The DNA‐damage‐induced transcriptional suppression of cell cycle regulatory genes correlates with a reduction in histone H3‐Thr 11 phosphorylation (H3‐pThr 11) on their promoters that is partly mediated by the dissociation of Chk1 from chromatin. In this study, we identify protein phosphatase 1γ (PP1γ) as a phosphatase responsible for DNA‐damage‐induced H3‐pThr 11 dephosphorylation. PP1γ is activated after DNA damage, which is mainly mediated by a reduction in Cdk‐dependent phosphorylation of PP1γ at Thr 311. The depletion of PP1γ sensitizes HCT116 cells to DNA damage. Our results suggest that the ataxia telangiectasia, mutated and Rad3‐related–Chk1 axis regulates H3‐pThr 11 dephosphorylation on DNA damage, at least in part by the activation of PP1γ through Chk1‐dependent inhibition of Cdks.
Eukaryotic cells have coordinated systems to respond to DNA damage, such as cell cycle arrest mechanisms, DNA repair pathways and the apoptotic response. Together, these maintain genomic integrity (Sancar et al, 2004). These systems are partly regulated by transcriptional activation and repression (Zhan et al, 1993; Engelberg et al, 1994). We and others have recently identified histone H3‐Thr 11 phosphorylation (H3‐pThr 11) as a new transcriptional marker (Metzger et al, 2008; Shimada & Nakanishi, 2008; Shimada et al, 2008) that rapidly decreases after DNA damage. Under normal conditions Chk1 associates with chromatin (Smits et al, 2006; Niida et al, 2007) and phosphorylates H3‐Thr 11. In response to DNA damage, Chk1 is rapidly released from chromatin and H3‐pThr 11 is reduced.
Mammalian serine (Ser)/threonine (Thr)‐specific protein phosphatases (PPs) are represented by eight distinct prototypes: PP1, PP2A, PP2B, PP2C, PP4, PP5, PP6 and PP7 (Moorhead et al, 2007; Swingle et al, 2007). Of these, PP1, PP2A and PP4 have all been identified as histone phosphatases: PP1 dephosphorylates H1, which is phosphorylated in a cell‐cycle‐dependent manner (Paulson et al, 1996). The PP1 homologue dephosphorylates H3‐pSer 10 at mitotic exit in yeast and worms (Hsu et al, 2000), and PP2A contributes to its dephosphorylation in Drosophila (Nowak et al, 2003). Phospho‐H2AX (γ‐H2AX) is immediately dephosphorylated after DNA repair by PP2A and PP4 in mammals and yeasts (Chowdhury et al, 2005, 2008; Keogh et al, 2006; Nakada et al, 2008). However, the phosphatases that catalyse H3‐pThr 11 dephosphorylation in response to DNA damage have yet to be identified. In this study, we identify PP1γ as a phosphatase responsible for DNA‐damage‐induced dephosphorylation of H3‐pThr 11.
Results And Discussion
Dephosphorylation of pThr 11 is okadaic acid sensitive
To identify phosphatases that might be capable of DNA‐damage‐induced dephosphorylation of H3‐pThr 11, we treated HeLa cells with different concentrations of okadaic acid (OA) and examined the status of H3‐pThr 11 after ultraviolet (UV) irradiation. H3‐pThr 11 was reduced after UV irradiation as previously reported (Shimada et al, 2008). Treatment with ⩾25 nM OA resulted in an increase in H3‐pThr 11 in the absence of UV, making it difficult to evaluate the effect of OA treatment on DNA‐damage‐induced dephosphorylation of H3‐pThr 11 (Fig 1A). We have previously reported that H3‐pThr 11 is strongly enhanced at mitosis (Shimada et al, 2008). To establish whether the induction of H3‐pThr 11 on OA treatment simply reflected an increase in the number of mitotic cells, we stained OA‐treated cells with 4’,6‐diamidino‐2‐phenylindole (DAPI) and calculated the mitotic index (Fig 1B). The mitotic index value increased more than threefold after OA treatment. However, in interphase cells, H3‐pThr 11 is detected throughout the nuclear region with some exclusion of the peri‐centromeric heterochromatin foci (represented by DAPI foci in Fig 1C). By contrast, at mitosis, strong signals for H3‐pThr 11 are detected in condensed chromosomes, as reported previously (Preuss et al, 2003).
To evaluate the precise effect of OA treatment on H3‐pThr 11 independent of mitotic entry, we concomitantly treated cells with RO3306. Treatment with 9 μM RO3306 preferentially inhibits Cdk1 and thus prevents mitotic entry (Vassilev et al, 2006). The OA treatment did not result in an increase in the number of mitotic cells in the presence of 9 μM RO3306 (Fig 1D). Cell cycle arrest at G2 phase was induced as expected (supplementary Fig S1 online). DNA‐damage‐induced H3‐pThr 11 dephosphorylation was obvious at <100 nM OA, but almost completely abrogated at ≧100 nM OA in the presence of RO3306 when bands responsible for H3‐pThr 11 were quantified (Fig 1E). We observed that dephosphorylation of H3‐pThr 11 was not reduced after DNA damage, even at concentrations of 500 nM fostriecin—a specific inhibitor of PP2A—and its closely related PP4 and PP6 isotypes (Walsh et al, 1997; Fig 1F). At 100 nM, cyclin B1 had accumulated, demonstrating that inhibition was effective (Cheng et al, 1998). These results suggest that PP2A and its related phosphatases are not involved in DNA‐damage‐induced dephosphorylation of H3‐pThr 11.
PP1γ is a H3‐pThr 11 phosphatase
The OA sensitivity and fostriecin insensitivity in DNA‐damage‐induced dephosphorylation of H3‐pThr 11 suggested that the corresponding phosphatase might be a PP1 family member. We depleted PP1 by the using small interfering RNA (siRNA) that could suppress the expression of all three isoforms (PP1α, PP1β/δ and PP1γ) equally. Cells were treated with control, PP1 or PP2A siRNAs for 36 h in the presence of 0.5 mM hydroxyurea, which prevents mitotic arrest by PP1 or PP2A depletion. After 36 h, cells were released from hydroxyurea arrest and incubated with RO3306 for 16 h. Finally, cells were irradiated with UV and sampled 2 h later (Fig 2A, upper panel). The depletion of PP1, but not of PP2A, resulted in compromised H3‐pThr 11 dephosphorylation after UV (Fig 2A).
We further demonstrated that PP1γ depletion specifically compromised DNA‐damage‐induced H3‐pThr 11 dephosphorylation in both HCT116 (Fig 2B) and HeLa (supplementary Fig S2A,B online) cells. Depletion of individual PP1 subunits did not result in gross changes in cell cycle distribution, although PP1γ depletion mitotic index resulted in a slight increase in the mitotic index value (supplementary Fig S3 online). It is interesting to note that PP1γ phosphatase activity towards H3‐pThr 11 was activated as early as 1 h and maintained for up to 4 h after DNA damage (Fig 2C). Changes in the expression of cell cycle regulatory genes after PP1γ depletion were examined in cells treated with RO3306 to prevent low‐level induction of mitotic cells. Treatment with RO3306 did not affect the expression of these genes (supplementary Table S1 online). The depletion of PP1γ suppressed DNA‐damage‐induced transcriptional repression of cdk1, cyclin B1 and cyclin A2 (Fig 2D). Importantly, PP1γ depletion sensitized HCT 116 cells to UV when cell survival was evaluated by MTT (3‐[4,5‐dimethylthiazol‐2‐y1]‐2,5‐diphenyltetrazolium bromide) assay (Fig 2E), suggesting the physiological importance of PP1γ‐dependent transcriptional repression after DNA damage. Although the molecular mechanism by which PP1γ regulates H3‐pThr 11 dephosphorylation after DNA damage has remained elusive, isoform‐specific functions for PP1 have been reported in the regulation of SRp38 dephosphorylation (Shi & Manley, 2007) and in the control of chromosomal architecture (Trinkle‐Mulcahy et al, 2006).
Regulation of PP1γ activity in response to DNA damage
The activity of PP1α is reported to be downregulated by Cdk‐dependent Thr 320 phosphorylation (Dohadwala et al, 1994). The equivalent threonine residue is conserved in all three PP1 isoforms (Thr 316 in PP1β/δ and Thr 311 in PP1γ). Indeed, cyclin B1–Cdk1‐phosphorylated PP1γ‐Thr 311 in vitro (supplementary Fig S4A online). Cdk activity is strongly inhibited after DNA damage, in a manner dependent on Chk1 (Harper & Elledge, 2007). We therefore examined changes in the phosphorylation status of PP1 subunits after UV irradiation by using a combination of isoform‐specific antibodies and a phospho‐specific Thr 320 (PP1α) antibody (Fig 3A). All PP1 isoforms were similarly localized in both chromatin and soluble fractions of HCT116 cells. As a control, IkB kinase α (IKKα)—a typical cytosolic protein—and lamin B1—a marker of the nuclear soluble fraction—were predominantly detected in the soluble fraction, indicating that the subcellular fractionation was successful. The PP1‐pThr 320‐specific signal for each isoform was rapidly dephosphorylated in response to UV, particularly in the chromatin fraction, with kinetics that were similar to that of H3‐pThr 11. As the PP1α‐pThr 320‐specific antibody can recognize the corresponding phosphorylations of PP1β/δ and PP1γ, we further examined whether PP1γ‐pThr 311 was dephosphorylated after UV treatment. We observed that PP1γ‐pThr 311 in PP1γ immune complexes was reduced (Fig 3B). We separated the three isoforms of PP1 using sodium dodecyl sulphate–polyacrylamide gel electrophoresis containing Phos Tag (Kinoshita et al, 2009). Treatment of HCT116 cells with each isoform‐specific siRNA or their combinations revealed that PP1γ migrated fastest, followed by PP1α and PP1β (Fig 3C), although the amounts of each isoform and their phosphorylations at Thr 311 were induced after depletion of the other isoforms, presumably due to a compensatory mechanism. The PP1γ‐pThr 311‐specific signal was consistently decreased in response to UV irradiation when bands responsible for PP1γ‐pThr 311 were quantified. Similar kinetics were observed when cells were treated with bleomycin (supplementary Fig S4B online) or ionizing radiation (supplementary Fig S4C online), and the extent of PP1γ‐pThr 311 dephosphorylation was found to be dependent on the UV dose (supplementary Fig S4D online). Importantly, the phosphorylation‐defective mutant of PP1γ (T311A) was more active in vitro than in the wild‐type PP1γ (Fig 3D).
ATR–Chk1 axis regulates PP1γ activity
Treatment with caffeine—an inhibitor of the ataxia telangiectasia, mutated and Rad3‐related (ATR) and ataxia telangiectasia, mutated kinases—partly suppressed DNA‐damage‐dependent dephosphorylation of PP1γ‐pThr 311 (Fig 4A). The knockdown of ATR by siRNA compromised UV‐dependent PP1γ‐pThr 311 dephosphorylation (Fig 4B). Treatment with purvalanol A, a specific Cdk inhibitor, resulted in almost complete reduction in PP1γ‐pThr 311 and H3‐pThr 11 in the absence of DNA damage (Fig 4C). Both PP1γ‐pThr 311 and H3‐pThr 11 were significantly reduced when Cdk1 was depleted, and further reduced when both Cdk1 and Cdk2 were depleted (Fig 4D). By contrast, ectopic expression of a constitutively active Cdk1 mutant (Cdk1AF) inhibited UV‐induced dephosphorylation of PP1γ‐pThr 311 and H3‐pThr 11 (Fig 4E). These results suggest that ATR/Chk1‐dependent inhibition of Cdk activity results in dephosphorylation of PP1γ‐pThr 311 and activation of PP1γ, leading to dephosphorylation of H3‐pThr 11 (Fig 5).
In this study, we demonstrate that the dephosphorylation of H3‐pThr 11 and the subsequent transcriptional repression of cell cycle regulatory genes after DNA damage are regulated at least in part by the ATR–Chk1–Cdk–PP1γ axis. This pathway seems to have an important role in cell survival after DNA damage. Although the precise mechanism by which PP1γ regulates cell viability remains unknown, levels of cyclins and Cdks might be a crucial determinant of cell viability after DNA damage.
Cell culture and adenovirus infection. HCT116 cells were cultured in McCoy's 5a medium containing 10% fetal bovine serum. HeLa cells and mouse embryonic fibroblasts were cultured in Dulbecco's modified Eagle medium containing 10% fetal bovine serum. All cells were cultured at 37°C in 5% CO2. Adenovirus‐Cdk1AF (Niida et al, 2005) or LacZ (control) was used to infect HCT116 cells at multiplicity of infection 40.
Immunoblotting. Subcellular fractionation was performed as previously described (Niida et al, 2007). To solublize the chromatin fraction, pellets were suspended in immunoprecipitation kinase buffer (50 mM HEPES (pH 8.0), 150 mM NaCl, 2.5 mM EGTA, 1 mM dithiothreitol, 0.1% Tween‐20, 10% glycerol and protease inhibitors). For preparation of whole‐cell extracts, cells were lysed in immunoprecipitation kinase buffer.
Phosphatase assay. Chromatin‐bound PP1γ was solubilized and immunoprecipitated with PP1γ antibody. The precipitates were incubated with chromatin at 30 °C for 1 h in phosphatase buffer (10 mM HEPES, 10 mM MgCl2, 1 mM MnCl2 and 1 mM DTT). His‐tagged wild‐type PP1γ or PP1γ‐Thr 311A was purified from Sf9 cells and phosphatase assays were performed using a chromatin fraction as a substrate.
Knockdown experiments by siRNA transfection. HCT116 or HeLa cells were transfected with either a control siRNA (Silencer Negative Control, Ambion) or an siRNA for ATR, PP1, PP1α, PP1β, PP1γ, PP2A or Cdk1 and Cdk2 using Lipofectamine 2000 (Invitrogen). The sequences of these siRNAs were described in the supplementary Table S2 online.
Immunofluorescence. Cells were grown on glass coverslips or chamber slides, washed with phosphate buffered saline (PBS) and fixed with 2% paraformaldehyde in PBS for 15 min, and stained with the indicated antibodies diluted in 5% blocking buffer (Convance, normal serum block) and DAPI.
Quantitative real‐time RT–PCR for measurement of Cdk1, cyclin B1 and cyclin A2 mRNA expressions. The RNA was extracted using ISOGEN (Wako) according to the manufacturer's protocol. Total RNAs were subjected to reverse transcription. Cdk1, cyclin B1 and cyclin A2 mRNA levels were measured by Taqman quantitative PCR, normalized to GAPDH gene expression and expressed relative to the un‐irradiated sample.
MTT assay. HCT116 cells were incubated with medium containing MTT solution (0.5 mg/ml) for 4 h. After removing the medium, cells were incubated with DMSO and the optical density of the resultant supernatant was measured at 535 nm.
Antibodies. The antibodies used for immunoblotting were ATR (sc‐1887; Santa Cruz Biotechnology), β‐actin (ab6276; Abcam), Chk1 (sc‐8408 and sc‐56291; Santa Cruz Biotechnology), Myc (sc‐789 and sc‐40; Santa Cruz Biotechnology), Cdk1 (sc‐54; Santa Cruz Biotechnology), Cdk2 (sc‐163; Santa Cruz Biotechnology), FLAG (M2; Sigma), H3 (ab1791; Abcam), IKKα (sc‐7182; Santa Cruz Biotechnology), H3 (9715; Cell Signaling Technology), pT11 (ab5168; Abcam), PP1 (sc‐7482; Santa Cruz Biotechnology), pThr320‐PP1 (2581; Cell Signaling Technology), PP2A (05‐421; Upstate), PP1α (06‐221; Upstate), PP1β (ab53315; Abcam) and PP1γ (sc‐6108; Santa Cruz Biotechnology, 07‐1298; Millipore).
Supplementary information is available at EMBO reports online (http://www.emboreports.org).
Conflict of Interest
The authors declare that they have no conflict of interest.
We thank P.M. Carr, D. Zineldeen and M. Delhase for critical reading of this paper; and Y. Chiba, S. Tsubaki, Y. Kawada, T. Misaki, Y. Koshiyama and C. Yamada‐Namikawa for technical assistance. This study was supported in part by a Grant‐in‐Aid for Scientific Research (B) and the project for realization of regenerative medicine, by the Mitsubishi Foundation, by the Naito Memorial Foundation and by the Toyoaki Foundation (to M.N.), by the Mochida Memorial Foundation, Shiseido Female Researcher Science Grant and by a Grant‐in‐Aid for Young Scientists (A; to M.S.).
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