degradation by Ubr1, and traditional degron alleles encode proteins capped by an arginine followed by a temperature-sensitive murine dihydrofolate reductase (DHFR^ts) peptide fused to the target protein. As translation initiates at an ATG codon, an ATG-initiated segment encoding a ubiquitin moiety is placed before the arginine codon. Once synthesized, this segment is cleaved in cells by ubiquitin C-terminal proteases to expose the N-terminal arginine. Such degron systems also place the modified allele of interest under the control of a regulatable promoter, which traditionally has been the copper-inducible promoter pCUP1.

We constructed pCUP1UBI4DHFR^tsc-mycSTH1 and pCUP1UBI4DHFR^tsc-mycREB1 alleles in strains that had UBR1 under the control of the pGAL1 promoter. We were unable to achieve substantial degradation of these proteins or growth arrest under degron-inducing conditions, although such an allele for STH1 has been reported (Parnell et al., 2008). We achieved more complete degradation of Reb1 and Sth1 under degron-inducing conditions with a different construct (pMET3UBI4DHFR^ts3xHA) that utilized the methionine-repressed pMET3 promoter (Fig. 1C). Yeast strains carrying Reb1-degron or Sth1-degron alleles were inviable under degron-inducing conditions (Fig. 1D). While an Abf1 DHFR^ts degron has been reported in the W303 strain background (Reed et al., 1999), we were unable to construct a viable pMET3UBI4DHFR^ts3xHAABF1 degron allele in our S288C background despite several attempts and strategies. We, however, successfully created a pMET3UBI4abf1(M1R) allele in a strain carrying a pGAL1UBR1 allele. As described above, the ubiquitin moiety of the translated protein is cleaved off soon after translation, leaving an Abf1 protein capped with a destabilizing N-terminal arginine. Under inducing conditions, this Abf1-degron yielded growth arrest and displayed Abf1 depletion (Fig. 1C, 1D). We also constructed an Abf1 Reb1 “double degron” strain in order to simultaneously deplete both factors from cells (Fig. 1C, 1D). For the studies described below, we determined nucleosome positions by hybridizing mononucleosomal versus genomic DNA-derived probe with in-house printed custom tiling arrays that span S.cerevisiae chromosome III at 20bp resolution (Yuan et al., 2005). The arrays also included oligonucleotides that tiled sequences corresponding to the PRM1 gene to allow us to observe nucleosome positions programmed by Reb1:dT₇ (see Experimental Procedures for details). Unless otherwise specified, all experiments represent averages of four independent, biological replicates.

NFR formation mediated by a Reb1 binding site requires Reb1 and the chromatin remodeling complex RSC, but not H2A.Z

As expected from our previous study (Raisner, et al., 2005), nucleosome position analysis of the PRM1 ORF with or without a Reb1:dT₇ insertion revealed that the insertion produces an NFR (Fig. 2A). NFR formation was unaffected in strains carrying the Reb1-degron or Sth1-degron alleles under conditions in which the degron system was inactive (Fig. 2B). We mapped nucleosome positions in strains carrying the Reb1-degron or the Sth1-degron five hours after activating the degron system, and these positions were compared to nucleosome positioning data from a control strain isogenic to the degron strains except that it lacked the Reb1 or Sth1 degron allele. The latter control strains were subjected to the identical culture growth protocol as the experimental strains. Depletion of Reb1 resulted in loss of the NFR programmed by Reb1:dT₇ inserted into the PRM1 ORF, consistent with a direct role for Reb1 (Fig. 2B). Likewise, inactivation of the RSC complex through depletion of Sth1 resulted in complete loss of the NFR programmed by the Reb1:dT₇ sequence (Fig. 2B), supporting the hypothesis that Reb1 functions by recruiting RSC.

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Figure 2

Tiling array analysis of nucleosome positions in the PRM1 ORF containing the Reb1:dT₇ insertion

A. Analysis of the effect of Reb1:dT₇ sequence insertion on nucleosome positioning. Shown are line traces of a moving average of mononucleosome/genomic probe signals across the PRM1 gene with and without the indicated sequence insertion. Triangles indicate the insertion site.

B. Analysis of effects of Reb1 and Sth1 depletion on NFR formation mediated by Reb1:dT₇. Indicated strains containing the Reb1:dT₇ insertion in the PRM1 gene were analyzed as described in A.

C. Analysis of effects of htz1Δ and swr1Δ mutations on NFR formation mediated by Reb1:dT₇. Indicated strains containing the Reb1:dT₇ insertion in the PRM1 gene were analyzed as described in A.

Model II above proposes that NFR formation requires the prior deposition of H2A.Z into chromatin, which is mediated by the Swr1 chromatin remodeling complex. We tested whether H2A.Z or Swr1 is required for NFR formation induced by the Reb1:dT₇ signal. We constructed htz1Δ and swr1Δ strains and mapped the nucleosome positions in these strains. NFR formation meditated by insertion of Reb1:dT₇ at PRM1 did not require H2A.Z or Swr1 (Fig. 2C).

To test whether NFR formation at PRM1 might be a consequence of transcription, we examined whether NFR formation induced by Reb1:dT₇ was associated with the production of transcripts. Transcript levels were examined using RT-PCR analysis of extracted total RNA. In a control strain in which the PRM1 gene was induced with mating pheromone for 1hr, a specific signal was obtained (Fig. S10A). We examined strains containing the insert and the Sth1 degron under both degron-inducing and permissive conditions as well as a strain lacking the insert. In these three cases, multiple peaks rather than a single peak were obtained in the QPCR melting curves indicating that cross-reacting cDNAs, instead of products specific to the PRM1 locus, were being detected. Moreover, no quantitative differences were observed (Fig. S10A). Taken together, these data suggest that NFR formation was not associated with transcription that could be detected by these methods.

Reb1 is required for the formation of a subset of NFRs

We next examined the chromosome-wide requirement for Reb1 in NFR formation. We mapped and compared nucleosome positions in the Reb1-degron strain and an isogenic control strain that lacked the Reb1-degron under conditions described above. Fig. 3A shows a gene-by-gene “difference map” of the positioning data in which genes were aligned to each other based on the position +1 nucleosome downstream of the NFR in control strains and then the mutant signal subtracted from the control signal (yellow indicates more nucleosomal DNA signal in the mutant than in wild-type cells). The data were organized by K-means clustering. Two clusters are evident, one in which positioning was affected (Cluster I affecting 12% of assayed promoters), and another where no effect was evident (Cluster II). Line traces of the average signals of the control and degron strains for these two clusters are shown in Fig. 3B. Inspection of Cluster I indicates that the two NFR-flanking nucleosomes move inward towards the center of the NFR, and this movement propagates further such that other flanking nucleosomes also shift their positions (Fig 3B) Figure 3C shows that the changes in the size of the trough signal representing the NFR between degron and control strains were dependent on the induction of the degron.

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Figure 3

Chromosome-wide tiling array analysis of nucleosome positions in cells depleted of Reb1

A. Difference map analysis of effects of Reb1 depletion on nucleosome positions. Map represents nucleosome positioning data from a control strain lacking the Reb1-degron subtracted from a strain with the Reb1-degron. See Experimental Procedures for further details. Nucleosome positioning data 1kb upstream and downstream of the ATG of 150 genes are shown and orientated such that the direction of transcription is to the right. Asterisks indicate center of the +1 nucleosome downstream of the NFR in the control strain. The x-axis represents the distance (in bp) from the center. Data are organized into two clusters using the k-means method.

B. Line traces of average nucleosome positions of the two clusters shown in A The indicated strains were grown under degron-inducing conditions.

C. Scatter plots of the lowest probe signal in NFRs. Points indicate the lowest probe signal in the NFR for a locus in control versus degron strains grown under the indicated conditions.

D. Correlation between Reb1 binding and changes in nucleosomal enrichment at NFRs at promoters. The significance values (log10 p-value) of Reb1 binding at promoters (Harbison, et al. 2004) are compared against the highest fold changes of nucleosome positioning signals at the associated NFR.

E. Correlation between Reb1 consensus sites in promoters and the highest fold change of nucleosome enrichment at the associated NFR under the indicated promoters.

F. Enrichment of Reb1 sites in clusters. Shown are the p-values (hypergeometric testing) of the significance of the indicated Reb1 motifs in the indicated clusters.

A previous study of transcription factor association at promoters assigned likelihood scores for a given transcription factor binding to a given promoter (Harbison et al., 2004). We compared these scores for Reb1 to the highest fold-change probe in the NFRs of promoters we assayed that were assigned scores. As shown in Fig. 3D, there was a significant correlation (p=7.81×10^-7 based on a hypergeometric test using a likelihood score cutoff of p<0.05). Consistent with this finding, promoters that contained at least one copy of the most conserved Reb1 binding site (TTACCCG, (Liaw and Brandl, 1994)) tended to experience changes in NFR structure (Fig. 3E). Cluster I was enriched for the most conserved Reb1 consensus site; indeed, 14 out of the 18 promoters in Cluster I contained this motif (p= 5.07×10^-13, Fig. 3F). Relaxing the consensus to reflect the poorer conservation of the first two residues of the consensus still yielded highly significant enrichments (Fig. 3F).

Abf1 is required for the formation of a subset of NFRs

We next performed the same analysis on Abf1 degron strain. Difference map analysis and clustering (Fig. 4A) show that 9.3% of promoters were affected, and these promoters were distinct from promoters affected by Reb1 depletion (see below). As with the Reb1 degron, the affected cluster displays a smaller NFR and movement of flanking nucleosomes towards the NFR (Fig. 4B), and these changes were dependent on induction of the degron (Fig. 4C). Likewise, affected NFRs were enriched for Abf1 binding (Fig. 4D) and for an Abf1 consensus site (Fig. 4E, F). The latter correlations are weaker than for the Reb1 site, perhaps because of the higher degeneracy of the Abf1 consensus site (Beinoraviciute-Kellner et al., 2005).

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Figure 4

Chromosome-wide tiling array analysis of nucleosome positions in cells depleted of Abf1

A-F. These panels are analogous to those of Figure 3 except that a strain with the Abf1-degron was compared to the same control strain used for analysis of nucleosome positions upon Reb1 depletion.

RSC is required for proper positioning of NFR-flanking nuclesomes

We next examined the effects of Sth1 depletion on nucleosome positioning using strains carrying the Sth1-degron as described above. Strikingly, our analysis showed that Sth1 depletion affected a majority (55%) of promoters (see Cluster I in Fig. 5A). The affected cluster displayed shrinking of the NFR and movement of flanking nucleosomes, whereas little change in nucleosome position was apparent for members of Cluster II (Fig. 5B). As with the Reb1 and Abf1 degron strains, growth under degron-inducing conditions was required to observe these differences (Fig. 5C). Fig. S10B shows a superposition of a histogram of the locations of mapped TSSs (Nagalakshmi, et al., 2008) and the positioning data. Consistent with previous studies, TSSs tend to lie just inside the downstream nucleosome and the movement observed in Sth1-depeleted cells moves these sites further into the nucleosome core (Fig. S10B). This may explain why RSC depletion has been reported to cause cessation of transcription by RNA polymerase II (Parnell et al., 2008).

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Figure 5

Chromosome-wide tiling array analysis of nucleosome positions in cells depleted of Sth1

A-C. These panels are analogous to those of Figure 3 except that a strain with the Sth1-degron was compared to the same control strain used for analysis of nucleosome positions upon Reb1 depletion.

D. Gene expression analysis. Shown is the correlation between mRNA and NFR changes in cells depleted of Sth1. Ploted are the values for genes on chromosome III. Shown on right is the p-values (hypergeometric testing) of the significance of the enrichments in the indicated gene groups using a 1.5-fold cutoff for decreases in mRNA levels. Expression data were not available for all genes in the clusters; hence, Cluster I N=79, Cluster II N=64.

E. NPS signature averages. Lines traces of NPS predictions (Ioshikes et al., 2006) for genes in the indicated gene clusters are shown in green. These predictions were smoothed using a 51bp moving average window. Experimental nucleosome position averages (control is blue, Sth1-degron is red) are shown as in panel B.

F. Effect of transcription on nucleosome positioning. Shown are the average nucleosome positions for the indicated gene clusters in rpb1-1 strains grown under permissive conditions or for 1 hr under nonpermissive conditions.

When RSC, Abf1, or Reb1 were depleted, NFRs shrank but were not eliminated. We hypothesized that intrinsic positioning sequences might explain the positions of nucleosomes under these conditions. Therefore, we compared the positions we observed in Cluster I of Sth1-depleted cells with those predicted by Pugh and colleagues (Ioshikhes et al., 2006) based on AA/TT dinucleotide periodicity enrichment. As shown in Figure 5E, the average nucleosome position of the +1 and -1 nuclesomes of Cluster I relax towards positions specified by the NPS signature. For the largely unaffected cluster (Cluster II), a discrepancy between the NPS-predicted and observed positions is still apparent for the +1 nucleosome, whereas the -1 nucleosome is poorly aligned in this despite the sharp NPS prediction peak (Fig. 5E).

To test whether Sth1 depletion results in changes in gene expression, we performed expression profiling of the Sth1 degron strain against a control strain under degron-inducing conditions. As we expected global changes in gene expression, we incorporated external spiked-in RNA controls into our normalization procedure (see Experimental Procedures). We then asked whether there was an enrichment for genes whose expression was reduced in Sth1-depleted cells in Cluster I vs. Cluster II. As shown in Figure 5D, Cluster I is indeed highly enriched for genes whose expression requires Sth1, indicating that that the changes in positioning correlate with changes in expression. Given this result, we tested whether loss of transcription might be responsible for changes in positioning. We mapped nucleosome positions in a temperature-sensitive RNA polymerase II strain (rpb1-1) which ceases transcription within minutes upon shift into restrictive conditions (Nonet et al., 1987). The rpb1-1 mutant was grown at either 25°C or shifted for 1 hour to 37°C. The average nucleosome positions in these conditions were determined for Clusters I and II of the Sth1 degron difference map. There were were no detectable differences in nucleosome positions in either cluster (Fig. 5F, Fig. S10C). Hence, the changes in NFR structure observed upon Sth1 depletion appear to be due to the action of RSC rather than from cessation of transcription per se.

Abf1 and Reb1 are required for NFR formation at distinct sets of promoters

To identify promoters that are redundantly controlled by Abf1 and Reb1, we examined a double degron strain that carried the Abf1-degron and Reb1-degron alleles (Fig.1C, 1D). The resulting difference map was clustered together with difference maps for the Abf1 single degron strain, the Reb1 single degron strain, and the Sth1 degron strain (Fig. 6A; Fig. 6B shows that NFR changes in the double degron are dependent on degron induction). K-means clustering revealed that the promoter NFRs affected by loss of either Abf1 or Reb1 were reproducibly affected in the double degron strain, but there were no other promoter NFRs that were significantly affected in the double degron strain. It is also evident that most NFRs affected by loss of Reb1 also required Sth1 for proper positioning of nucleosomes (Fig. 6A), consistent with the data obtained with the synthetic NFR described above (Fig. 2B). The NFRs affected by loss of Abf1 appeared to have a somewhat lesser degree of dependence for Sth1 for nucleosome positioning (Fig. 6A). Likewise, there clearly are many NFRs that require RSC, but not Abf1 or Reb1, for proper nucleosome positioning (Fig. 6A), suggesting the existence of additional RSC recruitment mechanisms. Analysis of average nucleosome positions for each cluster indicates that the the changes observed (Fig 6A) are due to shifts in nucleosome positions (Fig. S14).

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Figure 6

Comparison of the roles of Abf1, Reb1, and RSC in nucleosome positioning

A. Clustering analysis of difference maps. Shown are difference maps for the indicated strains including the Abf1-Reb1 “double degron” strain. K-means (K=15) clustering was applied. Line traces of cluster averages are shown in Fig. S14.

B. Scatter plots of double degron. Analysis was performed as in Fig. 3C.

C. Gene expression analysis. Shown is the correlation between mRNA and NFR changes in cells depleted of both Reb1 and Abf1. The p-value (hypergeometric testing) of the significance of the enrichments in the group of genes (N=20) consisting of those affected in NFR size by Abf1-or Reb1-depletion (Cluster I in Fig. 3 and Cluster I in Fig. 4; corresponding points are colored red in this figure) using a 1.5-fold cutoff for decreases in mRNA levels was 1.2×10^-6. The p value for the remaining, unaffected genes (N=123) was 0.99.

As with the Sth1 degron strain, we examined the Abf1-Reb1-double-degron strain for changes in transcript levels using whole genome microarrays and spiked-in external controls. As shown in Figure 6C, we found a significant correlation between decreases in NFR size and decreases in transcript accumulation.

H2A.Z deposition is generally dispensable for nucleosome positioning

To complete our analysis of positioning, we used cells lacking H2A.Z (htz1Δ) or lacking the ATPase subunit of its deposition complex (swr1Δ) to determine whether H2A.Z exchange was required for nucleosome positioning chromosome-wide (Fig. 7). Based on the results with the synthetic NFR (Fig. 2C), we expected to see no differences in positioning. Indeed, as shown by line traces of the average positions of aligned promoter nucleosomes, H2A.Z deposition resulted in no detectable changes. Of course, we cannot rule out the possibility that there could be changes too subtle to observe using our 20bp resolution tiling arrays.

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Figure 7

Chromosome-wide tiling array analysis of nucleosome positions in cells defective in H2A.Z nucleosomes

Panels A and B are analogous to Fig. 3B and 3C except that data were derived from cells lacking H2A.Z (htz1Δ) or the Swr1 ATPase (swr1Δ).

Development an inducible H2A.Z deposition system

Nonetheless, these data argue against Model II (Fig. 1B), which proposed that H2A.Z deposition is essential for NFR formation. We then considered the two remaining models: (1) the deposition of H2A.Z at a promoter requires the presence of an NFR at that promoter (Model II), or (2) H2A.Z deposition occurs independently of NFR formation (Model III). In principle, these models could be distinguished through development of a system in which NFR loss is induced under conditions where H2A.Z is not deposited into chromatin, but then H2A.Z is induced and its deposition examined. The Sth1 degron provides a tool to trigger abrogation of the synthetic NFR programmed by Reb1:dT₇ and a shrinkage of bona fide promoter NFRs. However, since global transcription is shut off in RSC-depleted cells, we sought a posttranslational method to control H2A.Z deposition.

We utilized an engineered M. tuberculosis RecA intein whose intrinsic protein splicing is controlled by the human estrogen receptor ligand binding domain (Buskirk et al., 2004). This construct was used previously to interrupt several coding sequences in yeast, and its splicing was shown to be activated in vivo using the estrogen agonist 4-hydroxytamoxifen (4-HT). The chemistry of splicing requires cysteine cleavage sites and leaves a single cysteine residue at the splice junction.

We initially targeted the Swr1 and Swc2 subunits of the Swr1 complex by inserting the intein construct before the codons for several native cysteine residues in the corresponding genes. These alleles abrogated H2A.Z deposition in vivo, but addition of 4-HT did not restore H2A.Z deposition (unpublished observations), suggesting that the placement of the intein was incompatible with protein stability and/or intein splicing. We next attempted engineering a spliceable HTZ1 allele under the assumption that the smaller size of H2A.Z relative to the intein construct would make the protein context less likely to interfere in proper structural formation of the intein. H2A.Z, however, lacks cysteine residues, so such a spliced allele would by necessity contain a cysteine point mutation. Four H2A.Z residues (Ala46, Thr68, Thr88 and Asp100) that did not confer a significant growth defect in high precision measurements when mutated (S. Braun, D. Breslow, J. Weissman, H. D. M., unpublished observations) were replaced with the intein construct. These alleles initially replaced wild-type HTZ1 at its native chromosomal locus, but none displayed detectable spliced product in the presence of 4-HT (unpublished observations). We therefore placed these alleles under the control of a pGAL1 promoter on a high-copy 2 micron plasmid vector. The allele in which Ala46 was replaced with the intein construct yielded a protein that was spliced in vivo when cultures were treated with 4-HT; Ala46 is a residue in the core histone fold domain (Fig. 8A, 8B). As splicing produced somewhat higher levels of H2A.Z than in that found in wild-type cells (Fig. 8B), we placed the construct on a low-copy CEN-ARS plasmid. Regulated splicing of the H2A.Z intein was also observed (Fig. S13A), and this construct was used in further experiments. We refer to this pGAL1htz1(A46intein) allele on a CEN-ARS plasmid as the H2A.Z intein construct.

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Figure 8

Analysis of H2A.Z deposition requirements using a steroid-regulated intein

A. Schematic of H2A.Z intein constructs. An engineered M. tuberculosis RecA intein controlled by the human estrogen receptor ligand binding domain was placed at a chosen site in HTZ1 gene (encoding H2A.Z). The HTZ1 promoter was replaced with the galactose-inducible pGAL1 promoter. Protein splicing would occur in the presence of 4-hydroxytamoxifen (4-HT) and leave a cysteine residue (“scar”) at the splicing junction.

B. Splicing of the H2A.Z intein construct occurs with an allele that replaces Ala46 with the intein construct. The 4-HT-regulated intein was inserted in place of four different residues in H2A.Z, and these constructs each were placed on a 2μ plasmid under the control of a pGAL1 promoter and transformed into a htz1Δ strain. Strains were grown to mid-log phase in the presence of 2% galactose and, when indicated, 10 μM 4-HT. Shown is a Western using polyclonal antibody specific to the C-terminus of H2A.Z. A strain with a chromosomal-based, wild-type copy of HTZ1 and a htz1Δ strain are included as controls.

C-F: Analysis of H2A.Z deposition requirements. A strain carrying the Sth1 degron was shifted to degron-inducing conditions which also induces synthesis of unspliced H2A.Z intein construct. After 5 hrs, 4-HT was added to 10μM to induce splicing. Cells were collected after 3 hrs of further incubation, and H2A.Z enrichment at select loci was determined relative to histone H3 enrichment and normalized to a locus in the middle of the large BUD3 ORF. H2A.Z/H3 enrichment profiles under Sth1-depleted conditions were compared against control profiles. The promoters of DCC1/BUD3 are analyzed in C; the PRM1 ORF containing the Reb1:dT₇ insertion is analyzed in D; the promoter of YCR016W is analyzed in E, and the promoter of YCRO23C is analyzed in F. Top panels of C-F indicate nucleosomal DNA enrichment in various conditions (blue: degron-OFF conditions, red: 5 hrs of degron-ON, green: 8 hrs of degron-ON, with the last 3 hrs in the presence of 4-HT). Bottom panels of C-F represent normalized H2A.Z/H3 enrichment values at the indicated amplicons.

We examined the deposition of H2A.Z whose synthesis was directed by this construct using chromatin immunoprecipitation (ChIP). Although splicing in the H2A.Z intein is regulated, a small amount of H2A.Z deposition was observed in the absence of 4-HT, presumably due to low levels of background splicing (Fig. S13C); however, the H2A.Z enrichment signal steadily increases over time in response to 4-HT treatment (Fig S13D), indicating stimulation of H2A.Z deposition by the activation of splicing.

(1) RSC displaces NFR-flanking nucleosomes away from their average NPS-predicted positions

A striking result presented here is that at a majority of promoters, the normal positioning of NFR-flanking nucleosomes requires the essential multisubunit chromatin modeling complex RSC. Such a central role for RSC in generating promoter chromatin architecture is consistent with several of its properties: 1) RSC, unlike most chromatin remodeling enzymes in yeast, is essential for viability (Cairns et al., 1996; Cairns et al., 1999), 2) RSC slides nucleosomes in vitro (Lorch et al., 2001), and 3) RSC is required globally for RNA polymerase II transcription (Parnell et al., 2008). Our studies are also consistent with a recent lower-resolution study that concluded that RSC affected histone density at a number of promoters (Parnell et al., 2008). A recent study indicated changes in the positioning nucleosomes at ~12% of promoters in cells lacking the Isw2 chromatin remodeling complex (Whitehouse et al., 2007). The primary function of Isw2 appears to be in transcriptional repression and in suppressing antisense transcription (Whitehouse et al., 2007). Interestingly, in contrast to RSC, Isw2 appears to move nucleosomes in vivo toward the NFR, raising the possibility that it antagonizes the action of RSC at some promoters. The potential for dynamic involvement of multiple ATPases at promoters further underscores the active nature of mechanisms that position nucleosomes in vivo.

The finding in this study and in the previous study that the final resting positions of nucleosomes are strongly influenced by ATP-dependent chromatin remodeling mechanisms argues that that the intrinsic affinity of the octamer for underlying DNA sequences is not determinative for the final positioned state. However, our observation that depletion of Sth1 causes nucleosome positions to relax on average closer to those predicted by an NPS signature strongly suggests that sequence properties play a role in a stepwise mechanism for NFR formation. That is, NPS-mediated positioning exposes binding sites for factors such as Reb1 and Abf1, which in turn induce the action of RSC to move nucleosomes to their steady-state average positions in wild-type cells. Such a model is also consistent with in vitro and in vivo observations that suggest that the Isw2 remodeling enzyme moves nucleosomes into energetically unfavorable sites (Whitehouse and Tsukiyama, 2006). We speculate that, compared to a purely “hard-wired” system, this more dynamic, ATP-dependent mechanism may facilitate binding of DNA binding proteins to nucleosomal sites and transcription initiation. It is important to note that NPS predictions vary in their accuracy considerably at the level of individual genes, suggesting they likely do not predict with full accuracy the intrinsic thermodynamics of octamer-DNA interactions. A histogram of predictions (Ioshikhes et al., 2006) reveals that NPS-predicted positions for individual genes deviate significantly from experimental positions even in the Sth1 degron strain (Fig S15). Nonetheless, the close correspondence of the average profiles supports the two-step model proposed above.

(2) Sequence-specific DNA binding proteins are required for positioning of NFR-flanking nucleosomes at a significant fraction of promoters

Using a signal for NFR formation/H2A.Z deposition we identified previously, we demonstrated a role for the Reb1 protein and RSC for NFR formation programmed by this isolated signal. Given the previously reported biochemical interactions between Reb1 and subunits of RSC, the simplest interpretation is that recruitment of RSC by Reb1 generates the NFR. Our examination of the generality of this mechanism across chromosome III suggests that a subset of promoters, enriched for Reb1 binding sites, use this mechanism in a nonredundant fashion. Abf1, another essential Myb family member, operates at a distinct subset of promoters. These observations are consistent with studies that show that Reb1 and Abf1 sites are highly enriched in NFRs compared to the binding sites for nearly all other studied DNA binding proteins (Lee et al., 2007). The remaining promoters presumably target RSC and other remodeling mechanisms through other means. In this regard, it is interesting to note that four subunits of RSC contain potential DNA binding domains. Using standard ChIP protocols as well as ones using additional crosslinking agents, we have been unable to detect either wild-type Sth1 or an induced catalytically-dead version of Sth1 at the Reb1:dT₇ signal inserted into PRM1, suggesting transient binding of RSC to this site (unpublished data). Likewise, only a fraction of intergenic regions display RSC binding in published ChIP-chip experiments (Ng et al., 2002), despite the global requirement for RSC in pol II transcription (Parnell et al., 2008). We suggest that at many sites of action the off-rate of the RSC complex in vivo may be too high to allow detection by ChIP.

Antibodies

H2A.Z-specific polyclonal antisera were generated against a peptide specific for the C-terminus of S. cerevisiae H2A.Z (custom-generated). The HA epitope tag in the degron alleles was detected using monoclonal antibody HA.11 (Covance). Abf1 was detected using polyclonal antibodies directed towards the Abf1 C-terminus (yC-20, Santa Cruz). Histone H3-specific polyclonal antibodies were directed towards the C-terminus of human histone H3 (ab1791, abcam).

Microarray data

Microarray data can be obtained from NCBI GEO at series accession GSE13446.

Chromatin immunoprecipitation and QPCR

Chromatin immunoprecipitation and subsequent analysis by QPCR was performed as previously described (Raisner, et al., 2005, Meneghini, et al., 2003).

Gene expression profiling

For each strain, total RNA from four independently grown cultures was prepared using a TRIZOL procedure and spiked with RNA from the Agilent Dual-color RNA Spike-in Kit. Aminoallyl-dUTP-labeled probe was generated by reverse transcription, and hybridizations were carried out using 4×44k Agilent microarrays that cover 6256 S. cerevisiae features, each of which are replicated 7 times on the array (Agilent design ID 015072). Dye swaps were incorporated such that for each experiment, there were 2 arrays of one dye configuration, and vice-versa. Data normalization was performed using a composite loess procedure that used 1:1 DCP probes for the spike-in loess curve (Yang, et al. 2002). Expression ratios for each gene per array then were derived by calculating the mean of up to 7 technical replicates, while discarding any replicates that were not within 2 standard deviations.

Assaying the requirement of essential genes with degron technology

The essential genes ABF1, REB1, and STH1 were studied by regulated degradation of their encoded protein via degron alleles. Each degron allele was under the control of the pMET3 promoter, which is repressed by methionine. The REB1 and STH1 degron alleles had an arginine-capped N-terminal fusion of DHFRts and a triple-HA tag, while the ABF1 degron allele was an abf1(M1R) allele. UBR1, the N-end rule pathway E3 ubiquitin ligase, was placed under the control of a pGAL1 promoter.

To study phenotypes arising from loss of Abf1, Reb1 or Sth1, degron cultures were grown at 30°C to mid-log phase in synthetic complete media lacking methionine and cysteine with 2% raffinose and 0.1% dextrose as carbon sources. Activation of the degron was achieved by first adding galactose to a final concentration of 2% for 30 min, followed by centrifugation at room temperature to collect the cells. These cells were next grown at 37°C in rich media prewarmed at 37°C and supplemented with 2% galactose (YPAG).

Preparation of DNA for mapping nucleosome positions

Cultures in mid-log phase (which ranged from an OD₆₀₀ of 0.7-0.9) were crosslinked with 1% formaldehyde for 15 min at the same temperature used for growth, followed by a quenching step for 5 min at room temperature with 0.125M glycine. Cells were washed twice with cold ddH₂O prior to storage.

Approximately 20 OD₆₀₀ units of cells were spheroplasted with 0.25 mg Zymolyase 100-T (Seikagaku) in 2 ml Buffer Z (1M sorbitol, 50mM Tris-Cl pH 7.4, 10mM β-mercaptoethanol) at 30°C with shaking. The spheroplasting time ranged from 30 min to 75 min, depending on the strain and media conditions used for growth. The ideal spheroplasting time was one that yielded appropriately digested chromatin (~90% mononucleosomal-sized DNA) after 20 min of micrococcal nuclease (MNase) treatment. Spheroplasts were collected by centrifugation at 4°C and resuspended in 500μl MNase digestion buffer (0.075% NP-40, 50mM NaCl, 10mM Tris-Cl pH 7.4, 5mM MgCl₂, 1mM CaCl₂). Chromatin was digested with 3 units of MNase (Worthington) for 20 min at 37°C. Digestions were quenched with 50mM EDTA, and spheroplasts were lysed with 0.1% SDS and centrifuged to transfer the supernatant away from insoluble material. The supernatant containing solublized chromatin was incubated at 65°C overnight with 0.4 mg/ml proteinase K to deproteinize DNA and reverse methylene crosslinks. DNA was recovered by two extractions with phenol and one extraction with chloroform, followed by ethanol precipitation and resuspension in Tris-EDTA (TE) pH 8.0 supplemented with 10 μg/ml RNase A. After a 30 min treatment at 37°C, the DNA was ready for probe generation by linear amplification.

Preparation of reference genomic DNA

Genomic DNA was prepared by purification with a Qiagen 100 column after treating spheroplasts with RNase A and proteinase K. Purified DNA was digested with MNase at room temperature to obtain DNA that ranged in size from 100-300bp. This digested DNA was phenol-extracted twice, chloroform-extracted once, and then ethanol-precipitated and resuspended in TE pH 8.0. Genomic reference probe was then prepared by linear amplification in the same manner as mononucleosomal-sized probe.

Linear amplification of DNA to generate microarray RNA probe

Probes for use in microarray experiments were prepared by linear amplification (Liu et al., 2003) but instead of preparing aminoallyl-cDNA probe, aminoallyl-RNA probe was prepared. In brief, DNA obtained by MNase digestion was first treated with calf intestinal phosphatase (New England Biolabs (NEB)) and then thymidine-tailed using terminal dideoxytransferase (NEB). A T7 promoter was adapted to these T-tailed DNA via second strand synthesis with Klenow exo-polymerase (NEB). RNA was next generated using a MegaScript T7 RNA polymerase kit (Ambion) with a 2.3:1 ratio of aminoallyl-UTP to UTP.

Mapping nucleosome positions using tiling microarrays

Nucleosome positions were mapped using a 20bp resolution tiling microarray with the majority of the probes being identical to a previously described microarray version (Yuan et al., 2005). We designed the remaining probes, which included coverage of the PRM1 ORF. This tiling microarray was printed using a custom arrayer and consisted of 32 print blocks with a total of 16,429 bona fide probes. Unless otherwise specified, nucleosomes were mapped in four independently grown cultures for each strain by hybridizing 5μg of mononucleosomal-sized probe and 5μg of reference genomic DNA probe. Hybridizations were conducted at 65°C for at least 12 hr prior to scanning. Dye swaps were incorporated into the experiments in a balanced manner such that with four mononucleosome:reference replicates, two were labeled as Cy3:Cy5 and the remaining two Cy5:Cy3.

Microarray data processing

Raw microarray data was processed and analyzed using custom-written software implemented with Python. Algorithms for statistical analysis were provided by the R statistics software package. A mononucleosome:reference ratio was calculated for each feature by first subtracting the feature median background intensity from the feature foreground intensity and then taking the log base 2 transformed ratio. The feature ratios in each print block were normalized for intensity-dependent bias using a LOESS regression algorithm with a smooth value of 0.4 (LOESS function provided by the R statistics package).

Preparing nucleosome position data for analysis

Analysis of nucleosome enrichment data was done using custom-written software implemented with Python that integrated statistical algorithms from the R statistics package. Nucleosome enrichment values for each experiment were determined by taking the mean of the experiment replicates and applying a centered moving average with a window of 5 features (100 bp). This moving average was strictly implemented such that no missing values were permitted within the window, and each feature in the window must be offset by 20 bp (the array resolution) from its immediate neighbor.

Generation of difference maps

While results from two experiments can be compared side-by-side, generation of a difference map that represents differences between experiments is useful to highlight how the datasets differ from one another. As nucleosome enrichment data are in log2 space, difference maps were generated by subtracting probe data for a control experiment from corresponding probe data for the second experiment. For example, to generate the difference map shown in Fig. 3A, probe data from a control experiment that lacked the Reb1 degron were subtracted from corresponding probe data from an experiment that contained the Reb1 degron. The strains used for both experiments were grown under the same conditions.

Analysis of nucleosome position data as a one-dimensional line trace

Nucleosome positioning data shown in the Figures were generated by the Matplotlib plotting engine module for Python using a coordinate system with probe data representing nucleosome positions overlaid on the positions of features in the genome.

Generation of two-dimensional stacks of nucleosome positioning data and alignment of data at the boundary between NFRs and their downstream nucleosomes

Comparison of nucleosome positions among a set of ORFs is easily achievable by generating a two-dimensional stack in which each row represents data associated with one ORF and each column represents probe data that is relative to a defined reference point in each ORF. These two-dimensional stacks were generated using custom-written Python software for graphical visualization with Java TreeView.

For the specific purposes of this work, all two-dimensional stacks were generated for ORFs that were associated with probe data. As the resolution of the tiling microarray platform is 20bp, bins of 20bp were defined relative to the translational start site of each ORF, and available probe data 1000bp upstream and downstream of each ORF were assigned to the appropriate bin by the coordinate representing the last nucleotide of the probe. This process generated a two-dimensional stack of data arranged by ORF, but centered at the translational start site. This is not an ideal arrangement for analyzing nucleosome-free regions as the distance between translational start sites and NFRs can vary among ORFs. Therefore we developed a method of analyzing nucleosome positions about NFRs in a two-dimensional stack of data in which data for each ORF was aligned at the nucleosome downstream of the NFR.

We thank past and present members of our laboratory for discussions and advice over the course of this research. We thank Leslie Chu and Joachim Li for the pMET3 degron construct. We are especially grateful to Allen Buskirk and David Liu for providing the 4-hydroxytamoxifen-regulated intein construct. This work was supported by a National Institutes of Health Kirschstein predoctoral fellowship (1F31GM083619) to P.D.H. and NIH grant 5R01GM071801 to H.D.M. H.D.M. is a scholar of the Leukemia and Lymphoma Society.