Antagonistic Gcn5-Hda1 interactions revealed by mutations to the Anaphase Promoting Complex in yeast
© Islam et al; licensee BioMed Central Ltd. 2011
Received: 20 January 2011
Accepted: 8 June 2011
Published: 8 June 2011
Histone post-translational modifications are critical for gene expression and cell viability. A broad spectrum of histone lysine residues have been identified in yeast that are targeted by a variety of modifying enzymes. However, the regulation and interaction of these enzymes remains relatively uncharacterized. Previously we demonstrated that deletion of either the histone acetyltransferase (HAT) GCN5 or the histone deacetylase (HDAC) HDA1 exacerbated the temperature sensitive (ts) mutant phenotype of the Anaphase Promoting Complex (APC) apc5 CA allele. Here, the apc5 CA mutant background is used to study a previously uncharacterized functional antagonistic genetic interaction between Gcn5 and Hda1 that is not detected in APC5 cells.
Using Northerns, Westerns, reverse transcriptase PCR (rtPCR), chromatin immunoprecipitation (ChIP), and mutant phenotype suppression analysis, we observed that Hda1 and Gcn5 appear to compete for recruitment to promoters. We observed that the presence of Hda1 can partially occlude the binding of Gcn5 to the same promoter. Occlusion of Gcn5 recruitment to these promoters involved Hda1 and Tup1. Using sequential ChIP we show that Hda1 and Tup1 likely form complexes at these promoters, and that complex formation can be increased by deleting GCN5.
Our data suggests large Gcn5 and Hda1 containing complexes may compete for space on promoters that utilize the Ssn6/Tup1 repressor complex. We predict that in apc5 CA cells the accumulation of an APC target may compensate for the loss of both GCN5 and HDA1.
Eukaryotic genetic information is packaged into chromatin, a highly organized and dynamic protein-DNA complex. The fundamental unit of chromatin, the nucleosome, is an octameric structure composed of two copies of each of the four core histones (an H3/H4 tetramer and two H2A/H2B dimers), surrounded by approximately 146 bp of DNA [1, 2]. Many cellular processes depend on modifications of both DNA and histones within nucleosomes [3, 4]. Modification of chromatin by histone acetyltransferases (HATs) and histone deacetylases (HDACs) play key roles in transcriptional regulation [5–9]. Post-translational acetylation of the highly conserved lysines within the N-terminal tail domains of the core histones is strongly correlated with transcriptional activation [5, 10]. Although the precise mechanisms by which histone acetylation alters transcription are poorly understood [9–12], there is tremendous pressure to understand these mechanisms, as impaired histone modification is linked to many disease states .
The study of HAT and HDAC recruitment to promoters and their interaction with activators and repressors are essential for a better understanding of gene regulation. HATs and HDACs modify histones enzymatically throughout the genome . Histone acetylation potentially regulates transcription by manipulating the higher-order folding properties of the chromatin fiber [15–17]. General control nonderepressible 5 (Gcn5)  was the first identified HAT and exists as the catalytic subunit in multiple high molecular weight complexes in yeast, including SAGA (Spt-Ada-Gcn5-Acetyltransferase), SLIK (SAGA-like), ADA (transcriptional ADAaptor), and the smaller HAT-A2 complex [19–23]. As part of the evolutionarily conserved SAGA complex, Gcn5 predominantly acetylates nucleosomal H3 lysines K9, K18, and K27 . Defects in human SAGA subunits are associated with multiple disorders, including neurological diseases and aggressive cancers [25, 26]. Gcn5 is a direct target for recruitment by transcriptional activators in vitro [27, 28] and in vivo , which results in the acetylation of nearby histones . Elongation of the transcripts initiated by Gcn5-containing complexes is carried out by the Elongator complex, which utilizes Elp3 as its primary HAT [30, 31]. Cell cycle specific roles for Gcn5 have been reported, as recruitment of Gcn5 to a set of genes that are expressed in late mitosis requires SWI/SNF remodelling activity . Furthermore, Gcn5 displays an overlapping pattern of localization with several HDACs [24, 33, 34]. Acetylation microarrays have shown that Rpd3 and Hda1 are the principal HDACs in yeast, affecting numerous promoters throughout the genome with little overlap between promoters [10, 35]. Hda1, an evolutionary conserved HDAC, which deacetylates mainly histones H2B and H3 [36, 37], is recruited to promoters via utilization of different Tup1/Ssn6 domains [38–40], resulting in local deacetylation. HDAC recruitment may form a positive feedback loop to repress transcription locally and facilitate the spreading of Tup1 into adjacent regions . Tup1-mediated repression requires the deacetylation of histones within promoters [42–44], which may require direct recruitment of HDACs [36, 45, 46]. Overall, the mechanisms of Tup1/Ssn6-mediated transcriptional repression can be classified into 3 classes: (i) direct interaction with the activator; (ii) repression by changing chromatin structure; and (iii) interaction with the general transcription machinery [47, 48]. It appears that different groups of genes have developed different strategies to utilize Tup1/Ssn6, enabling it to function as a global repressor.
Our work has linked the Anaphase Promoting Complex (APC), an evolutionarily conserved 13 subunit complex in yeast that is critical for mitotic progression and G1 maintenance [49–52], with chromatin assembly and histone acetylation through genetic interactions with chromatin assembly factor (CAF), HAT and HDAC mutants [53–57]. The APC is a ubiquitin-protein ligase (E3) that targets proteins that block the initiation of anaphase (Pds1) and mitotic exit (Clb2) for degradation. Various regulators govern APC activity in positive and negative manners, from phosphorylation and transcriptional control of APC subunits, to sequestration of APC activators [58–63]. For example, protein kinase A (a complex of Bcy1, Tpk1, Tpk2 and Tpk3) and Mad2 inhibit APC activity through phosphorylation and subunit sequestration, respectively. Activating phosphorylation is supplied by the polo-like kinase (Cdc5) and Cdc28. Furthermore, Cdc20, inhibited by a Mad2-dependent mechanism, binds and activates the APC to promote the metaphase/anaphase transition, while Cdh1, another APC-binding partner, drives APC-dependent mitotic exit. Previous studies by our group have expanded the APC's functional repertoire by showing that the mutant APC subunit allele, apc5 CA , genetically interacted with deletions of the HAT encoding genes GCN5 and ELP3 . Strains harboring the apc5 CA gcn5 Δ or the apc5 CA elp3 Δ mutations had severely restricted growth at elevated temperatures compared to the single mutants. This interaction implies that the APC and these HATs positively interact, but a negative feedback loop appears apparent, as G1-specific Gcn5 instability was reduced in APC mutant cells. An additional synergistic genetic interaction between hda1 Δ and apc5 CA was also observed, suggesting that the APC interacts positively with the HDAC Hda1 . The study presented here focuses on a novel antagonistic relationship between gcn5 Δ and hda1 Δ that is revealed in apc5 CA , but not APC5 cells. We provide further evidence that the APC works with multiple histone modifiers to drive cell cycle progression.
gcn5 Δ/hda1 Δ interactions revealed in an APC mutant background
To examine whether Hda1 positively interacted with the APC, we expressed galactose driven HDA1 carrying a C-terminal HA tag (GAL pro HDA1-HA) at low levels in WT, apc5 CA and gcn5 Δ cells by using glucose as a carbon source (Figure 1B). Recently, we observed that mRNA levels of GAL pro GCN5-HA were elevated 100-fold when grown on 2% glucose and 900-fold when grown on 2% galactose . However, Gcn5-HA protein expression remained low even though GCN5-HA mRNA was 100-fold elevated when grown on 2% glucose. As shown with GCN5 , low-level GAL pro HDA1-HA expression improved apc5 CA growth (Figure 1B). This is not necessarily a general feature of histone modifying proteins, as deletion or overexpression of the HAT HPA2 had little effect on apc5 CA cells (Figure 1B) . Although the yeast Hpa2 has not yet been shown to acetylate histones in vivo, a bacterial acetyltransferase that does acetylate eukaryotic histones is most closely related to Hpa2, and Hpa2 does acetylate H3 in vitro [65, 66]. Moreover, Hpa2 appears to be active, as overexpression reduces growth of gcn5 Δ cells, whereas expression on glucose improves growth of apc5 CA cells (Figure 1B).
A further connection between Gcn5 and Apc5 was observed by the rescue of GAL pro APC5-HA overexpression toxicity by deletion of GCN5 (Figure 1B). It is unlikely that Apc5 protein levels induced from the GAL promoter are compromised in gcn5 Δ cells, as expression of HPA2 and HDA1 from the GAL promoter reduces gcn5 Δ growth. Overexpression of APC5 from the CUP1 promoter also reduced yeast replicative lifespan . Rescue of APC5 toxicity by GCN5 deletion is consistent with our recently proposed hypothesis that Gcn5 is required for APC activity, and may provide an explanation as to why GCN5  and HDA1 (Figure 1B) overexpression is toxic, considering that overabundance of Apc5 is detrimental to cells.
Next, we asked whether mutations to APC5 influenced acetylation of histone H3 lysine 9 or 14 (H3K9/14) in gcn5 Δ and hda1 Δ cells. Gcn5 appears to play a greater role on H3K9, compared to H3K14, whereas loss of HDA1 results in increased acetylation of both H3K9 and H3K14 (Figure 1C). The apc5 CA background did not change the acetylation status of H3K9/14 in gcn5 Δ or hda1 Δ cells, suggesting the apc5 CA background may be revealing an effect other than global histone H3 acetylation. H3K9Ac was reduced in gcn5 Δ, apc5 CA gcn5 Δ and apc5 CA gcn5 Δ hda1 Δ cells, but not in gcn5 Δ hda1 Δ cells. The ability to acetylate H3K9 in gcn5 Δ hda1 Δ cells indicates that on a global level, other HATs can use H3K9 as a substrate. However, at the gene level, deletion of GCN5 was previously shown to reverse histone hyperacetylation at the PHO5 promoter when HDA1 was deleted . Therefore, we tested whether transcript levels are influenced by apc5 CA in gcn5 Δ or hda1 Δ cells.
The apc5 CA allele increases transcript levels in hda1 Δ cells
Increased PDS1 transcripts in apc5 CA hda1 Δ cells correlates with increased promoter acetylation
Consistent with our observations that transcript levels of BCY1 and PDS1 increase in apc5 CA hda1 Δ cells, we detected increased BCY1 and PDS1 promoter acetylation in these cells, specifically at 37°C. Transcript levels and promoter acetylation are both increased with PDS1 at 37°C in apc5 CA hda1 Δ cells. However, we note some differences in the patterns observed. For example, BCY1 transcripts are not elevated in apc5 CA hda1 Δ cells at 37°C while promoter acetylation is. This may reflect the complex nature of the factors assembled at promoters that is not addressed in this study.
Gcn5 promoter occupancy increases in the absence of Hda1
Next we asked whether promoter occupancy by Gcn5 correlated with gene expression and promoter acetylation. GAL pro GCN5-HA was induced in gcn5 Δ and gcn5 Δ hda1 Δ cells so that the only Gcn5 expressed was HA tagged. gcn5 Δ cells expressing GAL pro GCN5-HA grew like WT (data not shown), and were considered the WT control for this experiment. ChIP was performed in lysates prepared from these cells. Control ChIPs were performed using untagged lysates (data not shown), and reactions without antibody, neither of which produced PCR products. Primers against the 5', middle, and 3' regions of CDC20 demonstrated that Gcn5-HA recruitment was most prominent at the promoter and was reduced 5' to 3' (data not shown). We found that in HDA1 cells expressing GCN5-HA, very little Gcn5-HA was present at the promoters tested compared with the RDN1 promoter (Figures 4C and 4D). In hda1 Δ GCN5-HA cells, however, increased Gcn5-HA promoter recruitment was observed. The increases observed were slight except for the CDC20 promoter. Promoter acetylation also increased in hda1 Δ cells, consistent with increased recruitment of Gcn5. These observations present the possibility that i) increased promoter H3K9/14 acetylation in hda1 Δ cells is due to increased Gcn5-HA promoter recruitment; and/or ii) Hda1 may block access of Gcn5 to promoters.
Tup1 occludes Gcn5 promoter occupancy
To distinguish between these possibilities, we predicted that if Tup1 and Hda1 work together, then deletion of TUP1 in apc5 CA cells should have the same synergistic effects as an HDA1 deletion. Our results show that deletion of TUP1 impairs the apc5 CA phenotype (Figure 6C), similar to an hda1 Δ mutation. This suggests that both Hda1 and Tup1 perform a function that is beneficial to APC activity. However, it does not necessarily indicate they work together to perform this task.
Hda1 and Tup1 likely interact at promoters, which can be inhibited by Gcn5
Novel Gcn5/Hda1 antagonistic functional interactions are revealed when APC activity is compromised
The work presented here provides evidence to support a model in which the HAT Gcn5 and the HDAC Hda1 functionally interact at promoters to determine transcriptional readouts (Figure 8). In otherwise WT cells, mutations to GCN5 or HDA1 do not create significant growth defects, whereas in apc5 CA cells, these same mutations produce severe ts growth defects (Figure 1A). The focus of this study was to characterize an antagonistic functional gcn5 Δ/hda1 Δ interaction revealed in the apc5 CA background, as the severe apc5 CA gcn5 Δ and apc5 CA hda1 Δ ts defects are suppressed in apc5 CA gcn5 Δ hda1 Δ cells. Growth phenotypes associated with deletion of GCN5 have been shown in two separate Synthetic Genetic Array (SGA) genome-wide screens to be suppressed by deletion of HDA1 [76, 77]. However, spot dilution analysis of the gcn5 Δ and hda1 Δ cells on YPD did not reveal any phenotypes , as shown in our study (Figure 1A). Thus, the gcn5 Δ hda1 Δ antagonistic interaction is not apparent under normal growth conditions, such as on YPD, but under conditions imposed by the SGA screen (selective media, for example), the antagonistic interaction can be exposed. The influence of the apc5 CA allele on this interaction was investigated. The apc5 CA allele had little effect on global histone H3 acetylation status in gcn5 Δ and hda1 Δ cells, but did cause the increase of BCY1 and PDS1 transcripts in hda1 Δ cells (Figures 1C, 3). Both Bcy1 and Pds1 proteins antagonize APC activity and may be involved in the enhanced growth defect when APC is mutated. Therefore, in apc5 CA cells, it may be the inappropriate expression of inhibitory transcripts that are paramount to synergistic apc5 CA gcn5 Δ and apc5 CA hda1 Δ phenotypes.
A molecular mechanism explaining the Gcn5/Hda1 interaction likely involves competition for Tup1 binding. We observed that in cells lacking HDA1 or TUP1, Gcn5 recruitment at our tested promoters was increased (Figures 4 and 6). On the other hand, deletion of GCN5 increased Hda1-Tup1 physical interactions at promoters (Figure 7). A competition between Hda1 and Gcn5 for Tup1 binding is a possibility worth considering, as both Hda1 and Gcn5 have been shown to physically interact with Tup1 [36, 73–75]. However, in gcn5 hda1 Δ cells this mechanism would not be possible. In addition to the accumulation of Gcn5 in apc5 CA cells, we observed that Elp3 also accumulates when the ubiquitin system is compromised (Figures 4A, B). We previously demonstrated that gcn5 Δ and elp3 Δ deletions impair apc5 CA defects, that GCN5 and ELP3 overexpression stalls the cell cycle in G1, and that Gcn5 G1-specific instability is reversed in APC mutants . Thus, when apc5 CA is combined with gcn5 Δ hda1Δ, an APC target likely accumulates that creates novel transcripts that allow bypass of the severe ts defects observed in the double mutants. Elp3 is an attractive candidate since it is involved in elongating transcripts initiated by Gcn5 containing complexes . A global transcript analysis is likely required to follow this further. Our previous work suggests that the apc5 CA phenotype is sensitive to global transcript levels .
Hda1-dependent occlusion of Gcn5 from promoters requires Tup1
Several reports describe the recruitment of the Tup1/Ssn6 repressor complex to DNA via interactions with multiple partners [41, 48, 68]. Once recruited, Tup1 then contacts H3 and H4 N-terminal tails . Mechanisms employed to recruit Tup1/Ssn6 to promoters by the various individual interacting partners appears to be complex, seems to vary, and may have overlapping roles. Gcn5-HA recruitment to the tested promoters was increased in hda1 Δ, tup1 Δ and ssn6 Δ cells (Figure 6; data not shown), indicating that the interaction of Hda1 with the Tup1/Ssn6 repressor complex is necessary to block access to Gcn5. Tup1 and Hda1 did indeed co-immunoprecipitate while bound to the same promoters, as shown by sequential ChIP (Figure 7). We find it unlikely that Tup1 and Hda1 are simply associating independently at adjacent sequences within the 200-basepair DNA PCR fragment, since they have been shown to interact previously , and are part of large complexes [19–23], but we cannot discount this possibility. However, we observed that in gcn5 Δ cells, Hda1-Tup1 association increased at some promoters (PDS1 and BCY1), suggesting Gcn5 opposes complex formation. The mechanism of action that Gcn5 uses to block Hda1-Tup1 association remains unclear. Previous reports indicating that Tup1 is capable of recruiting and interacting with Gcn5/SAGA at promoters [73–75] suggest it is possible that Gcn5 and Hda1 may compete for Tup1 interaction. The scenario for recruiting either Gcn5 or Hda1 would differ, implying other proteins may be involved in deciding whether Gcn5 or Hda1 gain access. We were unable to observe complex formation between Gcn5-TAP and Hda1-HA in whole cell lysates (data not shown), indicating possible exchange of Gcn5 and Hda1 at Tup1 complexes does not require Gcn5-Hda1 association. It is also possible that Gcn5-Hda1 physical interactions are transient and promoter specific, therefore may not be detectable using the methods applied here. Nonetheless, support for our model was provided by reports describing recruitment of Gcn5 to promoters by the Tup1/Ssn6 complex under osmotic stress conditions [40, 74], indicating that Tup1/Ssn6 may be a transcriptional activator under certain conditions.
The results presented in this manuscript provide evidence for a complex network of interactions between a mitotic/G1 cell cycle regulator (the APC), and antagonistic interplay between a HAT (Gcn5), and an HDAC (Hda1). Gcn5 is known to function during mitosis [32, 57, 79, 80]. Data on the role Hda1 plays in cell cycle progression is limited, but Hda1 may provide some function to ensure histones are deacetylated prior to passage through mitosis . It is noteworthy that Gcn5 and Hda1 expression is temporally regulated during the cell cycle (microarray data compiled at Saccharomyces Genome Database), providing insight into how the potential competition for Tup1 binding could be regulated. APC mutations cause cell cycle progression to stall during mitosis, potentially skewing the equilibrium between Gcn5 and Hda1 promoter recruitment if the cell cycle does indeed influence Hda1 and Gcn5 recruitment. Future work will focus on identifying the molecular mechanisms regulating how cell cycle progression influences chromatin dynamics. Chromosome synthesis and segregation defects are widely associated with human disease, thus continued work into furthering our understanding of this process is vital.
Media, yeast strains, plasmids and general methods
Yeast strains used in this study
MATα ade2 his3 Δ200 lys2 Δ201 ura 3-52
MATa ade2 his3 Δ200 lys2 Δ201 ura3-52
MATa his3 Δ1 leu2 Δ met15 Δ ura3 Δ tup1 Δ::kanMX6
MATa his3 Δ1 leu2 Δ met15 Δ ura3 Δ ssn6 Δ::kanMX6
MATa ade2 his3 Δ200 lys2 Δ201 ura3-52
MAT(?) ade2 his3 leu2 lys2(?) ura3 apc5 CA -PA::His5 + tup1 Δ::kanMX6
MAT(?) ade2 his3 leu2 lys2(?) ura3 hda1 Δ::kanMX6
MAT(?) ade2 his3 leu2 lys2(?) ura3 apc5 CA -PA::His5 + hda1 Δ::kanMX6
MAT(?) ade2 his3 leu2 lys2(?) ura3 gcn5 Δ::kanMX6
MAT(?) ade2 his3 leu2 lys2(?) ura3 apc5 CA -PA::His5 + gcn5 Δ::kanMX6
MAT(?) ade2 his3 leu2 lys2(?) ura3 gcn5 Δ::kanMX6 hda1 Δ::kanMX6
MAT(?) ade2 his3 leu2 lys2(?) ura3 apc5 CA -PA::His5 + gcn5 Δ::kanMX6 hda1 Δ::kanMX6
MATa his3 Δ1 leu2 Δ met15 Δura3 Δ rpn10 Δ::kanMX6
MATa his3 Δ1 leu2 Δ met15 Δ ura3 Δ GCN5-TAP::HIS3
as YTH1235, with GCN5-TAP::HIS3
as YTH5, with tup1 Δ::kanMX6
as YTH5, with ssn6 Δ::kanMX6
MAT(?) ade2 his3 leu2 lys2(?) ura3 gcn5 Δ::kanMX6 ssn6 Δ::kanMX6
MAT(?) ade2 his3 leu2 lys2(?) ura3 gcn5 Δ::kanMX6 tup1 Δ::kanMX6
MATa his3 Δ1 leu2 Δ met15 Δ ura3 Δ APC5-TAP::HIS3 rpn10 Δ::kanMX6
Plasmids used in this study
URA3 CEN ARS
GAL pro -APC5-HA
2μ GAL10 pro -APC5-HA URA3
GAL pro -GCN5-HA
2μ GAL10 pro -GCN5-HA URA3
GAL pro -HDA1-HA
2μ GAL10 pro -HDA1-HA URA3
GAL pro -HPA2-HA
2μ GAL10 pro -HDA1-HA URA3
2μ CUP1 pro -TUP1 URA3
2μ CUP1 pro -TUP1 URA3
Primers generated for the Northern analysis
Reverse transcriptase PCR (rtPCR)
Primers generated for the ChIP analysis
Chromatin immunoprecipitation (ChIP)
ChIP was performed essentially as described elsewhere [82, 83] with the following modifications: DNA fragment size achieved by sonication was 500-1000 bp, and 100 μg of protein lysate was used for each IP. Protein concentration was determined by a Bradford protein assay. 5 μg of ChIP grade rabbit polyclonal anti-acetyl-H3K9/14 (Upstate Biotechnology), rabbit polyclonal anti-H3 (Abcam), rabbit polyclonal HA antibody (Abcam), and rabbit polyclonal GST antibody (Abcam) were used for IP. One-tenth of the total volume of lysate was used as input for each sample. Sequential ChIP was performed as previously described . In sequential ChIP experiements, the immune complexes were eluted by incubation for 30 minutes at 37°C in 10 mM DTT. After centrifugation, the supernatant was diluted 25 times with ChIP dilution buffer (1% Triton X-100, 2 mM EDTA, 150 mM NaCl, 20 mM Tris-HCl [pH 8.1]) and subjected again to ChIP using a different antibody. In this experiment, HA antibody was applied first, followed by GST antibody. Cross-linking of the immune complex was reversed by adding NaCl to a final concentration of 0.3 M and incubated overnight at 65°C. Samples were treated first with 1 μg/μl RNaseA (Millipore [formerly Upstate]) for 30 minutes at 37°C, followed by 1 μg/μl proteinase K (Millipore [formerly Upstate]) at 45°C for 1 hour. DNA was purified by chromatography on QIAquick columns, and eluted with elution buffer (PCR purification kit, Qiagen). PCR was performed for semiquantitative determination by standard end point PCR. 1 μl DNA was used for PCR, and the reaction continued to the predetermined mid-linear range for each primer set. The end point PCR product was resolved on a 1% agarose gel and visualized by ethidium bromide. Two independent experiments were performed for each ChIP. The gel bands from each experiment were analyzed by ImageJ, and the means and standard error were plotted for graphical representation. For time course experiments, 200 ml cultures were induced at a final concentration of 4% galactose. Samples (20 ml) were immediately removed, and again after 1, 3 and 5 hours. The 20 ml samples were in duplicate for Western and ChIP analysis.
AI was supported by Post-Doctoral Fellowships from the Saskatchewan Health Research Foundation (SHRF), and from the Canadian Institutes for Health Research-Regional Partnership Program (CIHR-RPP). ELT was supported by Graduate Scholarships from the College of Graduate Studies at the U of S. Deletion mutants and plasmids were kindly provided by Dr. W. Xiao (University of Saskatchewan). We thank members of the Harkness lab, Ata Ghavidel and Spike Postnikoff, for careful reading of the manuscript and for providing insightful suggestions. Funding was generously provided to TAAH through a CIHR Operating Grant and a New Investigator Award from the Canadian Foundation for Innovation (CFI).
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