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. 2006 May 2;39(3):231–240. doi: 10.1111/j.1365-2184.2006.00380.x

Synchronization in the cell cycle by inhibitors of DNA replication induces histone H2AX phosphorylation: an indication of DNA damage

A Kurose 1,2,3, T Tanaka 1,4, X Huang 1, F Traganos 1, Z Darzynkiewicz 1
PMCID: PMC6496703  PMID: 16672000

Abstract

Abstract.   Several methods to synchronize cultured cells in the cell cycle are based on temporary inhibition of DNA replication. Previously it has been reported that cells synchronized this way exhibited significant growth imbalance and unscheduled expression of cyclins A and B1. We have now observed that HL‐60 cells exposed to inhibitors of DNA replication (thymidine, aphidicolin and hydroxyurea), at concentrations commonly used to synchronize cell populations, had histone H2AX phosphorylated on Ser‐139. This modification of H2AX, a marker of DNA damage (induction of DNA double‐strand breaks; DSBs), was most pronounced in S‐phase cells, and led to their apoptosis. Thus, to a large extent, synchronization was caused by selective kill of DNA replicating cells through induction of replication stress. In fact, similar synchronization has been achieved by exposure of cells to the DNA topoisomerase I inhibitor camptothecin, a cytotoxic drug known to target S‐phase cells. A large proportion of the surviving cells ‘synchronized’ by DNA replication inhibitors at the G1/S boundary had phosphorylated histone H2AX. Inhibitors of DNA replication, thus, not only selectively kill DNA replicating cells, induce growth imbalance and alter the machinery regulating progression through the cycle, but they also cause DNA damage involving formation of DSBs in the surviving (‘synchronized’) cells. The above effects should be taken into account when interpreting data obtained with the use of cells synchronized by inhibitors of DNA replication.

INTRODUCTION

Often, there is a need to obtain populations of cells synchronized in the cell cycle, and numerous approaches have been developed to accomplish this goal (reviews: Grdina et al. 1987; Merril 1998; Davis et al. 2001; Amon 2002). Each of these methods offers some advantages but also has limitations. One of the classical and widely used approaches is based on selection of mitotic cells that detach from flasks during culturing (Terasima & Tolmach 1963). Its advantage stems from the fact that the procedure does not perturb cell progression through the cycle. However, the technique is applicable to relatively few cell lines, for example, HeLa or Chinese hamster ovary (CHO) cells, that grow attached in cultures. A further technique that selects mitotic cells has been designed for cells that otherwise grow in suspension; their enforced attachment to polylysine‐coated membranes leads to production of daughter cells that detach from the membranes during mitosis (Thornton et al. 2002; , Edward et al. 2004). This method has only recently been introduced and it is unknown whether it can be applied to wide range of cell lines as several lines do not strongly attach to polycation surfaces (Huang et al. 2005a). Some cell lines can be synchronized in G1 by inhibitors of protein farnesylation and geranyl‐geranylation such as statins (Jakóbisiak et al. 1991), or protein kinase inhibitors such as staurosporine (Crissman et al. 1991; Bruno et al. 1992). These techniques also are not universally applicable because many cell lines do not respond to statins or kinase inhibitors by arrest in G1.

Non‐tumour cells can be synchronized in G0 or early G1 by withdrawal of either growth factors (‘serum starvation’) (Griffin 1976), or a particular amino acid (Tobey et al. 1990), or by contact inhibition (Holley & Kiernan 1968). These approaches generally fail to synchronize most tumour lines. Furthermore, the metabolism of cells synchronized this way is often perturbed which affects their rate of progression through the cycle (Pardee & Keyomarsi 1992). It should also be noted that when cell populations are synchronized in early G1 or at mitosis they lose synchrony during progression through G1 and therefore become less synchronous while in S or G2.

Centrifugal elutriation offers the possibility to select uniformly sized cells, and because cell size generally correlates with cell cycle stage, these cells are synchronized with respect to their position in the cycle (Mitchell & Tupper 1977). This procedure, like mitotic detachment, also does not perturb cell cycle progression. It requires, however, rather complex and expensive instrumentation and an experienced operator. Furthermore, the synchrony of cell populations obtained through elutriation is not always sufficiently narrow. Density gradient centrifugation has several problems and generally yields cell populations less synchronized than the elutriation procedure (Cymerman & Beer 1980). Once again complex and expensive instrumentation (a cell‐sorting flow cytometer) is needed to select cells based on differences in their DNA content after staining with Hoechst 33342 (Juan et al. 2002). The dye Hoechst 33342 elicits long‐term toxicity (Zhang et al. 1999), and undergoes redistribution from labelled to unlabelled cells in mixed cell populations (Mohorko et al. 2005).

Synchronization of cell populations at mitosis by using mitotic spindle poisons such as colchicines, vinca alkaloids or nocodazole has been widely used (Samake & Smith 1996; Harper 2005). The advantage of these methods is simplicity, low cost and high degree of synchrony when one desires to obtain mitotic or immediately post‐mitotic cell populations (Darzynkiewicz et al. 1982). As mentioned, however, the populations become less synchronous after progression through G1. Cytotoxicity and other undesirable effects of such synchronization procedures, including growth imbalance, have been reported (Urbani et al. 1995; Chou & Chou 1999; Piotrowska et al. 2000).

The use of agents that inhibit DNA replication such as hydroxyurea (HU), high concentration of thymidine (TH), methotrexate or aphidicolin (AP) are the most commonly used approaches to obtain populations of synchronized cells, particularly in S phase (Tobey & Crissman 1972; Vogel et al. 1978; Fox et al. 1987; Matherly et al. 1989). A combination of these agents with other treatments that synchronize cells is also widely used (Kues et al. 2000; , Harper 2005). The virtue of cell synchronization by DNA replication inhibitors is, similar to mitotic blockers, their simplicity and low cost. A plethora of undesirable side effects, however, plague this methodology. The major drawback of all methods utilizing DNA synthesis inhibitors is the induction of unbalanced growth (Cohen & Studzinski 1967; Urbani et al. 1995). While cells exposed to these inhibitors are arrested in their progression through S, their growth, in terms of RNA and protein synthesis, continues which leads to severe perturbation of metabolic functions. Cell cycle regulatory machinery of cells arrested in this way is also severely perturbed, as reflected by unscheduled expression of cyclin proteins (Gong et al. 1995a).

We have recently observed that inhibitors of DNA replication such as hydroxyurea and aphidicolin induce phosphorylation of histone H2AX on Ser‐139 in S‐phase cells (Kurose et al. 2006). Phosphorylated H2AX is defined as γH2AX and this modification is considered to be an indicator of DNA damage that involves formation of DNA double‐strand breaks (DSBs) (Rogakou et al. 1998; Sedelnikova et al. 2002). The aim of the present study was to explore in more detail the effects of HU, AP, and TH at concentrations commonly used in synchronization protocols, on histone H2AX phosphorylation. For comparison, we also treated cells with the DNA topoisomerase I inhibitor camptothecin (CP), an agent which selectively induces apoptosis of S‐phase cells leaving the live subpopulation of cells ‘synchronized’ in G1 (Del Bino et al. 1992). Multiparameter cytometry was used to correlate the induction of DSBs with the cell's position in the cycle. The data indicate that not only DNA replicating cells but also the cells arrested at the G1/S boundary by these inhibitors have phosphorylated histone H2AX.

MATERIALS AND METHODS

Cell lines and culture conditions

HL‐60 cells were purchased from American Type Culture Collection (ATCC; Manassas, VA, USA) They were grown in 25‐ml Falcon flasks (Becton Dickinson Co., Franklin Lakes, NJ, USA) in RPMI 1640 supplemented with 10% fetal calf serum, 100 units/ml penicillin, 100 µg/ml streptomycin and 2 mm l‐glutamine (all from Gibco/BRL Life Technologies, Inc., Grand Island, NY, USA) at 37 °C in an atmosphere of 5% CO2 in air. At the onset of the experiments, there were fewer than 5 × 105 cells per millilitre in culture and the cells were at an exponential and asynchronous phase of growth. HeLa cells were obtained from ATCC and were grown in Dulbecco's minimum essential medium (DMEM) supplemented with 10% fetal bovine serum, 100 units/ml penicillin, 100 µg/ml streptomycin and 2 mm l‐glutamine (Gibco/BRL) at 37 °C in an atmosphere of 5% CO2 in air. The cultures were diluted and re‐plated every 4 days to maintain them in an asynchronous and exponential phase of growth. For experiments, the cells were trypsinized and seeded in 2‐chambered Falcon Culture Slides (Beckton Dickinson Labware).

Evaluation of DNA damage after using synchronizing drugs

HL‐60 cells were incubated with various concentrations of HU, AP, TH or CP (Sigma Chemical Co., St. Louis, MO, USA) for different time intervals, as described in the figure legends. The cells were then fixed with 1% methanol‐free formaldehyde (Polysciences, Inc., Warrington, PA, USA) in PBS for 15 min on ice followed by suspension in 80% ethanol in which they were stored at −20 °C for 2–24 h.

G1/S cell synchronization of HeLa cells using double thymidine block

HeLa cells were grown in 2‐chambered Falcon Culture Slides to about 40% confluency. TH was added to a final concentration of 2 mm and the cells were incubated for 19 h. Cells were then washed twice with PBS, and incubated with regular medium for 9 h before a second incubation in 2 mm TH for 16 h. The cells were then fixed on slides with 1% methanol‐free formaldehyde in PBS for 15 min on ice followed by suspension in 80% ethanol, where they were stored at −20 °C for 2–24 h.

Immunocytochemical detection of γH2AX phosphorylation

The fixed HL‐60 cells were washed twice in PBS and suspended in 0.2% Triton X‐100 (Sigma) in a 1% (w/v) solution of bovine serum albumin (BSA; Sigma) in PBS for 30 min to suppress non‐specific antibody (Ab) binding. The cells were then incubated in 100 µl of 1% BSA containing 1 : 100 diluted antiphospho‐histone H2AX (Ser‐139) mAb (Upstate, Lake Placid, NY, USA) for 2 h at room temperature, washed twice with PBS and resuspended in 100 µl of 1 : 30 diluted FITC‐conjugated F(ab′)2 fragment of goat anti‐mouse immunoglobulin (DAKO, Carpinteria, CA, USA) for 30 min at room temperature in the dark. The cells were then counterstained with 5 µg/ml PI in the presence of 100 µg/ml of RNase A (Sigma) for 30 min. The fixed HeLa cells were washed twice in PBS and suspended in 0.2% Triton X‐100 (Sigma) in a 1% (w/v) solution of BSA in PBS for 30 min. The cells were then incubated in 100 µl of 1% BSA containing 1 : 100 diluted antiphospho‐histone H2AX (Ser‐139) mAb, and were then incubated for 2 h at room temperature. The cells were washed twice in PBS and incubated with 1 : 30 diluted FITC‐conjugated F(ab′)2 fragment of goat anti‐mouse immunoglobulin for 30 min at room temperature in the dark. The cells were then counterstained with 1 µg/ml 4,6‐diamidino‐2‐phenylindole (DAPI; Molecular Probes, Eugene, OR, USA) in PBS for 10 min.

Fluorescence measurements

Cellular green (FITC) and red (PI) fluorescence of HL‐60 cells in suspension were measured using a FASCcan flow cytometer (Becton‐Dickinson, San Jose, CA, USA). The red (PI) and green (FITC) fluorescence from each cell were separated and quantified using standard optics and CELLQuest software (Becton‐Dickinson). Ten thousand cells were measured per sample. Cellular green (FITC) and blue (DAPI) fluorescence of HeLa cells attached to slides was measured using an iCys laser scanning cytometer (LSC) (CompuCyte, Cambridge, MA, USA) utilizing standard filter settings; fluorescence was excited with 488‐nm argon ion and violet diode lasers, respectively. The intensities of maximal pixel and integrated fluorescence were measured and recorded for each cell. At least 3000 cells were measured per sample. All experiments were repeated at least three times. Other experimental details are presented elsewhere (2005a, 2005b, 2004; Kurose et al. 2006).

RESULTS

Figure 1 illustrates the effect of HU at 0.1–3.0 mm concentration on expression of γH2AX in HL‐60 cells. It is evident from this set of γH2AX IF versus DNA content bivariate distributions that exposure of cells to HU led to apoptosis preferentially of S‐phase cells. Apoptotic cells, as shown previously (Huang et al. 2004, , 2005b), were characterized by markedly increased intensity of γH2AX IF reflecting phosphorylation of this histone on Ser‐139 (Rogakou et al. 1998; Sedelnikova et al. 2002), and in the cultures treated with HU for 2 h had an S‐DNA content. The presence of apoptotic cells in these cultures was confirmed visually by fluorescence microscopy of DAPI‐stained cells (not shown). Apoptosis of S‐phase cells was also reflected by the decline in percentage of mid‐S‐phase cells among the non‐apoptotic cell population, from 12.9% in the untreated culture to 0.5% at 3 mm HU after 4 h of treatment. Due to progressive DNA fragmentation and partial extraction of the fragmented DNA (Gong et al. 1995b), some apoptotic cells had a sub‐G1 DNA content, which was apparent after 4 h. Also, as a result of exposure to HU, the remaining non‐apoptotic S‐phase cells, as well as some cells with a G1 DNA content (i.e. arrested at G1/S boundary), showed an increased degree of H2AX phosphorylation. Thus, a distinct subpopulation of cells with a G1‐DNA content and expression of γH2AX higher than that of G1 cells in untreated specimens (control) culture was apparent in the HU‐treated cultures (Fig. 1; within rectangular dashed boxes).

Figure 1.

Figure 1

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Changes in expression of γH2AX in relation to cell cycle phase in HL‐60 cells treated with 0.1, 1.0 or 3.0 mm HU for 2 or 4 h. Subpopulations of cells in G1, S and G2M phases are distinguished based on differences in their DNA content, as shown in the left panel (control). Apoptotic cells (Ap), identified based on their very high intensity of γH2AX IF (2005b, 2004), are located within the oval dashed gates. The dashed‐line horizontal threshold shows the upper limit of γH2AX immunofluorescence (IF) for the 95% of G1 cells in the untreated culture (control). The cells with a G1‐DNA content in the HU‐treated cultures that express γH2AX IF above this threshold are located within the rectangular dashed gates. The percentage of cells within the mid‐S DNA‐content gate (DI = 1.3–1.7) as marked in control sample by the vertical dashed lines (g) was calculated among the non‐apoptotic cell population for each culture (shown at the bottom of panels). The inset shows DNA content histogram of the cells in untreated culture.

The effect of cell treatment with AP on H2AX phosphorylation, in relation to the cell cycle phase of HL‐60 cells, is presented in Fig. 2. Exposure of cells to AP led to an increase in γH2AX IF and to the appearance of apoptotic cells. The AP‐induced increase in γH2AX IF was most pronounced for early S‐phase cells, though, compared with cultures treated with HU, the frequency of apoptotic cells was lower in AP‐treated cultures. Following the treatment with AP, as in cultures treated with HU, a subpopulation of cells that had higher intensity of γH2AX IF than G1 cells of the untreated culture was present at the G1/S boundary.

Figure 2.

Figure 2

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Expression of γH2AX in relation to cell cycle phase in HL cells treated with 1 or 4 µm AP for 2 or 4 h. Subpopulations of G1, S and G2M phase cells are identified as shown in the control. The presence of apoptotic cells (Ap) is apparent at 4 µm AP. As in Fig. 1, the dashed‐horizontal threshold defines the upper limit of γH2AX IF for cells in G1 in control. Note the presence of cells with a G1 DNA content having γH2AX IF increased above this threshold in AP‐treated cultures. The inset presents DNA content histogram of the cells in untreated culture. See legend to Fig. 1 for further explanation.

Exposure of HL‐60 cells to TH also led to apoptosis of S‐phase cells and accumulation of cells with a G1 DNA content having increased content of phosphorylated H2AX (Fig. 3). After 24‐h exposure to TH nearly all cells at G1/S boundary had γH2AX IF higher than G1 cells from the untreated culture. The two right panels in Fig. 3 present γH2AX IF versus DNA content bivariate distributions of HeLa cells, control, and those subjected to double thymidine block, used to synchronize them at G1/S phase. Unlike HL‐60 cells that were measured in suspension by flow cytometry, HeLa cells were grown on microscope slides and their fluorescence was measured by LSC. The absence of apoptotic HeLa cells in the scattergrams reflects their selective detachment in cultures, known to occur at an early stage of apoptosis (Darzynkiewicz et al. 2001). It is quite evident that the double thymidine block was successful in ‘synchronizing’ HeLa cells at G1/S interphase, with rather few cells remaining in S. However, it is also apparent that a large percentage of these ‘synchronized’ cells had increased levels of phosphorylated H2AX.

Figure 3.

Figure 3

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Expression of γH2AX in relation to the cell cycle phase in HL cells treated continuously with 2 mm TH, for 4, 8 or 24 h (4 left panels), and in HeLa cells synchronized in G1 by double TH block (TH DB, 2 right panels). See legend to Fig. 1 for further explanation. The dashed‐horizontal thresholds mark the upper limit of γH2AX IF for untreated cells. The presence of cells with a G1 DNA content having γH2AX IF increased above this threshold (gated in rectangular windows) is apparent in all TH‐treated cultures. The insets show DNA content histograms in the respective cultures. Fluorescence of HL‐60 cells was measured by flow cytometry (γH2AX IF plotted on log scale), of HeLa cells, which were growing on slides, by LSC (γH2AX IF plotted on linear scale).

There was a striking similarity in the pattern of response of HL‐60 cells to the DNA topoisomerase I inhibitor CP (Fig. 4), as compared to the changes induced by HU (Fig. 1). Namely, concurrent with phosphorylation of H2AX in S‐phase cells, subsequent loss of S‐phase cells within the non‐apoptotic cell population, and with the appearance of apoptotic cells, there was induction of H2AX phosphorylation in cells arrested at G1/S boundary.

Figure 4.

Figure 4

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Effect of CP at 50 or 250 nm concentration on H2AX phosphorylation in HL‐60 cells. See legend to Fig. 1 for further explanation. Note the increase in expression of γH2AX in S‐phase cells as well as in the cells ‘synchronized’ at G1/S boundary (gated in rectangular windows), concurrent with the appearance of apoptotic cells (AP), in the CP‐treated cultures.

DISCUSSION

The present data show that exposure of cells to concentrations of HU, AP and TH, at which these agents are being generally used to synchronize cells in the cell cycle, induced phosphorylation of histone H2AX on Ser‐139. Most affected were the S‐phase cells, as well as the cells that arrested at the G1/S boundary. S‐phase cells underwent apoptosis, and thus the ‘synchronization’, to a large extent, was generated by a selective kill of DNA replicating cells and by the arrest of cells at the entrance to S phase. Of all four inhibitors used in the present study, the frequency of apoptosis was lowest in cultures treated with AP, while HU and CP were the most effective killers of S‐phase cells. The above findings are consistent with our earlier observation that HU and AP induces H2AX phosphorylation, followed by apoptosis, preferentially affecting S‐phase cells (Kurose et al. 2006). The novelty of the present findings is that histone H2AX also is phosphorylated in cells ‘synchronized’ by these agents at the G1/S boundary. Because H2AX phosphorylation is considered a specific marker of induction of DSBs (Rogakou et al. 1998; Sedelnikova et al. 2002), the data indicate that the synchronization procedure causes DNA damage that involves formation of DSBs in the population of synchronized cells. In fact, the exposure of cells to CP, a cytotoxic agent which targets DNA topoisomerase I, produced a response similar to that observed following cell treatment with DNA replication inhibitors. Therefore, inhibitors of DNA replication not only selectively kill DNA replicating cells (Kurose et al. 2006), induce growth imbalance (Cohen & Studzinski 1967; Urbani et al. 1995), and alter machinery regulating progression through the cycle (Gong et al. 1995a), but also cause DNA damage involving formation of DSBs in the surviving (‘synchronized’) cells. The damage to DNA in all probability triggers the signalling pathways activating the G1/S DNA damage checkpoint (Lisby & Rothstein 2005). Cells synchronized by DNA replication inhibitors, thus, are significantly altered and in many respects do not represent cells at the G1/S boundary from untreated cultures. These changes should be taken into an account when interpreting data obtained with the use of cell populations synchronized by these compounds.

Although different agents were used in the present study, the pattern of cell response in terms of H2AX phosphorylation and apoptosis vis‐à‐vis cell cycle phase was similar. HU is an inhibitor of ribonucleotide reductase, the enzyme which by converting ribonucleotide diphosphates to deoxyribonucleotide diphosphates, is essential for the de novo synthesis of all the DNA precursors (Yarbro 1992). By altering the ribo‐ and deoxyribonucleotide diphosphate pools, HU stalls DNA replication. AP is a specific inhibitor of DNA polymerase α (Ikegami et al. 1978). TH at the concentrations used to synchronize cells blocks DNA replication by altering the pool of deoxynucleotides. CP is a DNA topoisomerase I inhibitor (Thomas et al. 2004). The common trait of all these agents was inhibition of DNA replication. As shown by Ward & Chen (2001), replicative stress, such as caused by DNA replication inhibitors, induces DNA damage involving formation of DSBs, which is reflected by phosphorylation of H2AX. In the case of CP, the collisions of DNA replication forks with the DNA‐topoisomerase complexes stabilized by CP lead to formation of DSBs (Deptala et al. 1999; Liu et al. 2000). As the present data indicate, DNA was damaged not only in the cells that were already progressing through S phase at the time of administration of these agents but also in the cells that were initiating DNA replication but become arrested at the G1/S boundary.

Our team has previously shown (Deptala et al. 1999) that the G1 checkpoint is activated in MCF‐7 cells arrested by CP, as manifest by induction of p53 and subsequent up‐regulation of p21WAF1/CIP1, which led to prolongation of cell progression arrest at the boundary. In the present study we have observed that the cells arrested at G1/S by CP were entering S phase, but only 4 h after removal of the drug, when H2AX was already de‐phosphorylated (data not shown). Apparently this was the minimal time during which DNA was repaired to the point that the cells could be released from checkpoint arrest. One would expect therefore that the cells with damaged DNA arrested at G1/S boundary by HU, AP or TH, if they have a functional G1/S checkpoint, also will be delayed before they resume progression through S. Indeed, such a delay was observed following prolonged cell arrest by HU or AP (Borel et al. 2002).

ACKNOWLEDGEMENT

Supported by NCI CA RO1 28 704.

A.K. and T.T. contributed equally to this study.

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