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Promoter-specific p53-dependent histone acetylation following DNA damage

Abstract

We have used chromatin immunoprecipitation (ChIP) to measure p53-dependent histone acetylation at the p21, MDM2, and PUMA promoters. The pattern of histone acetylation was different at each promoter. H3 and H4 acetylation increased at both the p21 and PUMA promoters in response to p53 activation, whereas there was only a minimal increase in H4 acetylation and no increase in H3 acetylation at the MDM2 promoter. The high p53 occupancy of the p21, MDM2 and PUMA promoters has been attributed to the presence of two p53 binding sites in these promoters, but mutation of the p53 binding sites in integrated p21 promoter constructs showed that the two sites in the p21 promoter do not cooperate to stabilize p53 binding. Despite 10-fold higher p53 binding to the proximal than the distal site in the p21 promoter, both sites showed similar patterns of H3 and H4 acetylation. Mutation of the binding sites showed that acetylation of the proximal, low-affinity site requires p53 binding to that site but not to the distal, high-affinity site. Since low-affinity p53 binding sites can confer strong acetylation, the DNA binding affinity in vitro is an unreliable guide to the likely importance of p53 in regulating candidate target genes in vivo.

Introduction

The p53 tumour suppressor gene is mutated in over 50% of human tumours and plays an important role in the response to genotoxic stress (Vogelstein et al., 2000). It responds to upstream signals by activating transcription of genes important for cell cycle arrest, DNA repair, and apoptosis. The spectrum of target genes induced depends on the type of stress and the tissue analysed (Zhao et al., 2000). Studies examining the binding of p53 to promoters by chromatin immunoprecipitation (ChIP) have shown that there are large differences in the kinetics and extent of p53 binding to target gene promoters in vivo (Szak et al., 2001; Kaeser and Iggo, 2002). Differences in the affinity of p53 for its binding sites have been invoked to explain why some cells undergo growth arrest and others apoptosis following p53 activation (Vousden and Lu, 2002). High-affinity sites are predominantly found in cell cycle arrest genes, which would allow selective transactivation of these genes when low levels of p53 are present in the cell. In contrast, low-affinity sites are predominantly found in proapoptotic target genes, which thus should require higher levels of p53 for transactivation (Chen et al., 1996). ChIP analysis has demonstrated that the p21, MDM2, and PUMA promoters show higher p53 occupancy in vivo than the AIP1, bax, and PIG3 promoters (Kaeser and Iggo, 2002). This is broadly consistent with the affinity model, except for PUMA, which is a proapoptotic BH3-family protein. As well as the affinity of individual DNA sequences for p53, the number of binding sites in the promoter appears to be an important factor in determining p53 occupancy in vivo. Recent NMR studies suggest that p53 can link noncontiguous DNA binding sites in a sandwich-like manner, resulting in more efficient DNA binding when multiple well-separated binding sites are present, as in the p21, MDM2, and PUMA promoters (Klein et al., 2001). Since p53 is present at the promoter of these genes even in the absence of genotoxic stress (Kaeser and Iggo, 2002), it is likely that additional mechanisms regulate p53 activity in vivo. Many studies have shown that p53 undergoes phosphorylation and acetylation in response to genotoxic stress (reviewed by Anderson and Appella, 2003). One function of these modifications is to regulate p53 stability, for example, through regulation of MDM2 binding (Chehab et al., 1999). Another possibility is that p53 modification controls the access of p53 to basal transcription factors (Thut et al., 1995), mediator (Ito et al., 1999), chromatin remodelling enzymes (Lee et al., 2002), or histone acetyltransferases (HATs). p53 interacts with components of multiple different HAT complexes, including p300/CBP (Dumaz and Meek, 1999), TRRAP (Barlev et al., 2001), GCN5 (Candau et al., 1997), and P/CAF (Sakaguchi et al., 1998), and p53-mediated transactivation of p21 transcription correlates with increased histone acetylation (Barlev et al., 2001; Espinosa and Emerson, 2001; Espinosa et al., 2003). p53 itself is acetylated at the carboxy-terminus by p300/CBP and P/CAF (Sakaguchi et al., 1998). An intriguing model for the function of this region of the p53 protein is that it acts like a histone tail, progressively accumulating modifications that permit the sequential recruitment of chromatin modifying enzymes, for example through binding of acetylated p53 residues to bromodomain proteins (Agalioti et al., 2002).

We have used ChIP to measure the effect of p53 activation on histone acetylation at the p21, MDM2, and PUMA promoters. We show that p53 activation has different effects at each promoter tested.

Results

Endogenous p53 induces promoter-dependent H3 and H4 acetylation

To test the kinetics and p53-dependence of histone acetylation at the p21, MDM2, and PUMA promoters, HCT116 colon cancer cells and p53 knockout derivatives were treated with 5-fluorouracil (5FU), and ChIP was performed with anti-p53 and antiacetylated histone antibodies. Western blotting showed that the p53, MDM2, and p21 levels all rose in the parental HCT116 cells after treatment (Figure 1a). Consistent with previous observations (Kaeser and Iggo, 2002), p53 was already present at promoters in unstressed cells, resulting in several hundred-fold higher recovery of p53 target gene promoter DNA in the cells containing p53 (Figure 1b, left panels). The maximal recovery of input chromatin by anti-p53 antibodies in 5FU-treated cells was 5, 3, and 1.8% for the p21, MDM2, and PUMA promoters, respectively, confirming that these promoters show high p53 occupancy after genotoxic stress. The amount of histone acetylation was measured by ChIP using antibodies recognizing multiple acetylated forms of H3 or H4. In the absence of 5FU, the level of H4 acetylation was two- to threefold higher at the p21 and PUMA promoters in the parental than the p53 knockout cells (Figure 1b, middle panels). After 5FU treatment, there was no change in H4 acetylation in the p53 knockout cells but a two- to threefold increase in the parental cells. H4 acetylation remained high 12 h after treatment at the p21 promoter, but peaked at 3 h and then decreased with time on the PUMA promoter. H3 acetylation showed a similar pattern to H4 acetylation on both promoters, though with a higher p53-independent basal level and a smaller increase following 5FU treatment. In contrast to the p21 and PUMA promoters, the MDM2 promoter showed little, if any, induction of acetylation by 5FU, and H3 acetylation was higher at all time points in the p53-deficient cells (Figure 1b, bottom panels). These results indicate that the pattern of acetylation is specific to each promoter.

Figure 1
figure1

p53 binding and histone acetylation at the p21, MDM2, and PUMA promoters. HCT116 colon cancer cells were treated with 375 μ M 5FU to activate p53. (a) Western blot for p53, MDM2, and p21. The loading control is an invariant Ponceau S stained band. (b) ChIP for p53, H3, and H4 acetylation at the indicated times after 5FU treatment. wt, parental HCT116 cells containing wild-type p53; ko, p53 knockout HCT116 cells. The antihistone antibodies recognize multiple acetylated forms of H3 and H4

p53 binds preferentially to the distal response element in the p21 promoter

The p21 promoter was studied in detail because it showed the greatest p53-dependent histone acetylation. It contains proximal and distal p53 response elements, located 1.4 and 2.3 kb upstream of the transcription initiation site, respectively. Both are located in regions conserved between human, mouse, and rat (el-Deiry et al., 1995). The distal p53 binding site is a better match to the consensus p53 binding sequence (el-Deiry et al., 1992). In addition to the p53 binding sites, the promoter contains response elements for several other transcription factors, including the vitamin D3 receptor, Sp1, myoD, and MIZ-1 (Halevy et al., 1995; Biggs et al., 1996; Liu et al., 1996; van de Wetering et al., 2002). Six equally spaced amplicons spanning 3 kb of the promoter and 0.6 kb of the transcribed region were used to map p53 binding in vivo (Figure 2). p53 with an S to F mutation at codon 121 has previously been shown to have an altered DNA binding sequence specificity, most likely through changes in hydrogen bonding of K120 to DNA (Freeman et al., 1994). Compared to wild-type p53, 121F has a sevenfold lower affinity for the distal p21 site, but a sixfold higher affinity for the sequence GGG CATG CCC, which resembles the proximal p53 binding site in the p21 promoter (Freeman et al., 1994). Wild-type p53, the 121F mutant, and the non-DNA binding 175H mutant were expressed from a tetracycline-regulated promoter in p53-null H1299 cells (Saller et al., 1999). The p53 protein level was the same for all three clones after induction (data not shown, Kaeser and Iggo, 2002). ChIP for wild-type p53 showed a twofold higher occupancy of the distal than the proximal site (Figure 2). In contrast, 121F bound preferentially to the proximal site, indicating that the ChIP assay can resolve differences in the location of p53 within the p21 promoter. It also shows that the ChIP assay measures direct p53 binding to p21 promoter DNA and confirms that differences in affinity measured in vitro have corresponding effects on DNA binding in vivo.

Figure 2
figure2

Sites of p53 binding within the p21 promoter. Wt p53 and p53 mutants were expressed from the tet-off promoter in p53-null H1299 cells (Saller et al., 1999). p53 expression was induced by removal of tetracycline from the medium. The negative control cells contain an integrated empty vector (no p53) or express the tumour-derived mutant 175H from the tet-off promoter. p53 binding was measured by ChIP using amplicons located from −3.1 to +0.6 kb relative to the transcription start site. The amplicons labelled upstream, VitD, TATA, and intron do not contain known p53 binding sites. Wt p53 preferentially binds to the distal p53 binding site; the sequence specificity mutant 121F preferentially binds to the proximal site

Damage-induced H3 and H4 acetylation of the p21 promoter

5FU-inducible p53-dependent histone acetylation at the p21 promoter was tested in HCT116 cells (Figure 3). H4 acetylation increased threefold 9 h after 5FU treatment. H4 acetylation of both p53 binding sites was p53-dependent (Figure 3b, compare wt and ko lanes). Despite a ninefold difference in the amount of p53 at the proximal and distal binding sites after 5FU treatment (Figure 3a), the level of H4 acetylation was the same. It was centred on the p53 binding sites and did not extend to the TATA box. H3 acetylation showed a similar but weaker pattern over the p53 binding sites, but was strongest in the transcribed region, where it was independent of p53 status or the level of p21 expression (Figure 3c).

Figure 3
figure3

p53 binding and histone acetylation at the p21 promoter after 5FU treatment. ChIP from HCT116 wt and p53 knockout cells was used to map p53 binding and histone acetylation at the sites shown in Figure 2. Cells were treated with 375 μ M 5FU for 0, 3, 6, and 9 h before crosslinking. (a) ChIP for p53 binding. (b) ChIP for H4 acetylation. (c) ChIP for H3 acetylation. H4 acetylation is p53-dependent and centred on the p53 binding sites; H3 acetylation is weaker and is seen in the transcribed DNA (intron) even in the absence of p53

The lack of difference in the level of acetylation at the two p53 binding sites despite the large difference in the amount of bound p53 was unexpected. To determine whether the high relative acetylation of the proximal site was dependent on p53 binding to the distal site, the sites were mutated in a 2.8 kb p21 promoter fragment and the DNA was stably integrated into the genome of mouse F9 teratocarcinoma cells using a lentiviral vector (Figure 4a). The mutations disrupt the invariant C–G pairs of the p53 binding site (Figure 4b). F9 cells contain wild-type p53 and respond to DNA damage by stabilizing p53 (data not shown, Lutzker and Levine, 1996). The cells were treated with doxorubicin, and ChIP was performed with human-specific primers to avoid amplifying endogenous mouse p21 promoter DNA. Despite the artificial nature of the construct, p53 binding was strictly dependent on the presence of the previously defined binding sites, showed the same pattern of binding as on the p21 promoter in human cells and increased appropriately following DNA damage (Figure 4c). The reduction following mutation of either site was 30- to 60-fold. Mutation of the proximal site had no effect on binding to the distal site, and mutation of the distal site had no effect on binding to the proximal site, indicating that the two sites do not cooperate to stabilize p53 binding.

Figure 4
figure4

Mutation of p53 binding sites in the p21 promoter. A lentiviral vector containing a fragment of the human p21 promoter was integrated into mouse F9 cells, which contain wild-type p53. (a) Schematic diagram showing the structure of the lentiviral vector. The construct contains the proximal, distal, and VitD amplicons shown in Figure 3. (b) Residues mutated to abolish p53 binding to the proximal and distal p53 response elements. Residues matching the p53 consensus are in capitals, with the invariant C/G basepairs in bold. (ce) ChIP from transduced F9 cells treated for 9 h with 200 ng/ml doxorubicin. The PCR primers are human-specific and do not amplify endogenous murine DNA. (c) ChIP for p53 binding. (d) ChIP for H4 acetylation. (e) ChIP for H3 acetylation

Histone acetylation of the integrated DNA could be influenced by the presence of other control elements in the construct. The LTRs in the integrated DNA lack enhancer elements because of a deletion in the 3′-LTR in the vector (Zufferey et al., 1998), but the SV40 promoter, which is used to express the pac gene for puromycin selection, could recruit factors that acetylate the p21 promoter DNA. Despite these potential limitations, the pattern of histone acetylation was remarkably similar to that of the endogenous p21 promoter in human cells: H4 acetylation was stronger than H3 acetylation, and each histone showed similar acetylation at the two p53 binding sites (Figure 4d). The level of acetylation increased following doxorubicin treatment, and this increase was p53 dependent because it did not occur in similarly treated p53 null MEFs harbouring the same construct (data not shown). Mutation of the distal p53 binding site, which abolished p53 binding, had a small effect on H4 acetylation of this site and no effect on H4 acetylation elsewhere in the promoter. Mutation of the proximal p53 binding site had no effect on H4 acetylation at the distal site but reduced acetylation of the proximal site and downstream VitD site by >50%. H3 acetylation on all three sites increased several fold after doxorubicin treatment, but the changes following mutation of the p53 binding sites were small and may not be significant, possibly because the lower level of acetylation reduced the signal-to-noise ratio (Figure 4e).

In summary, we can see no evidence of cooperativity between the proximal and distal sites to promote p53 binding to the p21 promoter, H4 acetylation is more responsive to p53 activation than H3 acetylation of this promoter, and acetylation of the proximal p53 binding site is disproportionate given the amount of p53 bound to this site.

Amino-terminal phosphorylation is not required for histone acetylation

The p53 amino-terminus is phosphorylated on residues 6, 9, 15, 18, 20, 33, 37, and 46 in response to stress (reviewed by Anderson and Appella, 2003). This phosphorylation is postulated to have multiple effects, including stabilization of p53 protein and recruitment of basal transcription factors. To test the importance of phosphorylation of residues 6, 9, 15, 20, 33, and 37 for promoter acetylation, we mutated these residues to alanine and transduced p53 knockout HCT116 cells with the p53 mutant (‘N6A’) using a lentivirus. The vector used allows expression of p53 at physiological levels that respond appropriately to DNA damage signals by increasing in level and activating transcription of p21 and MDM2 (Figure 5a and Kaeser et al., 2004). ChIP was performed on 5FU-treated cells infected with viruses expressing wild-type p53, N6A, and empty vector. The pattern of H3 and H4 acetylation in the infected cells (Figure 5b, c) was identical to that in cells expressing wild-type p53 from the endogenous gene (Figure 3). Mutation of p53 phosphorylation sites had no effect on the pattern of either H3 or H4 acetylation. We conclude that p53 phosphorylation on residues 6, 9, 15, 20, 33, and 37 is not required for p53-dependent acetylation of H3 and H4 in the p21 promoter.

Figure 5
figure5

Mutation of p53 phosphorylation sites. Lentiviral vectors were used to express wild-type p53 and phosphomutant p53 in p53 knockout HCT116 cells. The N6A mutant contains alanine substitutions of six phosphorylation sites at the p53 N-terminus. ChIP was performed from cells treated with 375 μ M 5FU for 9 h. (a) Western blot for p53, p21, and MDM2. (b) ChIP for H4 acetylation. (c) ChIP for H3 acetylation

Discussion

The major conclusions from this study are that the pattern of p53-dependent histone acetylation depends on the promoter studied, and that the magnitude of the acetylation response depends both on the amount of p53 bound and the identity of the site. All three promoters tested showed greater p53-dependent changes in H4 than H3 acetylation. Several HATs previously implicated in p53 function can acetylate both H3 and H4, including p300/CBP, GCN5/PCAF, TAFII250, and TIP60 (reviewed by Sterner and Berger, 2000). The most selective for H4 is TIP60 (Ikura et al., 2000), which can be recruited to the p53 transactivation domain by TRRAP (Barlev et al., 2001; Ard et al., 2002). In the conditions of our quantitative ChIP experiments, we were unable to precipitate credible amounts of promoter DNA using antibodies against HATs, so the identity of the complexes responsible for the observed histone acetylation could not be defined.

The MDM2 P2 promoter showed the least change in acetylation after p53 activation, despite a large increase in the amount of p53 bound. This indicates that p53 can regulate transcription by mechanisms that do not involve histone acetylation. For example, it could modify promoter structure by recruitment of chromatin remodelling complexes (Lee et al., 2002), or communicate directly with basal factors to cause release of RNA polymerase from preinitiation complexes (Espinosa et al., 2003). The lack of effect of p53 on H3 acetylation at the MDM2 P2 promoter may be a consequence of pre-existing high basal H3 acetylation, caused by the passage of RNA polymerase-elongator complexes initiating at the constitutive P1 promoter (Winkler et al., 2002).

Relative to the distal p53 binding site in the p21 promoter, acetylation of the proximal site was disproportionate to the amount of p53 bound. This could be related to a fundamental difference in the way p53 binds to the two sites, or depend on the presence of other proteins binding to nearby sequences. The possibility that affinity dictates the choice between cell cycle arrest and apoptosis, with high-affinity sites regulating cell cycle arrest genes and low-affinity sites regulating proapoptotic genes, has been widely discussed (Rowan et al., 1996; Vousden and Lu, 2002). The p21 promoter can be used to address these issues because it contains one high- and one low-affinity site. Our ChIP data support the idea that the two sites have a different affinity for p53 in vivo. What we did not expect to see was the high level of acetylation at the proximal site despite the presence of 10-fold less p53 at this site. The acetylation appears to be due to p53 binding to the proximal site directly, rather than spreading of acetylation from the distal site, because mutation of the distal site did not reduce it. A possible model to explain this observation is that rapid turnover of p53 on DNA is a better mechanism to load coactivators than stable binding of p53 to DNA. An alternative explanation could be that the structure of the bound p53 is different when there are only two subunits in the p53 tetramer bound tightly to the DNA, allowing different p53 surfaces to be exposed for interaction with coactivators. Whatever the explanation, a practical consequence is that measurement of p53-dependent histone acetylation may provide useful insight into the biological relevance of p53 to transactivation of a putative target gene, even if the known p53 binding site is a very poor match to the consensus.

It has been reported previously that widely spaced p53 binding sites in promoters allow the formation of loops bridged by p53 tetramers, resulting in synergistic activation of transcription (Stenger et al., 1994; Jackson et al., 1998; Klein et al., 2001). Mutation of either the proximal or the distal p53 binding site in the p21 promoter in an integrated lentiviral vector showed that p53 binds independently to the two sites. Use of a lentivirus to integrate promoter DNA for ChIP studies has not been described previously. Given the artificial nature of the construct it is remarkable that both p53 binding and histone acetylation were still responsive to stress.

It is widely assumed that p53 phosphorylation selectively regulates the recruitment of coactivator complexes to promoters. The differences in the pattern of histone acetylation at the p21, PUMA, and MDM2 promoters in our study, together with many reports in the literature describing interactions between p53 and HAT-containing complexes, suggest a simple mechanism for p53 to regulate promoters selectively in response to diverse signals. Our failure to see differences in histone acetylation at the p21 promoter following reintroduction of p53 with alanine at six well-characterized amino-terminal phosphorylation sites cautions that proving this model in vivo will be a very difficult task. There is no doubt that p53 is phosphorylated in response to stress signals (reviewed by Anderson and Appella, 2003), but until there is a simple experimental model that shows strong phenotypes for p53 phosphorylation site mutants, it will be difficult to prove that phosphorylation of particular residues regulates the recruitment of specific coactivators to individual promoters.

Materials and methods

Cell lines and antibodies

HCT116 human colorectal cancer and p53 knockout derivatives were supplied by Dr B Vogelstein (Bunz et al., 1998). H1299 cells expressing p53 from a tetracycline-regulated promoter were described by Saller et al. (1999). p53 expression was induced as described by Saller et al. (1999). Murine teratocarcinoma F9 cells were obtained from ATCC and cultured on gelatin-coated dishes. DO7 and PAb248 (Yewdell et al., 1986) murine hybridoma cells were supplied by Dr D Lane. SMP-14, C-19, and FL-393 were purchased from Santa Cruz Biotechnology. Antiacetylated H3 (#06-599) and H4 (#06-866) were purchased from Upstate Biotechnology. Western blotting was performed as previously described (Kaeser and Iggo, 2002).

Plasmids and viruses

pMDK36 containing a 1 kb fragment of the distal p21 promoter was constructed by BP-cloning (Invitrogen) a PCR fragment from human placental DNA into pSP161 as described by Bridge et al. (2003). The fragment was amplified using primers oMDK7 (IndexTermGGGGACAAGTTTGTACAAAAAAGCAGGCTctgaggcagaattgcttgaa) and oMDK8 (IndexTermGGGGACCACTTTGTACAAGAAAGCTGGGTgcatttcaaatagactagac). To obtain 2.8 kb of the p21 promoter, a BamHI–SacI fragment of pMDK36 and a SacI–HinDIII fragment of WW-pLUC (el-Deiry et al., 1993) were cloned into BamHI–HinDIII of pUC19. Site-directed mutagenesis of the distal site was performed by iPCR using oMDK15 (IndexTermATcTCCCAAgATtTTGagctctggcatagaagag) and oMDK16 (IndexTermcTTCctgacggccagaaagcc), or oMDK17 (IndexTermaGtaATtTCTgggcagagatttccagac) and oMDK18 (IndexTermAGTaTTCTTCctctaacgcagctga) for the proximal site. Fragments of wild type or mutated p21 promoter were amplified using oMDK7 and oMDK19 (IndexTermGGGGACCACTTTGTACAAGAAAGCTGGGTaagcttctgacttcggcag) and BP-cloned into pSP161. Lentiviruses were produced as described previously (Bridge et al., 2003). The p53-expressing lentiviruses were described by Kaeser et al. (2004). F9 cells were transduced at low multiplicity (approx. 20% efficiency) and selected in puromycin.

Chromatin immunoprecipitation

ChIP with DO7, antiacetylated H3 and antiacetylated H4 antibodies was performed as described by Kaeser and Iggo (2002), except that for Figure 3, DNA was sonicated to an average length of 300 bp. For Figure 4, DNA was sonicated to an average length of 500 bp and 1 μg PAb248 was used to precipitate murine p53. SYBR real-time quantitative PCR was performed on a PE5700, using the primers described below at 500 nM each and 60°C, except 5′up.fw and 5′up.rv, which were used at 62°C. Integrated provirus mutated at the proximal site was measured using prox(MUT).fw and prox.rv. Primer sets and probes for MDM2, p21 (distal) and PUMA were described by Kaeser and Iggo (2002). Error bars indicate standard deviation of triplicate measurements.

5′up.fw:

IndexTermCGGCTGATTTTTGTATTTTTAATG

5′up.rv:

IndexTermTCACAGGGTCAGGAGTTTTGAGA

prox.fw:

IndexTermGAGGAAGAAGACTGGGCATGTCT

prox.rv:

IndexTermGCTTGGAGCAGCTACAATTACTGAC

VitD.fw:

IndexTermGGTTGCCCTTTTTTGGTAGTCTC

VitD.rv:

IndexTermTGAGGAAATTGAGGTCCACTGA

TATA.fw:

IndexTermGGGCGGTTGTATATCAGGGC

TATA.rv:

IndexTermCGGCTCCACAAGGAACTGACTT

int.fw:

IndexTermAGTGGAGTAAGTTCGTCTAGGATCG

int.rv:

IndexTermTGGCGTAAAGGACCTGAACC

prox(MUT).fw:

IndexTermTCAGCTGCGTTAGAGGAAGAATACTA

References

  1. Agalioti T, Chen G and Thanos D . (2002). Cell, 111, 381–392.

  2. Anderson CW and Appella E . (2003). Handbook of Cell Signaling, Bradshaw RA, Dennis E (eds). Vol. 3. Academic Press, pp 237–247.

    Book  Google Scholar 

  3. Ard PG, Chatterjee C, Kunjibettu S, Adside LR, Gralinski LE and McMahon SB . (2002). Mol. Cell Biol., 22, 5650–5661.

  4. Barlev NA, Liu L, Chehab NH, Mansfield K, Harris KG, Halazonetis TD and Berger SL . (2001). Mol. Cell, 8, 1243–1254.

  5. Biggs JR, Kudlow JE and Kraft AS . (1996). J. Biol. Chem., 271, 901–906.

  6. Bridge AJ, Pebernard S, Ducraux A, Nicoulaz AL and Iggo R . (2003). Nat. Genet., 34, 263–264.

  7. Bunz F, Dutriaux A, Lengauer C, Waldman T, Zhou S, Brown JP, Sedivy JM, Kinzler KW and Vogelstein B . (1998). Science, 282, 1497–1501.

  8. Candau R, Scolnick DM, Darpino P, Ying CY, Halazonetis TD and Berger SL . (1997). Oncogene, 15, 807–816.

  9. Chehab NH, Malikzay A, Stavridi ES and Halazonetis TD . (1999). Proc. Natl. Acad. Sci. USA, 96, 13777–13782.

  10. Chen X, Ko LJ, Jayaraman L and Prives C . (1996). Genes Dev., 10, 2438–2451.

  11. Dumaz N and Meek DW . (1999). EMBO J., 18, 7002–7010.

  12. el-Deiry WS, Kern SE, Pietenpol JA, Kinzler KW and Vogelstein B . (1992). Nat. Genet., 1, 45–49.

  13. el-Deiry WS, Tokino T, Velculescu VE, Levy DB, Parsons R, Trent JM, Lin D, Mercer WE, Kinzler KW and Vogelstein B . (1993). Cell, 75, 817–825.

  14. el-Deiry WS, Tokino T, Waldman T, Oliner JD, Velculescu VE, Burrell M, Hill DE, Healy E, Rees JL and Hamilton SR . (1995). Cancer Res., 55, 2910–2919.

  15. Espinosa JM and Emerson BM . (2001). Mol. Cell, 8, 57–69.

  16. Espinosa JM, Verdun RE and Emerson BM . (2003). Mol. Cell, 12, 1015–1027.

  17. Freeman J, Schmidt S, Scharer E and Iggo R . (1994). EMBO J., 13, 5393–5400.

  18. Halevy O, Novitch BG, Spicer DB, Skapek SX, Rhee J, Hannon GJ, Beach D and Lassar AB . (1995). Science, 267, 1018–1021.

  19. Ikura T, Ogryzko VV, Grigoriev M, Groisman R, Wang J, Horikoshi M, Scully R, Qin J and Nakatani Y . (2000). Cell, 102, 463–473.

  20. Ito M, Yuan CX, Malik S, Gu W, Fondell JD, Yamamura S, Fu ZY, Zhang X, Qin J and Roeder RG . (1999). Mol. Cell, 3, 361–370.

  21. Jackson P, Mastrangelo I, Reed M, Tegtmeyer P, Yardley G and Barrett J . (1998). Oncogene, 16, 283–292.

  22. Kaeser MD and Iggo RD . (2002). Proc. Natl. Acad. Sci. USA, 99, 95–100.

  23. Kaeser MD, Pebernard S and Iggo RD . (2004). J. Biol. Chem., 279, 7598–7605.

  24. Klein C, Planker E, Diercks T, Kessler H, Kunkele KP, Lang K, Hansen S and Schwaiger M . (2001). J. Biol. Chem., 276, 49020–49027.

  25. Lee D, Kim JW, Seo T, Hwang SG, Choi EJ and Choe J . (2002). J. Biol. Chem., 277, 22330–22337.

  26. Liu M, Lee MH, Cohen M, Bommakanti M and Freedman LP . (1996). Genes Dev., 10, 142–153.

  27. Lutzker SG and Levine AJ . (1996). Nat. Med., 2, 804–810.

  28. Rowan S, Ludwig RL, Haupt Y, Bates S, Lu X, Oren M and Vousden KH . (1996). EMBO J., 15, 827–838.

  29. Sakaguchi K, Herrera JE, Saito S, Miki T, Bustin M, Vassilev A, Anderson CW and Appella E . (1998). Genes Dev., 12, 2831–2841.

  30. Saller E, Tom E, Brunori M, Otter M, Estreicher A, Mack DH and Iggo R . (1999). EMBO J., 18, 4424–4437.

  31. Stenger JE, Tegtmeyer P, Mayr GA, Reed M, Wang Y, Wang P, Hough PV and Mastrangelo IA . (1994). EMBO J., 13, 6011–6020.

  32. Sterner DE and Berger SL . (2000). Microbiol. Mol. Biol. Rev., 64, 435–459.

  33. Szak ST, Mays D and Pietenpol JA . (2001). Mol. Cell Biol., 21, 3375–3386.

  34. Thut CJ, Chen JL, Klemm R and Tjian R . (1995). Science, 267, 100–104.

  35. van de Wetering M, Sancho E, Verweij C, de Lau W, Oving I, Hurlstone A, van der Horn K, Batlle E, Coudreuse D, Haramis AP, Tjon-Pon-Fong M, Moerer P, van den Born M, Soete G, Pals S, Eilers M, Medema R and Clevers H . (2002). Cell, 111, 241–250.

  36. Vogelstein B, Lane D and Levine AJ . (2000). Nature, 408, 307–310.

  37. Vousden KH and Lu X . (2002). Nat. Rev. Cancer, 2, 594–604.

  38. Winkler GS, Kristjuhan A, Erdjument-Bromage H, Tempst P and Svejstrup JQ . (2002). Proc. Natl. Acad. Sci. USA, 99, 3517–3522.

  39. Yewdell JW, Gannon JV and Lane DP . (1986). J. Virol., 59, 444–452.

  40. Zhao R, Gish K, Murphy M, Yin Y, Notterman D, Hoffman WH, Tom E, Mack DH and Levine AJ . (2000). Genes Dev., 14, 981–993.

  41. Zufferey R, Dull T, Mandel RJ, Bukovsky A, Quiroz D, Naldini L and Trono D . (1998). J. Virol., 72, 9873–9880.

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Acknowledgements

We thank Drs B Vogelstein, D Lane, D Trono, C Prives, and E Saller for supplying cell lines, antibodies and plasmids. We thank Dr V Simanis for critical reading of the manuscript. We thank the Swiss National Science Foundation for financial support.

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Correspondence to Richard D Iggo.

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Kaeser, M., Iggo, R. Promoter-specific p53-dependent histone acetylation following DNA damage. Oncogene 23, 4007–4013 (2004). https://doi.org/10.1038/sj.onc.1207536

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Keywords

  • chromatin immunoprecipitation
  • p53
  • p21
  • MDM2
  • PUMA
  • histone acetyltransferase

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