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Zidovudine (3′-azido-3′-deoxythymidine; AZT), which is currently used in the treatment of acquired immunodeficiency syndrome, has been shown to have anticancer properties. In the present study, we examined the mechanisms contributing to increased sensitivity of cancer cells to the growth-inhibitory effects of AZT. This was accomplished by incubating a hepatoma cell line (HepG2) and a normal liver cell line (THLE2) with AZT in continuous culture for up to 4 weeks and evaluating the number of viable and necrotic cells, the induction of apoptosis, cell cycle alterations, and telomerase activity. In HepG2 cells, AZT (2–100μM) caused significant dose-dependent decreases in the number of viable cells at exposures > 24 h. During a 1-week recover period, there was only a slight increase in the number of viable cells treated with AZT. The decrease in viable cells was associated with an induction of apoptosis, a decrease in telomerase activity, and S and G2/M phase arrest of the cell cycle. During the recovery period, the extent of apoptosis and telomerase activity returned to control levels, whereas the disruption of cell cycle progression persisted. Western blot analysis indicated that AZT caused a decrease in checkpoint kinase 1 (Chk1) and kinase 2 (Chk2) and an increase in phosphorylated Chk1 (Ser345) and Chk2 (Thr68). Similar effects, to lesser extent, were observed in THLE2 cells given much higher concentrations of AZT (50–2500μM). These data show that HepG2 cells are much more sensitive than THLE2 cells to AZT. They also indicate that a combination of a delay of cell cycle progression, an induction of apoptosis, and a decrease in telomerase activity is contributing to the decrease in the number of viable cells from AZT treatment, and that checkpoint enzymes Chk1 and Chk2 may play an important role in the delay of cell cycle progression.
Zidovudine (3′-azido-3′-deoxythymidine; AZT) was the first drug approved by U.S. Food and Drug Administration (FDA) for the treatment of acquired immunodeficiency syndrome (AIDS), and it is still extensively used in combination with other antiretroviral drugs in the treatment of patients with AIDS and AIDS-related complex and in the prevention of the mother-to-child transmission of human immunodeficiency virus-1 (HIV-1) (American Hospital Formulary Service, 2007; Fischl et al., 1990; Ingrand et al., 1995; McGowan and Shah, 2000; Physicians' Desk Reference, 2007). The antiretroviral activity of AZT is dependent upon the formation of AZT-triphosphate, which efficiently inhibits HIV-1 reverse transcriptase by acting as a competitive inhibitor of normal nucleotides (i.e., thymidine triphosphate) and causing proviral DNA termination (Brinkman and Kakuda, 2000; Furman et al., 1986; Jones and Bischofberger, 1995). In addition to inhibiting viral reverse transcriptase, AZT-triphosphate also is a weak inhibitor of mammalian DNA polymerases α, β, and γ, which catalyze DNA replication in vivo (Brinkman and Kakuda, 2000; Huang et al., 1990; Nickel et al., 1992).
AZT was originally developed as an antineoplastic agent (Horwitz et al., 1964; International Agency for Research on Cancer, 2000), and it has been shown to inhibit growth of a variety of human cancer cells (Collier et al., 2003; Falchetti et al., 2005; Humer et al., 2008; Melana et al., 1998; Olivero et al., 2005; Roskrow and Wickramasinghe, 1990; Wu et al., 2004). In addition, Tejera et al. (2001) reported a reduced tumor incidence and increased survival in syngeneic BALB/c mice inoculated with AZT-treated F3II mouse mammary carcinoma cells, and in other studies, AZT has been shown to reduce tumor growth of 518A2 melanoma cell xenografts in severe combined immunodeficiency mice (Humer et al., 2008). When AZT has been combined with other chemotherapeutic agents, such as 5-fluorouracil and cisplatin, significant antiproliferative activity has been observed in cancer cells in vitro (Andreuccetti et al., 1996). These results have led to the use of AZT, in combination with other chemotherapeutic agents, in phase I and II clinical studies in patients having colorectal cancer, leukemia/lymphoma, and other advanced malignancies (Falcone et al., 1996, 1997; Hermine et al., 2002; Morgan et al., 2003).
AZT can be incorporated into cellular DNA of human cancer cells (Brunetti et al., 1990; Escobar et al., 2007; Tosi et al., 1992; Vazquez-Padua et al., 1990), which can lead to DNA strand breaks and cytotoxicity. Cancer cells have higher growth rates than normal cells, with a concomitant higher rate of thymidine turnover, which could contribute to their increased sensitivity to AZT (Humer et al., 2008; Melana et al., 1998). Humer et al. (2008) reported that AZT impaired the proliferation of melanoma cell lines at concentrations where the proliferation of normal human skin fibroblasts and melanocytes was not affected. Likewise, Melana et al. (1998) reported that four human breast cancer cell lines were more sensitive than a normal breast cell line to the antiproliferative activity of AZT.
AZT-dependent inhibition of proliferation is accompanied by a significant S-phase arrest of the cell cycle (Humer et al., 2008; Roskrow and Wickramasinghe, 1990; Wu et al., 2004), the induction of apoptosis (Collier et al., 2003; Humer et al., 2008), or both. In addition, normal somatic cells generally express little or no detectable telomerase activity, whereas most malignant cancer cells express telomerase, which being a reverse transcriptase, is susceptible to the inhibitory effects of AZT. These effects could contribute to the differential sensitivity of cancer cells to AZT (Falchetti et al., 2005; Humer et al., 2008; Melana et al., 1998).
In the present study, we examined the mechanisms contributing to increased sensitivity of cancer cells to the growth-inhibitory effects of AZT. This was accomplished by incubating a hepatoma cell line (HepG2) and a normal liver cell line (THLE2) with AZT in continuous culture for up to 4 weeks and evaluating the number of viable and necrotic cells, the induction of apoptosis, cell cycle alterations, and telomerase activity. We demonstrate that the hepatoma cell line HepG2 is much more sensitive to AZT than the immortalized normal liver cell line THLE2, and that AZT interferes with cell growth by slowing cell cycle progression, inducing cell death by apoptosis, and inhibiting telomerase activity. We further demonstrate that the checkpoint kinases Chk1 and Chk2 may play an important role in the delay of cell cycle progression.
AZT was obtained from Cipla, Ltd. (Mumbai, India). The reported purity, as assessed by high-performance liquid chromatography (HPLC), was 99.7%. [3H]AZT (12.7 Ci/mmol, radiochemical purity > 99.8% by HPLC) was obtained from Moravek Biochemicals, Inc. (Brea, CA). Williams’ Medium E, propidium iodide (PI), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), 5-bromo-2′-deoxyuridine (BrdU), phosphoethanolamine, bovine serum albumin, and RNase A were acquired from Sigma (St. Louis, MO). Penicillin-streptomycin and 2.5% trypsin were purchased from Fisher Scientific (Pittsburgh, PA). Dulbecco's phosphate-buffered saline (minus calcium-chloride and magnesium-chloride) (PBS), epidermal growth factor (culture grade), and LHC-8 medium were obtained from Invitrogen (Carlsbad, CA). Fetal bovine serum was purchased from Atlanta Biologicals (Lawrenceville, GA). The BCA Protein Assay kit was purchased from Pierce Chemical Co. (Rockford, IL). All other chemicals and biochemicals were of analytical grade and used without further purification.
Monoclonal antibodies to Chk2 and β-actin were purchased from Sigma, and monoclonal antibody to Chk1 was obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Rabbit monoclonal antibody to phospho-Chk1 (ser345, pChk1) and rabbit antisera against phospho-Chk2 (Thr68, pChk2) were purchased from Cell Signaling Technology (Danvers, MA). The fluorescein isothiocyanate (FITC)–conjugated anti-BrdU monoclonal antibody (clone B44) was purchased from BD Biosciences (San Jose, CA).
HepG2 cells, a cell line derived from a human hepatocellular carcinoma, and THLE-2 cells, a cell line derived from normal human liver cells, were purchased from American Type Culture Collection (Manassas, VA). THLE2 cells express phenotypic characteristics of normal adult liver epithelial cells and retain phase I and II enzyme activities, including the ability to metabolize carcinogens to their ultimate carcinogenic metabolites capable of binding DNA (Pfeifer et al., 1993). HepG2 cells were grown in Williams’ Medium E supplemented with 10% fetal bovine serum and antibiotics. THLE-2 cells were cultured in LHC-8 medium supplemented with 70 ng/ml phosphoethanolamine, 5 ng/ml epidermal growth factor, 10% fetal bovine serum, and antibiotics. Both cell lines were incubated at 37°C in a humidified atmosphere containing 5% CO2 in air. AZT was dissolved in PBS and the concentration was confirmed spectrophotometrically at 266 nm, using a molar extinction coefficient of 11,500 cm−1 M−1 (Borojerdi et al., 2009). Cells were plated at a density of 8 × 104, 6 × 104, and 5 × 103 cells/cm2 and cultured for 24 h, 48 h, or 4 weeks. AZT was added at the beginning of the cultures. The final concentrations were 2, 20, and 100μM AZT for the HepG2 cells and 50, 500, and 2500μM AZT for the THLE2 cells. These concentrations were selected based upon the results from preliminary experiments in which AZT produced both a dose-response in decrease in the total numbers of viable cells and no effects on the necrotic cell death. By comparison, the therapeutic maximum AZT concentration (Cmax) in plasma is about 6μM (Wang et al., 1999). When appropriate, fresh medium and AZT were added every other day. For the 4-week incubation, the cells were subcultured weekly. Following the 4 weeks of culturing in the presence of AZT, the cells were then allowed to recover in drug-free complete medium for 1 week. Control cells were fed with complete culture medium free of AZT. Each of the incubations was performed three separate times, and all the measurements described below were conducted independently for each of the experiments.
Cells were cultured, as described above, for 48 h using 0, 2, 20, or 100μM [3H]AZT (diluted to a specific activity of 32 mCi/mmol with unlabeled AZT) for HepG2 cells, and 0, 50, 500, or 2500μM [3H]AZT (diluted to a specific activity of 20 mCi/mmol with unlabeled AZT) for THLE2 cells After the treatment, the cells were washed three times in PBS. They were then harvested and DNA was isolated using a conventional phenol-chloroform extraction method. The radioactivity incorporated into the DNA was determined by liquid scintillation counting.
The number of viable cells was assessed by a MTT reduction assay, as previously described (Fang et al., 2009). The number of necrotic cells was assessed by measuring the amount of lactate dehydrogenase (LDH) released into the medium using an LDH assay kit (Sigma) according to the manufacturer's instructions.
After treatment, floating and adherent cells were collected and washed twice in PBS. Aliquots (2 × 106 cells) were then fixed in 4 ml of freshly prepared paraformaldehyde (1% wt/vol) in PBS. After a 1 h fixation on ice, the cells were washed twice in PBS, centrifuged, and the pellet was resuspended in 100 μl of ice-cold PBS followed by the addition of 4.0 ml of ice-cold 70% (vol/vol) ethanol. The fixed cell suspensions were stored at −20°C until analyzed.
Apoptotic cells were double stained using an APO-BrdU kit (BD Biosciences) and subjected to flow cytometric analysis as described previously (Fang et al., 2009). Briefly, the fixed cells (2 × 106) were filtered through a 35-μm nylon mesh to remove any clumped cells. After filtration, aliquots of fixed cells (1 × 106) were washed once with PBS and resuspended in 51 μl of a DNA labeling solution that contained BrdU-triphosphate and terminal deoxynucleotidyl transferase. After incubation for 1 h at 37°C, the cells were rinsed twice with PBS, resuspended in 100 μl of an antibody solution containing FITC-labeled anti-BrdU monoclonal antibody, and incubated for 30 min at room temperature. Cell suspensions were mixed with 500 μl of PBS containing 2 μg/ml PI and 50 μg/ml RNase A, and analyzed by flow cytometry.
After treatment, cells were trypsinized and washed twice in PBS. Approximately, 0.5 × 106 cells were lysed in 200 μl of CHAPS buffer (10mM Tris-HCl, pH 7.5, 1mM ethylene glycol-bis(2-aminoethylether)-N,N,N′,N′-tetraacetic acid, 1mM MgCl2, 5mM β-mercaptoethanol, 50 μg/ml phenylmethylsulfonyl fluoride, 0.5% CHAPS, and 10% glycerol) and incubated at 4°C for 30 min. The lysate was then centrifuged at 12,000 × g for 30 min at 4°C, and the supernatant was collected and stored at −80°C. Protein concentrations were measured using a BCA protein assay kit.
All of the lysates were normalized to the same protein concentration of 100 μg/ml. Telomerase activity was determined using a real-time PCR-based, telomeric repeat amplification (TRAP) assay, as described by Wege et al. (2003), with minor modifications. Briefly, the total volume of the reaction mixture was 25 μl and contained 1 × IQ SYBR Green Supermix (BioRad, Hercules, CA), 0.1 μg each of primers TS (5′-AATCCGTCGAGCAGAGTT-3′) and ACX (5′-GCGCGGCTTACCCTTACCCTTACCCTAACC-3′), and 100 ng of cell lysate protein. The PCR was performed in a 96-well microplate on a BioRad iCycler iQ Detection System. The reaction mixture was first incubated at 25°C for 30 min to allow the telomerase in the cell lysate to elongate the TS primer by adding TTAGGG repeat sequences. The PCR was then started at 95°C for 10 min, followed by a 40-cycle amplification (95°C for 30 s, 53°C for 30 s, and 72°C for 90 s). The threshold cycle values (Ct) were determined from semi-log amplification plots (log increase in fluorescence vs. cycle number). The amount of telomerase was determined through comparison to a calibration curve generated from serial dilutions of a pooled HepG2 cell lysate (1.6–500 ng of protein). All samples were run in triplicate, and heat-inactivated cell lysates (by heating at 90°C for 10 min prior to the telomerase activity assay) and the lysis buffer were used as negative controls. The telomerase values were normalized based on the control value at each time point.
At the end of treatment, cells were trypsinized and washed once in complete medium. Aliquots (2 × 106) cells were resuspended in 15 ml of complete medium containing 10μM BrdU and incubated for 1 h at 37°C. The cells were then washed twice in PBS containing 1% bovine serum albumin and fixed with 70% (vol/vol) ice-cold ethanol. Following an overnight fixation at 4°C, the cells were collected by centrifugation and incubated with 4 ml of 2N HCl containing 0.5% Triton X-100 for 30 min at room temperature to denature the DNA. This process caused the cells to lose most of their cytoplasm. The cells were isolated by centrifugation and neutralized with 4 ml of 0.1M Na2B4O7 at pH 8.5 for 10 min at room temperature. After resuspending in 2 ml of PBS containing 1% bovine serum albumin, the cells were filtrated through a 35-μm nylon mesh to remove any clumps. The cells were then stained with FITC-conjugated anti-BrdU monoclonal antibody and PI, and analyzed on a FACScan flow cytometer as previously described (Fang et al., 2009).
After treatment, cell lysates were prepared and Western blot analysis was performed as described previously (Fang et al., 2009), with minor modifications. Briefly, the protein concentration was determined by using a Pierce BCA protein assay kit. Fifty micrograms of cell lysate protein was subjected to electrophoresis in a 12% sodium dodecyl sulfate–polyacrylamide gel. The resolved proteins were electrophoretically transferred onto an Immun-Blot polyvinylidene difluoride membrane (BioRad). Both electrophoresis and blotting were performed with a Mini-PROTEAN 3 electrophoresis system (BioRad). Blots were blocked with 5% milk and probed with anti-Chk1 (1: 300 dilution), anti-Chk2 (1:1000 dilution), anti-pChk1(Ser345) (1:1000 dilution), anti-pChk2 (Thr68) (1:1000 dilution), or anti-β-actin (1:2500 dilution), followed by a secondary antibody to IgG conjugated to horseradish peroxidase (HRP). The blots were then detected by chemiluminescence using Immobilon Western HRP Substrate (Millipore Corporation, Billerica, MA). The intensity of each band was quantified by densitometry (UVP BioSpectrum AC Imaging System using VisionWorksä LSD Image Acquisition & Analysis Software, Upland, CA), and the relative protein levels were calculated using β-actin as an internal reference. All primary antibodies were incubated with the same membrane after consecutive stripping using Restore Western Blot Stripping Buffer (Pierce). Densitometric results were always obtained at values below saturation levels as indicated by the densitometer software.
Data are expressed as mean ± SD. Comparisons amongst doses were conducted by one-way ANOVA, with multiple comparisons versus control group being performed by Dunnett's method. When necessary, the data were log transformed to maintain an equal variance or normal data distribution. The results were considered significant at p < 0.05.
In order to insure that the experimental conditions would permit the incorporation of AZT into DNA, an experiment was conducted in which HepG2 cells and THLE2 cells were incubated with various concentrations of [3H]AZT for 48 h. After isolation of the DNA, the extent of radioactivity was assessed by liquid scintillation counting. The levels of AZT incorporation into DNA are summarized in Table 1. There was a dose-dependent incorporation of AZT into the DNA in both cell lines. The level of AZT incorporation into DNA in HepG2 cells incubated with 100μM AZT was 268 AZT molecules/106 nucleotides, which was about 50% higher than that in THLE2 cells incubated with 2500μM AZT.
Treatment of HepG2 cells with AZT caused significant dose-dependent decreases in the number of viable cells, with the effect being greatest at the 3- and 4-week time points (Fig. 1A). Compared with the control cultures, growth rates at 2, 20, and 100μM AZT were 76.7, 64.4, and 19.4%, respectively, after 3 weeks of exposure, and 77.4, 64.6, and 9.8%, respectively, after 4 weeks of exposure. In cells allowed a 1-week recovery after being treated for 4 weeks with AZT, there was still a significant decrease in the number of viable cells at the two highest doses.
AZT also affected cell growth of the normal human liver cell line THLE2 cells. At concentrations of 50–2500μM AZT, which was 25 times higher than that given to HepG2 cells, significant concentration-dependent decreases in the number of viable cells were observed, with the effect being greatest at the 4-week time point (Fig. 1B). As with the HepG2 cells, the decrease in the number of viable THLE2 cells persisted upon the 1-week recovery period. These results indicate that THLE2 cells are more tolerant than HepG2 cells to AZT treatment.
The decrease in the number of viable cells could be due to a number of reasons, including an increase in necrotic cell death, an induction of apoptosis, a decrease in telomerase activity, and/or a prolongation in the progression of the cells transiting the cell cycle. Each of these possibilities was investigated.
To establish whether the decrease in the number of viable cells was due to necrotic cell death, an LDH release assay was conducted. LDH release assays have been used extensively as a marker for cell death (Allen et al., 1994). The release of LDH into culture medium accurately reflects necrotic cell death in vitro. In HepG2 cells treated with AZT, there was a dose-dependent decrease in LDH release (Fig. 1C), with the maximum decrease being observed after 2 weeks of treatment. A similar decreased release of LDH was also observed in THLE2 cells (Fig. 1D). These findings of a decreased release of LDH indicated that the doses used in the study were not toxic and that the decrease in viable cells resulting from the AZT treatment was not due to necrotic cell death.
To evaluate whether apoptosis was induced upon treatment with AZT, cells were cultured continuously with AZT (2–100μM for HepG2 cells and 50–2500μM for THLE2 cells) for 4 weeks and apoptosis was measured at selected intervals by an APO-BrdU flow cytometry assay. HepG2 cells showed a significant increase in apoptosis with respect to the controls at the two highest doses, with the maximum response being observed after 2 weeks of incubation (Fig. 2A). No significant difference in apoptosis levels between control and AZT-treated cells was observed at the lowest concentration of the drug at any time point. After the 1-week recovery period, the induction effect of apoptosis was completely abrogated (Fig. 2A). With THLE2 cells, there was an increase in apoptotic cell death only at 2500μM AZT after 3 weeks of incubation (Fig. 2B). This effect disappeared upon the 1-week recovery period (Fig. 2B).
These data indicate that apoptosis might contribute to the decrease in the number of viable cells resulting from AZT treatment.
Incubation with AZT caused a dose-dependent decrease in telomerase activity in HepG2 cells (Fig. 3). After 48 h of exposure, there was a significant reduction in telomerase activity with 100μM AZT. With 20μM, AZT a significant reduction occurred after 3 weeks of incubation. There was no decrease in activity with 2μM AZT. Upon the 1-week recovery period, the telomerase activity was restored to normal levels (Fig. 3).
Telomerase activity was not detected in THLE2 cells.
The effect of AZT upon cell cycle kinetics was assessed by flow cytometry. Representative flow scatter plots depicting the cell cycle distribution in HepG2 cells after a 1-week exposure to 0, 2, 20, and 100μM AZT are shown in Figure 4A. As shown by the regional gates applied to the PI versus FITC-BrdU dot plots, flow cytometric analysis of cells stained with PI and FITC-BrdU allowed discrimination of the cells residing in the G1/G0, S, and G2/M phases of the cell cycle.
In HepG2 cells treated with AZT, there was a significant dose-dependent decrease in the percentage of G1/G0 phase cells at each time point (Fig. 4B). The maximum decrease occurred after 4 weeks. Upon removal of the AZT, there was still a significant decrease in the percentage of G1/G0 phase cells. Concomitant with decrease in G1/G0 phase cells, treatment with 2 and 20μM AZT caused a significant increase in the percentage of cells in the S phase (Fig. 4C) and a slight increase in the percentage of cells in the G2/M phase (Fig. 4D) at all time points. Treatment with 100μM AZT caused an increase in S-phase cells and G2/M-phase cells at early treatment time (< 2 weeks), followed by a substantial increase in G2/M phase cells after prolonged exposures (≥ 2 weeks) (Fig. 4D). These changes persisted upon removal of the AZT. These data indicate that the AZT-mediated growth inhibition of HepG2 cells is associated with S and G2/M phase cell cycle arrest.
The effect of AZT upon the cell cycle distribution of THLE2 cells treated with AZT is shown in Figures 4E–4G. AZT caused a decrease in the percentage of cells in the G1/G0 phase (Fig. 4E), although the decrease was attenuated when compared with HepG2 cells. The decrease in cells in the G1/G0 phase was accompanied by an increase in S-phase cells at all time points (Fig. 4F), followed by an increase in G2/M phase cells at 4 weeks (Fig. 4G). These changes persisted upon removal of the AZT.
Cell cycle checkpoint kinases Chk1 and Chk2 play a crucial role in regulating cell cycle progression. The activation of Chk1 and Chk2, in response to DNA damage, involves phosphorylation of Ser345 and Thr68, respectively. Because AZT had a very pronounced effect upon the cell cycle in HepG2 and THLE2 cells, the effects of AZT on Chk1 and Chk2 were analyzed by Western blotting. As shown in Figure 5A, antibodies to Chk1, Chk2, pChk1 (Ser345), and pChk2 (Thr68) were specific and gave a distinct band for each protein.
In HepG2 cells treated with AZT, there was a dose-dependent decrease in the relative level of Chk1 after 48 h of exposure (Fig. 5B). AZT treatment also decreased the relative level of Chk2 (Fig. 5C), with the decrease being significant at doses ≥20μM AZT after 48 h of exposure. The decrease in the relative levels of Chk1 and Chk2 persisted during the 1-week recovery culture (Figs. 5B and 5C). In THLE2 cells, long-term (≥3 weeks) exposures to ≥500μM AZT resulted in a decrease in relative level of Chk1 (Fig. 5F). THLE2 cells showed a decrease in the relative level of Chk2 only at 4-week time point and only at the highest concentration of AZT (2500μM; Fig. 5G). After the 1-week recovery period, the relative levels of Chk1 and Chk2 in AZT-treated THLE2 cells approached those of the controls (Figs. 5F and 5G). These data indicated that the effect of AZT on the decrease in the relative levels of Chk1 and Chk2 is greater in HepG2 cells than in THLE2 cells.
Western blot analysis of HepG2 cells showed an increase in the relative levels of pChk1 (Ser345) at all time points (Fig. 5D). AZT treatment also increased the relative level of pChk2 (Thr68) (Fig. 5E), with the increase being significant at doses ≥20μM AZT after 24 h of exposure. After a 1-week recovery period, the response of pChk1 (Ser345) persisted at the two highest doses while the relative level of pChk2 (Thr68) was returned to the values of control cells (Figs. 5D and 5E). The effect of AZT on the relative levels of pChk1 (Ser345) and pChk2 (Thr68) in THLE2 cells is shown in Figures 5H and 5I. Exposing THLE2 cells to AZT led to an increase in the relative levels of pChk1 (Ser345) and pChk2 (Thr68), although the increase was much smaller when compared with HepG2 cells and occurred after prolonged exposures (≥1 week for pChk1 (Ser345) and ≥3 weeks for pChk2 (Thr68)).
In this study we have demonstrated that AZT impairs the growth of both the hepatoma cell line HepG2 and the immortalized normal liver cell line THLE2, with much higher concentrations being required in the THLE2 cells. The inhibitory effect on cell growth was not due to necrotic cell death, but rather was associated with a combination of factors, including the induction in apoptosis, the inhibition in telomerase activity, and S and G2/M phase cell cycle arrest. Further investigation demonstrated that checkpoint kinases Chk1 and Chk2 may play a role in the cell cycle arrest.
AZT is a prodrug that is phosphorylated intracellularly to its active metabolite AZT-triphosphate. AZT-triphosphate can be incorporated into replicating DNA, which leads to DNA chain termination. Several studies have demonstrated the incorporation of AZT into cellular DNA in vitro (Fang et al., 2009; Olivero et al., 1994b; Sommadossi et al., 1989; Vazquez-Padua et al., 1990) and in vivo (Meng et al., 2007; Olivero et al., 1994a, 1997). In the present study, the incorporation of AZT into DNA of the HepG2 cells and THLE2 cells occurred in dose-dependent manner, ranging from 20.1 to 267.5 AZT molecules per 106 nucleotides. The resulting damaged DNA by AZT might either delay cell cycle progression until the damage is repaired or cause apoptotic cell death.
Specific checkpoints exist to restrict the progression of damaged cells through the cell cycle. Chk1 and Chk2 are the components of the DNA damage checkpoint pathway that prevent transmission of altered genetic information to progeny; as such, they are involved in regulating cell cycle arrest. The activation of Chk1 and Chk2, in response to DNA damage, involves phosphorylation of Ser345 and Thr68, respectively. Our data clearly indicate that the levels of pChk1 (Ser345) and pChk2 (Thr68) are elevated in AZT-treated cells, and the maximum increase in pChk1 (Ser345) and pChk2 (Thr68) seems to occur at the time when there is a substantial decrease in S and an increase in G2/M. Moreover, the increase in the active forms of Chk1 and Chk2 is accompanied by a decrease in the levels of Chk1 and Chk2. Cell cycle analysis revealed a dose-dependent increase in the percentage of cells in the S and G2/M phases, with a concomitant depletion in the percentage of cells in the G1/G0 phase, indicating S and G2/M cells arrest. These finding agree with our previous report in which the arrest of S and G2/M phase cells was observed in NIH 3T3 cells after the extended exposures (> 48 h) (Fang et al., 2009). We suggest that the reduction of the number of viable cells seen following AZT treatment is due, in part, to AZT-mediated S and G2/M phase cell cycle arrest.
Perturbation of cell cycle progression was not the sole mechanism by which AZT affected cell growth. Previous studies have shown that AZT induces apoptosis in cancer cells and primary explant cultures from term and first trimester human villous placentas (Collier et al., 2003; Humer et al., 2008), and we found an increase in apoptotic cell death occurred in both cell lines, with the effect being more pronounced in HepG2 cells.
Telomerase activity is present in many types of tumor cells and is essential for the continued growth of malignant cells (Hahn et al., 1999). In contrast to cancer cells, most human somatic cells lack telomerase (Hahn et al., 1999; Kim et al., 1994). Because telomerase is a reverse transcriptase, several investigators have tried to inhibit its activity using nucleoside reverse transcriptase inhibitors (Falchetti et al., 2005; Melana et al., 1998). Recently, Falchetti et al. (2005) showed a significant inhibition of telomerase activity by 100μM AZT in human parathyroid cancer cells after 48 h of incubation, and the effect was correlated with inhibition of cell proliferation (Falchetti et al., 2005; Hahn et al., 1999). We also observed a reduction in telomerase activity, and this occurred only in the HepG2 cells and required greater than 20μM AZT and extended exposures (> 48 h).
Two facts from our study suggest that there was removal of AZT from DNA. The first is the activation of a checkpoint pathway that arrests the cell cycle to permit DNA repair and the transcription of genes that facilitate DNA repair, and the second is that AZT effects were partially or totally restored during the recovery period. A study by Slameňová et al. (2006) indicated that AZT-induced DNA damage is repairable. To date, only a few studies have been performed to address the mechanisms underlying the removal of AZT from DNA (Slamenova et al., 2006; Vazquez-Padua et al., 1990). Additional studies to understand the mechanisms of AZT-DNA repair are clearly warranted.
Although the effects on cell growth exerted by AZT on HepG2 cells and THLE2 cells appear similar, the responses of HepG2 to AZT treatment tended to be greater and occurred at much lower concentrations than were used in THLE2 cells. Whole genome human microarray studies are underway in our laboratory to investigate the mechanisms for the differential response in these cells, particularly with regard to genes involved the metabolic activation and deactivation of AZT, cell growth, cell cycle checkpoint regulation, and DNA repair.
In conclusion, we have demonstrated that hepotoma-derived HepG2 cells are more sensitive to AZT treatment than the immortalized normal liver THLE2 cells, and that AZT affects cell growth due to a combination of factors, including a delay of cell cycle progression, the induction in apoptosis, and a decrease in telomerase activity, and that checkpoint enzymes Chk1 and Chk2 may play an important role in delay of cell cycle progression.
Interagency Agreement (224-07-0007) between the National Center for Toxicological Research, U.S. Food and Drug Administration, and the National Institute for Environmental Health Sciences/National Toxicology Program.