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Previous Article | Next Article 
Molecular and Cellular Biology, October 2006, p. 7575-7586, Vol. 26, No. 20
0270-7306/06/$08.00+0 doi:10.1128/MCB.01887-05
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Department of Pharmacology and Therapeutics,1 McGill Program for the Study of Behavior, Genes and Environment, McGill University, 3655 Sir William Osler Promenade, Montréal, Québec H3G 1Y6, Canada,3 Douglas Hospital Research Center, 6875 LaSalle Blvd, Montréal, Québec H4H 1R3, Canada2
Received 26 September 2005/ Returned for modification 2 November 2005/ Accepted 21 July 2006
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ABSTRACT |
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H2AX focus formation, and cell division control protein 25a (CDC25a) degradation, in an ataxia telangiectasia mutated-Rad3-related (ATR)-dependent manner. siRNA knockdown of ATR blocks the response to DNMT1 depletion; DNA synthesis continues in the absence of DNMT1, resulting in global hypomethylation. Similarly, the response to DNMT1 knockdown is significantly attenuated in human mutant ATR fibroblast cells from a Seckel syndrome patient. This response is sensitive to DNMT1 depletion, independent of the catalytic domain of DNMT1, as indicated by abolition of the response with ectopic expression of either DNMT1 or DNMT1 with the catalytic domain deleted. There is no response to short-term treatment with 5-aza-deoxycytidine (5-aza-CdR), which causes demethylation by trapping DNMT1 in 5-aza-CdR-containing DNA but does not cause disappearance of DNMT1 from the nucleus. Our data are consistent with the hypothesis that removal of DNMT1 from replication forks is the trigger for this response. |
INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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A loss of DNMT1 during replication would result in a loss of epigenetic information. It has therefore been proposed that cells have developed multiple ways to coordinate the transfer of genetic and epigenetic information from mother to daughter cell during cellular division (40). Proposed mechanisms include cell cycle regulation of DNMT1 expression at both the transcriptional and posttranscriptional levels (41). However, such mechanisms only explain how cells are able to synthesize sufficient amounts of DNMT1 under normal conditions. We have shown that a knockdown of DNMT1 leads to a reduction in firing of origins of DNA replication (20). Using double labeling with propidium iodide to mark S-phase cells and bromodeoxyuridine to label cells actively synthesizing DNA, we showed that DNMT1 depletion triggers intra-S-phase cell cycle arrest (26). The mechanisms responsible for DNA replication inhibition following DNMT1 knockdown have been unknown.
This type of response is also seen when replication forks are stalled during cell division. Cells respond to the appearance of single-stranded DNA (48) that arises from stalled replication forks during DNA replication or DNA damage by activating ataxia telangiectasia mutated (ATM) and ATM-Rad3-related (ATR) effector kinases (1) to initiate a signaling pathway (7) that involves activation of the checkpoint kinases (ChK) leading to phosphorylation and degradation of cell division control protein 25a (CDC25a). As a consequence, the downstream effect is the decreased capacity to load CDC45 onto replication origins, therefore leading to impaired recruitment of replication complexes and DNA replication arrest. In this paper, we define the pathway responsible for DNA replication arrest in response to loss of DNMT1 and show that it is similar to the pathway elicited by hydroxyurea, a classic inducer of the DNA replication stress checkpoint. We also show that once this response is blocked, loss of DNMT1 leads to genomic hypomethylation.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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CAT (catalytic domain deletion) were created by digestion of pBluescript DNMT1 and pBluescript DNMT1 CAT plasmids with SmaI/BssHII and insertion of the DNMT1 fragments into pEF6/HisB vector (Invitrogen).
Cell culture, transfections, and drug treatments.
T24, a human bladder transitional carcinoma-derived cell line, A549, a human non-small-cell lung carcinoma cell line, and NIH 3T3, a mouse fibroblast cell line, were obtained from the ATCC (HTB-4, CCL-185, and CRL-1658, respectively). Normal human fibroblasts and Seckel syndrome fibroblasts were obtained from the CCR (GM01887 and GM18366). T24 cells were maintained in McCoy's medium, A549 cells were grown in Dulbecco's modified Eagle's medium (low glucose), and NIH 3T3 cells were grown in Dulbecco's modified Eagle's medium (high glucose). Normal human fibroblasts and ATR mutant fibroblasts were grown in minimum essential medium. The cells were supplemented with 10% fetal calf serum, 2 mM glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin. Eighteen to 24 h prior to either antisense or small interfering RNA (siRNA) treatment, cells were plated at a concentration of 4 x 105 cells in a 100-mm tissue culture dish, 5 x 104 cells in a 6-well plate, or 1 x 104cells in a 24-well plate. The phosphorothioate oligodeoxynucleotides used in this study were previously described (26): human DNMT1 antisense oligonucleotide (DNMT1 AS; 5'-AAGCATGAGCACCGTTCTCC-3') and its mismatch control (Con AS, 5'-AACGATCAGGACCCTTGTCC-3'), which has a 6-bp difference from DNMT1 AS (9). The following siRNAs, obtained from Dharmacon, were used in this study: human DNMT1 siRNA (siDNMT1; antisense strand, 5'-GCAUGAGCACCGUUCUCCdTdT-3' [sequence taken from reference 32]), siRNA mismatch control (siCon; antisense strand, 5'-UGGAGAGCACCGUUCUCCdTdT-3'), human ATR siRNA (siATR; SMARTpool siRNA), and mouse 3' UTR DNMT1 siRNA (simDNMT1; antisense strand, 5'-AAUAAAGCGUUCUGUCAACC-3'). Antisense oligonucleotides were transfected using Lipofectin (Invitrogen) in serum-free Opti-MEM. The oligonucleotide-containing Opti-MEM was removed from the cells and replaced with regular growth medium after 4 h. The cells were harvested after 24 h. siRNA was transfected using Lipofectamine (Invitrogen) in serum-free Opti-MEM. The oligonucleotide-containing Opti-MEM was removed from the cells and replaced with regular growth medium after 4 h. Stable cell lines were produced using NIH 3T3 cells transfected with pBabe.puro and either pEF6, pEF6-DNMT1 WT, or pEF6-DNMT1 CAT by the CaPO4 method, at a pBabe.puro/pEF6 construct ratio of 1:20. Cells were selected using Puromycin (3 µg/ml) for 2 weeks. Individual clones were isolated and seeded to 24-well dishes and were screened for protein expression. For 5-aza-deoxycytidine (5-aza-CdR) treatment, cells were grown in regular culture medium in the presence of 10–6 M 5-aza-CdR (Sigma) dissolved in dimethyl sulfoxide (DMSO). The cells were treated for 24 h.
Western blot analysis.
Fifty micrograms of nuclear protein, whole-cell or chromatin-bound extracts, and 25 µg of extracted histones were fractionated on a 7.5% to 12% sodium dodecyl sulfate-polyacrylamide gel and transferred to polyvinylidene difluoride membrane. Nuclei were isolated following lysis of the cells with buffer A (10 mM Tris, pH 8, 1.5 mM MgCl2, 5 mM KCl, 0.5 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride [PMSF], 0.5% NP-40), followed by centrifugation. Free nuclear proteins were extracted from isolated nuclei with buffer B (1 M Tris, pH 8, 25% glycerol, 1.5 mM MgCl2, 0.5 mM PMSF, 0.2 mM EDTA, 0.5 mM dithiothreitol, 0.4 M NaCl). The chromatin pellet was precipitated by centrifugation. Chromatin-bound proteins were extracted by sonication of the chromatin pellet in protein loading buffer. Histones were extracted from isolated nuclei by acid hydrolysis in buffer HE (0.2 N HCl, 10 mM Tris, pH 8, 1.5 mM MgCl2, 100 nM PMSF). The following antibodies were used: human DNMT1 (New England Biolabs), mouse DNMT1 (ab5208-100; Abcam), CDC25a (F-6; Santa Cruz), CDC25b (C-20; Santa Cruz), ß-actin (A5316; Sigma), PCNA (PC10; Santa Cruz), phospho-Chk2 (Thr68) (sc-16297-R; Santa Cruz), Chk2 (Ab-1; Oncogene), phospho-Chk1 (Ser 317) (DR-1025; Calbiochem), Chk1 (sc-8408; Santa Cruz), Xpress (46-0528; Invitrogen), H2A.X (05-636; Upstate), and pan-histone 3 (30374; Upstate). Membranes were blocked in 5% milk in Tris-buffered saline (TBS) with 0.05% Tween (TBST) for 1 h and then exposed to antibody in the TBST-5% milk solution overnight at 4°C. The membranes were washed with TBST and then exposed to either horseradish peroxidase-conjugated anti-rabbit immunoglobulin G (IgG) or anti-mouse IgG in TBST-5% milk or TBST, respectively, for 1 h at room temperature. The signal was then developed using an ECL enhanced chemiluminescence kit from Amersham (catalog no. 135,136). Quantification was performed by measurement of relative optical density (ROD). To calculate the final signal for each sample, the ROD value of the antibody signal was divided by the ROD value of the band within the actin lane. To calculate the normalized final signal of the pChk2 samples, the ROD value of the pChk2 band was divided by the ROD value of the Chk2 band.
Immunocytochemistry and confocal microscopy.
T24 cells were plated on coverslips in 24-well plates. After treatment, the cells were washed with phosphate-buffered saline (PBS) and fixed with 4% paraformaldehyde. Coverslips were washed with PBS and blocked with 10% horse serum and 0.1%Triton X-100 (Sigma) in PBS (PBST) for 1 h. Coverslips were stained with either H2AX antibody (JBW103; Upstate), DNMT1 antibody (New England Biolabs), or ATR antibody (N-19; Santa Cruz Biotechnology) in PBST and 5% horse serum for 1 h. The coverslips were washed with PBST and stained with either Alexa Fluor 488 anti-rabbit IgG, Alexa Fluor anti-rabbit 548, Alexa Fluor 633 anti-rabbit IgG, or Alexa Fluor 568 anti-goat IgG (Molecular Probes). The coverslips were then washed with PBST and mounted on slides using Immuno-mount. 5-Methylcytosine staining was performed by fixation with 4% paraformaldehyde for 30 min and two washes with PBS, and staining was blocked with 10% fetal bovine serum in PBST for 2 h. DNA was denatured in 2 N HCl and 0.1%Triton X-100 in PBS for 45 min at 37°C. The cells were neutralized with 0.1 M sodium borate solution, washed with PBS, and stained with a mouse monoclonal anti-5-methylcytosine antibody (a gift from Alain Niveleau) in PBST. The cells were washed with PBS and stained with Alexa Fluor 488 anti-mouse IgG (Molecular Probes) in PBS. The coverslips were washed with PBS and were mounted on slides using Immuno-count. DAPI (4',6'-diamidino-2-phenylindole) staining was performed by staining cells with DAPI (Sigma) in PBS for 5 min and washing them with PBS. The staining was analyzed using LSM 520 laser scanning microscopy, version 2.5 (Zeiss). The intensity of staining for ATR, DNMT1, H2AX, and 5-methylcytosine was quantified (MCID Elite image analysis software; Imaging Research, Inc.) from an average of three independent fields. Signal intensity represented the strength of fluorescence from stained cells as determined by MCID Elite 7.0.
Thymidine incorporation. The rate of DNA synthesis was determined 24 h after initiation of treatment with either ATR or DNMT1 siRNAs by measuring [3H]thymidine incorporation into DNA following a 6-h pulse with 66 µCi/ml [3H]thymidine. Pulsed cells were washed five times with PBS and incubated in 10% trichloroacetic acid at 4°C for 12 h. Cells were washed twice with 5% trichloroacetic acid for 30 min. The cells were lysed with 1% sodium dodecyl sulfate in 1 N NaOH, and the level of [3H]thymidine incorporation was measured in a liquid scintillation counter.
5-Methylcytosine quantification by nearest-neighbor analysis. Levels of 5-methylcytosine were quantified by nearest-neighbor analysis as described previously (30). The intensity of 5-methylcytosine and cytosine mononucleotide spots was measured using a PhosphorImager screen and the Image Quant image analysis program. Levels of unmethylated cytosine content are expressed as a percentage of [cytosine]/[cytosine + methylcytosine].
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RESULTS |
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To determine whether this cascade of events is common to cells derived from different cancer tissues, we examined whether DNMT1 depletion would increase levels of Chk2 phosphorylation in the non-small-cell lung carcinoma A549 cell line. Treatment of A549 cells with DNMT1 antisense oligonucleotide decreased levels of DNMT1 protein in comparison to those in the cells treated with the mismatch oligonucleotide control (Fig. 2). Although, DNMT1 antisense oligonucleotide increased levels of Chk2 threonine 68 phosphorylation, absolute levels of ChK2 were not altered. These results show that the causal relationship between knockdown of DMNT1 and checkpoint activation is not unique to T24 cells.
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H2AX focus formation.
A hallmark in the cell's response to replication stress is the formation of histone variant H2AX foci, a characteristic marker of cells undergoing replication stress (10). We first examined the effect of DNMT1 depletion on histone variant H2AX focus formation. Treatment of T24 cells with either DNMT1 AS or siDNMT1 led to a significant induction (P < 0.05) of H2AX foci in comparison to that in cells treated with either the Con AS or siCon, respectively (Fig. 3a, b, c, and d). H2A.X induction in DNMT1-depleted T24 cells was confirmed by a Western blot analysis (Fig. 3e and f). These results lead us to conclude that induction of H2AX focus formation occurs as a consequence of depletion of DNMT1, which is consistent with other studies showing that germ cells lacking DNMT3l are positive for H2AX foci (3).
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H2AX (5, 35). We tested whether ATR is required for the effect of DNMT1 depletion on histone variant H2AX focus formation. Treatment of T24 cells with 100 nM siATR significantly (P < 0.05) decreased ATR immunoreactivity in comparison to that of the cells treated with siCon (Fig. 4a and b). T24 cells treated with siATR did not elicit H2AX focus formation (Fig. 4c). Treatment of cells with siDNMT1 led to H2AX foci (Fig. 4C); however, when DNMT1 was depleted in cells treated with siATR, H2AX focus formation was significantly (P < 0.05) reduced (Fig. 4d), suggesting that ATR is required for the DNMT1 knockdown response.
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H2A.X when treated with siDNMT1 compared to siCon, confirming that the response to DNMT1 knockdown is not limited and occurs in many cell types (Fig. 4f). However, Seckel syndrome cells treated with siDNMT1 exhibited a markedly reduced response (Fig. 4f). Together, these results indicate that ATR is required for the replication arrest induced by DNMT1 depletion. Loss of DNMT1 leads to global hypomethylation once the DNA stress response pathway is blocked. We then tested the hypothesis that the response elicited by DNMT1 depletion serves to protect the genome from disruption of the DNA methylation pattern by replication in the absence of maintenance methylation. Thus, we hypothesized that blocking the response elicited by DNMT1 knockdown will affect the integrity of the DNA methylation state. We therefore examined the effect of a double knockdown of ATR and DNMT1 on the global state of methylation. Using a nearest-neighbor analysis to determine the cytosine levels in the siATR- and siDNMT1-treated T24 cells (Fig. 4g and h), the global cytosine levels were revealed to be unaltered in the siATR-treated cells as compared to those in control-treated cells. DNMT1 depletion did not result in global hypomethylation as well. However, the double knockdown of ATR and DNMT1 led to a significant increase in the fraction of unmethylated cytosines in the genome (P < 0.05) as compared to those in the control-treated cells or single knockdowns of either ATR or DNMT1. Our data imply that the arrest of DNA replication following DNMT1 depletion prevents loss of DNA methylation and that blocking this response results in a global loss in DNA methylation. These data reveal the functional significance of the ATR-mediated response to depletion of DNMT1.
DNMT1 depletion response is insensitive to global hypomethylation. To determine whether DNMT1 depletion in the nucleus or demethylation of DNA signals the response described here, we took advantage of the classic DNA methylation inhibitor 5-aza-2'-deoxycytidine (5-aza-CdR) (26). The nucleoside analogue is incorporated in the DNA and inhibits the DNA methylation transfer activity of DNMT1 and other DNMTs by entrapment, but it does not inhibit the de novo synthesis of DNMT1 protein or its residence in the nucleus (Fig. 5d and e) (18). 5-Aza-CdR causes an immediate loss of global methylation. If the signal that triggers the response to DNMT1 depletion is global demethylation, then 5-aza-CdR should elicit a response similar to the DNMT1 knockdown. We first compared the effects of DNMT1 knockdown and 5-aza-CdR on genomic methylation using previously characterized anti 5-methylcytosine antibodies. Levels of genomic methylation in T24 cells treated with siDNMT1 did not significantly differ in comparison to the cells treated with siCon (Fig. 5a and b), as discussed in the previous section, whereas, T24 cells treated with 1 µM 5-aza-CdR showed significant (P < 0.05) DNA demethylation, as indicated by the loss of 5-methylC immunoreactivity (Fig. 5a and b), which has been shown previously (26). The cells were then harvested, and nuclear protein levels were measured by Western blot analysis. As expected, protein extracts of T24 cells treated with 5-aza-CdR showed no immunoreactivity to the DMNT1 primary antibody (Fig. 5c), implying that the free DNMT1 had been significantly depleted. However, when DNMT1 levels were examined in extracts prepared from chromatin bound protein, DNMT1 was not depleted by 5-aza-CdR treatment (Fig. 5c). These results are consistent with the classical mechanism of action of 5-aza-CdR, where it traps DNMT1 in the nucleus (34) rather than depletes DNMT1 protein levels through mRNA degradation like siRNA or antisense. Since it has recently been shown that 5-aza-CdR could cause degradation of DNMT1 protein in addition to its trapping on the DNA (12), we further confirmed the presence of DNMT1 in the nuclei of 5-aza-CdR-treated cells by immunostaining (Fig. 5d) for DNMT1. Although there was some small reduction of DNMT1 in T24 cells treated with 5-aza-CdR, as compared to the level in control cells, DNMT1 was clearly present in 5-aza-CdR-treated cells, in contrast to T24 cells treated with DNMT1 siRNA, where it was essentially absent.
Levels of Chk2 threonine 68 phosphorylation remained undetected following 5-aza-CdR treatment, while absolute levels of ChK2 protein were not significantly (P < 0.05) altered by the treatment (Fig. 5c). Thus, although 5-aza-CdR causes global hypomethylation, it does not trigger the cellular response elicited by DNMT1 knockdown. These results confirm previous findings that 5-aza-CdR retards replication but does not initiate intra-S-phase arrest (26). This is consistent with the hypothesis that depletion of DNMT1 in the nucleus rather than DNA hypomethylation initiates the replication arrest response described here.
Ectopic expression of DNMT1 and a catalytic domain mutant blocks the cellular response to endogenous DNMT1 knockdown. To verify that knockdown of DNMT1 expression rather than a nonspecific side effect of siRNA is responsible for the cellular response described here, we generated mouse NIH 3T3 transfectants ectopically expressing either an empty expression vector, a murine DNMT1 cDNA, or a catalytic domain-deficient murine DNMT1 cDNA (Fig. 5f). We then selectively targeted the endogenous DNMT1 with siRNA, which is specific to the 3' UTR present in the endogenous DNMT1 but not in the ectopic constructs. If reduction in DNMT1 levels triggers the response, then the expression of ectopic DNMT1, which is not downregulated by the treatment, should rescue the loss of endogenous DNMT1 and block the cellular response to reduction in DNMT1 siRNA. A catalytic DNMT1 mutant was used to verify whether loss of DNA methylation triggered the response. If loss in DNA methylation caused by knockdown of DNMT1 enzymatic activity triggered the response, then this response could only be blocked by a catalytically active DNMT1. However, if other functional domains of DNMT1 were involved, then the catalytic mutant would rescue the downregulation of endogenous DNMT1 and block the response. Stable NIH 3T3 cell lines expressing these constructs were generated and validated by examination of the expression of the Xpress epitope included in the ectopic constructs (Fig. 5f). We utilized siRNA specific for mouse DNMT1 3' UTR, simDNMT1, (Fig. 5f) which depleted the cells of endogenous DNMT1 (Fig. 5g) but not exogenous Xpress-tagged DNMT1 (Fig. 5f).
We first determined whether nontransformed NIH 3T3 cells exhibited the same response to the DNMT siRNA as that observed above in cancerous human cells. DNMT1 depletion in empty vector transfectants responded with an induction of H2A.X, which corresponded to our observations seen in T24 cell lines (Fig. 3). This result demonstrates that the effects of DNMT1 knockdown are not limited to humans or cancer cells such as T24 or A549 and occur in a nontransformed mouse cell line as well. The response is not an idiosyncrasy of a specific siRNA sequence since the siRNA used here targets a totally different region of the DNMT1 mRNA from the siRNA used in Fig. 1 to 3 (Fig. 5f). Cells expressing either pEF6-DNMT1 WT or pEF6-DNMT1 CAT did not exhibit an induction of H2A.X. The fact that expression of exogenous DNMT1 rescues the cells from the cellular response to DNMT1 siRNA suggests that a reduction in DNMT1 levels rather than other effects of the sequence is responsible for this response. If the siRNA per se were toxic, a similar response would be expected in the ectopic DNMT1 transfectants as well as in the controls since they were all exposed to the same siRNA. The fact that the rescue by ectopic DNMT1 takes place even in the absence of the catalytic domain suggests that this response is independent of the catalytic function of DNMT1. These data are consistent with the notion that DNMT1 plays a role in the replication fork, which is independent of its catalytic activity as previously proposed (40).
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DISCUSSION |
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The response to DNMT1 depletion is dependent on ATR, since ATR depletion abolishes the response; moreover, the response is markedly attenuated in cells derived from a patient with Seckel syndrome who bears highly defective ATR. The ATR-dependent cellular response described here is sensitive to DNMT1 depletion rather than demethylation per se since a similar response is not triggered by a DNA demethylation agent (Fig. 5c). In addition, the response is blocked by both catalytically active and inactive DNMT1, suggesting that loss of catalytic activity of DNMT1 is not driving this cellular response (Fig. 5f). ATR is known to be activated by a wide variety of genotoxic challenges as well as uncoupling of DNA replication proteins from the replication fork (28).
DNMT1 depletion might cause loss of DNA methylation as previously observed in dnmt1–/– mice (21). We have recently shown, however, that DNMT1 depletion leads to very limited genomic demethylation (26). We previously proposed that the intra-S-phase arrest in DNA replication protected the cells from demethylation upon reduction in DNMT1 levels (26). Similarly, DNMT1 knockout by homologous recombination in human colorectal cancer cells resulted in very limited inhibition of DNA methylation, suggesting that other DNMTs compensate for the loss in DNMT1 (31, 42). Nevertheless, these data do not exclude the possibility that a low level of hemimethylated DNA generated in the course of synthesis of nascent DNA in the absence of DNMT1 signaled activation of checkpoint signaling pathway. This hypothesis is not supported, however, by the lack of Chk2 activation in response to the DNA methylation inhibitor 5-aza-CdR (Fig. 5c). Although 5-aza-CdR induced extensive global hypomethylation, as shown in Fig. 4f and g and 5a, it did not elicit the phosphorylation of Chk2, similar to DNMT1 knockdown (Fig. 5c). This is consistent with our previous observations that short-term 5-aza-CdR treatment did not induce an intra-S-phase cell cycle arrest (26). Additionally the fact that this response could be completely blocked by ectopic expression of a catalytic mutant of DNMT1 suggests that this effect is not caused by the lack of DNMT1 catalytic activity. Thus, our data suggest that it is the absence of the DNMT1 protein from the fork, not the loss of its DNA methylation activity, that elicits this response. This might be a general rule for other epigenetic proteins such as CAF1 and histone-modifying enzymes (11, 15, 45, 46). It is possible that a global mechanism exists that senses the absence of the full complement of chromatin-modifying enzymes in the fork and activates ATR, leading to arrest of DNA replication in the S phase of the cell cycle. However, the simplest interpretation is that removal of DNMT1 from replication forks disrupts fork progression and causes stalled forks and DNA double-strand breaks which elicit this response. We show that this response protects the DNA from global loss of methylation. If this response is inhibited by knockdown of ATR, replication is not blocked and DNA synthesis proceeds in the absence of DNMT1: as a result, the DNA is globally hypomethylated.
Our study unravels many of the components of the pathway leading from DNMT1 depletion to S-phase arrest. DNMT1 depletion in the replication fork leads to ATR activation, and as a consequence, its downstream effectors such as Chk1 and Chk2 are activated by phosphorylation, leading to phosphorylation and degradation of CDC25a phosphatase and therefore decreased capacity of its substrate, CDC45, to load onto replication origins, leading to an overall arrest of replication and S phase (Fig. 6). The model proposes a potential two-phase effect of DNMT1 depletion, in which the immediate effect of DNMT1 depletion causes an arrest in DNA replication through the response outlined in this study, as well as a transcriptional effect previously described (25, 26), whereby DNMT1 depletion causes induction of stress response genes. Malfunctioning of this response such as in the case of an ATR knockdown could lead to replication in the absence of DNMT1 and loss of DNA methylation (Fig. 4h).
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ACKNOWLEDGMENTS |
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This study was supported by a grant from the Canadian Institute of Health Research to M.S.
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FOOTNOTES |
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REFERENCES |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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2. Bigey, P., S. Ramchandani, J. Theberge, F. D. Araujo, and M. Szyf. 2000. Transcriptional regulation of the human DNA methyltransferase (dnmt1) gene. Gene 242:407-418.[CrossRef][Medline]
3. Bourc'his, D., and T. H. Bestor. 2004. Meiotic catastrophe and retrotransposon reactivation in male germ cells lacking Dnmt3L. Nature 431:96-99.[CrossRef][Medline]
4. Chaturvedi, P., W. K. Eng, Y. Zhu, M. R. Mattern, R. Mishra, M. R. Hurle, X. Zhang, R. S. Annan, Q. Lu, L. F. Faucette, G. F. Scott, X. Li, S. A. Carr, R. K. Johnson, J. D. Winkler, and B. B. Zhou. 1999. Mammalian Chk2 is a downstream effector of the ATM-dependent DNA damage checkpoint pathway. Oncogene 18:4047-4054.[CrossRef][Medline]
5. Cortez, D., S. Guntuku, J. Qin, and S. J. Elledge. 2001. ATR and ATRIP: partners in checkpoint signaling. Science 294:1713-1716.
6. Falck, J., N. Mailand, R. G. Syljuasen, J. Bartek, and J. Lukas. 2001. The ATM-Chk2-Cdc25A checkpoint pathway guards against radioresistant DNA synthesis. Nature 410:842-847.[CrossRef][Medline]
7. Falck, J., J. H. Petrini, B. R. Williams, J. Lukas, and J. Bartek. 2002. The DNA damage-dependent intra-S phase checkpoint is regulated by parallel pathways. Nat. Genet. 30:290-294.[CrossRef][Medline]
8. Florensa, R., O. Bachs, and N. Agell. 2003. ATM/ATR-independent inhibition of cyclin B accumulation in response to hydroxyurea in nontransformed cell lines is altered in tumour cell lines. Oncogene 22:8283-8292.[CrossRef][Medline]
9. Fournel, M., P. Sapieha, N. Beaulieu, J. M. Besterman, and A. R. MacLeod. 1999. Down-regulation of human DNA-(cytosine-5) methyltransferase induces cell cycle regulators p16(ink4A) and p21(WAF/Cip1) by distinct mechanisms. J. Biol. Chem. 274:24250-24256.
10. Furuta, T., H. Takemura, Z. Y. Liao, G. J. Aune, C. Redon, O. A. Sedelnikova, D. R. Pilch, E. P. Rogakou, A. Celeste, H. T. Chen, A. Nussenzweig, M. I. Aladjem, W. M. Bonner, and Y. Pommier. 2003. Phosphorylation of histone H2AX and activation of Mre11, Rad50, and Nbs1 in response to replication-dependent DNA double-strand breaks induced by mammalian DNA topoisomerase I cleavage complexes. J. Biol. Chem. 278:20303-20312.
11. Gaillard, P. H., E. M. Martini, P. D. Kaufman, B. Stillman, E. Moustacchi, and G. Almouzni. 1996. Chromatin assembly coupled to DNA repair: a new role for chromatin assembly factor I. Cell 86:887-896.[CrossRef][Medline]
12. Ghoshal, K., J. Datta, S. Majumder, S. Bai, H. Kutay, T. Motiwala, and S. T. Jacob. 2005. 5-Aza-deoxycytidine induces selective degradation of DNA methyltransferase 1 by a proteasomal pathway that requires the KEN box, bromo-adjacent homology domain, and nuclear localization signal. Mol. Cell. Biol. 25:4727-4741.
13. Goodship, J., H. Gill, J. Carter, A. Jackson, M. Splitt, and M. Wright. 2000. Autozygosity mapping of a Seckel syndrome locus to chromosome 3q22.1-q24. Am. J. Hum. Genet. 67:498-503.[CrossRef][Medline]
14. Hassepass, I., R. Voit, and I. Hoffmann. 2003. Phosphorylation at serine 75 is required for UV-mediated degradation of human Cdc25A phosphatase at the S-phase checkpoint. J. Biol. Chem. 278:29824-29829.
15. Hoek, M., and B. Stillman. 2003. Chromatin assembly factor 1 is essential and couples chromatin assembly to DNA replication in vivo. Proc. Natl. Acad. Sci. USA 100:12183-12188.
16. Ishimi, Y., Y. Komamura-Kohno, H. J. Kwon, K. Yamada, and M. Nakanishi. 2003. Identification of MCM4 as a target of the DNA replication block checkpoint system. J. Biol. Chem. 278:24644-24650.
17. Issa, J. P., P. M. Vertino, J. Wu, S. Sazawal, P. Celano, B. D. Nelkin, S. R. Hamilton, and S. B. Baylin. 1993. Increased cytosine DNA-methyltransferase activity during colon cancer progression. J. Natl. Cancer Inst. 85:1235-1240.
18. Jones, P. A. 1985. Effects of 5-azacytidine and its 2'-deoxyderivative on cell differentiation and DNA methylation. Pharmacol. Ther. 28:17-27.[CrossRef][Medline]
19. Kanai, Y., S. Ushijima, Y. Nakanishi, M. Sakamoto, and S. Hirohashi. 2003. Mutation of the DNA methyltransferase (DNMT) 1 gene in human colorectal cancers. Cancer Lett. 192:75-82.[CrossRef][Medline]
20. Knox, J. D., F. D. Araujo, P. Bigey, A. D. Slack, G. B. Price, M. Zannis-Hadjopoulos, and M. Szyf. 2000. Inhibition of DNA methyltransferase inhibits DNA replication. J. Biol. Chem. 275:17986-17990.
21. Li, E., T. H. Bestor, and R. Jaenisch. 1992. Targeted mutation of the DNA methyltransferase gene results in embryonic lethality. Cell 69:915-926.[CrossRef][Medline]
22. MacLeod, A. R., J. Rouleau, and M. Szyf. 1995. Regulation of DNA methylation by the Ras signaling pathway. J. Biol. Chem. 270:11327-11337.
23. MacLeod, A. R., and M. Szyf. 1995. Expression of antisense to DNA methyltransferase mRNA induces DNA demethylation and inhibits tumorigenesis. J. Biol. Chem. 270:8037-8043.
24. Manke, I. A., A. Nguyen, D. Lim, M. Q. Stewart, A. E. Elia, and M. B. Yaffe. 2005. MAPKAP kinase-2 is a cell cycle checkpoint kinase that regulates the G2/M transition and S phase progression in response to UV irradiation. Mol. Cell 17:37-48.[CrossRef][Medline]
25. Milutinovic, S., S. E. Brown, Q. Zhuang, and M. Szyf. 2004. DNA methyltransferase 1 knock down induces gene expression by a mechanism independent of DNA methylation and histone deacetylation. J. Biol. Chem. 279:27915-27927.
26. Milutinovic, S., Q. Zhuang, A. Niveleau, and M. Szyf. 2003. Epigenomic stress response. Knockdown of DNA methyltransferase 1 triggers an intra-S-phase arrest of DNA replication and induction of stress response genes. J. Biol. Chem. 278:14985-14995.
27. O'Driscoll, M., V. L. Ruiz-Perez, C. G. Woods, P. A. Jeggo, and J. A. Goodship. 2003. A splicing mutation affecting expression of ataxia-telangiectasia and Rad3-related protein (ATR) results in Seckel syndrome. Nat. Genet. 33:497-501.[CrossRef][Medline]
28. Pohler, J. R., M. Otterlei, and E. Warbrick. 2005. An in vivo analysis of the localisation and interactions of human p66 DNA polymerase delta subunit. BMC Mol. Biol 6:17. [Online.] doi:10.1186/1471-2199-6-17.[CrossRef][Medline]
29. Ramchandani, S., A. R. MacLeod, M. Pinard, E. von Hofe, and M. Szyf. 1997. Inhibition of tumorigenesis by a cytosine-DNA, methyltransferase, antisense oligodeoxynucleotide. Proc. Natl. Acad. Sci. USA 94:684-689.
30. Ramsahoye, B. H. 2002. Nearest-neighbor analysis. Methods Mol. Biol. 200:9-15.[Medline]
31. Rhee, I., K. E. Bachman, B. H. Park, K. W. Jair, R. W. Yen, K. E. Schuebel, H. Cui, A. P. Feinberg, C. Lengauer, K. W. Kinzler, S. B. Baylin, and B. Vogelstein. 2002. DNMT1 and DNMT3b cooperate to silence genes in human cancer cells. Nature 416:552-556.[CrossRef][Medline]
32. Robert, M. F., S. Morin, N. Beaulieu, F. Gauthier, I. C. Chute, A. Barsalou, and A. R. MacLeod. 2003. DNMT1 is required to maintain CpG methylation and aberrant gene silencing in human cancer cells. Nat. Genet. 33:61-65.[CrossRef][Medline]
33. Robertson, K. D., E. Uzvolgyi, G. Liang, C. Talmadge, J. Sumegi, F. A. Gonzales, and P. A. Jones. 1999. The human DNA methyltransferases (DNMTs) 1, 3a and 3b: coordinate mRNA expression in normal tissues and overexpression in tumors. Nucleic Acids Res. 27:2291-2298.
34. Schermelleh, L., F. Spada, H. P. Easwaran, K. Zolghadr, J. B. Margot, M. C. Cardoso, and H. Leonhardt. 2005. Trapped in action: direct visualization of DNA methyltransferase activity in living cells. Nat. Methods 2:751-756.[CrossRef][Medline]
35. Shiloh, Y. 2001. ATM and ATR: networking cellular responses to DNA damage. Curr. Opin. Genet. Dev. 11:71-77.[CrossRef][Medline]
36. Shiloh, Y. 2003. ATM and related protein kinases: safeguarding genome integrity. Nat. Rev. Cancer 3:155-168.[CrossRef][Medline]
37. Slack, A., N. Cervoni, M. Pinard, and M. Szyf. 1999. Feedback regulation of DNA methyltransferase gene expression by methylation. Eur. J. Biochem. 264:191-199.[Medline]
38. Szyf, M. 1991. DNA methylation patterns: an additional level of information? Biochem. Cell Biol. 69:764-767.[Medline]
39. Szyf, M. 1994. DNA methylation properties: consequences for pharmacology. Trends Pharmacol. Sci. 15:233-238.[CrossRef][Medline]
40. Szyf, M. 2001. The role of DNA methyltransferase 1 in growth control. Front. Biosci. 6:D599-D609.[Medline]
41. Szyf, M., V. Bozovic, and G. Tanigawa. 1991. Growth regulation of mouse DNA methyltransferase gene expression. J. Biol. Chem. 266:10027-10030.
42. Ting, A. H., K. W. Jair, H. Suzuki, R. W. Yen, S. B. Baylin, and K. E. Schuebel. 2004. CpG island hypermethylation is maintained in human colorectal cancer cells after RNAi-mediated depletion of DNMT1. Nat. Genet. 36:582-584.[CrossRef][Medline]
43. Ward, I. M., and J. Chen. 2001. Histone H2AX is phosphorylated in an ATR-dependent manner in response to replicational stress. J. Biol. Chem. 276:47759-47762.
44. Wu, J., J. P. Issa, J. Herman, D. E. Bassett, Jr., B. D. Nelkin, and S. B. Baylin. 1993. Expression of an exogenous eukaryotic DNA methyltransferase gene induces transformation of NIH 3T3 cells. Proc. Natl. Acad. Sci. USA 90:8891-8895.
45. Ye, X., and P. D. Adams. 2003. Coordination of S-phase events and genome stability. Cell Cycle 2:185-187.[Medline]
46. Ye, X., A. A. Franco, H. Santos, D. M. Nelson, P. D. Kaufman, and P. D. Adams. 2003. Defective S phase chromatin assembly causes DNA damage, activation of the S phase checkpoint, and S phase arrest. Mol. Cell 11:341-351.[CrossRef][Medline]
47. Zhou, B. B., and S. J. Elledge. 2000. The DNA damage response: putting checkpoints in perspective. Nature 408:433-439.[CrossRef][Medline]
48. Zou, L., and S. J. Elledge. 2003. Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 300:1542-1548.
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