Histone H2AX is one of the heteromorphous variants of family of at least eight protein species of the nucleosome core histone H2A (1–3). Induction of DNA double strand breaks (DSBs) in live cells triggers its phosphorylation (4, 5). The phosphorylation is mediated by ATM- (4–7), ATR- (8), and/or DNA-dependent protein kinase (DNA-PK) (9), affects H2AX molecules flanking the DSBs in chromatin, and occurs on Ser 139 (4, 5) at the C terminus. The phosphorylated form of H2AX has been defined as γH2AX (10). Shortly after induction of DSBs by ionizing radiation, the appearance of γH2AX in chromatin can be detected immunocytochemically in the form of discrete nuclear foci (5, 10), each focus being presumed to represent a single DSB (5). The frequency of foci per nucleus, thus, is considered to reflect the incidence of DSBs. Checkpoint and DNA repair proteins such as Rad50, Rad51, and Brca1 colocalize with γH2AX (11). In addition, the translocation of the p53 binding protein 1 (53BP1) to irradiation-induced foci is mediated by H2AX (7–9). However, it was recently shown that while the migration of repair and signaling proteins to DSBs is not abrogated in H2AX (–/–) cells, these proteins fail to form irradiation-induced foci (12). The loss of H2AX in mice leads to genomic instability (13). The H2AX −/− mice are radiation sensitive, growth retarded, and immune deficient (13). H2AX haploinsufficiency compromises genetic integrity, and in the absence of p53, enhances susceptibility to cancer (14). H2AX, thus, appears to be a caretaker of genomic integrity that requires the function of both alleles for optimal protection against carcinogenesis (15).
H2AX is also phosphorylated in healthy, nontreated cells, in response to the formation of DSBs occurring in V(D)J and class-switch recombination during immune system development (16–20), as well as in association with DNA replication (21) in S phase cells. Likewise, DSBs generated in the course of DNA fragmentation in apoptotic cells also induce H2AX phosphorylation (21).
The intensity of γH2AX immunofluorescence (IF) measured by cytometry was reported to correlate with the dose of ionizing irradiation used to induce DSBs (21). However, since normal cells, particularly the cells replicating DNA, express γH2AX (21, 22), to obtain a stoichiometric relationship between DSBs and the intensity of γH2AX IF resulting from drug-induced (DI) DNA damage, compensation must be used to account for the extent of this “programmed” H2AX phosphorylation. Following compensation, the γH2AX IF measured by cytometry offers a sensitive and convenient means to detect and measure DSBs in individual cells (21, 23), and has been proposed as a surrogate for cell killing in viability tests for drugs that generate DSBs (24). These elegant studies by MacPhail et al. (21, 23) and Banath and Olive (24), provided evidence that DSBs induced by radiation can be detected and conveniently measured using multiparameter flow and image cytometry of γH2AX IF.
Inhibitors of DNA topoisomerase I and II (topo1 and topo2) are among the most clinically effective antitumor drugs. They work by stabilizing otherwise transient “cleavable complexes” formed between topo1 or topo2 and DNA (25). In DNA replicating cells this leads to the collision between the progressing DNA replication fork and the stabilized complex, and in turn, to conversion of the complex into secondary lesions that consist of DSBs (25). Collisions with the cleavable complexes also occur during transcription, between the progressing RNA polymerase molecule and the inhibitor-stabilized topo1 or topo2 cleavable complex located on the template strand within the DNA region being transcribed (26). The RNA polymerase collisions, similar to the collisions of the DNA replication fork, are also converted into DSBs (27). In both instances, it is presumed that the secondary DSB lesions are recognized by the cell as lethal and that they trigger apoptosis (25–27). Predominantly S-phase cells undergo apoptosis upon exposure to topo1 or topo2 inhibitors (28–30). In contrast to topo inhibitors, which generate DSBs, cisplatin (CP) induces primarily DNA intrastrand cross-links. Single-strand DNA breaks, however, appear during the nucleotide excision repair of the CP-induced lesions (31, 32).
In a previous study (33), we reported that H2AX was phosphorylated in response to DNA damage induced by topo1 (camptothecin [CPT] and topotecan [TPT]) and topo2 (mitoxantrone [MTX]) inhibitors. Furthermore, using multiparameter cytometry, we correlated the induction of γH2AX with cell cycle position. We also observed that induction of γH2AX in response to DNA lesions preceded apoptosis, as detected by caspase-3 activation and DNA fragmentation (33). The aim of the present study was to develop the means to identify the DI H2AX phosphorylation from the subsequent, apoptosis-associated (AA) phosphorylation of H2AX, which occurs in response to DNA fragmentation during apoptosis. Analyzing activation of caspase-3, the enzyme that activates the DNase that fragments DNA during apoptosis (34), along with induction of γH2AX, and correlating these events with cell cycle position and chromatin condensation (33), all measured by laser scanning cytometry (LSC) (36, 37), or iCyte, the more advanced version of LSC, we have been able to characterize the DI γH2AX IF and AA γH2AX IF in HL-60 cells treated with TPT, MTX, or CP.
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Figure 1 illustrates labeling with γH2AX Ab of HL-60 cells, untreated (Fig. 1A and B) or treated with TPT for 1.5 h (Fig. 1E and F) or 3 h (Fig. 1C and D). The nonapoptotic cells from the untreated culture show minimal γH2AX IF; they are marked with arrows in Figure 1A. The fluorescence intensity of the single apoptotic cell (about 2–4% of cells in HL-60 cultures undergo spontaneous apoptosis) in this field, compared with the nonapoptotic cells, was so strong that the photograph had to be overexposed to show both the apoptotic and nonapoptotic cells. Because of the overexposure, the γH2AX IF in the apoptotic cell appears blue-white rather than green in color. Figure 1C and D show another apoptotic cell, this time in a photograph that was properly exposed. It is quite evident that the intensity of γH2AX IF was uneven, appearing much brighter and forming a ring on the periphery of the condensed chromatin of the nuclear fragments compared to staining in their center.
Interestingly, the cytoplasmic areas not overlapping with DAPI fluorescence (DNA) in the apoptotic cells, i.e., outside of chromatin structures, are distinctly labeled with the γH2AX Ab (Fig. 1C and D). Apparently, during apoptosis, γH2AX dissociates from DNA and is translocated into the cytosol. This may be a consequence of AA DNA fragmentation.
In the case of cells treated with TPT (Fig. 1E and F), two subpopulations of cells, some strongly labeled with the γH2AX Ab, and others showing weak fluorescence (Fig. 1E, arrows) can be identified. The labeling appears to have an uneven, “dotted” pattern, which is more easily discerned in the cells that do not have very strong labeling. The cells showing strong γH2AX IF were generally larger than the weakly labeled ones (Fig. 1E and F).
The scattergrams shown in Figures 2 and 3 (parts A and B, in both figures) reveal the cell cycle phase specificity of γH2AX IF following HL-60 cell treatment with TPT and MTX, respectively. The slopes of the line plots in Figures 2C and 3C illustrate the relationship between drug concentration and induction of the γH2AX IF with respect to the cell cycle phase. It is apparent that the increase in γH2AX IF in response to TPT treatment was the most pronounced in S-phase cells. In contrast, following MTX treatment, the highest increase in γH2AX IF was manifest in G1 phase cells. The population of S phase cells, however, was the population that preferentially underwent apoptosis, as detected by the TUNEL assay (Figs. 2D and 3D) in both sets of cultures, regardless of whether they were treated with TPT or MTX.
In the next set of experiments using the multiparameter LSC and iCyte, we correlated, in the same cells, induction of γH2AX with the activation of caspase-3, chromatin condensation and cell cycle phase (Figs. 4–6). Figure 4 shows images of HL-60 cells treated with TPT for 3 h that were differentially immunostained for γH2AX and activated caspase-3 (green fluorescence), while their DNA was counterstained with DAPI. It is quite evident that the cells expressing activated caspase-3 have very strong γH2AX IF. In contrast, the cells with weak γH2AX IF are caspase-3 negative. Large numbers of cells are both caspase-3 and γH2AX negative.
Figure 5 presents the results of an experiment in which γH2AX IF of HL-60 cells, grown in the absence or presence of TPT for 1 and 3 h, was measured by multiparameter LSC, concurrent with DNA content and activation of caspase-3. Thus, activation of caspase-3 could be quantitatively correlated with induction of γH2AX in the same cells, both in relation to cell cycle position. It is quite evident that a subpopulation of predominantly S phase cells showed a moderate increase in γH2AX IF after 1 h of the treatment. As expected, there was no evidence of apoptosis as manifested by caspase-3 activation at that time. A distinct population of cells strongly expressing γH2AX IF (Ap; Fig. 5, bottom left panel) arose after 3 h treatment with TPT, concomitant with the appearance of cells expressing caspase-3 activation (Fig. 5, bottom middle panel). The majority of both γH2AX- and activated caspase-3-expressing cells had an S phase DNA content. By using gating analysis of γH2AX IF and caspase-3 activation, it is evident that nearly all cells demonstrating strong γH2AX IF also had activated caspase-3, i.e., they were undergoing apoptosis. In contrast, relatively few cells that showed a moderate increase in γH2AX IF contained activated caspase-3 (Fig. 5, bottom right).
The direct relationship between expression of γH2AX IF versus caspase-3 activation versus chromatin condensation, all in turn related to the cell cycle position, is illustrated in Figure 6. In this experiment, HL-60 cells were treated with TPT for 3 or 4 h, to examine possible differences between the early apoptotic cells, which prevail after 3 h of treatment with TPT, and the late-apoptotic cells, which appear with increased length of exposure (4 h). Condensation of chromatin, known to occur during apoptosis (35, 41), manifests as increased hyperchromicity of DNA, which in turn is detected by LSC as increased maximal pixel (i.e., concentration) of DNA-associated (DAPI) fluorescence (42, 43). The gating analysis shown in the top panels of Figure 6 revealed that the cells strongly expressing γH2AX IF (Ap; marked in blue in Fig. 6A and D), all had activated caspase-3 (Fig. 6C). After 4 h treatment with TPT, a population of cells with intermediate expression of γH2AX became apparent (marked in red in Fig. 6D). Concurrently, there was an increase in the number of cells with elevated values of DAPI (blue) maximal pixel fluorescence (Fig. 6E). The gating analysis revealed that among the cells with increased DAPI maximal pixel fluorescence (above an arbitrary threshold marked with a dashed line in Fig. 6E) there was a higher percentage of cells with intermediate γH2AX IF (marked in red) than with maximal γH2AX IF (blue; 70% versus 30%). This proportion is reversed among the cells that do not yet show increased chromatin condensation, i.e., with maximal pixel values of DAPI-fluorescence below the threshold indicated in Figure 6. Thus, in the later stages of apoptosis (characterized by increased levels of chromatin condensation), a decrease in γH2AX IF was observed. Interestingly, the progression of apoptosis was also paralleled by a decrease in expression of activated caspase-3. This was evident by the fact that the cells with moderate expression of γH2AX (red) also had distinctly lower expression of activated caspase-3 compared with the blue-marked cells (Fig. 6F).
Administration of the pan-caspase inhibitor z-VAD-FMK concurrent with the cell treatment with TPT precluded the appearance of cells strongly expressing γH2AX (Fig. 7). It did not affect, however, the low level of induction of γH2AX IF by S and a fraction of G1 cells.
In the presence of phosphatase inhibitor calyculin A (44), the TPT-induced increase in γH2AX IF was little affected after 1 h of the treatment, but was markedly higher 3 h after administration of the drug (Fig. 8). After the subtraction of γH2AX IF due to the “programmed” H2AX phosphorylation represented by the untreated (Ctrl) cells, the mean intensity of γH2AX IF was 3.4-, 3.2-, or 2.7-fold higher for the population of G1, S, or G2/M cells compared to the cells treated with TPT alone in the same phases of the cell cycle, respectively.
We also tested whether the antitumor drug that does not directly induce DSBs can affect the expression of H2AX. Such a drug is CP, which is known to generate the DNA intrastrand, and to a lesser degree interstrand cross-links, but no DSBs as the primary lesions (31, 32). Following 1 h treatment with CP, a rather small percentage of cells (<16%) showed γH2AX IF elevated above the control level (Fig. 9). After 3 h, however, the numbers of cells with the increased γH2AX IF was quite substantial, particularly at 1.0 or 2.0 μM concentration of this drug (Fig. 9). However, there was no evidence of the cell cycle–phase specificity in the increase γH2AX IF following treatment with CP.
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We have recently reported (33) that exposure of HL-60 cells to DNA topo1 (camptothecin or TPT), or topo2 (MTX) inhibitor leads to an increase in binding of the γH2AX Ab by the treated cells. This Ab is specific towards the H2AX phosphorylated on Ser-139, and its binding by permeabilized cells, therefore, is being considered to report histone phosphorylation. Phosphorylation of H2AX was shown to occur in response to induction of DSB in chromatin by ionizing radiation (21–24), and, as mentioned, in the case of topo1 or topo2 inhibitor-treated cells, DSB are generated by collisions between the progressing replication fork or RNA polymerase complex, and the topo1 or topo2 stabilized cleavable complex (25–27). The multiparameter flow or laser-scanning cytometry that combines immunocytochemical detection of γH2AX with other parameters such as cell cycle phase, caspase-3 activation, or chromatin condensation, thus, offers a convenient tool to study DNA damage by these inhibitors. Such an approach has been recently used to investigate DSB induced by ionizing radiation (21, 23, 24).
Confirming the observations of McPhail et al. (21), we observed a low level of γH2AX IF in untreated HL-60 cells (Fig. 1). The untreated Jurkat and transitional cell carcinoma T24 cells showed low intensity γH2AX IF as well (not shown). In addition, the foci of γH2AX IF in the untreated cells were of a smaller size than in the drug-treated ones, also in agreement with the observation of these authors (21). For quantitative analysis of DI γH2AX IF, this programmed (or scheduled) level of histone H2AX phosphorylation, which appears to be primarily associated with DNA replication (21), was subtracted from the mean values of the drug treated cells, for each cell cycle phase, separately. Thus, the plots in Figures 2C and 3C represent the DI increase in γH2AX immunofluorescence, for each phase of the cell cycle (see Materials and Methods). This increase is expressed as percent of the IF of the untreated cells in the same phase of the cycle.
Histone content doubles during the cell cycle along with the doubling of DNA. Unlike other proteins, however, whose content may vary in individual cells with respect to DNA content, histone synthesis is coupled with DNA synthesis and therefore the ratio of histone to DNA content remains invariable throughout the cell cycle for all cells (44). As a consequence of their higher histone content, therefore, with the same degree of H2AX phosphorylation (the same percent of phosphorylated H2AX molecules within the total H2AX molecules), the cells in S and G2/M have 1.5 and 2.0 times higher γH2AX IF compared to cells in G1. To assess the degree of H2AX phosphorylation, and thus to make γH2AX IF independent of histone doubling during the cycle, we have normalized the data by presenting them per unit of DNA (histone). This was accomplished by multiplying the mean S-phase and G2/M-phase γH2AX IF of the untreated cells as well as the Δ-γH2AX of the drug-treated cells, by 0.75 and 0.5, respectively (Figs. 2C and 3C). The data shown in these plots, thus, represent not the cellular content of γH2AX in relation to the cell cycle, but after compensating for the change in total H2AX content, the degree of H2AX phosphorylation, representing a ratio of phosphorylated H2AX molecules per total number of H2AX within each cell. In fact, because the background fluorescence, i.e. the fluorescence of the untreated cells, was subtracted, per population of G1, S, or G2/M cells, respectively, the ΔγH2AX IF shown on these plots represents the DI increase in the degree of H2AX phosphorylation.
The following attributes allow one to distinguish the cells with DI H2AX phosphorylation from the cells that have an additional, phosphorylation of this histone, triggered by DNA fragmentation during apoptosis: 1) The DI γH2AX IF is seen very early (during the initial two hours) during the treatment, i.e., well prior to caspase-3 activation, which is the prerequisite for the apoptotic endonuclease activation and DNA fragmentation (34). 2) The intensity of DI γH2AX IF is several-fold lower than the intensity of AA γH2AX IF (Fig. 5). It should be noted, however, that because the intensity of the AA γH2AX IF decreases during progression of apoptosis (Fig. 6), at late stages of apoptosis this attribute may fail to discriminate between DI- versus AA-γH2AX IF. 3) The difference in sensitivity of AA versus DI phosphorylation of histone H2AX to the caspase inhibitor zVAD-FMK; in its presence only DI γH2AX is detected (Fig. 7). 4) The AA H2AX phosphorylation occurs in parallel with the concurrent activation of caspase-3 in the same cells. Multiparameter (activated caspase-3 versus γH2AX IF) cytometry, thus, appears to be the most direct approach to distinguish the cells in which DNA strand breaks (histone H2AX phosphorylation) were induced by the studied drugs, from the cells that have H2AX phosphorylation additionally triggered in response to apoptotic DNA fragmentation.
There is an equilibrium between the rate of phosphorylation of histone H2AX after DNA damage, and its dephosphorylation that occurs when DNA repair progresses (45). We observed that the intensity of γH2AX IF was elevated when the cells were treated with TPT in the presence of the protein (serine/threonine) phosphatase inhibitor calyculin A for 3 h but was essentially unchanged after 1 h of the treatment. These data indicate that while no significant dephosphorylation of H2AX occurs during the first hour of treatment, significant degree of dephosphorylation occurs between 1 and 3 h. Our data conform with the observation of Nazarov et al. (45), who reported that following DNA damage by ionizing radiation 50% of H2AX is already dephosphorylated after 3 h. In fact, by comparing the mean values of γH2AX IF of the cells treated with TPT in the presence and absence of calyculin A for 3 h (Fig. 8), one may conclude that dephosphorylation of H2AX during TPT treatment, at a 3-h time interval, was even more extensive (3.4- to 2.7-fold) than after ionizing radiation, as observed by Nazarov et al. (45). To obtain a measure of the cumulative H2AX phosphorylation (as a yardstick of the total number of DSBs) in response to progressive DNA damage (e.g., as occurs during continuous cell treatment with a DNA damaging drug), one has to incubate the cells in the presence of the protein phosphatase inhibitor, to prevent γH2AX dephosphorylation. It should be noted, however, that calyculin A is cytotoxic and prolonged (>3 h) cell incubation with this inhibitor leads to extensive chromatin condensation followed by apoptosis (unpublished results).
A plateau and a decline in the intensity of γH2AX IF was seen at the higher ranges of concentration of TPT or MTX (Figs. 2 and 3). It is possible that at high concentration these intercalating drugs alter DNA topology by inducing torsional stress on double helical DNA in the closed loops of chromatin (46). Such a change is expected to inhibit progression of the replication forks or RNA polymerase molecules along the DNA molecule, thereby lowering the incidence of their collisions with the cleavable complexes and induction of DSB.
The pattern of response of HL-60 cells to the topo1 inhibitor TPT versus the topo2 inhibitor MTX vis-à-vis cell cycle position was distinctly different. Namely, whereas TPT induced H2AX phosphorylation preferentially in S-phase cells, this selectivity was not apparent in the case of MTX treated cells. In fact, after compensation for the increase in total histone content that occurs during S, the G1 cells demonstrated a higher degree of H2AX phosphorylation than S or G2/M cells after exposure to MTX. Yet, the S-phase cells preferentially underwent apoptosis after treatment with MTX (Fig. 3). This data indicates that, at least in the case of treatment with MTX, notwithstanding the same or even lower frequency of DSB, DNA replicating cells are more prone to undergo apoptosis than G1 or G2/M cells. It appears, therefore, that with a comparable extent of DNA damage, the cells arrested at the G1 or G2/M checkpoint remain at the checkpoint alive for extended period of time, apparently in an attempt to repair the damage, while the S-phase cells succumb to apoptosis.
We observed the increase in phosphorylation of H2AX following the cell treatment with CP, which was more pronounced 3 h after drug administration than after 1 h (Fig. 9). The CP-induced H2AX phosphorylation cannot be attributed to apoptosis, because during the initial 3 h of the treatment no caspase activation was detected (data not shown). Furthermore, the AA rise in γH2AX IF is of much higher degree (Fig. 5) than that observed after treatment (Fig. 8). The observed increase in γH2AX IF in cells treated with CP, thus, was definitely drug-induced. However, in contrast to topo inhibitors, CP cross-links DNA rather than induces DSB (31, 32). It is unlikely, therefore, that the primary lesions induced by CP (DNA cross-links), contributed to the observed H2AX phosphorylation. The repair process of these lesions, however, involves the NER mechanism, known to generate ss DNA breaks. A fraction of ss DNA breaks, in turn, is known to be converted in the cell to ds DNA breaks (47). Furthermore, repair of cisplatin-induced damage also involves NHEJ (48). This mechanism may additionally contribute to the formation of ds DNA breaks and to the observed H2AX phosphorylation. Thus, when the primary drug-induced lesions do not involve ds DNA breaks, but ds DNA breaks are formed during DNA repair, as in the case of CP, analysis of H2AX phosphorylation may indicate efficiency of the repair process.