This Article Abstract Full Text (PDF) Other Versions of this Article: MCB.01799-07v1 28/2/752    most recent Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Palii, S. S. Articles by Robertson, K. D. Search for Related Content PubMed PubMed Citation Articles by Palii, S. S. Articles by Robertson, K. D.

 Previous Article  |  Next Article 

Molecular and Cellular Biology, January 2008, p. 752-771, Vol. 28, No. 2
0270-7306/08/$08.00+0     doi:10.1128/MCB.01799-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

DNA Methylation Inhibitor 5-Aza-2'-Deoxycytidine Induces Reversible Genome-Wide DNA Damage That Is Distinctly Influenced by DNA Methyltransferases 1 and 3B

Department of Biochemistry and Molecular Biology and University of Florida Shands Cancer Center Program in Cancer Genetics, Epigenetics and Tumor Virology, University of Florida, Box 100245, Gainesville, Florida 32610

Received 2 October 2007/ Accepted 25 October 2007

	ABSTRACT

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Genome-wide DNA methylation patterns are frequently deregulated in cancer. There is considerable interest in targeting the methylation machinery in tumor cells using nucleoside analogs of cytosine, such as 5-aza-2'-deoxycytidine (5-azadC). 5-azadC exerts its antitumor effects by reactivation of aberrantly hypermethylated growth regulatory genes and cytoxicity resulting from DNA damage. We sought to better characterize the DNA damage response of tumor cells to 5-azadC and the role of DNA methyltransferases 1 and 3B (DNMT1 and DNMT3B, respectively) in modulating this process. We demonstrate that 5-azadC treatment results in growth inhibition and G₂ arrest—hallmarks of a DNA damage response. 5-azadC treatment led to formation of DNA double-strand breaks, as monitored by formation of -H2AX foci and comet assay, in an ATM (ataxia-telangiectasia mutated)-dependent manner, and this damage was repaired following drug removal. Further analysis revealed activation of key strand break repair proteins including ATM, ATR (ATM-Rad3-related), checkpoint kinase 1 (CHK1), BRCA1, NBS1, and RAD51 by Western blotting and immunofluorescence. Significantly, DNMT1-deficient cells demonstrated profound defects in these responses, including complete lack of -H2AX induction and blunted p53 and CHK1 activation, while DNMT3B-deficient cells generally showed mild defects. We identified a novel interaction between DNMT1 and checkpoint kinase CHK1 and showed that the defective damage response in DNMT1-deficient cells is at least in part due to altered CHK1 subcellular localization. This study therefore greatly enhances our understanding of the mechanisms underlying 5-azadC cytotoxicity and reveals novel functions for DNMT1 as a component of the cellular response to DNA damage, which may help optimize patient responses to this agent in the future.

	INTRODUCTION

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
DNA methylation is an essential epigenetic modification required for normal mammalian development, gene regulation, genomic imprinting, and chromatin structure (9). Methylation occurs at the C-5 position of cytosine within the CpG dinucleotide and is carried out by a family of DNA methyltransferases (DNMTs), DNMT1, DNMT3A, and DNMT3B (32). DNMT1 acts primarily as a maintenance methyltransferase by copying existing methylation patterns following DNA replication (53). DNMT3A and DNMT3B exhibit de novo methyltransferase activity and are required for establishing new methylation patterns during embryonic development (72). Deregulation of the DNA methylation machinery has been identified in several disease states, particularly cancer (78), and leads to global DNA hypomethylation of repetitive DNA, which has been linked to genomic instability (21) and, concomitantly, hypermethylation at the promoter regions of certain genes (tumor suppressors), leading to their aberrant silencing (43).

The process of cytosine methylation is reversible and may be altered by biochemical and biological manipulation, making it an attractive target for therapeutic intervention. Demethylation and consequent reactivation of tumor suppressor genes are rational approaches being used in the treatment of cancer (38, 39, 61). Currently available nucleoside-based DNMT inhibitors 5-azacytidine (5-azaC), 5-aza-2'-deoxycytidine (5-azadC), and zebularine (ZEB) are analogues of cytosine that are believed to have a similar mechanism of inhibition (16, 96). 5-azaC, 5-azadC, and ZEB are metabolically activated in vivo and readily incorporated into DNA during replication, and as a result of the chemistry of the methyltransferase reaction, the DNMT becomes covalently linked to DNA, in effect creating genome-wide protein-DNA cross-links (16, 57, 61, 103). This results in depletion of soluble DNMT protein levels, leading to replication-dependent global demethylation and gene reactivation (15, 100). Both 5-azaC and 5-azadC are used clinically for the treatment of myelodysplastic syndrome (MDS) and other leukemias, where the drugs have the approval of the Food and Drug Administration (38). Clinical trials combining aza drugs with other agents, such as histone deacetylase inhibitors and interleukin-2, have also recently been reported (33, 34). Promising results were observed with hematologic malignancies, particularly MDS. In phase II and III clinical trials, 5-azaC treatment yields a better response rate than supportive care alone (91). Similarly encouraging results were obtained with clinical trials using 5-azadC (38).

Although there is considerable literature on the possible antitumor mode of action of aza drugs, their exact in vivo mechanism remains unclear. One model for their effects involves the reactivation of aberrantly silenced growth-regulatory genes accompanied by cell cycle arrest and/or apoptosis (36). A second model for their antitumor activity is related to formation of covalent DNMT-DNA adducts in aza-containing DNA, leading to DNA damage and cytotoxicity (44). While the former model has been extensively studied, the latter has not. 5-azadC treatment is perceived as DNA damage and leads to activation of the G₁ checkpoint regulator p53 (49). Furthermore, 5-azadC treatment results in inhibition of cell proliferation due to p53-dependent activation of p21^Waf1/Cip1 (49, 104). 5-azaC also induces apoptosis, either in a p53-dependent (88) or p53-independent manner (66). Aside from the role of p53 and p21^Waf1/Cip1, the cellular events activated by 5-azaC, 5-azadC, and ZEB that occur downstream of DNA methylation inhibition in DNMT-aza-DNA-containing cells are largely unknown, as is the means by which the cellular DNA repair machinery recognizes and responds to this form of damage. It is likely that cytotoxicity is, at least in part, mediating the antitumor effect of aza drugs since clinical responses do not necessarily correlate with gene-specific or global demethylation (46, 64, 87). Given the interest in using these drugs as part of an "epigenetic"-based chemotherapy regimen and their likely increased use in the clinic, it is essential to better understand how aza-DNA-DNMT adducts mediate their cytotoxic effects and how this contributes to the overall therapeutic outcome.

One of the first steps in the response of cells to DNA damage is activation of the phosphatidylinositol 3-kinase (PI3K) family of high-molecular-weight protein kinases, ATM (ataxia-telangiectasia mutated) and ATR (ATM-Rad3-related) (90). ATM is present in an inactive state and, in response to DNA damage, undergoes auto-phosphorylation at serine 1981 (4). Activated ATM then phosphorylates many downstream effectors, such as p53, NBS1, BRCA1, and SMC1 (90). Phosphorylation of ATM also results in activation of cell cycle checkpoints via the transducing kinases checkpoint kinase 1 (CHK1) and CHK2, which are substrates of ATR and ATM, respectively (6). There also appears to be active cross talk between ATM and ATR as ATM function is required for recruitment of ATR to sites of damage (1). The G₁/S checkpoint response is mediated primarily through a pathway involving ATM/CHK2-p53/MDM2-p21^Waf1/Cip1 (68). ATM directly phosphorylates p53 at serine 15, contributing to its stabilization and increased activity as a transcription factor (12), which in turn leads to activation of a key transcriptional target of p53, the cyclin-dependent kinase (CDK) inhibitor p21^Waf1/Cip1, causing G₁ arrest (58, 59). The G₂ checkpoint primarily targets the cyclin B/CDK1 kinase and is mediated by ATM/ATR, CHK2/CHK1, and/or p38 kinase, which inhibit the CDC25 family of phosphatases required for CDK1 activation (11, 68, 101). The recruitment and/or retention of DNA repair factors at sites of damage is mediated in part by ATM-dependent phosphorylation of the histone variant H2AX at serine 139 (phospho-Ser139 H2AX, or -H2AX), an early event known to occur in response to DNA double-strand breaks (81). H2AX phosphorylation is rapid and extensive, covering megabase chromatin domains adjacent to the break that are easily visualized as discrete foci by immunofluorescence microscopy (74, 82). Phosphorylation of ATM is observed upon radiation-induced DNA double-strand breaks and through other signal transduction events and leads to activation of the G₂/M checkpoint and the double-strand break response pathway (4, 90).

Our lack of knowledge of the mechanism of action of 5-azaC and 5-azadC, in particular the contribution of the cytotoxic DNMT-aza-DNA adduct to drug efficacy, prevents them from being used as effectively as possible. We therefore set out to better characterize the cytotoxic properties of these drugs and to determine the relative contribution of DNMT1 and DNMT3B to this process, since these DNMTs mediate, alone or in combination, most of the DNA methylation in cancer cells, and they are often aberrantly expressed (76, 80). Our results indicate that 5-azadC treatment results in growth inhibition, G₂ arrest, and reduced clonogenic survival. In addition, cells lacking functional DNMT1 are markedly deficient in cell cycle arrest and growth inhibition while DNMT3B-deficient cells generally show an intermediate effect. 5-azadC treatment also leads to robust induction of -H2AX, DNA fragmentation, and activation of DNA repair proteins in both the ATM and ATR pathways, such as CHK1 and CHK2, NBS1, BRCA1, and RAD51, and their relocalization to a focal staining pattern characteristic of cells with damaged DNA. Interestingly, DNMT1 hypomorphic mutant cells (1KO) were profoundly deficient in their ability to induce -H2AX and also exhibited altered induction of other DNA repair proteins. Further demonstration of the particularly critical role for DNMT1 was the finding that DNMT1 strongly colocalized with -H2AX in 5-azadC-treated cells, demonstrating for the first time the presence of DNMT1 at sites of DNA damage in 5-azadC-treated cells. We have also identified a novel interaction between DNMT1 and CHK1. The defective damage response of 1KO cells is, at least in part, due to altered subcellular localization of CHK1 in the absence of functional DNMT1. Furthermore, other clinically relevant DNMT inhibitors also cause induction of -H2AX staining and DNA fragmentation in a manner roughly proportional to their ability to reactivate aberrantly silenced genes. These studies therefore greatly enhance our understanding of how human tumor cells respond to 5-azadC and also reveal an important new role for DNMT1 as a component of the DNA damage response system.

	MATERIALS AND METHODS

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Cell lines, transient transfection, and quantitative reverse transcriptase PCR (RT-PCR). HeLa and parental HCT116 cells were purchased from the American Type Culture Collection. Isogenic HCT116 cell lines with knockouts of DNMT1 (1KO, a hypomorph) and DNMT3B (3BKO) were provided by Bert Vogelstein. Cells were cultured in McCoy's 5A medium (Cellgro Mediatech Inc.) supplemented with 10% heat-inactivated fetal bovine serum and 2 mM L-glutamine (Invitrogen) at 37°C in a 5% CO₂ incubator. The isogenic cell lines YZ5 and EBS are simian virus 40-transformed fibroblasts derived from an ataxia telangectasia patient; EBS cells are ATM deficient, and YZ5 cells stably express full-length wild-type ATM (105). EBS and YZ5 (Coriell Cell Repositories) were grown in Dulbecco's modified Eagle medium (Cellgro Mediatech Inc.) supplemented with 10% fetal bovine serum and 100 µg/ml G418. For immunofluorescence staining experiments, cells were transfected using the TransIT LT1 transfection reagent (Mirus) according to the manufacturer's protocol. Forty-eight hours later, transfected cells were trypsinized and replated at a 1:3 dilution in wells containing glass coverslips and then treated with 10 µM 5-azadC for 48 h.

For quantitative RT-PCR, 2 µg of RNA was reverse transcribed using Superscript III reverse transcriptase according to the manufacturer's protocol (Invitrogen). One microliter of cDNA was used per PCR. Real-time PCR was performed using the Bio-Rad MiniOpticon Real-Time PCR System (Bio-Rad Laboratories). Each 20-µl reaction mixture consisted of 1x Sybr Green PCR Master Mix (Applied Biosystems) and 200 nM concentrations of forward and reverse primers for either WIF1 (WIF1F, 5'-CAACCGTCAATGTCCCTCTGCT-3'; WIF1R, 5'-TCCACTTCAAATGCTGCCACC-3') or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (described previously in reference 97). The cycling conditions were as follows: 1 cycle at 95°C for 10 min and either 40 (WIF1) or 30 (GAPDH) cycles at 95°C for 30 s, followed by 65°C (WIF1) or 60°C (GAPDH) for 30 s. The threshold cycle (C_T) values were determined using the Opticon Monitor 3 software (Bio-Rad Laboratories). WIF1 expression was normalized to GAPDH expression (2^{[Ct(GAPDH) – Ct(WIF1)]}), and relative expression of treated versus untreated cells was determined by the ratio of normalized expression values for the treated cells to the normalized expression value of the calibrator (untreated cells).

Chemicals and plasmids. 5-azaC, 5-azadC, ZEB, bleomycin, hydroxyurea, doxorubicin, and wortmannin were purchased from Sigma. Plasmids expressing green fluorescent protein (GFP)-tagged DNMT1 and Dnmt3b1 have been described elsewhere (28, 77). The FLAG-CHK1 plasmid was provided by Yolanda Sanchez. General chemicals were purchased from Sigma. Protease inhibitors were purchased from Roche. 5-aza, 5-azadC, and ZEB were dissolved in 1x phosphate-buffered saline (PBS; pH 7.0). Cells were treated with 0.1 to 10 µM 5-azadC or 5-azaC and 50 to 500 µM ZEB for the times and doses described in Results. For recovery experiments, cells were treated with 5-azadC for 48 h (fresh drug was added every 24 h), after which the medium was changed. Cells were fixed or harvested and analyzed every 24 h for up to 4 days after drug removal. Cells were treated with 10 µg/ml bleomycin for 4 h, the medium was changed, and the cells were analyzed 20 h later. Wortmannin was used at a final concentration of 10 µM for 24 h of treatment. Hydroxyurea, doxorubicin, and bleomycin treatments for cell viability assays are described in Results.

Growth inhibition. Cells were plated, allowed to grow overnight and treated with 10 µM 5-azadC for 48 h (fresh drug was added every 24 h), and then the medium was changed. Cell growth was then monitored for 4 days by counting live cells following staining with trypan blue. Growth inhibition was calculated as the ratio of the number of viable treated cells versus untreated cells and plotted against the day posttreatment with 5-azadC as follows: [100 – (number of cells in treated sample/number of cells in untreated sample)] x 100.

Clonogenic assay. Cells were plated at a density of 200 to 300 cells per dish and treated with 5-azadC (10^–4 µM, 10^–3 µM, 10^–2 µM, 10^–1 µM, 1 µM, and 10 µM) or vehicle only (control) medium for 48 h, with fresh medium added after 24 h. After completion of the treatments all cells received fresh growth medium. The cells were allowed to grow for the next 10 to 12 days to allow colony formation; then the medium was removed, and the colonies were stained and fixed with 0.2% methylene blue in 50% methanol and counted. Each experimental treatment was performed in quadruplicate.

Cell cycle analysis. Cell cycle profiles were determined by analyzing DNA content using propidium iodide (PI) staining and flow cytometry. Cells were treated with various concentrations of 5-azadC (0.1 to 10 µM) for 24, 48, and 72 h. Cells were harvested by trypsinization, washed with 1x PBS, and fixed with ice-cold 70% ethanol overnight. Fixed cells were washed once with 1x PBS and resuspended at 1 x 10⁶ cells/ml in PI staining solution (0.1 mg/ml PI, 0.1 mg/ml RNase, and 0.1% Triton X-100 in PBS) and incubated in the dark at room temperature for 15 min before analysis. Cell cycle profiles were determined using fluorescence-activated cell sorting (FACS) with a Becton Dickinson FACSort. For each sample, 3 x 10⁴ events were recorded. Data were analyzed by ModFit cell cycle analysis software (Verity) to determine the percentage of cells in each phase. For quantitative assessment of G₂ arrest, a modification of an assay outlined previously was used (101). Briefly, cells were treated with 10 µM 5-azadC for 48 h; then demecolcine (Sigma-Aldrich) was added to the medium (0.4 µg/ml final concentration), and cells were incubated for an additional 24 h. Cells were harvested and fixed in ice-cold 70% ethanol. After fixation, cells were washed twice in 1x PBS, resuspended in 1 ml of 1x PBS containing 0.25% Triton X-100, and incubated on ice for 5 min. Cells were collected by centrifugation, and the cell pellet was resuspended in 100 µl of PBS containing 1% bovine serum albumin and 1 µg of anti-histone H3 phospho-Ser10 (H3 phosphorylated on serine 10) antibody (06-570; Upstate Biotechnology). Cells were incubated with this antibody for 3 h at room temperature, washed, and subsequently incubated with fluorescein isothiocyanate (FITC)-conjugated goat anti-rabbit secondary antibody. After a 30-min incubation at room temperature, cells were washed with PBS, resuspended in PBS containing 25 µg/ml PI and 100 µg/ml RNase A, and incubated for an additional 30 min at room temperature. Both FITC and PI fluorescence were simultaneously measured by flow cytometry.

Cell viability and apoptosis. To assess cell viability following drug treatment, a CellTiter 96 AQ_ueous One Solution kit (Promega) was used, and assays were performed as per the manufacturer's instructions. Cells were plated in 96-well plates, and drug treatments were typically initiated 24 h later. For all drugs tested, parallel cell-free reactions were set up, and the readings from these wells were used for background correction. For all dose-response experiments measurements were performed in quadruplicate. For time course experiments, parallel dishes were set up for each time point for the drug-treated and control cells, and the results were obtained from eight replicates per treatment per cell line. The UV treatments were performed with cells seeded in 96-well plates in a Bio-Rad GS Gene Linker UV chamber (10 mJ/cm²) without removing the growth medium from the plates. The dishes containing the control cells were removed from the incubator and kept outside for the same length of time as the cells receiving the UV treatments. Upon treatment completion, the growth medium was replaced for both control and UV-treated samples. Apoptosis was measured with the Annexin V-PI kit according to the manufacturer's protocol (Trevigen). Briefly, cells were harvested, washed with cold 1x PBS, and resuspended at 1 x 10⁶ cells/ml in 100 µl of annexin V-FITC/PI-containing binding buffer for 15 min in the dark. Apoptotic cells were determined by FACS using a Becton Dickinson FACSort. Data were analyzed using Cellquest software.

Analysis of DNA damage. A single-cell gel electrophoresis system (Comet assay kit; Trevigen) was used to quantitate DNA damage. Briefly, cells were harvested by trypsinization and mixed with 1% low-melting-point agarose. A total of 75 µl (500 to 1,000 cells) of the cell suspension was spread on a precoated glass slide and placed at 4°C for 30 min. Cells were lysed by submerging the slides in ice cold lysis solution (10 mM Tris, 100 mM EDTA, 2.5 M NaCl, with 1% Triton X-100 and 10% dimethyl sulfoxide added just prior to use) at 4°C for at least 1 h. After lysis, the slides were placed in freshly made alkaline buffer (300 mM NaOH and 1 mM EDTA, pH >13.0) for 45 min at room temperature. The samples were subjected to electrophoresis in 0.5x Tris-borate-EDTA buffer at 0.5 V/cm for 25 min. After electrophoresis, the slides were rinsed gently with 0.4 M Tris-HCl (pH 7.5), and the DNA was stained with Sybr Green (Molecular Probes). Fluorescently stained nucleoids were scored visually using an epifluorescence microscope (Zeiss Axioplan 3). Images were captured at a magnification of x20. The percentage of cells having a comet-like appearance was calculated by analyzing at least 100 cells from each treatment. The ratio of comet-positive cells in the 5-azadC-treated versus the mock-treated (1x PBS) sample was plotted against time posttreatment with 5-azadC. The tail moment, a measure of the amount of DNA damage, was calculated using CometScore from TriTek Corporation (a minimum of 50 cells was used for each condition).

Immunofluorescence staining. Cells were grown on 22-mm² glass coverslips in six-well plates and treated as described in Results. Treated and mock-treated cells were fixed with ice-cold methanol for 15 min at –20°C or 2% paraformaldehyde in 1x PBS, pH 7.0 (RAD51 and ATR only). Cells were washed with 1x PBS (pH 7.0), incubated with diluted primary antibody (in 1x PBS with 0.1% Tween-20 [PBST]) for 1 h at room temperature, washed three times with PBST, and subsequently incubated with appropriate fluorophore-conjugated secondary antibody in PBST. Cells were washed and counterstained for nuclear DNA using Hoechst 33342 (1 µg/ml) for 10 min before being mounted onto glass slides using Fluoromount G (Southern Biotech). Images were captured using a Zeiss Axioplan 3 upright fluorescent microscope and deconvolved using Openlab software.

Antibodies. Antibodies for Western blotting and immunofluorescence by source include the following: phospho-Ser139 H2AX (-H2AX), phospho-Ser1981 ATM, p53, CHK2, BRCA1, and phospho-Ser343 NBS1 from Upstate; phospho-Ser1981 ATM from Rockland; phospho-Ser15 p53, CHK2 phosphorylated at threonine 68 (phospho-Thr68), and phospho-Ser345 CHK1 from Cell Signaling; phospho-Ser317 CHK1, RAD51, p21^Waf1/Cip1, ATR, and DNA-dependent protein kinase (DNA-PK) from Calbiochem; DNMT3A (P16), Ku70, CHK1 (G-4), PCNA, and RNA polymerase II (Pol II) from Santa Cruz; anti-FLAG (M2) from Sigma; Mre11 from GeneTex; and GAPDH from Abcam. Epitope affinity-purified rabbit anti-DNMT3B antibody (number 79/80) (29), rabbit polyclonal anti-DNMT1 (74 or N1020) (77), and anti-ATM (pAb 354) (5) have been described previously. Generally, antibodies were used at a 1:50 dilution for immunofluorescence and a 1:500 to 1:1,000 dilution for Western blotting (except GAPDH, which was used at a 1:8,000 dilution). FITC-, tetramethyl rhodamine isocyanate-, and Texas Red-conjugated anti-mouse, anti-rabbit, and anti-goat secondary antibodies were from Southern Biotech. Horseradish peroxidase-conjugated secondary antibodies for Western blotting were used at a 1:2,000 dilution (Santa Cruz).

Immunoprecipitation and Western blotting. Typically, immunoprecipitations were performed with 1 to 2 mg of HeLa high-salt nuclear extracts or total cell lysates derived from one 150-mm dish of HeLa cells at 80 to 90% confluence per reaction. The immunoprecipitation controls were done with species-matched normal immunoglobulin G (IgG) purchased from Pierce (rabbit and mouse) or Santa Cruz (goat). Immunoprecipitations and nuclear extract preparations were performed as previously described (28). Briefly, for immunoprecipitation, the nuclear extract was supplemented with immunoprecipitation dilution buffer to a final concentration of 50 mM Tris (pH 7.5), 150 mM NaCl, 1 mM EDTA, and 0.5% NP-40 plus protease inhibitors. For each reaction, 200 to 300 µl of nuclear protein (1 to 2 mg) was mixed with immunoprecipitation dilution buffer and precleared with 20 µl of protein A/G agarose (Santa Cruz Biotechnology) for 30 min at 4°C with rotation, followed by centrifugation and collection of supernatants. For DNMT1 immunoprecipitations, each tube received 20 µl of DNMT1 (antibody 74) (79) or 10 µl of CHK1 (G-4; Santa Cruz) antibody, and reaction mixtures were incubated overnight at 4°C with rotation. Immune complexes were collected the next day by incubation with 20 µl of protein A/G beads for 2 h at 4°C with rotation. Washes were performed five times in 500 µl of wash buffer (50 mM Tris [pH 7.5], 150 mM NaCl, 1 mM EDTA, 0.5% NP-40, protease inhibitors) at 4°C for 5 min each. For whole-cell extracts, cells were washed with cold 1x PBS, and 0.5 to 1 ml of radioimmunoprecipitation assay buffer was added to each plate. Cells were scraped, collected, and incubated on ice for 30 min. Cells were then centrifuged at 12,000 rpm for 15 min. The supernatant was collected, and the protein concentration was determined using the Bio-Rad protein assay reagent. An equal amount of protein (60 µg) from each sample was loaded on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels, transferred to polyvinylidene difluoride membrane, and used for Western blotting by standard methods.

Subcellular fractionation. Fractionation was performed as described previously (60). Briefly, cells were scraped in 1x PBS, collected by brief centrifugation, and counted. Equal numbers of viable cells were suspended in 100 µl of solution A (10 mM HEPES [pH 7.9], 10 mM KCl, 1.5 mM MgCl₂, 0.34 M sucrose, 10% glycerol, 1 mM dithiothreitol, protease and phosphatase inhibitors) per 2 x 10⁶ cells. Triton X-100 was added to a final concentration of 0.1%, and the cells were incubated on ice for 5 min. The cytoplasmic fraction was harvested by centrifugation at 1,300 x g for 4 min. The nuclear pellet was washed once with solution A and lysed in 100 µl of solution B (3 mM EDTA, 0.2 mM EGTA, 1 mM dithiothreitol, protease and phosphatase inhibitors) per 2 x 10⁶ cells by incubating at 4°C with rotation for 15 min. The soluble nuclear fraction was harvested by centrifugation at 1,700 x g for 4 min. The chromatin pellet (P3) was then washed once in solution B, centrifuged at 10,000 x g for 1 min, and resuspended in 100 µl of sample buffer per 2 x 10⁶ cells. The P3 fraction was homogenized by sonication three times for 30 s each time.

	RESULTS

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Effects of 5-azadC on DNMT protein levels and cell growth. The effects of 5-azadC on DNMT protein levels (30, 71, 100) and cell growth (8, 49, 62, 88) have been examined by others in several different cell lines and experimental systems. As a starting point for defining the role of individual DNMTs in mediating the cytotoxic effects of 5-azadC, we determined its effects on soluble DNMT protein levels in HeLa cells. As has been reported (30, 100), treatment of cells with an increasing concentration of 5-azadC (0.1 to 10 µM) resulted in dose-dependent depletion of DNMTs from soluble cell extracts derived from treated HeLa cells. DNMT1 was affected to the largest extent, followed by DNMT3A and DNMT3B, which showed depletion only at the highest 5-azadC doses (Fig. 1A). Similar results were obtained with HCT116 cells (data not shown).

View larger version (47K): [in this window] [in a new window]   FIG. 1. Responses of HeLa and HCT116 cells to the DNA methylation inhibitor 5-azadC and the role of DNMT1 and DNMT3B. (A) Effect of increasing concentrations of 5-azadC on DNMT1, DNMT3A, and DNMT3B protein levels. HeLa cells were mock treated (–) or treated with 0.1, 0.5, 1.0, 5.0, and 10.0 µM 5-azadC for 48 h (indicated by the wedge); then soluble protein extracts were prepared and subjected to SDS-PAGE followed by Western blotting with the antibodies indicated at the left. GAPDH served as a loading control. (B) Effect of 5-azadC on cell viability. Cells were treated with indicted doses of 5-azadC for 48 h (fresh drug added every 24 h); the medium was changed, and then cell viability was determined using the MTT assay. (C) 5-azadC treatment results in decreased clonogenic survival. Cell lines were treated with 5-azadC for 48 h, followed by replacement of the medium and continued growth for 10 to 12 days. Results are presented as the average of quadruplicate measurements, and the bar is the standard deviation. (D) Effect of increasing doses of 5-azadC on the cell cycle summarized as the relative increase in the number of cells (n-fold) arrested in G2/M for each of the cell lines following drug treatment. Experiments were repeated three times and averaged. (E) HeLa cells treated with 10 µM 5-azadC for 48 h preferentially arrest in G2 as determined by DNA content (PI staining shown on the x axis) and staining cells with an antibody for the M phase specific marker phospho-Ser10 histone H3 (y axis) followed by flow cytometry.

Role of individual DNMTs in mediating the growth effects of 5-azadC on human tumor cells. We next sought to extend our work with HeLa cells to an experimental system in which we could directly examine the effects of individual DNMTs in mediating the growth-suppressive properties of 5-azadC. We chose to use the well-studied HCT116 colorectal cancer cell line because of the availability of isogenic variants in which the DNMT1 or the DNMT3B genes have been disrupted by homologous recombination (76). Recent studies have revealed that the DNMT1-deficient HCT116 line is actually a hypomorph, expressing significantly reduced levels of a DNMT1 protein lacking its PCNA binding domain (22). Previous studies employed murine embryonic stem cell lines carrying deletions in DNMT genes (44, 71); however, in this study we specifically sought to determine the effects of 5-azadC on human tumor cells, the target of a chemotherapeutic regimen containing 5-azadC. HCT116 parental, HCT116 DNMT1 knockout (1KO), and HCT116 DNMT3B knockout (3BKO) cells were treated with 0.05 to 10 µM 5-azadC for 48 h (fresh drug added every 24 h), and cell viability was assessed. Interestingly, while the parental HCT116 and 3BKO lines behaved similarly to HeLa cells, showing relatively minor reductions in cell viability; the HCT116 1KO cells displayed significantly reduced viability (Fig. 1B). The long-term survival of all three HCT116 lines was reduced with increasing doses of 5-azadC, and overall all lines were more sensitive to 5-azadC than HeLa cells. In contrast to previous experiments utilizing complete genetic knockouts of Dnmt1 in murine embryonic stem (ES) cells (44), however, we found that cells deficient (or hypomorphic) in DNMT1 or DNMT3B were significantly more sensitive to 5-azadC in a clonogenic survival assay (Fig. 1C). Cell cycle analysis showed that HCT116 parental and 3BKO cells arrested preferentially in G₂, like HeLa cells; however, 1KO cells were impaired in their ability to arrest in G₂ upon 5-azadC treatment (Fig. 1D). These results suggest that 1KO cells are markedly defective in their responses to 5-azadC treatment and that their reduced capacity to arrest in G₂ and potentially repair DNA damage may, in turn, contribute to the reduced viability of these cells we have observed. Given that reduced short- and long-term viability and G₂ arrest are characteristic responses of cells treated with traditional DNA damage-inducing chemotherapeutic agents, we next sought to examine in more detail the nature of the cellular DNA damage response to 5-azadC treatment.

5-azadC treatment induces formation of -H2AX foci, a marker of DNA double-strand breaks, in an ATM- and DNMT1-dependent manner. The data presented in the preceding sections clearly implicate DNMT1 in mediating much of the survival and cell cycle-related effects of 5-azadC. Interestingly, lack of a major target of 5-azadC, DNMT1, actually resulted in greater cell death and reduced G₂ arrest (Fig. 1), suggesting that reduced DNMT1 levels and/or the presence of the truncated hypomorphic form of DNMT1 may result in a defective DNA damage response. The type of DNA damage induced in 5-azadC-treated cells and how cells respond to this form of damage have not been well characterized. In order to investigate this in greater detail, we analyzed whether 5-azadC treatment caused induction of a well-known marker of DNA double-strand breaks, -H2AX. H2AX is rapidly phosphorylated by ATM and other PI3K family members (ATR and DNA-PK) in response to DNA double-strand breaks due to naturally occurring endogenous processes, such as V(D)J recombination, or exposure to DNA damaging agents from the environment (75). Treatment of HCT116 cells (Fig. 2A) with 10 µM 5-azadC for 48 h resulted in a large increase in cells positive for -H2AX, as monitored by immunofluorescence staining (cells were scored positive if they contained five or more -H2AX foci). 5-azadC-treated cells exhibited intense -H2AX staining in highly localized foci, a characteristic of many agents known to induce DNA double-strand breaks (26, 82). HeLa and parental HCT116 cells demonstrated a dose-dependent increase in -H2AX staining, with foci readily detectable following a 48-h exposure to 1 and 10 µM 5-azadC (Fig. 2B). Of notable significance, cells deficient in DNMT1 or ATM were completely devoid of a -H2AX response upon 5-azadC treatment (Fig. 2B). DNMT3B-deficient cells displayed an intermediate reduction in -H2AX staining when treated with 5-azadC (Fig. 2B). The lack of -H2AX staining we observed in 5-azadC-treated ATM-deficient cells is in keeping with previously published studies on the role of ATM in phosphorylating H2AX in response to DNA damaging agents (95). Interestingly, the basal level of -H2AX staining in untreated cells (not exposed to drug and grown under standard laboratory conditions) varied significantly, with DNMT1- and ATM-deficient cells showing fourfold or higher levels of staining compared to the respective line's wild-type for each of these proteins (Fig. 2C). In keeping with this result, 1KO cells also undergo apoptosis under standard growth conditions at approximately twice the frequency of all other cell lines (Fig. 2C, inset graph), suggesting that 1KO cells may have a chronically elevated level of DNA damage.

View larger version (25K): [in this window] [in a new window]   FIG. 2. Treatment of cells with 5-azadC results in induction of -H2AX, a marker of DNA double-strand breaks, that is dependent on DNMT1 and the PI3K ATM. (A) Representative immunofluorescence staining result for -H2AX (red) and DNA (blue) in untreated and 10 µM 5-azadC-treated HCT116 cells. A magnified view of the boxed cell is shown in the upper left corner of the 5-azadC-treated cells. (B) Dose-dependent increase in -H2AX-positive HeLa, HCT116 parental, 1KO, and 3BKO cells treated with 5-azadC (left graph). Induction of -H2AX staining in isogenic YZ5 and EBS cell lines following 5-azadC treatment (right graph). Results are plotted as the average relative change in cells showing five or more -H2AX foci compared to untreated cells. Data are graphed as the average increase (n-fold) in level of staining, relative to the respective parental cell lines (set at 1.0). (C) Quantitation of the percentage of -H2AX-positive cells and level of apoptosis (inset graph) in each cell line in the absence of any drug treatment (basal levels) showing that cell lines with reduced or absent induction of -H2AX staining upon 5-azadC treatment (1KO and EBS) exhibit elevated basal levels of -H2AX expression and apoptosis. (D) PI3Ks are required for H2AX phosphorylation in response to 5-azadC treatment. HeLa and parental HCT116 cells were mock treated (UT), treated with 10 µM 5-azadC alone for 24 h (5-azadC), or treated with both 10 µM 5-azadC and 10 µM wortmannin (a general PI3K inhibitor) for 24 h (A+W); then, cells were fixed and stained with a -H2AX antibody, and positive cells were quantitated (cells with five or more -H2AX foci). All values are an average of at least three experiments, and the error bar is standard deviation from the mean.

5-azadC treatment induces DNA double-strand breaks that are recognized and repaired differentially in DNMT-proficient versus -deficient cells. Our previous studies clearly showed that treatment of HeLa and HCT116 cells with 5-azadC results in G₂ arrest and induction of DNA double-strand breaks. We therefore wondered if cells may be accumulating DNA damage and then repairing it. To address this, we treated HeLa and HCT116 cells as shown in the experimental scheme in Fig. 3A. Cells were treated with 10 µM 5-azadC for 48 h (fresh drug added every 24 h), and the medium was changed. In this way, all cells in the population should have gone through at least one round of DNA replication in the presence of 5-azadC. Cells were then allowed to grow for an additional 96 h without drug, during which time growth inhibition and DNMT protein levels were measured. Data obtained from cell counting following trypan blue staining of cells revealed that HeLa cells show a peak of growth inhibition in this treatment scheme at 48 h following 5-azadC removal, while HCT116 cells and the KO lines peak at 72 h after drug removal (Fig. 3B). HCT116 1KO cells displayed reduced growth inhibition at early time points (0 to 48 h after drug removal) but caught up to the other HCT116 lines by 72 h (Fig. 3B). Levels of all DNMT proteins were heavily depleted after the end of the drug treatment but recovered to posttreatment levels by 48 h. DNMT1 protein levels rebounded by 48 h, increased further at 72 h to a level above the untreated cells, and then returned to posttreatment levels again by 96 h (Fig. 3C). We next analyzed -H2AX staining by immunofluorescence at 24-h intervals after 5-azadC removal. Both HeLa and HCT116 parental lines showed a five- to sixfold increase in -H2AX staining that peaked 24 to 48 h following withdrawal of 5-azadC from the medium (Fig. 3D). This level then steadily declined over the remainder of the experiment to nearly the levels of untreated cells at 96 h, suggesting that cells are accumulating DNA damage and then repairing it. The relatively low level of apoptosis, particularly in HCT116 cells (data not shown), and the 48-h treatment time (at least one cell doubling in the presence of drug) support the notion of repair rather than cell death and repopulation of the culture by cells that had not incorporated 5-azadC. Interestingly, similar to the results presented in Fig. 2, loss of normal DNMT1 function resulted in a profound deficiency in -H2AX induction following 5-azadC treatment, whereas cells deficient in DNMT3B showed an intermediate effect (Fig. 3D, right panel).

View larger version (48K): [in this window] [in a new window]   FIG. 3. 5-azadC treatment results in formation of DNA strand breaks that are repaired upon drug withdrawal. (A) Experimental scheme used for the assays in this figure. Cells were treated with 5-azadC for 48 h with fresh drug added each day. The medium was then changed, and the cells were allowed to recover for 4 days. Numbering in all graphs is relative to the day post-drug withdrawal, which starts at zero hours. (B) Growth inhibition of all cell lines after removal of 5-azadC. Results are presented as the average percent growth inhibition ([100 – (number of cells in treated sample/number of cells in untreated sample)] x 100) for quadruplicate experiments. (C) Soluble DNMT protein levels before and during drug treatment and recovery as monitored by Western blotting. Ku70 served as a loading control. Hrs, hours. (D) Time course (or recovery) of -H2AX staining in HeLa cells following withdrawal of 5-azadC (left). Time course of HCT116 parental, 1KO, and 3BKO cells following 5-azadC withdrawal from the growth medium (right). (E) Representative comet assay result showing formation of DNA strand breaks (formation of a "comet tail") in 5-azadC-treated HCT116 cells. The photograph is taken at a magnification of x20. (F) Time course of comet tail formation in HeLa cells (left) or HCT116, 1KO, and 3BKO cells (right) following withdrawal of 5-azadC from the medium. Results are presented as the relative change (n-fold) in cells displaying a comet tail compared to untreated cells. Experiments were repeated at least twice, and the results shown are representative.

5-azadC treatment induces a DNA damage response that is altered in DNMT-deficient cells, particularly in 1KO cells. Our previous studies strongly implicated ATM in the response to 5-azadC-mediated DNA damage. ATM is primarily activated by DNA double-strand breaks caused by ionizing radiation or other radiomimetic agents, whereas ATR is involved in the damage response to replicative stress or other forms of damage that result in formation of single-stranded DNA. Both ATM and ATR phosphorylate a large number of downstream substrates, such as the CHK2 and CHK1 kinases, respectively, which initiate the DNA damage response cascade (90). The exact structural nature of the DNA damage produced in 5-azadC-treated cells is unknown; however, we initiated a more detailed analysis of the cells' response to this damage, which may provide clues as to how cells respond to and repair it. For this purpose, we treated HCT116 parental, 1KO, and 3BKO cells with 10 µM 5-azadC for 72 h and monitored the changes in both the absolute levels of key DNA damage signaling molecules and induction of their active, phosphorylated forms, whenever possible, by quantitative Western blotting. Equal microgram amounts of protein from all cell lines and time points were loaded on SDS-PAGE gels, transferred, probed with a given antibody in a single experiment, and exposed to the same piece of film during Western blotting to allow for comparison of absolute levels of each protein between different cell lines. Upon 5-azadC treatment, we detected a time-dependent increase in the active, phosphorylated forms of ATM (at serine 1981), p53 (at serine 15), and CHK1 (at serines 317 and 345) in HCT116 parental cells (Fig. 4A, left). We also detected increases in the total levels of p53 and p21^Waf1/Cip1 in 5-azadC-treated HCT116 cells, consistent with previous studies in HCT116 and other cell lines (49, 66, 88, 104). The active form of CHK2 (phospho-Thr68) was only weakly induced by 5-azadC treatment. GAPDH levels served as a loading control. Interestingly, cells deficient in DNMT1 or DNMT3B exhibited a number of distinct differences in their DNA damage response compared to parental HCT116 cells. For example, basal and 5-azadC-induced levels of phospho-Thr68 CHK2 increased, relative to parental HCT116, in both the 1KO and 3BKO cells. The 1KO cells stood apart from parental and 3BKO cells in a number of respects. 1KO cells displayed reduced levels and/or delayed activation of phospho-Ser15 p53, phospho-Ser317 CHK1, and phospho-Ser1981 ATM but were constitutively activated for the phospho-Ser345 form of CHK1 (Fig. 4A, middle). The 3BKO cells behaved similarly to the parental HCT116 cells with regard to these markers. One response of 3BKO cells to 5-azadC that was particularly notable was the nearly complete lack of p21^Waf1/Cip1 induction despite an apparently normal p53 response. Total levels of the other PI3Ks, DNA-PK and ATR, were unchanged upon 5-azadC treatment except for ATR levels in 1KO cells and DNA-PK levels in 3BKO cells (Fig. 4A). As a control for these studies, we treated HCT116 cells with bleomycin, an agent known to induce DNA double-strand breaks and mimic many of the effects of irradiation (89). For the case of bleomycin-treated cells, all of the phosphorylated forms of the proteins we studied were robustly induced, and total levels of p53 and p21^Waf1/Cip1 also increased, as would be expected (Fig. 4B). CHK1, although generally considered an ATR target, can be phosphorylated by ATM (65). We therefore monitored CHK1 phosphorylation in 5-azadC-treated ATM-negative EBS cells. Quantitative Western blotting revealed that CHK1 was indeed phosphorylated in ATM-negative cells treated with 5-azadC, further supporting a role for ATR in the 5-azadC-mediated DNA damage response (Fig. 4C). Taken together, this analysis reveals that 5-azadC-induced DNA damage activates components of both the ATM and ATR pathways, suggesting that it involves the direct or indirect generation of DNA strand breaks and that other forms of damage or replicative stress are also created. Importantly, this study also demonstrates that DNMT1 (the loss of which blunts both p53 and alters CHK1 activation) and, to a lesser extent, DNMT3B (the loss of which enhances the CHK2 response and severely impairs p21^Waf1/Cip1 induction) or the adducts they form with 5-azadC play important roles in initiating the DNA damage response to this drug. Differential activation of proteins like ATM, CHK1, and CHK2 may also help explain the marked differences in induction of -H2AX staining we observed in 5-azadC-treated cells.

View larger version (71K): [in this window] [in a new window]   FIG. 4. DNA damage response to 5-azadC, as monitored by quantitative Western blotting, and the role of DNMT1 and DNMT3B. (A) The HCT116 cell lines indicated at the bottom of the Western panel were treated with 10 µM 5-azadC for 0, 24, 48, and 72 h (with fresh drug added every 24 h), and whole-cell extracts were prepared. Equal microgram amounts of extract were loaded onto SDS-PAGE gels, transferred to polyvinylidene difluoride membrane, and probed with the antibodies indicated at the right. Note that all cell lines and treatments were run on gels, probed with antibody, and exposed to film at the same time to allow for accurate and quantitative comparison between cell lines for a given antibody. GAPDH served as a loading control. (B) HCT116 cells untreated or treated with 10 µM 5-azadC (aza) for 48 h or 10 µg/ml bleomycin (bleo) for 4 h. (C) ATM-deficient cells show induction of the active phosphorylated form of CHK1 (phospho-Ser317) in response to 5-azadC treatment (10 µM), demonstrating involvement of the ATR pathway in response to 5-azadC-mediated DNA damage. Equal microgram amounts of whole-cell extract from EBS and YZ5 cells treated with 5-azadC for 0, 24, 48, and 72 h were used as described in panel A. pS, phospho-serine; pT, phospho-threonine. Hrs tx, hours of treatment; untx, untreated.

View larger version (50K): [in this window] [in a new window]   FIG. 5. 5-azadC treatment induces characteristic relocalization of DNA damage response proteins. Immunofluorescence staining of HeLa cells that were mock treated (UT), treated with 10 µM 5-azadC for 48 h, or treated with 10 µg/ml bleomycin (bleo) for 4 h. Cells were then fixed and stained with the indicated antibodies (red staining) and for DNA (blue staining), and representative images were collected on an upright fluorescence microscope at a magnification of x100. Note the formation of foci in drug-treated cells and the induction of phosphorylated (active) forms of select proteins.

View larger version (43K): [in this window] [in a new window]   FIG. 6. DNMTs relocalize in the presence of 5-azadC, and DNMT1 colocalizes with -H2AX DNA damage foci in 5-azadC-treated cells. (A) HeLa cells were transfected with GFP-DNMT1 or GFP-Dnmt3b1. Cells were then mock treated (untreated) or were treated with 10 µM 5-azadC for 48 h, fixed, and then examined for localization of GFP signal. Both GFP-DNMTs localize diffusely throughout the nucleoplasm and are also concentrated in DAPI-dense regions corresponding to heterochromatin in untreated cells (particularly Dnmt3b; green staining), as has been shown previously by us along with other investigators. Upon 5-azadC treatment, DNMT1 and Dnmt3b1 relocalize, becoming aggregated into more discrete foci and absent from DAPI-dense heterochromatin regions. 5-azadC-treated cells were also stained for -H2AX (red staining), showing the formation of characteristic foci in cells with damaged DNA. Overlaid images (far right) of the red and green signals show that DNMT1 is highly colocalized with -H2AX (yellow signal). (B) DNA staining of untreated and 5-azadC-treated HeLa cells to demonstrate that there is no gross disruption of heterochromatin or nuclear morphology in 5-azadC-treated cells. (C) Colocalization of activated ATM (phosphorylated at serine 1981) and -H2AX in 5-azadC-treated HeLa cells. Bar, 5 µm.

Other clinically relevant nucleoside-based DNMT inhibitors induce DNA damage. While 5-azadC and 5-azaC have been used in the laboratory and clinic for some time, renewed interest in their clinical use, particularly for the treatment of MDS and other leukemias, has increased in recent years (47, 64, 102). ZEB, a related analog, also inhibits DNMTs and induces expression of aberrantly hypermethylated genes, although it is considerably less potent. ZEB is, however, more stable in aqueous solution than 5-azaC and 5-azadC; therefore, there is interest in developing it for clinical use (14, 15, 96). While the demethylating properties of these agents have been directly compared (96), their capacity to induce DNA damage has not. We compared the ability of 5-azaC, 5-azadC, and ZEB to induce the well-known marker of damaged DNA, -H2AX, by immunofluorescence microscopy. Treatment of HCT116 cells with increasing doses of each drug (0.1, 1.0, and 10 µM for 5-azaC and 5-azadC and 50, 250, and 500 µM ZEB) for 48 h (fresh drug added every 24 h) resulted in a dose-dependent increase in -H2AX-positive cells, with 5-azadC being the most effective of the three drugs (Fig. 7A, left graph). The time course of -H2AX induction was also monitored over 3 days after the addition of 10 µM 5-azaC, 10 µM 5-azadC, and 250 µM ZEB. Treatment of HCT116 cells with each of the three agents caused a time-dependent increase in -H2AX staining that peaked between 48 and 72 h for 5-azaC and 5-azadC while ZEB reached its highest level at 72 h of treatment (Fig. 7A, right graph).

View larger version (39K): [in this window] [in a new window]   FIG. 7. Other clinically relevant nucleoside inhibitors of DNA methylation induce DNA damage in a manner related to their ability to demethylate genes. (A) Direct comparison of the ability of 5-azaC, 5-azadC, and ZEB to induce formation of -H2AX foci in HCT116 cells in a dose-dependent (left) and time-dependent (right) manner. Doses (indicated by the wedge) of 5-azaC and 5-azadC used were 0.1, 1.0, and 10 µM, while 50, 250, and 500 µM ZEB was used since it is less potent at inducing demethylation of aberrantly methylated genes. (B) Time- and dose-dependent induction of strand breaks in HCT116 cells treated with the same three drugs as monitored by comet assay. Values are the average of three independent determinations. (C) Calculation of the tail moment, a measure of the degree of DNA damage, from the data in panel B. The upper panel shows the tail moments for the dose-response experiment, and the lower panel shows the tail moments for the time course. –, mock-treated cells or zero hours. (D) Quantitative RT-PCR analysis for expression of the WIF1 gene, which is densely hypermethylated in parental HCT116 cells. RNA prepared from HCT116 cells treated with 10 µM 5-azaC, 10 µM 5-azadC, or 250 µM ZEB was reverse transcribed and PCR amplified with primers for WIF1 and GAPDH. WIF1 expression was normalized to GAPDH expression as described in Materials and Methods, and relative expression of treated versus untreated cells was determined by the ratio of normalized expression values for the treated cells to the normalized expression value of the calibrator (untreated cells) All drug treatments are significantly elevated over values of the mock-treated cells (paired t test, P < 0.04).

Defective DNA damage responses in HCT116 1KO cells are not limited to DNA methylation inhibitors. Our previous experiments indicated that HCT116 1KO cells were defective in their DNA damage response in a number of ways, including altered induction of the activated forms of -H2AX, CHK1, ATM, and p53, as well as reduced G₂ arrest and increased cell death following 5-azadC treatment. Our observations that HCT116 cells lacking a fully functional DNMT1, a major target of 5-azadC, were actually affected to a greater extent than parental or 3BKO cells, coupled with the defects just mentioned, led us to wonder if there was a more generalized defect in the DNA damage response in 1KO cells. To test this idea, we treated HCT116 parental and 1KO cells with four other well-characterized DNA damaging agents, doxorubicin, hydroxyurea, bleomycin, and exposure to UV light and measured cell viability. Following a 24-h drug exposure, 1KO cells showed significantly reduced cell viability, relative to parental HCT116 cells, at doses of 0.3 to 10 µM doxorubicin, 1 to 500 mM hydroxyurea, and 10 to 32 µM bleomycin (Fig. 8A to C). In addition, 1KO cells demonstrated significantly reduced viability following exposure to UV light (Fig. 8D). Since our previous results indicated that activation of CHK1 was markedly altered in 1KO cells after 5-azadC treatment, we used Western blotting of extracts from doxorubicin-treated cells to monitor activation of CHK1 (phosphorylation at both serines 317 and 345). This revealed that both phospho-Ser317 and phospho-Ser345 CHK1 were activated in parental HCT116 cells (Fig. 8E), as would be expected and consistent with the 5-azadC results presented earlier (Fig. 4A). In contrast, 1KO cells demonstrated constitutively activated phospho-Ser345 CHK1 that actually decreased following doxorubicin treatment, while levels of phospho-Ser317 were elevated, and no induction upon doxorubicin treatment was observed (Fig. 8E). We then compared the cell cycle dynamics of HCT116 and 1KO cells following treatment with 1 µM doxorubicin for 24 h. This experiment revealed that parental HCT116 cells arrested predominantly in G₂ and G₁ in response to doxorubicin treatment, consistent with published results (Fig. 8F) (20). 1KO cells, however, were markedly deficient in both G₁ and G₂ arrest after the identical doxorubicin treatment conditions and appeared to accumulate primarily in S phase (Fig. 8F). Taken together, these results show that the defective DNA damage response of 1KO cells is not restricted to agents that impact DNA methylation. Rather, 1KO cells are significantly more sensitive to other agents that directly or indirectly lead to DNA double-strand breaks. This increased sensitivity may, at least in part, be mediated by aberrant activation of CHK1 and subsequent defects in cell cycle checkpoints since CHK1 is critical for G₂ arrest in response to doxorubicin and other DNA damaging agents (37).

View larger version (43K): [in this window] [in a new window]   FIG. 8. Altered cellular responses of HCT116 1KO cells when exposed to other genotoxic agents. (A to D) MTT cell viability assays in parental HCT116 and 1KO cells after exposure to increasing doses of doxorubicin, hydroxyurea, bleomycin, and UV light. All values, presented as the percent viability relative to mock-treated cells, are the average of quadruplicate experiments, and the error bar is the standard deviation. Doxorubicin, hydroxyurea, and bleomycin treatments were for 24 h. (E) CHK1 activation following treatment of parental and 1KO HCT116 cells with 1 µM doxorubicin for the indicated times (in minutes), monitored by quantitative Western blotting. Equal amounts of whole-cell extract were used at each time point, and Western blotting was performed with the antibodies indicated at the right. Thin and thick arrows denote unmodified and phosphorylated forms of CHK1, respectively. (F) Representative cell cycle profiles of HCT116 and 1KO cells exposed to 1 µM doxorubicin for 24 h. The percentage of cells in each phase of the cell cycle is given above the G1, S, and G2/M peaks. Parental HCT116 cells arrest predominantly in G2/M and also in G1, while the G2/M and G1 checkpoints appear to be defective in 1KO cells after doxorubicin treatment. Untx, untreated; tx, treatment; Dox, doxorubicin.

View larger version (42K): [in this window] [in a new window]   FIG. 9. DNMT1 interacts with CHK1 and influences its subcellular distribution after DNA damage. (A) Demonstration that DNMT1 interacts with CHK1 in reciprocal coimmunoprecipitations. In the top panel, a CHK1 antibody (Ab) was used in the immunoprecipitation (IP), followed by Western blotting (WB) with a DNMT1 antibody. Immunoprecipitation with PCNA and p53 antibodies served as positive controls since they are known to interact with DNMT1, while preimmune IgG served as a negative control. In the bottom panel DNMT1 was used as the immunoprecipitating antibody followed by Western blotting with either CHK1 or CHK2 antibody. DNMT1 does not interact with CHK2 under these conditions. (B) Validation that the DNMT1 antibody used in panel A does indeed immunoprecipitate endogenous DNMT1 from HeLa nuclear extract. Untreated HeLa nuclear extract was used for panels A and B. (C) The DNMT1-CHK1 interaction is enhanced after DNA damage. HeLa cells were mock transfected (–) or transfected with GFP-DNMT1 and FLAG-CHK1. After 24 h, doxorubicin (Dox) was added to the cultures as indicated (+), and whole-cell extract was prepared 24 h later and used in immunoprecipitations with a DNMT1 antibody. CHK1 was detected in Western blotting with an antibody directed against FLAG (upper panel). Expression levels of DNMT1 and CHK1 were equal between untreated and drug-treated cultures (input). (D) Subcellular fractionation of untreated and 5-azadC-treated parental (WT) and 1KO HCT116 cells. Each cell line was mock treated (–) or treated with 10 µM 5-azadC for 48 h (+), followed by isolation of the cytoplasmic (S1/Cyto), soluble nuclear (S2/Nu-S), and chromatin-enriched (P/Chrom) fractions. Each protein fraction was used for Western blotting with the antibodies shown at the left. (E) Validation of the fractionation protocol for HCT116 cells by showing the expected localization of Ku70, GAPDH, and Pol II in the nuclear, cytoplasmic, and chromatin-enriched fractions, respectively. Similar results were obtained with the 1KO cells (data not shown).