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. Author manuscript; available in PMC: 2011 Jun 1.
Published in final edited form as: Cancer Res. 2010 May 11;70(11):4470–4480. doi: 10.1158/0008-5472.CAN-09-3028

Inhibition of histone deacetylase in cancer cells slows down replication forks, activates dormant origins and induces DNA damage

Chiara Conti 1,1,2, Elisabetta Leo 1,1, Gabriel S Eichler 1, Olivier Sordet 1,3, Melvenia M Martin 1, Angela Fan 1, Mirit I Aladjem 1, Yves Pommier 1,*
PMCID: PMC2880188  NIHMSID: NIHMS191439  PMID: 20460513

Abstract

Protein acetylation is a reversible process regulated by histone deacetylases (HDACs) that is often altered in human cancers. SAHA (suberoylanilide hydroxamic acid) is the first histone deacetylase inhibitor (HDACi) to be approved for clinical use as an anticancer agent. Given that histone acetylation is a key determinant of chromatin structure, we investigated how SAHA may affect DNA replication and integrity to gain deeper insights into the basis for its anticancer activity. Nuclear replication factories were visualized with confocal immunofluorescence microscopy and with single-replicon analyses conducted by genome-wide molecular combing after pulse labeling with two thymidine-analogues. Additionally, nascent strand real-time polymerase chain reaction (RT-PCR) in the human β-globin locus was used to assess the effects of SAHA on replication fork origin firing. We found that pharmacological concentrations of SAHA induce replication-mediated DNA damage, on the basis of single-cell and single-DNA molecule analyses. Molecular combing indicated slowdown in replication speed along with activation of dormant replication origins in response to SAHA. Similar results were obtained using siRNA-mediated depletion of HDAC3 expression, implicating this HDAC member as a likely target in the SAHA response. Activation of dormant origins was confirmed by molecular analyses of the β-globin locus control region. Our findings indicate that SAHA produces profound alterations in DNA replication that cause DNA damage, establishing a critical link between robust chromatin acetylation and DNA replication in human cancer cells.

Keywords: epigenetic, replication, HDAC, acetylation, DNA repair

Introduction

Abnormalities in DNA replication are a major cause of genomic instability, a hallmark of cancer cells. It is therefore important to understand the mechanisms regulating DNA replication, chromatin structure and cell cycle progression. DNA replication must be coupled with the cell cycle to coordinate the activation of thousands of origins with the velocity of replication forks to fully replicate the genome within each cell cycle (1, 2). In metazoans it is still not clear how origins of replication are selected, and a large number of potential origins remain quiescent. These dormant origins initiate replication when replication forks coming from neighboring origins experience a slowing-down or an arrest (3–6). Stalled forks must be stabilized and processed to avoid inappropriate recombination and genomic instability (7, 8), and these processes involve chromatin remodeling and histone modifications (9–11).

The accessibility of origins to the replication proteins is likely to be influenced by chromatin structure and compaction. Acetylation on lysines facilitates an “open” chromatin status by neutralizing the positive charges of lysines on histones (12). It also plays a regulatory role during DNA replication by facilitating both the removal of histones from the DNA template and the reloading of histone on newly replicated DNA (9, 10, 13–18). In addition, deacetylation of newly incorporated histones plays a major role in chromatin maturation (9, 18, 19).

HDACs [also called lysine deacetylases (KDACs)] and histone acetyl transferases are enzymes that ensure the homeostatic levels of histone acetylation. They also act reversibly on many other proteins in human cells. A recent analysis revealed at least 3600 acetylation sites in 1750 different human proteins (20). Human cells have eighteen zinc-dependent HDACs organized in four classes (21, 22). Class I HDACs are nuclear and consist of HDACs 1, 2, 3 and 8, which are orthologs of the Saccharomyces cerevisiae Rpd3. Yeast cells lacking Rpd3 display a delay in S phase progression, as measured by flow cytometry (23). 2D-gel analysis showed that rpd3Δ mutants have deregulated timing of origin firing, with late origins being activated early (17, 23, 24). The other HDACs (class IIa, IIb and IV) are primarily cytoplasmic and shuttle between the nucleus and cytoplasm. In contrast to the aforementioned zinc-dependent HDACs, class III HDACs (the sirtuins) are NAD-dependent and are not targeted by the HDAC inhibitors. All HDAC are components of large multiprotein complexes (22).

HDAC are important drug targets for cancer and neurodegenerative diseases (Parkinson and Alzheimer syndromes) and there are over 60 clinical trials currently under way. Given their broad regulatory role, HDACs are complex targets for cancer therapy. HDAC inhibitors (HDACi) generally target more than one class of zinc-dependent HDAC and several pathways are likely to be involved in their anti-cancer activity (22). Suberoylanilide hydroxamic acid (SAHA) (25), was the first HDACi approved by the FDA. SAHA is a broad spectrum HDACi targeting both class I and II HDACs. It is marketed as Vorinostat (Zolinza®) for the treatment of cutaneous T cell lymphomas (CTLC) (22). A number of clinical trials are ongoing to explore additional uses for SAHA as an anticancer drug in combination therapies and to evaluate other HDACi (22, 26). Although SAHA is known to induce histone hyperacetylation as early as 4 hours after exposure (27), most studies have focused on SAHA’s ability to induce cell differentiation and apoptosis after long exposures, typically 24 hours or longer (28).

Because of the growing importance of SAHA and HDACi as anticancer therapies (22, 26) and of our interest in DNA replication (1), genomic integrity (29) and molecular pharmacology (30, 31), we investigated the effects of pharmacological concentrations (32) of SAHA on replication fork progression, replication initiation and DNA integrity in human cells. We also studied the contribution of HDAC3 to the effects of SAHA on DNA replication.

Materials and Methods

Cell lines

MCF7 cells and HCT116 cells were cultured in DMEM and MDA-MD-231 cells in RPMI, respectively (Gibco, Carlsbad, CA) with 10% fetal calf serum (FCS) (Gemini Bioproducts, West Sacramento, CA). Human peripheral lymphocytes from healthy donors were obtained from the NIH Blood Bank and maintained in RPMI 1640 with 10% FCS.

siRNA transfections

siRNA was prepared in 750 μl RPMI (80 nM final concentration) and mixed with 750 μl RPMI containing 3 μl Lipofectamine RNAiMAX Reagents (Invitrogen, Carlsbad, CA) in a 6-well plate. The mixture was incubated for 30 min before addition of 105 MDA-MD-231 cells in 1.5 ml of RPMI supplemented with 10% FCS for an additional 72 hours.

Western blotting and antibodies

Cell pellets were resuspended in 100 μl lysis buffer (1% SDS, 10 mM Tris pH 7.4, 40 μl of 25X protease inhibitors [Roche, Indianapolis, IN], 10μl phosphatase inhibitors [Sigma, St. Louis, MO], 1 ml water). Total cell lysates (10–30 μg) were loaded in 4–20% Tris, Glycine gel (Invitrogen, Carlsbad, CA) and transferred to nitrocellulose membrane with a semi-dry apparatus overnight at 4 V. Membranes were blocked in 6% milk, 0.2% Tween20. The γH2AX antibody (Upstate-Millipore, Billerica, MA) was used at a 1:2000 dilution; the H3K9ac antibody (Upstate-Millipore, Billerica, MA) at 1:2500 dilution, and the thymidylate synthetase antibody (Thermo Scientific, Waltham, MA) at 1:100 dilution. Antibodies against ribonucleotide reductase RR1 and RR2 were from Chemicon International (Upstate-Millipore, Billerica, MA) and Santa Cruz (Santa Cruz, CA), respectively. Antibody signals were detected with SuperSignal West Pico Chemiluminescent Substrate (Pierce, Rockford, IL).

Immunofluorescence microscopy

Cells were grown with 1 ml medium in four-well chamber slides (Nalge-Nunc International, Rochester, NY) and staining was performed as described (33). γH2AX antibody was used at 1:2000 dilution followed by anti-mouse Alexa 488 (Invitrogen, Carlsbad, CA) diluted 1:500. Phospho-53BP1-Ser1778 was detected with a primary antibody from Cell Signaling (#2675) diluted 1:400, followed by secondary anti-rabbit Alexa 568 antibody (Invitrogen, Carlsbad, CA).

DNA replication foci were visualized by incorporation of chlorodeoxyuridine (CldU) and iododeoxyuridine (IdU) as described (33). Briefly, cells were pulse-labeled with 100 μM IdU and CldU (Sigma, St. Louis, MO), washed with PBS, fixed with cold 70% ethanol, and stored at 4°C. Primary anti-CldU (Accurate Chemical Scientific, Westbury, NY) and anti-IdU-FITC (BD PharMingen, San Diego, CA) antibodies were diluted 1:200, added to the slides, and incubated in a humid environment for 2 h. Secondary antibodies for CldU and IdU were donkey anti-rat Alexa-594 and goat anti-mouse Alexa-488, respectively (Molecular Probes), used at 1:500 dilution. Images were visualized with a Nikon Eclipse TE-300 confocal microscope (33).

COMET assays

Neutral COMET assays were performed (Trevigen, Gaithersburg, MD) after electrophoresisat 4°C. Tail lengths were measured with Comet IV program and data were transferred into SigmaPlot 9.0 software to generate the Box Plot histogram.

Single DNA molecule analyses of replicons

Molecular combing was performed as described (33, 34). Briefly, at the end of the CldU pulse, trypsinized cells were embedded in low-melting agarose plugs. After digestion with β–agarase (New England Biolabs, Ipswich, MA), DNA was combed on sylanised surfaces (Microsurfaces Inc., Minneapolis, MN) and replicons were detected with anti-IdU and anti-CldU antibodies (33). Images were captured with the software Attovision using the epifluorescence microscope Pathway (Beckton Dickinson, Franklin Lakes, NJ). Signals were measured using ImageJ (open source from NCI, NIH) with custom-made modifications.

Replication of nascent strand protocol

Nascent strand protocol was carried out as previously described with few variations (35). Briefly, MCF-7 cells were treated with 1.25 or 10 μM SAHA for an hour before harvesting. After harvesting cells, we followed our published protocol (35). Nascent strands were then analyzed by real-time polymerase-chain reaction (RT-PCR) for initiation activity near the known replication origin [the initiator of replication (IR)] or at a distal site that usually does not initiate replication [the locus control region (LCR)] on the human β-globin locus.

Results

Short exposure to SAHA induces DNA damage

Because γH2AX (phosphorylated histone H2AX on lysine 139) is selectively activated by DNA damage and is a sensitive biomarker (31), we determined whether SAHA induces γH2AX under conditions where it induces hyperacetylation. Exposure of human breast cancer MCF-7 cells to 1.25 μM SAHA revealed hyperacetylation within two hours, as shown by the increased acetylation of histone H3 at lysine 9 (H3K9ac) (Fig. 1A, left panel). Increased γH2AX was also detectable after 2 hours treatment but the signal was more robust and consistent after 4 hours (Fig. 1A, left panel). To investigate whether the γH2AX induction was dose-dependent, MCF7 cells were treated for 4 hours with increasing SAHA concentrations from 1.25 to 10 μM. On Western blot, H3K9ac increased with increasing SAHA concentration (Fig. 1B, left panel).

Figure 1.

Figure 1

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Time- and concentration-dependent induction of γH2AX in response to SAHA. A, Breast carcinoma MCF-7 cells were exposed to 1.25 μM SAHA for 1 to 24 hours. On the left, representative Western blots showing acetylation of histone H3 on lysine 9 (H3K9ac) and γH2AX induction. Total H2AX was examined in parallel. Tubulin was used as a loading control. On the right, representative images of γH2AX observed by immunofluorescence microscopy in interphase nuclei. Nuclear outlines are shown. B, MCF-7 cells were treated for 4 hours with the indicated SAHA concentrations. On the left, concentration-dependent induction of H3K9ac and γH2AX. Actin was used as a loading control. On the right, representative images of γH2AX foci observed by immunofluorescence microscopy. Nuclear outlines are shown. C, Quantitation of γH2AX induction by SAHA in MCF-7 cells. On the left, quantification of γH2AX foci induced by increasing concentrations of SAHA for 4 hours. In the middle and right, quantification of γH2AX foci induced by 1.5 μM SAHA for the indicated times. Error bars represent the SD of three independent experiments. D, Induction of γH2AX by SAHA in breast carcinoma MDA-231 cells (1.25 μM for 4 hours; left) and in colon carcinoma HCT116 cells treated with 1.25 μM SAHA for the indicated times (right). Experiments have been repeated at least 3 times with consistent results.

To further demonstrate the γH2AX DNA damage response induced by SAHA, we used immunofluorescence microscopy because it is more sensitive than Western blotting, as it can detect an isolated double-strand break (31). Figure 1A–B (right panels) shows that γH2AX was induced in focal pattern, which is typical of cells experiencing DNA-damage response (31). The number of γH2AX positive cells and the number of γH2AX foci per nucleus increased with increasing the SAHA concentration and exposure time (Fig. 1A–C).

Another breast cancer cell line, MDA-MB-231, which is p53 mutant and triple negative for estrogen receptor (ER), progesterone receptor (PR) and epidermal growth factor receptor 2 (Her2), also showed γH2AX induction in response to SAHA (Fig. 1D, left). Similarly, we found induction of γH2AX in human colon carcinoma HCT116 cells (Fig. 1D, right), confirming the induction of DNA damage by SAHA in different cancer cell lines.

SAHA can be released from its binding site on HDAC when removed from treated cells, thus allowing for deacetylation (36). To further characterize the SAHA-induced DNA damage response, histone acetylation and γH2AX were monitored after drug removal following 4-hour treatments with 1.25 or 10 μM of SAHA. Acetylation of H3K9 diminished as early as 20 min after SAHA removal and returned back to normal within one hour (Supplementary Fig. 1A and B). γH2AX levels also decreased within one or two hours (Supplementary Fig. 1C). Immunofluorescence analysis showed that the average γH2AX intensity per positive cell decreased rapidly after drug removal, which correlates with Western blotting results. However, γH2AX-positive cells persisted for at least four hours (Supplementary Fig. 1D), suggesting that some DNA damage repair requires a long time frame.

γH2AX was initially described as a marker for DNA double-strand breaks (DSBs) (37). However, a growing body of evidence suggests that γH2AX can also appear when the cells experience damages other than DSBs (31). To address whether the SAHA-induced γH2AX foci correspond to DSBs, we looked for colocalization of the γH2AX foci with phosphorylated 53BP1 foci, another marker for DSBs. MCF-7 cells were treated for 4 hours with 10 μM SAHA or 1 μM camptothecin [known to induce replication-associated DSBs (38) and used as positive control]. As for camptothecin, phospho-53BP1 colocalized with the γH2AX foci in SAHA-treated cells (Fig. 2A). To test directly for DSBs, we performed neutral COMET assays, which detect broken DNA. COMET tail length and tail moment increased in response to SAHA (Fig. 2B–D), thus confirming the induction of DSBs.

Figure 2.

Figure 2

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Induction of DNA double-strand breaks (DSB) by SAHA.

A, Colocalization of γH2AX foci (middle) with phospho-53BP1 (left) foci in MCF-7 cells treated with SAHA (10 μM) or camptothecin (CPT; 1 μM) for 4 hours. Nuclear outlines are shown with dotted lines. B, Representative images of COMET tails obtained in an untreated sample and in MCF-7 cells treated with 10 μM SAHA for 16 hours. C, Quantification of the COMET tail length in cells treated for 4 or 16 hours with 10 μM SAHA (a.u. = arbitrary unit; see Material and Methods). The assay was repeated three times. The graph shows the result of a representative experiment. The statistical significance was calculated with the Mann-Whitney test. D, Quantification of the COMET tail moment for samples including at least 35 cells each. In the Box Plot the gray area boxes represent the interval where the middle 50% of the data lies and the vertical line in the grey box represents the median. The horizontal bars extending from the boxes encompass the data set within a 95% confidence interval. The dots are outliers.

Replication dependence of SAHA-induced DNA damage

To assess the contribution of DNA replication to SAHA-induced DNA damage, we first tested whether the γH2AX foci colocalized with replication factories (7, 33). Cells were treated with SAHA for 4 hours and pulse-labeled with chlorodeoxyuridine (CldU) to detect replication factories by immunofluorescence microscopy (33). Eighty percent of the γH2AX foci colocalized with replication factories (labeled with CldU), and around 50% of the replication factories were γH2AX-positive in SAHA-treated cells (Fig. 3A).

Figure 3.

Figure 3

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Single cell analyses of DNA replication-associated γH2AX induction by SAHA. A, MCF-7 cells were treated with 1.25 μM SAHA for 4 hours and pulse-labeled with CldU during the last hour of treatment to label replication factories (33) (schematized at top). CldU and γH2AX were detected by immunofluorescence in red and green, respectively. Representative cells treated with SAHA show the colocalization of γH2AX with DNA replication factories (CldU). B, MCF-7 cells were pre-treated with 1μM aphidicolin (APH) followed by 4 hours co-exposure with APH and SAHA (scheme at the top). Bottom, quantification of the γH2AX response to 1.25 or 10μM SAHA in the absence or presence of APH. Bars represent SD for at least 3 independent determinations. C, FACS profile of cells treated for 4 hours with APH or SAHA at the indicated concentrations and pulse-labeled with IdU for the last 5 min of the treatment. The horizontal dotted lines show the decreased incorporation of IdU in APH- and SAHA-treated cells. D, Immunofluorescence analyses of DNA replication dynamics in individual cells treated with SAHA. MCF-7 cells were pulse-labeled for 45 min with IdU, treated for 4 hours or 45 min with SAHA and pulse-labeled with CldU during the last 45 min. IdU and CldU were then detected in green and red, respectively. Representative nuclei from untreated (control) and SAHA-treated cells are shown. Bottom left, Quantification of the CldU/IdU intensity ratio. The control data have been normalized to 1 and the error bar in the SAHA-treated sample represents the SD of three independent experiments.

The functional relationship between DNA replication and γH2AX was further examined under conditions where DNA replication was inhibited with aphidicolin (APH) (38) (see pulse-labeling experiment in Fig. 3C showing reduction of CldU incorporation and accumulation of cells at the G1-S boundary in the presence of APH). Co-treatment with 1 μM APH reduced the γH2AX signal after 4 h exposure to SAHA (Fig. 3B) under conditions where APH by itself did not induce γH2AX foci (7). These results provide further evidence for the induction of replication-associated DNA damage by SAHA.

Because we observed some γH2AX positive MCF-7 cells that did not stain for replication factories, we assessed γH2AX induction by SAHA in non-replicating cells. To do so, we treated post-mitotic circulating human lymphocytes with SAHA and looked for γH2AX by immunofluorescence microscopy (39). On average, two γH2AX foci per cell were visible after six hours treatment with 5 μM of SAHA (Supplementary Fig. 2A). FACS analysis also showed an increase in the percentage of γH2AX positive cells, which was reduced when cells were treated with the transcription-inhibitor flavopiridol (39) (Supplementary Fig. 2B). Together, these results show that SAHA can induce both replication- and transcription-dependent DNA damage. In the remaining part of this study, we chose to focus on the effects of SAHA on DNA replication.

SAHA alters DNA synthesis

Because our data demonstrate that SAHA induces DSBs and activates the DNA damage response in replication factories, FACS analyses with pulse-labeling were performed to monitor DNA replication in SAHA-treated cells. As before, MCF-7 cells were treated for 4 hours with SAHA but replication was also monitored by pulse-labeling with iododeoxyuridine (IdU), a thymidine analog, during the last 5 minutes of the SAHA treatment (see scheme in Fig. 4B). FACS analysis with anti-IdU antibodies detected the cells that were replicating during drug treatment. APH was used as positive control for replication inhibition (Fig. 4C, second panel from left). The FACS and IdU profiles of SAHA-treated cells showed a small but consistent decrease of the intensity of IdU incorporation. The effect was also observed with 3 minute IdU pulses (not shown), and was most obvious at 10 μM SAHA (Fig. 3C, compare IdU signals between the dotted red lines in the right and left panels).

Figure 4.

Figure 4

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Reduction of replication fork velocity by SAHA. A, Cell treatment protocol: MCF-7 cells were treated with 1.5 or 10 μM SAHA for 4 hours; the IdU pulse was performed in the last 20 minutes of treatment; after wash cells have been pulsed with further 20 minutes with CldU. B, Representative image and fork velocity analysis on individual combed molecules from untreated cells (top) and cells treated with 1.5 μM (middle) or 10 μM SAHA (bottom). Fork velocity was measured during the IdU (green) and CldU (red) pulses separately. n: number of signals scored; S.D.: standard deviation. For each treatment, at least three independent combing experiments have been performed, showing consistent results. C, Western blotting analyses demonstrate no detectable effect of SAHA on ribonucleotide reductase large and small subunits (RR1 and RR2, respectively) and thymidylate synthase (TS) after exposure to 10 μM SAHA for the indicated times. Histone hyperacetylation (H3K9ac) and γH2AX induced by SAHA are also shown at the same time points.

Because histone hyperacetylation (H3K9ac) and DNA damage (γH2AX) responses were apparent at 4 hours, and SAHA only minimally reduced deoxynucleotide (dNTP) incorporation under these conditions, we investigated the dynamics of DNA replication by studying the effects of SAHA on replication factories [i.e. the nuclear sites where replication origins fire together (7, 33, 40)]. To visualize those factories, cells were pulse-labeled with IdU for 45 min. After which, the cells were released in fresh medium without or with SAHA for either 4 hours or 45 minutes. During the last 45 min, replication factories were labeled again but this time with CldU (see scheme in Fig. 3D) (33, 41). In untreated cells (control), the same replication factories were labeled with both IdU and CldU when the two pulses immediately followed each other (45 min; yellow dots on the merge images in the right panels in Fig. 3D). On the other hand, each of the two pulses labeled different replication factories when the IdU label was conducted 4 hours before the CldU label because the replication factories labeled during the first pulse had completed their replication at the time of the second pulse. Figure 3D (right panels; 4 h time points) demonstrates that SAHA-treated cells show no detectable inhibition in the activation of new replication factories. However, we noticed a consistent increase in the intensity of the second pulse and the presence of yellow signals in the cells treated with SAHA for 4 hours (Fig. 3D), which suggested that SAHA might affect replication initiation and fork velocity (see below).

SAHA reduces fork velocity

We next tested the effects of SAHA on replication fork velocity and origin firing. To that effect, we used a single DNA molecule approach based on DNA combing (2, 4, 41–43). Asynchronous MCF-7 cells were treated with 1.5 or 10 μM SAHA for 4 hours and pulsed with IdU during the last 20 min of the treatment. After drug removal, cells were washed and pulsed for 20 min with fresh medium containing CldU (Fig. 4A). After combing the genomic DNA, newly replicated regions were detected with specific fluorescent antibodies against IdU and CldU. Figure 4B shows typical signals for three different replicons. The replication signals were then measured and fork velocity calculated for the signals that had similar length for the IdU and CldU signals (41).

Significant shortening of both the IdU and CldU-labeled tracks was observed in the SAHA-treated samples compared to untreated cells. The median velocity in MCF-7 cells went from approximately 0.8 kilobases per minutes (kb/min) in control cells to 0.54 and 0.4 kilobases per minutes in cells treated with 1.5 and 10 μM SAHA, respectively (Fig. 4B). These results demonstrate that SAHA reduces replication fork velocity.

Because previous studies showed decreased thymidylate synthetase (TS) expression after prolonged exposure to high doses of trichostatin A and SAHA (44), and reduction fork velocity upon inhibition of ribonucleotide reductase (RNR) (4), we looked at the levels of TS and RNR upon exposure of MCF-7 cells to SAHA (Fig. 4C). TS and the RNR subunits RR1 and RR2 remained relatively unaffected by SAHA during the 4-hour exposure conditions used to study replication fork velocity. These results are consistent with our prior FACS analysis showing only small reduction of IdU incorporation in MCF-7 cells treated with SAHA under similar conditions (see Fig. 3C). Therefore, the reduction in fork progression observed by molecular combing after 4 hour exposure to SAHA is not due to reduced dNTPs synthesis but rather to effects of SAHA on chromatin.

HDAC3 downregulation also reduces replication fork velocity

Because SAHA is a pan-HDAC inhibitor with multiple protein targets (22), we compared the effects of SAHA to those of histone hyperacetylation by silencing HDAC3 (Fig. 5A), which has recently been shown to be involved in DNA damage control and cell cycle progression (45). IdU incorporation and replication fork velocity were measured in cells transfected for 72 hours with siRNA targeting HDAC3 or with a negative siRNA. FACS analysis showed no detectable effect of HDAC3 silencing on the percentage of cells in S phase cells and dNTP incorporation (Fig. 5B), which is consistent with the small effects of SAHA on dNTP incorporation and cell cycle (see Fig. 3C). On the other hand, fork velocity was significantly reduced by HDAC3 siRNA (0.8 kb/min) compared to untreated controls (1.3 kb/min) (Fig. 5C), which is also consistent with the reduction of fork velocity in response to SAHA (see Fig. 4). These results implicate HDAC3 in the regulation of fork velocity and suggest that HDAC3 is one of the targets of SAHA with respect to replication fork slow down.

Figure 5.

Figure 5

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Reduction of replication fork velocity by HDAC3 downregulation. A, Western blotting showing the down-regulation of HDAC3 by siRNA in MDA-MB-231 cells. B, FACS analysis of cells transfected for three days with a control or HDAC3 siRNA. C, Fork velocity measured in cells treated with a control siRNA (top) and the HDAC3 siRNA (bottom). The median values of measured fork speeds are indicated for the two samples. Similar results were obtained in two independent transfection experiments. Representative image are shown at right.

SAHA treatment and HDAC3 depletion both activate dormant origins of replication

Inter-origin distance is generally positively correlated with fork velocity (2, 4, 41). When the distance between two neighboring origins increases, the speed of the two forks emanating from those origins also increases. Conversely, as fork velocity decreases the distance between origins tends to be shorter (2, 4, 41, 46).

Inter-origin distances were determined by molecular combing (Fig. 6A–B) (41). Figure 6B shows reduction of the average inter-origin distance in cells treated with SAHA compared with untreated cells, indicating that replication origins that do not normally initiate replication in untreated cells are activated in response to SAHA. siRNA-mediated depletion of HDAC3 also showed decreased inter-origin distance (Supplementary Fig. 3), confirming the induction of new origins in response to hyperacetylation.

Figure 6.

Figure 6

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Dormant origins of replication are activated and average inter-origin distance is reduced after SAHA treatment. A, MCF-7 cells were treated with 1.5 or 10 μM SAHA for 4 hours and replication forks were analyzed using a dual pulse with IdU and CldU. B, Inter-origin distance measured on individual molecules from untreated cells (No SAHA), and cells treated with 1.5 μM and 10 μM SAHA. C, Schematic representation of the human β-globin region (LCR: Locus Control Region) and position of the primers used for the RT-PCR. D, Origin activity analyzed in the LCR and human β-globin replicator (hBG) by RT-PCR on nascent strand DNA. The bars represent standard deviation.

To test the possibility that the increased frequency of initiation corresponded to the activation of dormant origins, we measured replication initiation in the well-characterized human β-globin locus by PCR-based quantification of nascent strand DNA (1, 35). The human β-globin locus contains an active origin of replication (IR) encompassing two replicators, Rep-P and Rep-I (Fig. 6C) (1). Nearby, the locus control region (LCR) is a silent region that does not contain active origins and is replicated passively by the forks traveling from IR. Comparison of the replication activity in the Rep-P and LCR region in untreated and SAHA-treated cells was performed by measuring the abundance of short, nascent DNA strands by real-time PCR (47). Figure 6D shows an increased abundance of nascent strands at both the β-globin Rep-P and LCR loci in SAHA-treated cells. Therefore, the open chromatin status induced by SAHA appears to facilitate the activation of dormant replication origins (such as the LCR locus).

Discussion

Here we show that SAHA induces replication-associated DNA damage within four hours of exposure, as demonstrated by the appearance of γH2AX and phospho-53BP1 foci colocalizing with replication foci, as well as by the appearance of COMET-positive cells indicative of DSBs. Those effects of SAHA are induced at therapeutically relevant concentrations, as oral administrations result in plasma concentrations between 1 and 2 μM for approximately 4 hours (32). The DSBs induced by SAHA are detectable within 4 hours of drug exposure and are not the consequence of apoptosis, which is generally observed at least one day after exposure to SAHA (22). Also, the SAHA-induced γH2AX response was characterized by well-defined nuclear foci within replication factories, whereas the γH2AX apoptotic signal does not form foci but rather an intense diffuse peripheral staining of the nuclei, which is referred to as the “apoptotic ring” (48, 49). Moreover, under our experimental conditions (4 hour treatment), we did not detect other common signs of apoptosis like PARP cleavage, the appearance of stress vesicles or detachment of the cells nor change in the sub-G1 area in the FACS cell cycle profiles (even at 24 hours after SAHA treatment) (Supplementary Fig. 4).

DNA replication-dependent damage occurs when replication forks are blocked and not efficiently stabilized and repaired. Fork arrests have been extensively studied with different agents such as hydroxyurea that inhibits ribonucleotide reductase and blocks replication by depriving DNA polymerase of new dNTPs (4), methyl methane-sulfonate (MMS) that reduces fork progression and origin firing by alkylating DNA (50), aphidicolin (APH) that directly inhibits replicative DNA polymerases (4, 7, 11), and camptothecins that induce fork arrest by trapping topoisomerase I and generating replication-associated DSBs with polymerase run-off (33, 38). Those agents, despite their different mechanisms of action, all inhibit DNA replication and induce γH2AX (31). Typically, cells treated with the above replication inhibitors show an enrichment of cells arrested in S and inhibition of thymidine incorporation. By contrast, SAHA induces γH2AX without detectable alteration in cell-cycle progression, without significant reduction of dNTP incorporation, and without alteration of thymidylate synthetase or ribonucleotide reductase suggesting lack of checkpoint response (33). Thus, SAHA-induced DNA replication damage is likely the consequence of chromatin hyperacetylation, which in turn interferes with replication initiation and fork progression.

The similarities between the effects of SAHA and HDAC3 silencing by siRNA indicate the relevance of HDAC3 for the cellular effects of SAHA. Our results are consistent with a recent study (45) showing the induction of S-phase- and replication-dependent γH2AX induction and DNA damage in HDAC3−/− murine embryo fibroblasts. In that study (45), SAHA was also shown to activate γH2AX at 24 hours. Our study extends those findings by showing that γH2AX activation takes place within 4 hours of SAHA exposure and is unrelated to apoptosis. We also performed the first single-molecule analyses of DNA replication in HDAC3-downregulated cells, and demonstrate that HDAC3 downregulation reduces replication velocity and increases origin firing.

Histone acetylation-deacetylation is a dynamic and reversible process that enables chromatin to adapt to DNA replication and transcription. Histone acetylation opens up chromatin by neutralizing the positive charges of lysines and by promoting the binding of bromodomain-containing proteins; both of which enable the loading of DNA processing enzymes. Hyperacetylation facilitates both the removal of histones from the DNA template and the reloading of histone on newly replicated DNA (9, 10, 13–18). Deacetylation of newly incorporated histones also plays a major role in chromatin maturation (9, 18, 19). A recent study demonstrates that a large number of chromatin proteins are acetylated (20). Histones represent only a small fraction of those proteins. Among the chromatin protein relevant to the present study, Mann and coworkers (20) found 52 replication proteins acetylated on 98 sites (including the replication helicases MCM2-6, DNA polymerase, Geminin, Timeless, and Claspin), 26 chromatin remodeling proteins acetylated on 46 sites (including SWI/SNF, NURD, INO80 and NURF), 46 DNA/RNA helicases acetylated on 105 sites, and 132 cell cycle proteins acetylated on 243 sites. Thus, it is likely that the replication defects induced by HDAC3 knockdown and SAHA result from targeting a broad range of chromatin-associated protein complexes.

Fork velocity and inter-origin distance are linearly correlated (2, 4), and it has generally been proposed that velocity regulates firing. Indeed, in the case of dNTP depletion, DNA polymerase and topoisomerase I inhibition, reduction of fork velocity is associated with activation of dormant origins (4, 33, 46). Since SAHA alters replication without affecting ribonucleotide reductase and thymidylate synthetase levels, its effects on fork velocity are likely to be due to chromatin changes rather than by alteration of the dNTP pools. Accordingly, the induction of DSBs and γH2AX foci in replication factories by SAHA suggests SAHA induces DNA damage that arrests replication forks. It is however not excluded that histone hyperacetylation by SAHA can activate dormant origins by facilitating chromatin opening and hyperacetylation of replication complexes (20), which could increase the risk of replication fork encounters leading to their collapse and thereby reduced fork progression.

From a clinical viewpoint, our study demonstrates that DNA damage can take place in response to SAHA and that such DNA damage needs to be considered for the clinical use and future development of HDAC inhibitors.γH2AX could also be used as a convenient pharmacodynamic biomarker for HDAC inhibitors (31).

Supplementary Material

1

Acknowledgments

Financial support: NIH Intramural Program, Center for Cancer Research, NCI, NIH (Grant number: Z01 BC 006150-19LMP). Elisabetta Leo was partially supported by the Bogue fellowship (UCL).

Footnotes

No conflict of interest.

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