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Previous Article | Next Article 
Molecular and Cellular Biology, November 2006, p. 7913-7928, Vol. 26, No. 21
0270-7306/06/$08.00+0 doi:10.1128/MCB.01220-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Markus Posch,2,
Iwona Sadzak,1,
Katrin Ramsauer,3
Gerda Egger,1,¶
Reinhard Grausenburger,1
Norbert Schweifer,2
Susanna Chiocca,4
Thomas Decker,3 and
Christian Seiser1*
Max F. Perutz Laboratories, Department of Medical Biochemistry, Medical University of Vienna, Vienna Biocenter, A-1030 Vienna, Austria,1 Boehringer Ingelheim Austria, A-1121 Vienna, Austria,2 Max F. Perutz Laboratories, Institute of Microbiology and Genetics, University of Vienna, Vienna Biocenter, A-1030 Vienna, Austria,3 European Institute of Oncology, Department of Experimental Oncology, 20141 Milan, Italy4
Received 6 July 2006/ Returned for modification 17 July 2006/ Accepted 21 August 2006
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ABSTRACT |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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A correlation
between histone acetylation and increased gene expression was
discovered earlier on (3).
According to the current model, the acetylation of lysine residues
within the histone tails neutralizes the positive charge of
-amino groups and thereby reduces the interaction between the
N-terminal tails of histones and the negatively charged DNA.
Acetylation at the N termini of core histones is therefore believed to
induce the local opening of chromatin structures. In addition,
acetylated histone tails are specifically recognized and bound by
bromodomain-containing proteins, such as components of the basal
transcription machinery or histone acetyltransferases (HATs)
(85). Reversible histone
acetylation is controlled by histone acetyltransferases, which usually
act as transcriptional coactivators, and histone deacetylases (HDACs),
which repress transcription. Activator complexes containing HAT
activity have been shown to contribute to transcriptional activation by
recruitment of general transcription factors and RNA polymerase II
(7,
74). In contrast,
recruitment of repressor complexes with HDAC activity is considered to
lead to deacetylation of histones, stabilization of nucleosome
structure, and formation of a repressive chromatin state.
During the last decade, more than a dozen histone deacetylases have been identified in mammalian cells. Based on sequence similarities, HDACs are divided into four functional classes: class I (HDAC1, HDAC2, HDAC3, and HDAC8), class II (HDAC4, HDAC5, HDAC6, HDAC7, HDAC9, and HDAC10), class III (SIRT1 to SIRT7), and the recently described class IV of HDACs, which consists of HDAC11-related enzymes (28, 29). The class I enzyme HDAC1 belongs to an ancient family of highly conserved enzymes and was the first protein shown to have histone deacetylating activity in mammals (reviewed in reference 46). Human HDAC1 was purified and cloned by an affinity purification approach (73) and was shown to share significant homology with the previously identified Saccharomyces cerevisiae transcriptional regulator Rpd3/Sdi2/Sds6 (51, 80-82).In mouse cells, expression of the HDAC1 gene is stimulated by growth factors (5) and controlled by its own product in a negative feedback loop (32, 65). The enzyme plays an important role in various biological processes, such as cell cycle progression, cell proliferation, and differentiation (46). The HDAC1 mouse knockout (KO) has also revealed the essential function of this deacetylase in embryonic development (36). HDAC1 is a nuclear protein and can heterodimerize with the closely related deacetylase HDAC2 (31, 72). Both enzymes are found in three major multiprotein complexes, named Sin3, NuRD, and CoREST (2, 29). HDAC1 can repress gene transcription either directly or as part of these multiprotein complexes when recruited by a variety of transcriptional regulators, including SP1/SP3, YY1, unliganded nuclear receptors, the pocket proteins pRB, p107, and p130, and the tumor suppressor p53 (13, 52).
HDAC inhibitors have been shown to induce cell cycle arrest, differentiation, or apoptosis in tumor cells, and some of these compounds are currently tested as antitumor drugs in clinical trials (19, 20, 44). These inhibitors affect the catalytic activity of most class I and class II deacetylases. However, little is known about the individual roles of mammalian deacetylases in transcriptional control and the relevant target enzymes for HDAC inhibitors as antitumor drugs. To unravel the role of HDAC1 as a transcriptional regulator, we identified putative HDAC1 target genes by comparing the gene expression profiles of wild-type and HDAC1-deficient embryonic stem (ES) cells. A restricted subset of mouse genes involved in biological processes, such as growth control, cell communication, and transcriptional regulation, was found to be reversibly deregulated upon the loss of HDAC1. By chromatin immunoprecipitation (ChIP) assays, we observed the presence of the HDAC1 enzyme associated with a reduction in histone acetylation levels at the regulatory regions, thereby providing compelling evidence for a direct regulation of these genes by HDAC1. In addition, our data revealed a delicate balance between the class I enzymes HDAC1 and HDAC2 in the regulation of common target genes. Finally, we also identified a novel function for HDAC1 as a positive regulator of gene expression.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Affymetrix analysis. Total RNA (approximately 5 µg) was used to synthesize double-stranded cDNA by using a custom Superscript double-stranded cDNA synthesis kit (Invitrogen, Karlsruhe, Germany). Biotin-labeled cRNA was then prepared from this template by using an Enzo BioArray high-yield RNA transcript labeling kit (Affymetrix, High Wycombe, United Kingdom), and unincorporated nucleotides were removed using RNeasy columns (QIAGEN, Hilden, Germany). Hybridization, washing, and fluorescence staining of the Affymetrix GeneChip murine genome MG-U74Av2 array (Affymetrix Inc., Santa Clara, CA) were carried out according to the manufacturer's instructions (GeneChip Expression Analysis Technical Manual; Affymetrix). All experiments were performed in triplicate with independently extracted RNAs. Data analysis was performed by means of a comparison matrix, with control (wild-type) experiments as a background, for generation of expression signal log ratios with basis 2 and subsequently changes (n-fold) between wild-type and knockout samples, using Microarray Suite software version 5.0 (Affymetrix). Next, mean changes (n-fold) and relative standard deviations between experiments were calculated and genes with changes (n-fold) of >2 or <0.5 and relative standard deviations of <0.6 were selected for further analysis. The association of genes with particular functions, pathways, and diseases was analyzed through the use of Ingenuity Pathways Analysis (Ingenuity Systems). Gene networks are ranked according to their scores. Calculations for network scores are based on the hypergeometric distribution calculated via the computationally efficient Fisher exact test for two-by-two contingency tables. The significance value associated with functions and pathways is a measure of how likely it is that genes from the data set file participate in that function. The significance is expressed as a P value, which is calculated using the right-tailed Fisher exact test.
Protein analysis and antibodies. Whole-cell protein extraction, histone isolation, and Western blot analysis were performed as previously described (5, 36, 59). The following antibodies were used for protein detection on immunoblots and for chromatin immunoprecipitation assays: HDAC1 (polyclonal rabbit antibody and monoclonal mouse antibody), HDAC2 (polyclonal rabbit antibody and monoclonal mouse antibody), acetyl histone-H3, acetyl histone-H4, acetyl K9-H3 from Upstate, and the C terminus of histone H3 (Abcam). Polyclonal trimethyl K9-H3 and trimethyl K27-H3 antibodies were kind gifts from T. Jenuwein (59). The ß-actin protein was visualized with a monoclonal antibody (AC-74; Sigma). The proliferation marker Ki67 was detected with the monoclonal Ki67 antigen antibody (Novo Castra) by indirect immunofluorescence microscopy (Zeiss Axiovert 135TV microscope) as previously described (72). Nuclear DNA was visualized with 4',6-diamidino-2-phenylindole (DAPI).
Immunoprecipitation assays. Immunoprecipitation assays were performed as previously described (18). For combined analysis of proteins and associated deacetylase activity, the precipitated material was resuspended in 50 µl of extraction buffer (20 mM Tris-HCl [pH 8.0], 100 mM NaCl, 1 mM EDTA, 0.5% Nonidet P-40, 1 mM phenylmethylsulfonyl fluoride, 2 mM dithiothreitol, Roche Diagnostics complete protease inhibitor cocktail), and 30-µl aliquots were examined for protein expression on Western blots. The remaining 20-µl aliquot was assayed for HDAC activity. Histone deacetylase activity assays were done as described previously (5, 37).
Chromatin immunoprecipitation assays. Chromatin immunoprecipitation assays were carried out as described previously (8, 11, 65) with some modifications. Chromatin was cross-linked for 10 min by using formaldehyde and then sonicated. Equal amounts of sonicated chromatin were diluted 10-fold and precipitated overnight with the following antibodies: HDAC1, HDAC2, acetyl histone-H3, acetyl histone-H4, acetyl K9-H3, trimethyl K9-H3, trimethyl K27-H3, and preimmune serum as a control. The chromatin-antibody complexes were isolated by incubation with 30-µl protein A-Sepharose beads (50% slurry, 100 µg/ml salmon sperm DNA, 500 µg/ml bovine serum albumin) while rocking at 4°C for 2 hours. The beads were harvested and washed as described previously (65).Chromatin-antibody complexes were eluted from the protein A-Sepharose beads by addition of 2% sodium dodecyl sulfate, 0.1 M NaHCO3, and 10 mM dithiothreitol. Cross-linking was reversed by addition of a 0.05 volume of 4 M NaCl and incubation of the eluted samples for 6 h at 65°C. The DNA was extracted with phenol-chloroform, precipitated with ethanol, and dissolved in water.
PCR analysis of immunoprecipitated DNA. All PCRs were performed on an iCycler (Bio-Rad) by using Promega PCR Master Mix. The linear range for each primer pair was determined empirically using different amounts of genomic DNA. PCRs with 1:40 dilutions of genomic DNA (input) were carried out along with the immunoprecipitated DNA. PCR products were resolved on 2% agarose-Tris-acetate-EDTA gels. Primer sequences are available upon request.
RNA isolation, Northern blotting, and real-time RT-PCR analysis. Total cellular RNA was isolated with TRIzol reagent (GibcoBRL) as recommended by the manufacturer. Northern blot hybridization was performed by the sandwich method as previously described (68). For cDNA, 1 µg of total RNA was reverse transcribed with an iScript cDNA synthesis kit (Bio-Rad). Real-time RT-PCRs were performed with 0.5 µl of the RT reaction mixture by the iCycler iQ system (Bio-Rad), using SYBR green (Molecular probes) for labeling. Primer sequences are available upon request.
Microarray data accession number. The microarray data have been deposited in the Gene Expression Omnibus (GEO) public database (http://www.ncbi.nlm.nih.gov/geo/) under the accession number GSE 5583.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Given the growth-inhibitory effects of HDAC inhibitors, the impaired proliferation of HDAC1 KO cells points towards HDAC1 as one of the possible targets for these tumor drugs (36). In this context, it is remarkable that several tumor-related genes, such as those encoding JunB, Prss11, Plagl1, Apc2, metallothionein 1, and metallothionein 2, were found in our screen (see also Discussion). In addition, a relatively high number of imprinted genes were found to be either upregulated (the Peg3, H19, and Plagl1 genes) or downregulated (the Igf2 and p57 genes) in this screen. Of the 35 imprinted genes present on the array, which represent about 55% of all known imprinted genes in the mouse (16, 49), 10 were deregulated in the absence of HDAC1 (data not shown). Most of these genes have been implicated in proliferation and growth control. Intriguingly, chromosome 7, which contains several clusters of imprinted genes, showed a significant deviation from the expected chromosomal distribution of deregulated genes, with 25% more deregulated genes than expected (Fig. 1D). In general, the chromosomal distribution of deregulated genes (i.e., the number of genes found to be deregulated on a specific chromosome relative to the total number of deregulated genes) was similar to the chromosomal distribution of genes present on the array and the chromosomal distribution of genes found in the whole mouse genome (Fig. 1D).
The deregulation of HDAC1 target genes is reversible. As a first step in the analysis of putative HDAC1 target genes, we wanted to validate the results from the DNA array screen. To this end, we isolated total RNA from wild-type and HDAC1 KO ES cells and analyzed a subset of randomly picked potential target genes by real-time RT-PCR and Northern blot analyses. Of the 33 deregulated genes examined, 16 upregulated and 12 downregulated genes were successfully confirmed (Fig. 2). Three genes (the Apobec2, Gata4, and Gjb3 genes) did not show the expected result, while two genes (the Rb1 and Rfx2 genes) were upregulated below the defined threshold level.
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Furthermore, mRNA levels of Igf2 and Dnmt3a, which were downregulated in KO cells, were restored almost to wild-type-cell levels after reintroduction of HDAC1 in both cell lines (Fig. 3D). The fact that p57 expression was reestablished in KO-reB cells but not in KO-reA cells might be due to the complex interplay of allele-specific repressive chromatin modifications that is linked to the regulation of imprinted genes (see Discussion). Taken together, these data suggest that HDAC1 can reversibly regulate the expression of a specific subset of mouse genes.
Cross talk between the class I deacetylases HDAC1 and HDAC2. Next, we wanted to know whether the enzymatic activity of HDAC1 is required for the regulation of putative target genes. For that purpose, we expressed an inactive HDAC1 protein in HDAC1-null cells. The mutation of histidines 140/141 into alanines in the catalytic domain leads to a complete loss of HDAC1 enzymatic activity without provoking conformational changes in the protein (31, 72). Wild-type and HDAC1 KO cells were stably transfected with a retroviral expression vector encoding the myc-tagged HDAC1-H140/141A mutant or the empty vector as a control. Interestingly, of several dozens of clones analyzed by indirect immunofluorescence analysis and immunoblotting (data not shown), only one showed a clearly detectable signal for the mutant HDAC1 protein. This clone, referred to as KO-mut, expressed HDAC1-H140/141A at a relatively low level compared to the levels of HDAC1 in the wild-type control (Fig. 4A, left panel), suggesting that constant high expression of the inactive HDAC1 mutant might interfere with proliferation of ES cells. Hence, this clone was taken for further analyses. Remarkably, the total HDAC activity in the HDAC1 mutant-expressing cells was about 10% lower than the activity in KO control cells (Fig. 4A, right panel). This might be due to the slightly reduced HDAC2 expression in HDAC1-H140/141A-expressing cells and/or to a negative effect of the mutant HDAC1 protein on HDAC2 activity. Alternatively, HDAC2 might act as an impostor in the absence of HDAC1, a mechanism that has previously been described for certain mitogen-activated protein kinases (40).
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To analyze how the HDAC1-H140/141A mutant affects the transcription of putative HDAC1 target genes, we examined the expression of selected target genes by quantitative real-time RT-PCR (Fig. 4C). In contrast to that of the HDAC1 wild-type protein, expression of the inactive mutant did not restore normal expression of putative HDAC1 target genes. Most of the genes analyzed, e.g., those encoding Mt2, Prss11, p21, Mt1, Plagl1, and Ass1, showed no change or a moderate increase in mRNA levels in HDAC1-H140/141A-expressing cells compared to those in the KO control. Strikingly, two of the genes analyzed, the Apc2 and JunB genes, had three- to four-times-higher expression levels in KO-mut cells. Given the fact that the inactive HDAC1 mutant reduces the enzymatic activity of HDAC2, the strong increases in the expression levels of these two genes suggest that, in the absence of HDAC1, HDAC2 might be involved in their regulation.
HDAC inhibitor treatment reveals two distinct groups of HDAC1 target genes. In order to determine whether other histone deacetylases are involved in the regulation of putative HDAC1 target genes, we next analyzed the responsiveness of putative HDAC1 target genes to the deacetylase inhibitor TSA, a general inhibitor of most class I and class II HDACs (43, 86). To this end, wild-type and HDAC1-deficient ES cells were treated for 12 h with the solvent dimethyl sulfoxide (DMSO) or increasing concentrations of TSA. The expression of a panel of putative HDAC1 target genes was then analyzed by real-time RT-PCR. The known TSA responsive gene HDAC1 (26, 32, 65) showed a dose-dependent increase in expression in wild-type ES cells (Fig. 5A). As expected for HDAC1 target genes, all tested genes displayed increased expression in response to TSA treatment in wild-type cells (Fig. 5B-I). Furthermore, in the absence of HDAC1, all tested target genes were less sensitive to the deacetylase inhibitor. These results confirm a crucial role for HDAC1 and its enzymatic activity in the regulation of these genes. Interestingly, in HDAC1 KO cells, genes such as those encoding Prss11, p21, Ass1, and Mt2 (Fig. 5B to E) showed only moderate response (two- to threefold) to TSA, suggesting that the major histone deacetylase involved in their regulation is HDAC1. In contrast, four other genes, the Mt1, JunB, Apc2, and Plagl1 genes, displayed a strong response to TSA (8- to 22-fold) in HDAC1 KO cells (Fig. 5F to I), indicating that these genes are regulated by other HDACs as well. As shown above, the expression of Apc2 and JunB was significantly enhanced in HDAC1 KO cells upon the expression of the enzymatically inactive mutant, i.e., under conditions where HDAC2-associated deacetylase activity was reduced (Fig. 4C). These findings strongly suggest that HDAC2 is involved in the regulation of Apc2 and JunB. Another putative HDAC1 target gene, the Plagl1 gene, showed a different response; it was not induced by the inactive HDAC1 mutant but showed a dramatic increase in expression upon TSA treatment. A possible explanation is that Plagl1 might be regulated by other histone deacetylases, which are sensitive to TSA but unaffected by the expression of the inactive HDAC1 mutant. Moreover, since the Plagl1 gene is an imprinted gene, the regulation of its expression might be more complex. Taken together, these data imply that HDAC1 putative target genes can be divided into two groups: genes that are regulated mainly by HDAC1 (e.g., the Prss11 and Ass1 genes) and genes, such as the JunB and Apc2 genes, that are regulated by HDAC1 and other histone deacetylases.
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In contrast, HDAC2 recruitment was highly increased at the JunB and Apc2 promoters in HDAC1 knockout cells. This is consistent with the finding that these genes respond robustly to TSA even in the absence of HDAC1 (Fig. 5G and H). We therefore conclude that both HDAC1 and HDAC2 are involved in the repression of these genes, probably within the same complexes, where the increased presence of HDAC2 can partially compensate for the loss of HDAC1 or act as an impostor. This idea is also supported by the fact that the expression of the inactive HDAC1 mutant activates JunB and Apc2 in KO cells, either by negatively affecting HDAC2 activity or by replacing HDAC2 in putative regulatory complexes.
To examine the effect of the loss of HDAC1 activity on local histone modifications, we next examined the acetylation levels of histones H3 and H4 at the chosen HDAC1 target genes in wild-type and HDAC1-deficient cells. In addition, we analyzed these genes for the presence of the repressive marks trimethyl K9 and trimethyl K27 at histone H3. To test whether the absence of HDAC1 and increased transcription affect the nucleosome density, we also performed ChIP analyses with an antibody that recognizes the C terminus of histone H3. As shown in Fig. 6, most of the target genes (with the exception of the Apc2 promoter) showed slightly reduced nucleosome densities at both the promoters and the intragenic regions in HDAC1 KO cells. Despite the small reduction in associated histone proteins upon the loss of HDAC1, we consistently observed increased acetylation levels of histones H3 and H4 on nucleosomes associated with the promoters and intragenic regions of all tested genes (Fig. 6). Furthermore, the analysis of modifications of specific lysine residues on the HDAC1 target gene promoters revealed a strong hyperacetylation of lysine 9 on histone H3 within all tested target gene regions. Taking into account the slight reduction in nucleosome density, we observed at the same time the loss of the trimethylation marks on both lysine 9 and lysine 27 on histone H3 in KO cells.
Taken together, these data show a strong correlation between the presence of HDAC1 and decreased histone acetylation at specific genes, indicating that these are indeed direct HDAC1 target genes. Our results also emphasize the role of HDAC1 as an epigenetic regulator, by showing that the loss of this enzyme can provoke changes not only on its direct substrates, acetylated histones, but also on substrates of other histone-modifying enzymes.
An HDAC1 target gene, the Prss11 gene, encodes a regulator of cell proliferation. An HDAC1 target gene, the Prss11 gene, is a putative tumor suppressor gene in ovarian cancer (12) and was shown to be consistently upregulated upon the loss of HDAC1 expression in a variety of cell types, including mouse fibroblasts and human tumor cells (G. Zupkovitz, S. Chiocca, and C. Seiser, unpublished observations). Remarkably, small interfering RNA-mediated ablation of HDAC1 impaired the proliferation of the human osteosarcoma cell line U2OS, suggesting that HDAC1 might be a relevant target for HDAC inhibitors as tumor drugs (S. Senese, K. Zaragoza-Dorr, S. Minardi, L. Bernard, G. F. Draetta, M. Alcalay, C. Seiser, and S. Chiocca, submitted for publication). To test whether enhanced expression of Prss11 would affect the proliferation rates of human tumor cells, U2OS cells were stably transfected with a Prss11 expression vector. Vector-transfected control cells and Prss11-overexpressing single clones were analyzed in proliferation assays and for expression of the proliferation marker Ki67. As shown in Fig. 7, both cell numbers and percentages of Ki67 positive cells were significantly reduced in Prss11-overexpressing tumor cells. Thus, the HDAC1 target gene Prss11 gene encodes a negative regulator of cell proliferation and its overexpression in the absence of HDAC1 might contribute to the proliferation phenotype observed in HDAC1 KO ES cells and HDAC1-deficient tumor cells.
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Four putative target genes, the Edg1, Efnb2, Ehd1, and Gja1 genes, showed significant downregulation in HDAC1 KO cells (2.5- to 5-fold) when tested by real-time RT-PCR analysis (Fig. 8A). The expression of three of these genes (the Efnb2, Ehd1, and Edg1 genes) was induced by TSA in HDAC1-null cells, indicating that these genes are negatively regulated by HDACs in the absence of HDAC1. Efnb2 and Ehd1 showed no significant response to the deacetylase inhibitor in wild-type cells, suggesting that their expression is normally not controlled by TSA-sensitive deacetylases. In contrast, higher concentrations of TSA stimulated Edg1 expression in wild-type cells and HDAC1 KO ES cells, indicating that this gene is negatively regulated by HDACs in both cell types.
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In contrast to those of the target genes described above, expression levels of Gja1 were reduced not only upon the loss of HDAC1 but also in a dose-dependent manner by TSA (Fig. 8A). As demonstrated by ChIP analyses, HDAC1 was present at the Gja1 promoter in wild-type cells, while HDAC2 was recruited mainly in HDAC1-null cells (Fig. 8B). Given that Gja1 expression is further reduced by TSA treatment of HDAC1 KO cells, HDAC2 might to some extent compensate for the loss of HDAC1. Analysis of chromatin modifications at the Gja1 promoter revealed that the acetylations of histone H4 and K9 at histone H3 were slightly reduced in the absence of HDAC1 (Fig. 8B), while the trimethylation of H3-K9 was increased in HDAC1 KO cells (Fig. 8B). Thus, the reduced expression of Gja1 correlated with the increased presence of the repressive K9 trimethylation mark on histone H3. The fact that histone acetylation on the Gja1 promoter is only slightly affected suggests that for the regulation of this gene, other proteins are the relevant targets for HDAC1. In summary, the Gja1 gene represents an unusual HDAC1 target gene that requires the presence of HDAC1 and its enzymatic activity for activation.
HDAC1 and its activity are required for the activation of IFN target genes.
Next, we asked whether the positive
role of HDAC1 in the regulation of gene expression is a more general
phenomenon. Several reports suggest that histone deacetylase activity
is necessary for activation of interferon (IFN) target genes, probably
through involvement of HDAC1
(9,
54,
63). Interferon
stimulation leads to activation of the JAK/STAT pathway, resulting in
tyrosine phosphorylation of STAT1 and STAT2. Homodimerization of
phosphorylated STAT1 or heterodimerization of phosphorylated STAT1 and
STAT2 induces their translocation into the nucleus and the activation
of their target genes (for a review, see reference
15). To test a direct
involvement for HDAC1 in the interferon response, we analyzed the
expression of IFN target genes in wild-type and HDAC1 KO ES cells.
Pilot experiments showed that IFN treatment induced STAT1
phosphorylation in both wild-type and HDAC1-null ES cells (data not
shown). We next stimulated wild-type and HDAC1 KO ES cells with
IFN-
in the presence or absence of TSA and analyzed the
expression of two known IFN- target genes, the Irf1 and Gbp2
genes. Real-time RT-PCR analysis demonstrated the transcriptional
activation of both genes in response to IFN- in wild-type
cells (Fig.
9A). In contrast, the presence of the deacetylase inhibitor TSA during
IFN- stimulation inhibited the induction of both genes.
Interestingly, the interferon-dependent activation levels of both Irf1
and Gbp2 were significantly reduced in HDAC1 KO ES cells. These data
strongly support the idea that histone deacetylase activity, and in
particular HDAC1 activity, is important for the induction of certain
interferon target genes.
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and in the
presence or absence of TSA, using specific antibodies for HDAC1, the C
terminus of histone H3, and acetylated histones H3 and H4. In addition,
we monitored the presence of trimethylated K9 and K27 at histone H3 at
both target promoters. The immunoprecipitated DNA was further analyzed
by semiquantitative PCR or real-time PCR with primers specific for the
IFN--activated site of the Irf1 promoter and the proximal
promoter region of Gbp2. As shown in Fig.
9B, HDAC1 was absent from
both promoters in unstimulated wild-type cells but was recruited in
response to IFN- stimulation. While TSA treatment abolished
the IFN--dependent induction of both genes, it did not affect
the presence of HDAC1 on the Irf1 promoter in IFN--treated
wild-type cells, suggesting that the activity of recruited HDAC1 is
essential for gene activation. Small amounts of HDAC1 were also found
on both promoters upon treatment with TSA alone. This might be due to
the previously demonstrated induction of HDAC1 expression by TSA
(32,
65) or to increased
recruitment caused by TSA-mediated acetylation of HDAC1
(60). As shown in ChIP assays with the C-terminal H3 antibody, the nucleosome density at the Irf1 promoter was slightly reduced in the presence of TSA and was unchanged at the Gbp2 promoter in both wild-type and HDAC1 KO cells under all conditions tested. Histone acetylation was mostly unaffected upon IFN treatment, with the exception of H4 acetylation at the Irf1 promoter in wild-type cells, which was decreased upon recruitment of HDAC1. Treatment with TSA led uniformly to hyperacetylation of histone H4 on both promoters. Since there was no consistent reduction of histone acetylation during the activation of these target genes (Fig. 9B), we conclude that the crucial substrates for HDAC1 seem to be nonhistone proteins.
Remarkably, IFN-dependent induction of both genes in wild-type cells correlated with reduced K9 trimethylation of histone H3 (Fig. 9B). In HDAC1 KO cells, levels of H3-K9 trimethylation at both promoters were relatively low and not affected by IFN or TSA treatments, with the exception of the Gbp2 promoter under TSA treatment. Methylation of H3-K27 was reduced on both promoters upon IFN or TSA treatment of wild-type cells and was generally lower in HDAC1 KO cells. Thus, activation of both target genes was consistently accompanied by the gradual loss of repressive chromatin marks. All together, these data strongly suggest that HDAC1 and histone deacetylase activity are necessary for the induction of certain IFN target genes.
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Biological function of HDAC1 target genes. Database analysis of HDAC1 target genes showed that the deacetylase controls the expression of genes involved in a variety of biological processes. Based on the phenotype of HDAC1-deficient embryos and ES cells, HDAC1 has been implicated in proliferation control. Along this line, a significant fraction of HDAC1 target genes is involved in growth control and cell communication. In particular, several genes with proposed tumor suppressor activity (the JunB, Plagl1, Apc2, metallothionein 1, metallothionein 2, and Prss11 genes) are regulated by HDAC1. For instance, JunB is downregulated in several human tumors (10, 57) and was shown to suppress cell proliferation by transcriptional activation of p16 (58). The Plagl1 gene, which encodes a growth suppressor, is frequently silenced in ovarian and breast cancer cells (1, 6), the adenomatous polyposis coli (APC)-like APC2 gene encodes a putative tumor suppressor (34), and the genes for Mt1 and Mt2 are repressed in some metastatic tumors (87).
The Prss11 gene was originally isolated as a gene whose expression was downregulated in a human fibroblast cell line after transformation with simian virus 40 (89). Repression of human Prss11 has been repeatedly observed in ovarian cancers (66) and melanomas (4), in close correlation with the malignant progression and metastasis of these tumors. We show here that Prss11 is negatively regulated by HDAC1 and that its overexpression significantly impairs proliferation in human tumor cells. These findings suggest that HDAC1 is one of the relevant target enzymes for HDAC inhibitors as tumor drugs. This idea is also supported by the finding that HDAC1 inactivation induced apoptosis in human tumor cells (Senese et al., submitted). We have recently established an HDAC1 mouse tumor model and will test the function of some of the above-mentioned genes for their relevance as HDAC1-regulated tumor suppressors.
Another interesting finding is the identification of several imprinted genes as HDAC1 targets. It is important to note that the allele-specific silencing of some of these imprinted genes occurs only upon differentiation. However, the observed deregulation of specific imprinted genes is not due to unscheduled ES cell differentiation in the absence of HDAC1, since virtually all wild-type and HDAC1 KO cells are positive for Oct4, a marker for undifferentiated ES cells (R. Brunmeir and C. Seiser, unpublished results). A hallmark of imprinted genes is the presence of differentially methylated CpG islands known as differentially methylated regions. Preliminary data suggest that HDAC1 might be required for the methylation of specific differentially methylated regions (G. Egger, unpublished observations). DNA methylation is closely linked to histone deacetylation, since class I deacetylases, including HDAC1, were found to associate with both methyl-binding proteins and methyltransferases (22, 23, 50, 88). In accordance with these findings, deacetylase inhibitors have been previously shown to affect the expression of several imprinted HDAC1 target genes, including the H19, Igf2, and p57 genes (25, 26). To better understand the role of HDAC1 in the regulation of imprinted genes, it will be necessary to establish a system suitable for the study of allele-specific gene expression in the presence and absence of HDAC1.
Consequences of HDAC1 recruitment. ChIP analysis revealed that HDAC1 is recruited to a specific set of target promoters. Similarly, the homologous yeast deacetylase Rpd3p was found to be associated with a specific class of target genes, while the HATs Gcn5 and Esa1 are generally recruited to the promoters of active protein-coding genes (61). In addition, HDAC1 was recruited to the 5' intragenic regions of all investigated target genes. A detailed ChIP analysis throughout the murine HDAC1 gene, whose expression is under the control of HDAC1, showed a predominant localization of HDAC1 at the 5' region of this gene (65; G. Zupkovitz, unpublished results). These findings corroborate the results from many laboratories showing the recruitment of HDAC1 by specific transcription factors to promoters and enhancers of target genes (46). Noticeably, the loss of HDAC1 led to reduced nucleosome density on most of the HDAC1 target promoters, suggesting a crucial role for the deacetylase for chromatin condensation at these genes. Accordingly, the occupancy of target genes by HDAC1, with the exception of positively regulated target genes (see below), resulted in the reduced acetylation of histones H3 and H4.
Interestingly, the presence of other epigenetic marks, namely, trimethylation at K9 and K27 of histone H3, was reduced at HDAC1 target genes in the absence of HDAC1. Concerning histone modifications on target promoters, these results are in agreement with several reports describing the association of HDAC1 with histone H3 methyltransferases, including the K9 methyltransferase Suv39h1 and the histone H3-modifying polycomb complex PRC2 (14, 77). However, in contrast to a previous report that showed a link between K9 trimethylation and transcriptional elongation (76), we observed reduced K9 trimethylation at intragenic regions at genes that were induced in the absence of HDAC1. This might be explained by the complex cross talk between enzymes that control histone acetylation and methylation. For instance, HDAC1 was shown to interact not only with a K9 methyltransferase but also with the recently identified K9-demethylating enzyme JMJD2A (83). In addition, the recruitment and expression of JMJD2A might be regulated in an HDAC-dependent manner (27). We are currently performing a chromosome-wide ChIP-on-chip analysis for mouse HDAC1 to understand in more detail the consequences of HDAC1 recruitment on histone acetylation and other chromatin modifications.
Functional links between HDAC1 and HDAC2. Expression of the related class I enzyme HDAC2 was reliably upregulated upon inactivation of HDAC1 in all cell systems that we tested, including mouse ES cells, fibroblasts, T cells, and human tumor cells (36; unpublished data; Senese et al., submitted). Interestingly, only 7% of the mouse HDAC1 target genes are found to be deregulated in human tumor cells missing HDAC1 (Senese et al., submitted). However, when HDAC1 and HDAC2 were simultaneously inactivated, more than 20% of the murine HDAC1 targets were differentially expressed in U2OS cells. Increased amounts of HDAC2 might partially compensate for the loss of HDAC1 as a transcriptional repressor. This idea is supported by several findings in the present study. For instance, a group of genes, including the JunB and Apc2 genes, showed a significant sensitivity towards the HDAC inhibitor TSA even in the absence of HDAC1 (Fig. 5), suggesting that they are regulated also by other HDACs. Accordingly, ChIP assays demonstrated increased recruitment of HDAC2 as a consequence of the loss of HDAC1 (Fig. 6). Such genes are most probably regulated by the recruitment of repressor complexes that contain increased amounts of HDAC2 in the absence of HDAC1, which mask the actual repressive capacity of HDAC1 for these target genes. In fact, HDAC2 might act as an impostor (40) by replacing HDAC1 as a component of certain repressor complexes in HDAC1 KO cells. In agreement with this idea, the expression of an enzymatically inactive HDAC1 mutant led to increased expression of JunB and Apc2 in HDAC1-null cells (Fig. 4C). It is likely that for some other genes, HDAC2 can fully compensate for the lost repressor function of HDAC1 in KO cells. These genes were not detected in our screen but might be identified in an HDAC1 ChIP-on-chip approach.
HDAC1 as a positive regulator of transcription. Finally, we have also characterized genes that display reduced expression levels in HDAC1-deficient cells. For one group of genes, the decrease in expression in the absence of HDAC1 was rescued by TSA treatment (Fig. 8A, Efnb2, Ehd1, and Edg1), indicating that these genes are repressed by other deacetylases in HDAC1 KO cells. In accordance with this idea, large amounts of HDAC2 were associated with these genes in the absence of HDAC1 but not in wild-type cells. In parallel with reduced expression levels, the acetylation levels of associated histones were also decreased. Thus, the loss of HDAC1 indirectly leads to the repression of this group of genes due to the enhanced presence of HDAC2. Efnb2, a ligand of the ephrin receptor tyrosine kinase that is required for hippocampal plasticity (30), was found to be a prognostic marker for neuroblastomas (70, 71). Ehd1, a member of the eps15 homology domain-containing family (47), was shown to participate in the endocytosis of the insulin-like growth factor 1 receptor (62), and Edg1 is a G-protein-coupled receptor for sphingosine-1-phosphate (33) that seems to stimulate cell migration and metastasis (84).
The most
surprising finding of this study was the identification of genes that
require HDAC1 directly for their activation. In contrast to all the
other HDAC1 target genes, these genes showed a negative response to TSA
(Fig. 8A, Gja1, and Fig.
9A). HDAC1 was recruited
either constitutively (Gja1) or, in the case of the Irf1 and Gbp2
IFN-responsive genes, in response to the IFN signal, strongly
suggesting that these genes are direct HDAC1 targets. Gja1 is the major
protein of gap junctions in the heart and seems to display differential
responses to deacetylase inhibitors in normal versus transformed cells
(55). The Irf1 gene was
shown to be a tumor susceptibility gene which encodes a protein with
tumor suppressor-like function (reviewed in reference
69). In contrast to those
of the tumor suppressors p21 and Prss11, which are negatively regulated
by HDAC1, Irf1 expression levels seem to be dependent on HDAC1. The
HDAC1 homologue Rpd3p was originally identified as a factor required
for both activation and repression of yeast genes
(82) and plays a positive
role in the activation of osmoresponsive promoters
(17). In mammalian cells,
HDAC1 was recently identified as a coactivator of the glucocorticoid
receptor (60).
Furthermore, histone deacetylase activity was shown to be required for
the proper activation of IFN-responsive genes and HDAC1 was implicated
in this process (reviewed in reference
53). We provide the first
evidence for the regulated HDAC1 recruitment to the promoters of Irf1
and Gbp2 during transcriptional induction by IFN-. The
mechanism of HDAC1-dependent activation of target genes is not entirely
clarified. On the one hand, HDAC1 might be required predominantly for
the deacetylation of specific transcription regulators that trigger
their activation. This idea is supported by the fact that histone
acetylation at the positively regulated Gja1, Irf1, and Gbp2 target
genes is mostly unaffected by the presence of HDAC1 (Fig.
8 and
9). On the other hand, the
stimulation of HDAC1 target genes could be accompanied by waves of
histone acetylation and deacetylation that are controlled by both HATs
and HDACs. Similar cyclic changes in the acetylation of core histones
have been previously observed during the hormone-dependent activation
of genes (45,
60). Taken together, the
results presented in our study demonstrate a major function for HDAC1
as a transcriptional regulator in mouse ES
cells.
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ACKNOWLEDGMENTS |
|---|
This work was supported by the Austrian Science Fund (FWF P16443-B04) and the GEN-AU project Epigenetic Plasticity of the Mammalian Genome (Federal Ministry for Education, Science and Culture). G.Z. was a fellow of the Vienna Biocenter Ph.D. program (Austrian Science Fund).
|
FOOTNOTES |
|---|
Published ahead of print on 28 August 2006.
Present
address: The Wellcome Trust Sanger Institute, Hinxton, Cambridge CB10
1SA, United Kingdom.
Present
address: Wellcome Trust Biocentre, University of Dundee, Dundee DD1 5EH, United Kingdom.
Present address: Max F. Perutz Laboratories, Institute of Microbiology and Genetics, University of Vienna, Vienna Biocenter, A-1030 Vienna, Austria.
¶ Present address: Department of Biochemistry and Molecular Biology, USC/Norris Comprehensive Cancer Center, Keck School of Medicine of the University of Southern California, Los Angeles, CA 90089-9181.
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1 levels and
shown as expression levels in HDAC1-null cells relative to expression
levels in wild-type cells (arbitrarily set to 1). (B) SYBR
green real-time RT-PCR analysis of downregulated genes. The analysis
was performed as described for panel
A.
