Gallic acid causes inactivating phosphorylation of cdc25A/cdc25C-cdc2 via ATM-Chk2 activation, leading to cell cycle arrest, and induces apoptosis in human prostate carcinoma DU145 cells

Prostate cancer is the second most common male malignancy and leading cause of deaths in men in the United States and Europe. The management and treatment of hormone-refractory prostate cancer (advanced stage) is a major problem, which increases morbidity and mortality in prostate cancer patients. Statistical predictions for 2006 show an estimated 234,460 new cases of prostate cancer with 27,350 deaths in the United States alone (1). Because the advanced-stage prostate cancer growth and development become independent of androgen and renders androgen ablation therapy ineffective, prostate cancer control through chemoprevention or intervention is highly desirable, which deals with suppression or reversal of both premalignant and malignant lesions by natural or synthetic agents (2–4). It is also important to emphasize here that the abnormal expression of growth factors and receptors, which is often associated with prostatic-intraepithelial neoplasia and invasive prostate cancer, leads to autocrine and paracrine loops for both mitogenic and antiapoptotic signaling leading to uncontrolled cell cycle progression followed by a growth advantage to cancer cells together with a loss of their apoptotic cell death (2–4). Accordingly and consistent with the above notion, the antiproliferative and apoptotic effects of nontoxic dietary agents could be of additional significance for the prevention, control, and/or management of prostate cancer, specifically that at an advanced and an androgen-independent stage of the malignancy. In this regard, various phytochemicals present in diet and those consumed as supplement are gaining increased attention in cancer chemoprevention research, including prostate cancer, focusing on both efficacy studies and associated mechanisms (refs. 2–6 and references therein).

Grape seed extract, also termed as grape seed polyphenols or procyanidins, is one such dietary supplement marketed in the United States as “grape seed extract” with 95% standardized procyanidins due to its several health benefits (7, 8). In addition to grape seeds, procyanidins are a diverse group of polyphenolic compounds that are also abundant in blackjack oak, horse chestnut, witch hazel, and hawthorn, as well as in apples, berries, barley, bean hulls, chocolate, rhubarb, rose hips, and sorghum (refs. 7–9 and references therein).

About the anticancer and cancer chemopreventive effects of grape seed extract, it inhibits the growth of human breast, lung, and gastric cancer cells but enhances the growth and viability of normal human gastric mucosal and normal murine macrophage cells (10). Our studies have shown that grape seed extract inhibits growth, induces cell cycle arrest, and causes apoptotic death of human breast carcinoma MDA-MB468, prostate carcinoma DU145, and LNCaP cells in culture (11–14). With regard to its anticarcinogenic effects in animal models, oral feeding of grape seed extract or procyanidin-rich fraction from grape seed extract has been shown to prevent azoxymethane-induced aberrant crypt foci formation in rats (15, 16), and that topical application of grape seed extract significantly prevents chemical and UVB radiation–induced skin tumorigenesis (17–19). With regard to the in vivo efficacy of grape seed extract against prostate cancer, we recently reported that oral feeding of grape seed extract strongly suppresses in vivo growth of advanced human prostate cancer DU145 xenograft in athymic nude mice (20). Together, these studies have convincingly documented the anticancer and chemopreventive efficacy of grape seed extract against various epithelial cancers, including prostate cancer, and have suggested the presence of biologically active phytochemicals in the crude mixture of grape seed extract.

In an effort to isolate and identify active compounds in grape seed extract, we recently found that gallic acid (Fig. 1) substantially contributes to the biological effects of grape seed extract in DU145 cells toward both cell growth inhibition and apoptotic cell death (21). The present study, therefore, was designed to examine efficacy and associated mechanism of gallic acid in advanced human prostate carcinoma DU145 cells; our objectives were (a) to study the effect of gallic acid on the growth and survival, cell cycle progression, and apoptotic death of DU145 cells; and (b) to study the molecular mechanisms of the biological effects of gallic acid. Our results provide the first evidence for the activation of ataxia telangiectasia-mutated (ATM)–checkpoint kinase (Chk) signaling as a central mechanism of gallic acid–induced cell cycle arrest in its biological effects in DU145 cells.

The human prostate carcinoma DU145 cell line was from American Type Culture Collection (Manassas, VA). Cells were cultured in RPMI 1640 with 10% fetal bovine serum (Hyclone, Logan, UT) and 1% penicillin-streptomycin under standard culture conditions (37°C, 95% humidified air and 5% CO₂). RPMI 1640 and other culture materials were from Life Technologies, Inc. (Gaithersburg, MD). Gallic acid used in the present study was from Sigma-Aldrich Chemical Company (St. Louis, MO). The primary antibodies for cyclin-dependent kinase (CDK) 4, CDK2, CDK6; cyclins D1, D3, E, A, B1; and cell division cycle 25 A (cdc25A) were from Santa Cruz Biotechnology (Santa Cruz, CA). Antibody for CDK inhibitor Kip1/p27 was from Neomarkers (Fremont, CA); antibody for phospho-cdc25A was from Abcam (Cambridge, MA); and antibody for actin was from Sigma-Aldrich. Primary antibodies for phospho- and total cdc25C and cdc2; Chk2 and Chk1; total cdc25B; and cleaved caspase-9, caspase-3, and poly(ADP)ribose polymerase were from Cell Signaling, Inc. (Beverly, MA). Antibody for phospho-ATM was from Rockland Immunochemicals (Gilbertsville, PA), and antibodies for Cip1/p21 and phospho-H2A.X were from Upstate Biotechnologies (Lake Placid, NY). Antibody for ataxia telangiectasia-rad3-related (ATR) was from Novus (Littleton, CO). Secondary anti-mouse antibody and enhanced chemiluminescence (ECL) detection system were from Amersham (Arlington Heights, IL). For all the gallic acid treatments, a 1,000-fold concentrated stock solution was prepared in DMSO and diluted directly into the medium immediately before the treatment of the cells.

DU145 cells were plated at 1 × 10⁵/60-mm plates under standard culture condition, and after 24 h, cells were fed with fresh medium and treated with DMSO alone (control) or varying doses of gallic acid (10–50 μmol/L, final concentrations in medium) dissolved in DMSO for different time points (24–72 or 6–24 h). The DMSO concentration was the same for all treatments and did not exceed 0.1% (v/v). After the desired treatment, cells were collected with brief trypsinization, washed with ice-cold PBS, and counted in duplicate using a hemocytometer. Trypan blue dye exclusion was used to determine cell viability.

DU145 cells at 60% confluency were treated with either DMSO alone or 50 μmol/L gallic acid in DMSO for 6, 12, and 24 h. Alternatively, cells were treated with DMSO, 50 μmol/L gallic acid, and 2.5 mmol/L caffeine without, or followed 2 h later with, 50 μmol/L gallic acid for 24 h. Following these treatments, the medium was aspirated, cells were trypsinized, and cell pellets were collected. Approximately 0.5 × 10⁵ cells in 0.5 mL of saponin-propidium iodide solution (0.3% saponin, 25 μg/mL propidium iodide, 0.1 mmol/L EDTA, and 10 μg/mL RNase in PBS) were incubated at 4°C for 24 h in the dark. Cell cycle distribution was then analyzed by flow cytometry using the fluorescence-activated cell sorting analysis core service of the University of Colorado Cancer Center (Denver, CO).

To quantify gallic acid–induced apoptotic death of DU145 cells, Annexin V and propidium iodide staining was done followed by flow cytometry, as recently described (21). Briefly, after treatment with DMSO or 50 μmol/L gallic acid in DMSO as a function of time for 6, 12, and 24 h, cells were collected by brief trypsinization and washed with PBS twice. Cells were then subjected to Annexin V and propidium iodide staining using Vybrant Apoptosis Assay Kit 2 (Molecular Probes, Eugene, OR) following the step-by-step protocol provided by the manufacturer. The kit contains recombinant Annexin V conjugated to fluorophores and the Alexa Fluor 488 dye, providing maximum sensitivity. After staining, flow cytometry was done for the quantification of apoptotic cells. In the caspase inhibitor study, cells were pretreated with 50 μmol/L pan-caspase inhibitor ZVAD.fmk (Enzyme Systems Products, Livermore, CA), 2 h before treatment with 50 μmol/L gallic acid.

Cells were treated with DMSO or 50 μmol/L gallic acid in DMSO for the desired time periods. In caffeine and gallic acid combination studies, cells were treated with caffeine (2.5 mmol/L) 2 h before gallic acid treatment. In pan-caspase inhibitor study, cells were treated with ZVAD.fmk (50 μmol/L) 2 h before gallic acid treatment at 50 μmol/L dose. Following desired treatments, cell lysates were prepared in nondenaturing lysis buffer [10 mmol/L Tris-HCl (pH 7.4), 150 mmol/L NaCl, 1% Triton X-100, 1 mmol/L EDTA, 1 mmol/L EGTA, 0.3 mmol/L phenylmethylsulfonyl fluoride, 0.2 mmol/L sodium orthovanadate, 0.5% NP40, 5 units/mL aprotinin]. Briefly, medium was aspirated and cells were washed with ice-cold PBS twice, followed by incubation in lysis buffer for 10 min on ice. Then, cells were scraped and kept on ice for 30 min, and finally cell lysates were cleared by centrifugation at 4°C for 30 min at 14,000 rpm. Protein concentrations in lysates were determined using Bio-Rad detergent-compatible protein assay kit (Bio-Rad Laboratories, Hercules, CA) by the Lowry method.

For immunoblot analysis, total cell lysates were denatured in 2× sample buffer; samples were subjected to SDS-PAGE on 4%, 12%, or 16% Tris-glycine gels; and separated proteins were transferred onto membrane by Western blotting. Membranes were blocked with blocking buffer for 1 h at room temperature and probed with primary antibodies against desired molecules overnight at 4°C followed by peroxidase-conjugated appropriate secondary antibody for 1 h at room temperature and ECL detection. In each case, blots were subjected to multiple exposures on the film to make sure that the band density is in the linear range. The bands were scanned with Adobe Photoshop 6.0 (Adobe Systems, Inc., San Jose, CA), and, as needed, the mean density of each band was analyzed by the Scion Image program (NIH, Bethesda, MD).

Statistical significance of differences between control and treated samples were calculated by Student's t test (SigmaStat 2.03). P < 0.05 was considered significant. All the results shown are representative of at least two to four independent experiments with reproducible findings.

To examine the biological effects of gallic acid, DU145 cells were treated with varying doses of gallic acid (10, 25, 40, and 50 μmol/L) for 24, 48, and 72 h, and both cell growth inhibition and cell death were assayed. The results of this experiment showed that gallic acid causes both cell-growth inhibition and cell death only at its two higher doses, 40 and 50 μmol/L, without any time-dependent response (data not shown). Based on these pilot observations, an early time-response study was next done to assess the effect of gallic acid at 40 and 50 μmol/L doses, which caused strong cell growth inhibition (Fig. 2A) and cell death induction (Fig. 2B) mostly in a dose-dependent, but not time-dependent (except 24 h), manner. Compared with DMSO-treated control cells, 50 μmol/L gallic acid treatment for 24 h resulted in 62% inhibition (P < 0.001) of cell growth (Fig. 2A) and caused 39% (P < 0.001) cell death (Fig. 2B). Based on these results, we selected the 50 μmol/L dose and next assessed whether the growth-inhibitory and cell death effects of gallic acid are accompanied by its effect on cell cycle progression and/or apoptotic cell death.

Gallic acid showed statistically significant cell cycle arrest at 50 μmol/L dose following its treatment for 6, 12, and 24 h (Fig. 2C). At early time points of 6 and 12 h, compared with vehicle controls, it caused an arrest in both S (32% versus 40% and 20% versus 38%, respectively, P < 0.001) and G₂-M (20% versus 23%, P = 0.001 and 17% versus 22%, P = 0.005, respectively) phases, which were at the expense of a strong decrease in G₁-phase cell population (Fig. 2C). When similar gallic acid treatment was done for 24 h, compared with vehicle control showing 15% cells in G₂-M phase, 28% of the gallic acid–treated cells showed their accumulation in G₂-M phase accounting for almost 1.87-fold increase (P = 0.004) compared with control without any change in S-phase population (Fig. 2C). The G₂-M accumulation of cells at this time point was only at the expense of a decrease in G₁-phase cell population (Fig. 2C). In other studies assessing whether gallic acid also causes apoptotic cell death, as shown in Fig. 2D, its treatment at 50 μmol/L dose for 6, 12, and 24 h resulted in a strong and statistically significant apoptotic cell death. The early time points of gallic acid treatment resulted in almost 3-fold increase (P < 0.001 and P = 0.012) in apoptotic cells compared with vehicle controls (Fig. 2D); however, a similar gallic acid treatment for 24 h resulted in 39.2% apoptotic cells compared with 5.4% in control accounting for 7.3-fold increase (P < 0.001) in apoptotic cells by gallic acid versus control (Fig. 2D). In other studies, 40 μmol/L gallic acid also showed a moderate S-phase arrest in the cell cycle progression and caused 6% to 16% apoptotic cell death at 6 to 24 h of treatment (data not shown). Together, the results shown in Fig. 2 clearly show the biological effects of gallic acid on DU145 cell growth inhibition and death, as well as cell cycle arrest and apoptosis induction. Accordingly, studies were next done to examine the mechanism of cell cycle arrest by gallic acid in DU145 cells.

Because gallic acid treatment of cells caused cell cycle arrest as early as 6 h after its treatment that sustained even at 24 h, a time kinetics study starting at 3 h was done to examine the effect of 50 μmol/L gallic acid on the protein levels of cell cycle regulatory molecules. In terms of the levels of CDKs, which drive the cell cycle by the phosphorylation of Rb, making transcription factor E2Fs free (2, 4), gallic acid caused a decrease in CDK4, CDK6, and CDK2 protein albeit at different levels following different treatment times. However, no effect was evidenced following 3-h gallic acid treatment (Fig. 3A). One of the two important regulators of CDKs is their regulatory subunit cyclins that bind to and positively regulate CDK activity (2, 4), and, therefore, we next assessed the effect of gallic acid on cyclin levels. As shown in Fig. 3A, gallic acid treatment of cells for varying time periods also resulted in a decrease in the protein levels of various cyclins. Interestingly, the levels of cyclin D1 and cyclin D3 decreased at early time points of gallic acid treatment, but those of cyclin A and cyclin B1 at later time points of 12 and 24 h, without any effect on cyclin E levels; the maximum gallic acid effect was evidenced on cyclin B1 decrease following 24-h treatment that is in accordance with a strong G₂-M arrest at this time point (Fig. 3A); densitometric analysis accounted for almost 50% reduction (data not shown). The other important regulator of CDKs is a family of their inhibitory proteins known as CDK inhibitors that bind to CDK-cyclin complex and negatively regulate CDK activity (2, 4). Based on our observed cell cycle arrest effect of gallic acid, we next assessed whether this agent also modulated the levels of CDK inhibitors. As shown in Fig. 3B, treatment of cells with gallic acid resulted in a strong increase in the protein level of Cip1/p21 at 12 and 24 h (∼2-fold induction versus control in densitometric analysis; data not shown), without any noticeable changes in Kip1/p27 levels. Together, these results clearly show a decrease in the protein levels of CDKs and cyclins by gallic acid in DU145 cells and a selective induction in Cip1/p21, suggesting their possible roles in the observed biological responses of gallic acid, including cell cycle arrest.

In addition to CDKs, cyclins, and CDK inhibitors examined above, the family of cdc25 phosphatases also play an important role specifically in S and G₂-M phases of the cell cycle in which their specific phosphorylation causes their inactivation toward dephosphorylation of cdc2 (CDK1); phosphorylated cdc2 is also inactive in driving cell cycle progression (refs. 22–25 and references therein). Based on our findings showing that gallic acid causes both S and G₂-M arrests, we next assessed its effect on both total and phosphorylated cdc25A, cdc25B, cdc25C, and cdc2 under identical experimental conditions as for other cell cycle regulators. As shown in Fig. 4A, gallic acid treatment of cells resulted in a decrease in total and phosphorylated levels of cdc25C and cdc25A at 12 and 24 h without any effect at early time points on these as well as cdc25B levels; however, a marginal increase in cdc2 phosphorylation without any change in its total level was also observed at early treatment times. Because these results were mostly not consistent with an anticipated increase in cdc25C/cdc25CA followed by cdc2 phosphorylation in terms of their involvement in the observed cell cycle arrest by gallic acid, we next conducted an early time point study. Interestingly, gallic acid treatment of cells resulted in a strong phosphorylation of both cdc25C at Ser²¹⁶ and cdc25A at Ser¹⁷ as early as 15 min without any changes in their total protein levels (Fig. 4B). The densitometric analyses of these blots showed ∼1.8- to 2.9-fold increase in the phosphorylation of cdc25C and cdc25A as a function of time up to 2-h after gallic acid treatment (data not shown). Consistent with these results, gallic acid treatment of cells also showed a strong phosphorylation of cdc2 at Tyr¹⁵ as early as 15 min without any change in its total protein levels (Fig. 4B). In terms of the quantitative estimation, densitometric analysis of the blot showed 1.3- to 1.7-fold increase in cdc2 phosphorylation at Tyr¹⁵ as a function of time up to 2 h after gallic acid treatment (data not shown). In other studies (data not shown), gallic acid treatment of cells at all the early time points and up to 24 h as well did not result in any noticeable changes in Wee1, which is a known kinase responsible for cdc2 phosphorylation at Tyr¹⁵. Together, these results clearly suggest that gallic acid causes an early inactivating phosphorylation of cdc25C and cdc25A, leading to an accumulation of inactive Tyr¹⁵-phosphorylated cdc2, and that these effects of gallic acid possibly drove the observed cell cycle arrest.

Inactivating phosphorylation of cdc25C/A followed by the accumulation of Tyr¹⁵-phosphorylated cdc2 and a resultant cell cycle arrest is known to occur via ATM/ATR-Chk1/2 activation in response to DNA damage (refs. 26–29 and references therein). In this regard, ATM and ATR are nuclear kinases recently identified as being activated in response to DNA damage/genotoxic stress in eukaryotic cells (26–29). Based on the results described above, we hypothesized that gallic acid activates ATM/ATR kinases that is known to activate cell cycle checkpoint kinases Chk1/2 (28–30). Accordingly, we next examined the effect of gallic acid on ATM/ATR and Chk1/2 levels and phosphorylation. Gallic acid treatment (50 μmol/L) of cells up to 24 h did not result in any change in total ATR and Chk1 levels as well as Chk1 phosphorylation at any of its known phosphorylation sites (data not shown). However, a strong Chk2 phosphorylation only at Thr⁶⁸ site was observed at 3 and 6 h of gallic acid treatment without any change in its total protein levels (Fig. 5A). Similar to this observation, gallic acid also showed a strong phosphorylation of ATM at Ser¹⁹⁸¹ following its treatment for 3 and 6 h; later time points did not show any effect on both Chk2 and ATM phosphorylation (Fig. 5A). H2A.X is a variant form of histone H2A that is directly phosphorylated at Ser¹³⁹ by an activated ATM kinase, marking an early event in response to DNA damage, and is also known to play a critical role in the retention of DNA repair factors at DNA-damaged sites (31). Because we observed a strong ATM phosphorylation at Ser¹⁹⁸¹ by gallic acid, we also examined the levels of H2A.X phosphorylation at Ser¹³⁹; we found that, indeed, gallic acid treatment causes strong levels of this effect (Fig. 5A). In additional studies examining whether the time of activation of ATM-Chk2 pathway corroborate the observed phosphorylation of cdc25C/A and accumulation of Tyr¹⁵-phosphorylated cdc2, as shown in Fig. 5B, gallic acid treatment caused a strong and a time-dependent increase in the phosphorylation of Chk2 at Thr⁶⁸, ATM at Ser¹⁹⁸¹, and H2A.X at Ser¹³⁹. In other studies, 40 μmol/L gallic acid also showed an increase in the phosphorylation of Chk2, ATM, and H2A.X in an early (15, 30, 60, and 120 min) time kinetics study (data not shown); however, these effects were moderate compared with those observed with 50 μmol/L gallic acid (Fig. 5B).

The results shown in Figs. 4 and 5 suggested that gallic acid causes a genotoxic stress leading to ATM-Chk2 pathway activation followed by inactivating phosphorylation of cdc25C/A and thereby accumulation of Tyr¹⁵-phosphorylated cdc2 in its inactive form as a central mechanism of the observed cell cycle arrest. To further substantiate this suggestion, cells were pretreated with caffeine, which is a known inhibitor of both ATM and ATR kinases (32), and then the effect of gallic acid on cell cycle progression and associated upstream events was examined. Pretreatment of cells with caffeine completely reversed gallic acid, induced G₂-M arrest (P < 0.001) observed following 24 h of its treatment (Fig. 5C), and strongly reduced gallic acid caused phosphorylation of both ATM and H2A.X (Fig. 5D). Together, these findings suggest that gallic acid causes DNA damage leading to ATM activation followed by Ser¹³⁹ phosphorylation of H2A.X, supporting the role of ATM activation in cell cycle arrest by gallic acid.

Based on our results showing that gallic acid causes strong apoptotic death (Fig. 2D), studies were also done to examine whether caspase activation is involved in this effect of gallic acid. As shown in Fig. 6A, treatment of cells with gallic acid resulted in a time-dependent increase in caspase-9, caspase-3, and poly(ADP)ribose polymerase cleavage with strongest effect at 24 h, suggesting a possible involvement of caspase activation in the apoptotic effect of gallic acid. To further address this issue, cells were pretreated with pan-caspase inhibitor ZVAD.fmk followed by gallic acid, and quantitative apoptotic cell population was measured. Interestingly, compared with gallic acid alone, pretreatment with caspase inhibitor resulted in more apoptotic cell death (Fig. 6B). To substantiate that caspase activation is indeed inhibited by caspase inhibitor pretreatment, under similar experimental conditions, samples were also analyzed for cleaved caspase-9 and cleaved caspase-3, which showed a complete reversal in their activation (Fig. 6C), suggesting that, at least under caspase-inhibiting conditions, additional pathway(s) is involved in the apoptotic effects of gallic acid. More studies are needed in the future to define both caspase-dependent and caspase-independent mechanisms of gallic acid–caused apoptotic cell death, and to address whether caspase activation initiates apoptosis or is part of terminal processing of death.

A wide range of naturally occurring substances have been shown to protect against experimental carcinogenesis, and numerous studies in the literature suggest that phytochemicals, particularly those in our daily diet, possess strong anticancer and cancer chemopreventive properties against various malignancies, including prostate cancer (refs. 3, 5, 6, 33–35 and references therein). In this regard, gallic acid is a ubiquitous dietary phytochemical being present in virtually all the fruits and vegetables, as well as in beverages such as green tea and red wine, either in its free or esterified forms (refs. 36, 37 and references therein). However, only few studies have shown the anticancer activity of gallic acid in limited cancer cell lines, including a study showing its selective cell death effect in various human and rodent cancer cells but not in normal cells (38, 39). With regard to prostate cancer, our study for the first time reports the biological activity of gallic acid in prostate cancer DU145 cells and associated mechanisms of its efficacy.

The central findings of the present study are that via ATM-Chk2 activation, gallic acid causes inactivating phosphorylation of cdc25C/A phosphatases, leading to accumulation of cdc2 in its Tyr¹⁵-phosphorylated inactive form and a resultant cell cycle arrest. Inhibition of ATM kinase by its specific inhibitor caffeine abrogated gallic acid–caused phosphorylation of both ATM and H2A.X and reversed gallic acid–induced cell cycle arrest, further supporting the role of gallic acid–caused DNA damage and ATM activation in its cell cycle arrest activity. Many DNA-damaging agents, especially those that generate DNA double-strand breaks, are known to activate ATM kinase and induce ATM-dependent apoptosis (22). Histone H2A.X, one of the target proteins of ATM, is phosphorylated by activated ATM at Ser¹³⁹ (31). In the present study, we also observed that gallic acid induces Ser¹³⁹ phosphorylation of H2A.X in DU145 cells, which is inhibited by caffeine, suggesting the involvement of ATM activation in gallic acid–induced DNA damage signaling.

The checkpoint functions of ATM and ATR are mediated by checkpoint effector kinases known as Chk1 and Chk2 (40), which are structurally distinct but functionally related kinases that phosphorylate an overlapping pool of cellular substrates (41). In mammalian cells, it has been shown that Chk1 and Chk2 play a central role in transducing DNA damage signals from ATR and ATM, respectively, in which Chk1 seems to be activated by ATR in response to replication inhibition and UV-induced damage, whereas Chk2 primarily functions through ATM in response to ionizing radiation (23, 42). The activation of Chk2 in response to DNA damage requires its phosphorylation at Thr⁶⁸ (43, 44). Accordingly, our data is consistent with other emerging information on the phosphorylation of Chk2 by ATM. For example, gallic acid treatment did not show any effect on ATR levels and also did not cause the phosphorylation of Chk1 at any of its known sites (data not shown). However, gallic acid showed a strong ATM activation in terms of its phosphorylation at Ser¹⁹⁸¹, together with Chk2 phosphorylation at Thr⁶⁸ as well as that of H2A.X at Ser¹³⁹. Furthermore, the time kinetics of these effects of gallic acid was also consistent, suggesting a close association between these events following gallic acid treatment of DU145 cells.

Activation of Chk1 and/or Chk2 causes the phosphorylation and thereby inactivation of cdc25 family of tyrosine phosphatases, which creates a binding site for 14-3-3 proteins and results in their export to and retention in the cytoplasm (45, 46). Nuclear cdc2 remains phosphorylated in the absence of active cdc25 phosphatases, and the cells remain arrested in specific phase. During S and G₂ phases of the cell cycle, the cdc2-cyclin B complex is kept in the inactive state through Tyr¹⁵ phosphorylation of cdc2 by Wee1/Mik1/Myt1 tyrosine kinases (32, 33); however, during the G₂-M transition, cdc2 is rapidly converted into the active form by Tyr¹⁵ dephosphorylation catalyzed by cdc25 tyrosine phosphatases (47, 48). Increased levels of Tyr¹⁵-phosphorylated cdc2 have been shown to be associated with cell cycle arrest following DNA damage in many cell culture systems (42, 49). Consistent with these reports and with the activation of ATM-Chk2 by gallic acid, our results also show that gallic acid induces the inactivating phosphorylation of cdc25C (Ser²¹⁶) and cdc25A (Ser¹⁷). Moreover, as a downstream effect, Tyr¹⁵-phosphorylated cdc2 did not get dephosphorylated by cdc25 phosphatases, keeping cdc2 (Tyr¹⁵) in an inactive form that leads to a resultant cell cycle arrest. In summary, the present study identifies gallic acid efficacy and associated mechanisms in an advanced and androgen-independent human prostate carcinoma DU145 cells, suggesting future in vivo efficacy studies with this ubiquitous dietary agent in preclinical prostate cancer models.