Generation of S phase-dependent DNA double-strand breaks by Cr(VI) exposure: involvement of ATM in Cr(VI) induction of
-H2AX
Linan Ha1,2,
Susan Ceryak1,3 and
Steven R. Patierno1,4
1 Department of Pharmacology and Physiology, 2 Program in Molecular and Cellular Oncology and 3 The George Washington University Cancer Institute, The George Washington University Medical Center, 2300 I Street NW, Washington, DC 20037, USA
4 To whom correspondence should be addressed at: Department of Pharmacology and Physiology, The George Washington University Medical Center, 2300 I Street NW, Washington, DC 20037, USA. Tel: +1 202 994 3286; Fax: +1 202 994 2870; Email: phmsrp{at}gwumc.edu
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Abstract
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Certain hexavalent chromium [Cr(VI)] compounds are implicated as occupational respiratory carcinogens. Cr(VI) induces a broad spectrum of DNA damage, but Cr(VI)-induced DNA double-strand breaks (DSBs) have not been reported. Previously we found that Cr(VI) activates the ataxia telangiectasia mutated (ATM) kinase. ATM is activated specifically in response to DSBs. Therefore, the objective of this study was to investigate DSB induction by Cr(VI) exposure with the overarching hypothesis that S phase-dependent DSBs are produced by Cr(VI) exposure. To test this hypothesis, normal human fibroblasts were treated with either Cr(VI) or neocarzinostatin (NCS). DSBs were analyzed by both comet assay under neutral conditions, which detects primarily DNA DSBs, and phosphorylation of histone H2AX (
-H2AX) and the resultant formation of nuclear foci, which are considered to be indicative of DSBs. Induction of DSBs was observed after Cr(VI) exposure, however, the Cr(VI)-induced DSBs were abrogated by G1 synchronization. Furthermore, our data showed that Cr(VI)-induced DSBs were only observed in the S phase population, whereas no significant DSBs were observed in Cr(VI)-treated G1 synchronized cells. In contrast, NCS-induced DSBs were equally distributed in all cell cycle phases in both asynchronous and G1 synchronized cells. Moreover, Cr(VI)-induced
-H2AX foci formation was restricted to PCNA-positive cells, whereas NCS-induced
-H2AX foci formed in both PCNA-positive and PCNA-negative cells. These results indicate that Cr(VI)-induced DSBs are S phase-dependent. Finally, our data showed that Cr(VI)-induced
-H2AX production was significantly decreased in ATM/ cells compared with ATM+/+ cells. Taken together, these results suggest that Cr(VI)-induced activation of ATM involves the formation of S phase-dependent DSBs. Examining the mechanism of Cr(VI)-induced DSBs will aid in understanding the interrelated mechanisms of Cr(VI) toxicity and carcinogenesis.
Abbreviations: ATM, ataxia telangiectasia mutated; DSBs, double-strand breaks; EMEM, essential medium with Earle's salts; FBS, fetal bovine serum; ICLs, interstrand cross-links; IR, ionizing irradiation; NCS, neocarzinostatin; PI, propidium iodide; ROS, reactive oxygen species; SSBs, single-strand breaks
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Introduction
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Hexavalent chromium [Cr(VI)] has been shown to be a complex genotoxin and to induce a broad spectrum of DNA damage. The spectrum of structural DNA damage includes DNACrDNA interstrand cross-links (ICLs) (13), CrDNA adducts (4,5), DNAprotein cross-links (5) and DNA single-strand breaks (SSBs) (6,7). Nevertheless, despite the wide range of DNA damage induced by Cr(VI), no evidence has linked Cr(VI) to direct DNA double-strand breaks (DSBs).
Cr(VI)-induced DNA damage is preferentially formed in nuclear matrix-associated DNA (8), which is the site of DNA replication and transcription in cells. DNA DSBs generated during DNA replication have been reported in bacteria (911) and eukaryotic cells (12). Replication fork progression can be impeded by DNA lesions, template blocks or mutations in replication proteins (13). Replication fork stress has been implicated as a major cause of genomic instability (14). A common intermediate in restoring replication forks after stalling is the DSB, which is thought to lead to recombination (14). The critical role of CrDNA interactions, involving the bifunctional adduction of Cr to DNA, in DNA polymerase arrest has been well established (15). One specific Cr(VI)-induced DNA lesion, the ICL, has been shown to interfere with the processivity of both prokaryotic and mammalian DNA polymerases and RNA polymerases (1,3,1618). Cr(VI) exposure has been shown to result in replication arrest (19). However, Cr(VI)-induced DNA DSBs, either induced directly by Cr(VI) or as intermediates during restoration of stalled replication forks, have not been reported.
Previously we found that Cr(VI) activates the ataxia telangiectasia mutated (ATM) kinase and that the physiological targets of ATM, p53 Ser15 and Chk2 Thr68, were phosphorylated on Cr(VI) exposure in an ATM-dependent fashion (19). Furthermore, our data strongly indicate that ATM is a major signal initiator for Cr(VI)-induced apoptosis but also contributes to cell survival by facilitating recovery/escape from terminal growth arrest (19). ATM is a 370 kDa serine/threonine kinase and belongs to a growing family of large proteins that contain the phosphatidylinositol 3-kinase-related domain (phosphatidylinositol 3-kinase-related kinases) (20). ATM protein has been shown to function specifically in multiple biochemical pathways linking the recognition and repair of DNA DSBs to downstream cellular processes, such as activation of cell cycle checkpoints, DNA repair and apoptosis, by phosphorylating numerous substrates (21). The activation of ATM by DNA DSBs has been extensively studied. However, the mechanism of activation of ATM by genotoxic agents not known to directly induce DSBs, such as Cr(VI) (19), CdCl2 (22) and UVA (23), needs further investigation.
H2AX is a member of the histone H2A family (24,25). H2AX has been shown to be involved in the maintenance of genomic stability in response to DNA DSBs (26). H2AX is phosphorylated on Ser139 after exposure to ionizing irradiation (IR) at a phosphatidylinositol 3-OH-kinase-related kinase motif in the C-terminus (27). This phosphorylated H2AX is referred to as
-H2AX.
-H2AX has been found at the site of DSBs in chromosomal DNA (28). Phosphorylation of H2AX and the resultant
-H2AX foci are formed within minutes after IR and are thought to be a sensitive signal for the existence of DNA DSBs (27).
-H2AX has been shown to be essential for recruitment of repair factors, such as the MRN (MRE11, Rad50 and NBS1) complex, RAD51 and BRCA1 (2931). H2AX phosphorylation has been reported to be ATM-dependent following direct DSB induction and ataxia telangiectasia and Rad3-related (ATR)-dependent following replication stress (32,33). However, it is not known whether ATM is involved in the phosphorylation of H2AX due to the DSBs generated during replication stress.
The objectives of this study were to: (i) determine whether there are DSBs generated during Cr(VI) exposure; (ii) clarify the mechanism of DSB induction by Cr(VI) exposure; (iii) investigate the involvement of ATM in Cr(VI)-induced phosphorylation of H2AX. DNA DSBs were studied by
-H2AX production and comet assay under neutral conditions. We showed that Cr(VI) exposure, within a toxicologically relevant dose range, results in generation of S phase-dependent DNA DSBs, which may, in turn, activate ATM. Furthermore, our data indicate the involvement of ATM in the Cr(VI)-induced S phase-dependent generation of
-H2AX.
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Materials and methods
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Cell culture
Normal human dermal fibroblasts (GM03440) and human ATM-null mutant fibroblasts (GM03395) (Coriell Institute, Camden, NJ) were maintained in minimum essential medium with Earle's salts (EMEM) (19). EMEM contained a 2x concentration of essential and non-essential amino acids, vitamins and 20% fetal bovine serum (FBS) (Hyclone Laboratories Inc., Logan, UT). Cells were incubated in a humidified 95% air/5% CO2 atmosphere at 37°C and the medium was replaced every 48 h.
Experimental treatment
Sodium chromate (Na2CrO4.4H2O) (J.T. Baker Chemical Co., Phillipsburg, NJ) was dissolved in deionized H2O and sterilized by passage through a 0.2 µm filter before use. Cells were treated with a final concentration of 0, 3 or 6 µM sodium chromate for the indicated times in complete medium. Neocarzinostatin (NCS) solution (0.6 mg/ml in phosphate-buffered saline) (a generous gift of Dr Irving Goldberg, Harvard Medical School, Cambridge, MA) was added to cells in the dark at a final concentration of 25 (
0.0023 µM) or 100 ng/ml (
0.0093 µM).
Comet assay under neutral conditions
The assay was carried out using a CometAssayTM Kit (Trevigen, Gaithersburg, MD) following the manufacturer's protocol for the neutral comet assay. Briefly, cells were seeded at 106/100 mm2 dish 24 h prior to treatment. Cells were treated with 6 µM Cr(VI) for 1 or 3 h or 100 ng/ml NCS for 30 min. At the indicated time period after the respective treatment, cells were gently harvested. The cells were then suspended in low melting point agarose at 3 x 105 cells/ml. The agarose was applied to CometSlidesTM and allowed to set at 4°C in the dark. After lysis of the agarose-embedded cells in lysis solution (2.5 M NaCl, 100 mM EDTA, pH 10, 10 mM Tris base, 1% sodium lauryl sarcosinate, 0.01% Triton X-100), the slides were placed in a DNA gel electrophoresis unit (Fisher, Pittsburgh, PA) and electrophoresed in TBE, pH 8 (0.089 M Tris, 0.089 M boric acid, 0.003 M EDTA). The samples were then fixed in 70% ethanol and dried overnight before staining with SyBr® Green (Molecular Probes, Eugene, OR) to visualize cellular DNA. SyBr Green stained samples were examined using an I x 70 Olympus inverted fluorescence microscope (Olympus, Lake Success, NY). For quantitation of the relative amount of DNA fragmentation in the different samples, the stained slides were scanned with a Meridian ACAS570 Interactive Laser Cytometer at an excitation wavelength of 488 nm using a laser power of 60 mW and a scan strength of 10%, with a 10% neutral density filter. The pseudocolor fluorescence images were analyzed using the Cell Image program to circumscribe the head and tail regions of each comet and the integrated fluorescence values of each defined area were recorded. The tail:head fluorescence ratio was used as a relative measure of DNA fragmentation for each sample (34). An average of 25 individual comets were scored per sample. The data were expressed as fold respective untreated controls.
Immunostaining of
-H2AX foci
Cells were seeded at 1 x 104 cells/well in 8-well chamber slides (Fisher) and incubated for 24 h prior to treatment. The cells were treated with either 6 µM Cr(VI) or 100 ng/ml NCS for 1 h. After treatment the cells were fixed with 100% methanol. The cells were incubated with a 1:1000 dilution of a polyclonal antibody (Trevigen) which specifically recognizes H2AX when phosphorylated at Ser139 (
-H2AX) at 4°C overnight. Cells were then incubated with goat anti-rabbit secondary antibody conjugated with the fluorochrome Alexa-488 (Molecular Probes) for 1 h at room temperature and viewed with a MRC1024 confocal laser scanning microscope (Bio-Rad, Hercules, CA) with excitation at 494 nm and a 520 nm emission filter.
Cell cycle analysis by propidium iodide (PI) single staining
Cells were seeded at 106/100 mm2 dish and incubated for 24 h. The cells were then rinsed twice with phosphate-buffered saline and incubated with EMEM containing 20% FBS or EMEM medium without FBS for an additional 24 h. The cells were harvested and fixed with 70% ethanol. After fixation, the cells were stained with PI (Sigma, St Louis, MO) and analyzed using a FACsort flow cytometer (BD Biosciences, Palo Alto, CA). Excitation was at 488 nm, with a 620 nm emission filter for PI. The percentage of cells in the G0/G1, S and G2/M phases was determined by Modfit I software (BD Biosciences) with 10 000 cells. The PI fluorescence signal (fluorescence pulse area versus pulse width) was used to exclude doublets and aggregates from analysis (19).
Flow cytometry with PI and
-H2AX double staining
Cells were seeded at 1 x 106/100 mm2 dish and incubated for 24 h. The cells were treated with 0, 1, 3, 6 or 25 µM Cr(VI) for 3 h or 25 or 100 ng/ml NCS for 30 min. After treatment the cells were harvested and fixed with methanol. After fixation, the cells were incubated with a monoclonal anti-
-H2AX antibody (Upstate Biotechnology, Lake Placid, NY) at a 1:500 dilution overnight. The cells were then stained with goat anti-mouse secondary antibody (1:1000 dilution) conjugated with Alexa-488 (Molecular Probes) for 1 h. The cells were stained with PI as described above and analyzed using a FACSort flow cytometer (BD Biosciences). Excitation was at 488 nm and the emission filters used were 530 nm for Alexa-488 and 620 nm for PI. The PI fluorescence signal (fluorescence pulse area versus pulse width) was used to exclude doublets and aggregates from analysis (19).
Immunofluorescence with PCNA and
-H2AX double staining
Cells were seeded at 1 x 104 cells/well in 8-well chamber slides and incubated for 24 h. The cells were then treated with 6 µM Cr(VI) for 3 h or 100 ng/ml NCS for 30 min. After treatment the cells were fixed with methanol and probed with a 1:1000 dilution of polyclonal anti-
-H2AX antibody at 4°C overnight. The cells were washed with phosphate-buffered saline and incubated with a 1:50 dilution of monoclonal anti-PCNA antibody for 2 h at room temperature. The cells were then stained with a 1:4000 dilution of goat anti-rabbit secondary antibody conjugated with Alexa-488 and a 1:1000 dilution of goat anti-mouse secondary antibody conjugated with the fluorochrome Texas Red for 1 h. After secondary antibody staining, the cells were viewed with a MRC1024 confocal laser scanning microscope (Bio-Rad). For Alexa-488 excitation was at 494 nm with a 520 nm emission filter. For Texas Red excitation was at 596 nm with a 615 nm emission filter.
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Results
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Generation of DNA DSBs by Cr(VI) exposure
Cr(VI) exposure results in a broad spectrum of DNA damage, but Cr(VI)-induced DSBs have not been reported. To investigate possible DNA DSBs generated during Cr(VI) treatment, we studied DSBs generated on Cr(VI) exposure using the CometAssayTM under neutral pH conditions, in which no DNA denaturation occurs. Therefore, it primarily detects DNA DSBs. Normal human fibroblasts (GM03440) were treated with 6 µM Cr(VI) or 100 ng/ml NCS (a radiomimetic agent) and analyzed by comet assay at the indicated time points. As shown in Figure 1A, DSBs were indicated by the production of comet tails in both Cr(VI)- and NCS-treated cells. No tail formation was observed in control cells. This result was further confirmed by studying Cr(VI)- and NCS-induced
-H2AX foci formation. Normal human fibroblasts were treated with 6 µM Cr(VI) or 100 ng/ml NCS and probed with anti-
-H2AX antibody at the indicated time points. As shown in Figure 1B,
-H2AX foci formation was observed in both Cr(VI)- and NCS-treated cells.
-H2AX foci formation was restricted to a fraction of the Cr(VI)-treated cells, whereas all cells treated with NCS exhibited
-H2AX foci. No foci formation was observed in control cells. Therefore, the results indicate that DNA DSBs are generated during Cr(VI) treatment.
Cr(VI)-induced DSBs are absent after serum starvation
The observation that
-H2AX foci formation was restricted to a fraction of Cr(VI)-treated cells (Figure 1B) suggested a difference in the mechanism of DSB generation between Cr(VI) and NCS. To investigate whether Cr(VI)-induced DNA DSBs are cell cycle dependent, we conducted a comet assay under neutral conditions using asynchronous and serum-starved GM03440 cells that were 85% arrested in G0/G1 phase. Cells were synchronized by incubation for 24 h in serum-free medium. Cells were then treated with 6 µM Cr(VI) for 3 h or 100 ng/ml NCS for 30 min. As shown in Figure 2A (lower panel), after serum starvation the S phase population decreased from 24% in asynchronous cells to 4% in serum starved-cells. Comet tail formation was observed in 6 µM Cr(VI)-treated asynchronous cells but not in Cr(VI)-treated serum-starved cells. However, when cells were treated with NCS, induction of comet tails was observed in both asynchronous and serum-starved cells. No comet tails were observed in untreated cells, regardless of serum status. As shown in Figure 2B, the tail: head ratio of the comets significantly increased to 3.2- and 7.1-fold of the control in asynchronous cells after 6 µM Cr(VI) and 100 ng/ml NCS treatment (P < 0.05), respectively. However, after serum starvation the tail:head ratio of the comets in Cr(VI)-exposed cells were not significantly different from the control and were significantly decreased compared with that of the asynchronous cells. In contrast, the tail:head ratio remained at 7.0-fold control in NCS-treated serum-starved cells. Therefore, the results show that Cr(VI)-induced comet tails, indicative of DNA DSBs, are abrogated by serum starvation, which suggests that Cr(VI)-induced DSBs are S phase-dependent.

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Fig. 2. Cr(VI)- and NCS-induced DNA damage in asynchronous and serum-starved cells. (A) Normal human dermal fibroblasts (GM03440) were incubated either with medium containing 20% FBS or medium without FBS for 24 h. After treatment with 6 µM Cr(VI) for 3 h or 100 ng/ml NCS for 30 min the cells were harvested and the cell membrane was permeabilized. The DNA damage was assessed by CometAssayTM under neutral conditions as described in the legend to Figure 1. (Lower) The cell cycle profile of asynchronous cells and serum-starved cells by flow cytometry at the time of exposure. (B) The fluorescently stained DNA comets were scanned using a Meridian ACAS570 Interactive Laser Cytometer for quantitative analysis of the extent of DNA damage. The head (cell body) and tail (fragmented DNA) were circumscribed and the integrated fluorescence values of each defined area were recorded for a minimum of 25 comets. The tail:head fluorescence ratio was used as a relative measure of DNA DSBs. Data are expressed as fold respective non-treated control and are the means ± SE of three independent experiments. *Statistically significant difference between asynchronous and serum-starved cells at P < 0.05. #Statistically significant difference versus untreated control at P < 0.05.
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Cr(VI)-induced
-H2AX production is S phase specific and is absent after serum starvation
To further characterize the cell cycle dependence of Cr(VI)-induced DNA DSBs, we studied Cr(VI)-induced
-H2AX production by flow cytometry in the presence of PI staining. Both asynchronous and serum-starved GM03440 cells were treated with 0, 1, 3, 6 or 200 µM Cr(VI) for 3 h and were then doubly stained for
-H2AX and PI. As shown in Figure 3A, the number of
-H2AX-positive cells increased in a dose-dependent manner after Cr(VI) exposure of asynchronous cells. Furthermore, as indicated by PI fluorescence, only cells in S phase had a high
-H2AX content, whereas no
-H2AX-positive cells were observed in cells in the G1 or G2/M phases of the cell cycle, even when asynchronous cells were treated with a very high dose of Cr(VI) (200 µM). However, in serum-starved cells
-H2AX expression induced by low doses (1, 3 and 6 µM) of Cr(VI) was nearly abrogated as the S phase population disappeared. As shown in Figure 3C, the percentage of
-H2AX-positive cells significantly increased to 2.0-, 3.3- and 5.6-fold non-treated control after 1, 3 and 6 µM Cr(VI) exposure, respectively, in asynchronous cells (P < 0.05). However, after serum starvation the percentage of
-H2AX-positive cells remained unchanged, at around 1- to 1.3-fold non-treated control after low dose (16 µM) Cr(VI) treatment and was significantly lower than that in the respective asynchronous cells (P < 0.05). In contrast, Cr(VI)-induced
-H2AX expression in G0/G1 and G2/M cells could only be observed when serum-starved cells were treated with a very high dose of Cr(VI) (200 µM) (Figure 3A). However, when GM03440 cells were treated with 100 ng/ml NCS for 30 min,
-H2AX was equally distributed in all cell cycle phases in both asynchronous and serum-starved cells (Figure 3B).
The dose of NCS used in this study, 100 ng/ml, was equitoxic with 6 µM Cr(VI) exposure, as analyzed by clonogenic lethality (data not shown). However, 100 ng/ml NCS induced a 66-fold increase in
-H2AX induction (data not shown), whereas only a 5.6-fold increase was observed after 6 µM Cr(VI) exposure (Figure 3C). This raises the possibility that cells exposed to a very high dose of Cr(VI), which might cause a greater increase in
-H2AX induction, may also have an equal cell cycle distribution of
-H2AX-positive cells. Likewise, cells exposed to a low dose of NCS, which would cause a similar level of
-H2AX induction as 6 µM Cr(VI), may have a S phase-specific distribution of
-H2AX-positive cells. The first possibility was ruled out as 200 µM Cr(VI) exposure still resulted in an increased
-H2AX content in asynchronous S phase cells, which was 10.7-fold untreated control (data not shown), whereas no
-H2AX-positive cells were observed in cells in the G1 or G2/M phase of the cell cycle (Figure 3A). In contrast,
-H2AX-positive cells were equally distributed in all cell cycle phases when cells were treated with 25 ng/ml NCS, which caused only a 3-fold increase in the percentage of
-H2AX-positive cells when compared with the untreated control (data not shown). This result strongly suggests that the differences between Cr(VI)-induced S phase-specific and the NCS-induced
-H2AX distribution are not due to a dose-related phenomenon.
Cr(VI)-induced
-H2AX foci formation is exclusive to PCNA-positive cells
PCNA is an auxiliary protein of DNA polymerase
that is essential for DNA replication during S phase (35). The level of PCNA is known to be highest in replicating cells (36). Therefore, to further confirm the Cr(VI)-induced S phase-dependent production of
-H2AX, GM03440 cells were doubly stained with both
-H2AX and PCNA. As shown in Figure 4,
-H2AX foci formation was restricted to PCNA-positive cells (Figure 4, middle panel), whereas in the case of NCS
-H2AX foci formed in both PCNA-positive and PCNA-negative cells (Figure 4, lower panel). Therefore, these results indicate that Cr(VI)-induced
-H2AX foci formation is DNA S phase dependent.
ATM is involved in Cr(VI)-induced S phase-dependent generation of
-H2AX
The phosphorylation of H2AX at Ser139 has been linked to both ATR (30,33) and ATM (32). To investigate the involvement of ATM in the induction of
-H2AX during Cr(VI)-induced replication-mediated DNA damage, we compared
-H2AX production in both normal human dermal fibroblasts (GM03440, ATM+/+) and fibroblasts with an ATM null mutation (GM03395, ATM/). Cells were treated with 6 µM Cr(VI) for 3 h, which we have previously shown to induce ATM activation (19), and
-H2AX induction was studied by flow cytometry. As shown in Figure 5, the percentage of
-H2AX-positive cells increased to 5.6-fold untreated control after 6 µM Cr(VI) exposure in ATM+/+ cells. However, the same Cr(VI) concentration induced only a 2.3-fold increase in the percentage of
-H2AX-positive cells in ATM/ cells, which was significantly less than that induced in ATM+/+ cells. We also studied the involvement of ATM in NCS-induced
-H2AX expression. After 30 min treatment with 100 ng/ml NCS the percentage of
-H2AX-positive cells increased to 66.1-fold untreated control in ATM+/+ cells, which was significantly lower at 36.4-fold control in ATM/cells (data not shown). Thus, these results indicate that ATM is required, in part, for both Cr(VI)-induced S phase-dependent formation of
-H2AX and NCS-induced
-H2AX production.
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Discussion
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We have previously shown that Cr(VI) activates ATM kinase (19). ATM kinase has been shown to be activated in response to DNA DSBs, which can be induced by IR or radiomimetic agents, but not by UVB/UVC or base-damaging agents (37), indicating that ATM may act specifically in response to DSBs. While there is no direct evidence for Cr(VI) DSB induction, the activation of ATM kinase by Cr(VI) may be due to either (i) ATM directly sensing ICLs or other lesions, (ii) chromatin structure changes induced by Cr(VI) or (iii) ATM sensing DSBs generated during the repair of Cr(VI)-induced DNA damage. The last hypothesis is supported by recent studies with the alkylating agent MNNG, which suggested that activation of ATM by MNNG was caused by the accumulation of strand breaks induced by the activity of repair-associated endonucleases (38). Alternatively, ATM has been reported to be a sensor of oxidative damage. Some studies have shown reactive oxygen species (ROS) to be generated during Cr(VI) reduction, however, in most of those studies experiments were conducted using extraordinarily high Cr(VI) doses (100 µM2 mM) and already malignant cells (3943). In the present study we employed non-malignant human cells with Cr(VI) doses (16 µM) that have not been shown to produce ROS and are more relevant to human exposure. Furthermore, the cytotoxicity of Cr(VI) within this dose range has been extensively studied by colony formation and apoptosis assays (19). The clonogenicity of a 24 h 1 µM Cr(VI) exposure was 59% of untreated control for ATM+/+ cells (GM03440) and 7% for ATM/ cells (GM03395). Although a 3 h exposure was used in the current study, our previous data have shown that the 3 h Cr(VI) exposure-induced cytotoxicity is
10-fold less than that of a 24 h Cr(VI) exposure (19).
In this study we have shown DSB generation by Cr(VI) using the comet assay under neutral conditions (Figure 1A) and
-H2AX foci formation (Figure 1B). These results support the hypothesis that ATM is activated by DSBs induced during Cr(VI) exposure. However, the observation that
-H2AX foci formation was restricted to a fraction of Cr(VI)-treated cells, whereas, all cells treated with NCS exhibited
-H2AX foci (Figure 1B), suggested a difference in the mechanism of DSB generation between Cr(VI) and the radiomimetic agent NCS.
Replicating cells are thought to be more susceptible to genotoxic agents, possibly because interference with replication fork progression by DNA lesions aggravates the consequences of genotoxic insult (44). Replication fork progression has been shown to be impeded by DNA lesions, DNA secondary structures and DNA-bound proteins (13). The arrested replication forks could be a target of nucleases, thereby converting relatively benign damage such as SSBs or modified bases into potentially cytotoxic DSBs or fixed mutations (14). Indeed, this is supported by recent studies which showed the induction of DSBs by nitrogen mustard treatment in replicating Chinese hamster ovary cells but not in confluent cells (45). Furthermore, Bessho (46) observed production of DSBs at or near the sites of psoralen-induced ICLs during replication. Cr(VI) treatment has been shown to rapidly inhibit DNA replication and secondarily block RNA and protein synthesis (15,18). Moreover, numerous in vitro studies have reported a markedly inhibited ability of Cr-damaged DNA to be replicated by purified recombinant mammalian and bacterial enzymes (1,16).
The inhibition of DNA replication by Cr(VI) could be related to the formation of replication-blocking genetic lesions, i.e. ICLs or bulky proteinCrDNA adducts (47). Therefore, the replication-blocking lesions induced by Cr(VI) may contribute to the generation of DSBs. In the present study, by using both the comet assay (Figure 2) and
-H2AX foci formation (Figure 3), we have shown that Cr(VI)-induced DSBs were nearly abrogated by serum starvation, during which the S phase population decreased from 24 to 4%. Furthermore, our data demonstrate that Cr(VI)-induced
-H2AX production was S phase-specific (Figure 3) and was exclusive to PCNA-positive cells (Figure 4). Indeed, Cr(VI)-induced
-H2AX expression in G0/G1 and G2/M cells could only be observed in serum-starved cells when they were treated with an exceedingly high dose of Cr(VI) (200 µM) (Figure 3A). It is likely that massive cell destruction caused by this high dose in the absence of serum may have facilitated the detection of Cr(VI)-induced DSBs. It has previously been reported that the absence of serum in human cell lines may provide an oxidative environment (48) that exacerbates xenobiotic toxicity. Nevertheless, our data clearly show that, within a toxicologically relevant dose range, Cr(VI)-induced DSBs are DNA S phase-dependent.
Here, we propose a model for production of DNA S phase-dependent DSBs by Cr(VI). As shown in Figure 6A, when the DNA replication machinery encounters a Cr(VI)-induced replication blocking lesion, such as ICLs or bulky proteinCrDNA adducts. DNA polymerase is stalled at the blocking site. This may result in formation of a Y-shaped DNA structure, which may be recognized by a specific endonuclease such as XPF-ERCC1 (46) or Mus81 (49), that, in turn, generates a nick in the template strand resulting in induction of a DSB near the replication-blocking lesion. An alternative model would relate the Cr(VI)-induced DNA S phase-dependent DSBs with SSBs generated during Cr(VI) exposure. The generation of SSBs during Cr(VI) treatment in cultured rodent and human cells as detected by alkaline elution techniques has been well documented and is considered one of the most commonly reported lesions arising from Cr(VI) treatment (6,7). A careful analysis of these studies shows that Cr(VI)-induced SSBs are alkali dependent, raising the possibility that the breaks are generated ex vivo, by the interaction of alkali with some other CrDNA lesions. Recognizing this unresolved question, we nevertheless point out that recent studies have demonstrated that S phase-dependent DSBs could be generated from SSBs. Studies by Kuzminov (50) revealed that SSB interruptions in replicating chromosomes cause DSBs and the DSBs were identified at the sites of SSBs. Furthermore, Merrill and Holm (51) showed the generation of DSBs in mec-srf mutated Saccharomyces cerevisiae, which has an accumulation of SSBs in response to hydroxyurea. Therefore, as shown in Figure 6B, if Cr(VI) exposure produces a SSB in a DNA template, when the DNA replication machinery encounters this SSB in the leading strand template the replication fork collapses, which may, in turn, produce a DSB.

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Fig. 6. Model for induction of DNA S phase-dependent DSBs by Cr(VI). (A) DSBs generated from a blocked replication fork. DNA replication encounters a Cr(VI)-induced replication blocking lesion, e.g. ICLs or bulky proteinCrDNA adducts. DNA polymerase is stalled at the blocking site. A Y-shaped DNA structure will be formed, which is recognized by a specific endonuclease. The endonuclease generates a nick in the leading template strand, resulting in induction of a DSB near the replication blocking lesion. (B) DSBs generated from replication fork collapse. Cr(VI) exposure may induce a SSB in a DNA template. When the DNA replication machinery encounters this pre-existing SSB in the leading strand template, the replication fork collapses, which may, in turn, produce a DSB.
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H2AX is phosphorylated at Ser139 as an early response to DSBs (27). The phosphatidylinositol 3-kinses, ATM, ATR and DNA-PK, have all been shown to be involved in the phosphorylation of H2AX (32,33,52). Numerous studies have been carried out to identify the specific kinase involved in this process. In the present study we have shown that ATM is required, in part, for NCS-induced phosphorylation of
-H2AX (data not shown). This result is consistent with studies by Burma et al., which established ATM as the major kinase involved in the phosphorylation of H2AX in response to DSBs induced by X-rays, NCS and etoposide (32). Furthermore, our data show that although Cr(VI) treatment significantly increased the phosphorylation of H2AX in ATM/ cells, this phosphorylation was significantly lower than that induced in ATM+/+ cells (Figure 5). Thus, these data indicate that ATM is also involved, in part, in Cr(VI)-induced S phase-dependent phosphorylation of H2AX (Figure 5). In contrast, a previous study by Ward and Chen. (33) using immunofluorescent imaging concluded that replication stress-induced
-H2AX after hydroxyurea exposure is ATR dependent but does not require ATM. We have previously shown that Cr(VI) activates ATM, causing p53 Ser15 phosphorylation as early as 1 h after exposure. The ATM dependence of p53 Ser15 phosphorylation was diminished over 24 h exposure, which we attributed to the contribution of ATR in this effect at a later time (19). In the present study we investigated Cr(VI)-induced phosphorylation of H2AX after 3 h Cr(VI) exposure, consistent with our reported Cr(VI)-induced ATM activation. Moreover, our flow cytometric method may provide a more quantitative report of
-H2AX induction versus immunofluorescent imaging alone. Finally, our data demonstrate that ATM is responsible for 60% of the observed
-H2AX production, thus suggesting an additional role of either ATR or DNA-PK in Cr(VI)-induced
-H2AX formation.
In summary, our data suggest that Cr(VI) exposure, within a toxicologically relevant dose range, induces one of the most lethal DNA lesions, DSBs. It further indicates that the generation of Cr(VI)-induced S phase-dependent DNA DSBs may be responsible for the reported Cr(VI)-induced ATM activation (19). Furthermore, our data reveal that ATM is also partially involved in Cr(VI)-induced S phase-dependent generation of
-H2AX. Examining the mechanism of Cr(VI)-induced DSBs will aid in understanding the interrelated mechanisms of Cr(VI) toxicity and carcinogenesis.
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Acknowledgments
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We would like to thank Dr Valerie Hu for her generous help with the comet assay and Dr Robyn Ruffner for her kind help in immunofluorescent imaging. We are grateful to Dr Travis O'Brien for critically reading the manuscript and Therese Lizardo for excellent technical assistance. We thank Dr Irving Goldberg for his generous gift of neocarzinostatin used in this study. This work was conducted in partial fulfillment of the requirements for a PhD degree in Molecular and Cellular Oncology, Columbian Graduate School of Arts and Sciences, The George Washington University (L.H.). L.H. is the recipient of a Scholar-in-Training Award of the 95th AACR Annual Meeting for this research. This work was supported by NIH grants ES05304 and ES09961 to S.R.P.
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Received May 7, 2004;
revised July 14, 2004;
accepted July 18, 2004.