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© 2006 American Society for Nutrition J. Nutr. 136:2728-2734, November 2006

Biochemical, Molecular, and Genetic Mechanisms

Benzyl Isothiocyanate-Induced DNA Damage Causes G2/M Cell Cycle Arrest and Apoptosis in Human Pancreatic Cancer Cells1

Ruifen Zhang, Sivakumar Loganathan, Ian Humphreys and Sanjay K. Srivastava*

Department of Pharmacology and University of Pittsburgh Cancer Institute, University of Pittsburgh School of Medicine, Pittsburgh, PA 15213

* To whom correspondence should be addressed. E-mail: srivastavask{at}upmc.edu.


   ABSTRACT
TOP
 ABSTRACT
Introduction
 Materials and Methods
 Results
 Discussion
 LITERATURE CITED
 
Benzyl isothiocyanate (BITC) has been shown to inhibit chemically induced pancreatic cancer in experimental animals. However, the mechanism responsible for the anticancer effects of BITC is not clearly understood. In this study, we tested whether BITC treatment would affect the growth of Capan-2 human pancreatic cancer cells. BITC (10 µmol/L) treatment caused marked phosphorylation of H2A.x (2.6-fold) and permanent damage to Capan-2 cells. BITC-mediated G2/M arrest was associated with up-regulation of cyclin dependent kinase inhibitor p21Waf1/Cip1 and the activation of checkpoint kinase 2, whereas the expressions of other G2/M regulatory proteins, including CyclinB1, Cdc2, and cell division cycle 25C (Cdc25C), were down-regulated by 19, 51, and 70%, respectively, compared with control. These changes resulted in a 55% inhibition of Cdc2 kinase activity. In addition, the decline in the expression of Cdc25C was completely blocked when the cells were treated with lactacystin (proteasome inhibitor) prior to BITC treatment. However, G2/M arrest and apoptosis induced by BITC were partially blocked by pretreatment of cells with lactacystin. Taken together, the results of this study suggest the involvement of multiple signaling pathways targeted by BITC in mediating G2/M cell cycle arrest and apoptosis in Capan-2 cells and warrant further investigation.


   Introduction
TOP
 ABSTRACT
 Introduction
Materials and Methods
 Results
 Discussion
 LITERATURE CITED
 
Epidemiological and pharmacological studies continue to support the notion that isothiocyanates present in cruciferous vegetables have substantial chemopreventive activity against a variety of malignancies including pancreatic cancer (1–4). The major source of isothiocyanates present in the diet comes from Brassica vegetables such as broccoli, watercress, and cabbage. Isothiocyanates exist as glucosinolates in these vegetables and are released by the action of the enzyme myrosinase once the plant cells are damaged either by cutting or chewing (5). Various studies have suggested that isothiocyanates are protective against chemically induced cancers in experimental animals by blocking the initiation process (6–10). For example, benzyl isothiocyanate (BITC)2 and phenethyl isothiocyanate (PEITC) inhibit lung and esophageal tumorigenesis by tobacco-derived carcinogens in rats and mice, respectively. Mechanistically, isothiocyanates exert chemopreventive effects by modulating phase I drug-metabolizing enzymes and phase II detoxification enzymes (6,11,12). In addition to preventing the initiation phase of the carcinogenesis process, isothiocyanates also inhibit the promotion phase by inducing cell cycle arrest and apoptosis in cancer cells.

We and others have recently shown that isothiocyanates are antiproliferative to cancer cells, with little or no toxicity toward normal cells, making this class of compounds ideal chemopreventive agents against various malignancies (13–22). Isothiocyanates have been shown to induce apoptosis in cancer cells with diverse mechanism of action. For example, PEITC induces apoptosis in both p53-dependent and -independent manner in JB6 and PC-3 cells (14,17). Similarly, activation of mitogen activated protein kinase pathways such as ERK, JNK, and p38 by PEITC and BITC were reported to be the possible mechanisms of growth arrest and apoptosis in PC-3, HL-60, and Jurkat cells (17,23,24). Likewise, activation of mitochondrial death pathway by BITC has also been documented in RL34 cells undergoing apoptosis (25).

Accumulating data suggest the involvement of cell cycle arrest by different mechanisms in inducing cell death by various isothiocyanates (18–28); however, not much is known about the induction of cell cycle checkpoint mechanisms by isothiocyanates. Cell cycle checkpoints are important growth arrest mechanisms that ensure the orderly progression of cell cycle events and prevent aberrant mitosis in response to DNA damage. Fragmented studies suggest the growth inhibitory effects of BITC in cancer cells. For example, cell cycle arrest of HL-60 cells in response to BITC treatment was mediated by up-regulating the expression of the G2/M cell cycle arrest-related genes (29). Similarly, induction of p21 by BITC was associated with the arrest of Caco-2 cells (30). A recent study also linked the involvement of p38 mitogen activated protein kinase in BITC-induced G2/M arrest in human T-cell leukemia cells (24). Nevertheless, the mechanism by which BITC causes DNA damage that leads to G2/M arrest and whether the effect is temporary or permanent remain unknown. This study therefore aimed to establish the molecular mechanism of the growth suppressive effects of BITC in Capan-2 human pancreatic cancer cells.


   Materials and Methods
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 ABSTRACT
 Introduction
 Materials and Methods
Results
 Discussion
 LITERATURE CITED
 
    Chemicals. BITC was obtained from Sigma-Aldrich. RNase A, propidium iodide, and antibodies against actin were from Sigma. Lactacystin was from Calbiochem. Electrophoresis reagents were from Bio-Rad Laboratories. Antibodies against CyclinB1, Cdc-2, phospho-Cdc-2 (Tyr-15), checkpoint kinase 2 (Chk2), phospho-Chk2 (Thr-68), cell division cycle 25C (Cdc25C), phospho-Cdc25C (Ser-216), phospho-H2A.x (Ser-139), Caspase-3, and poly(ADP-ribose)polymerase (PARP) were from Cell Signaling Technology. Cell culture medium, penicillin/streptomycin antibiotic mixture, and heat-inactivated fetal bovine serum were purchased from GIBCO BRL and Invitrogen.

    Cell culture and proliferation assays. Capan-2 cells were obtained from ATCC. This is a well-differentiated epithelial pancreatic adenocarcinoma cell line obtained from a male Caucasian donor having wild-type p53 and p16 and mutated K-ras. Acinar cells were isolated from normal human pancreas and provided by Dr. Massimo Trucco (University of Pittsburgh). Monolayer culture of Capan-2 cells were maintained in McCoy medium supplemented with 10% fetal bovine serum and antibiotics and mixture of acinar and ductal cells were cultured in CMRL 1066 medium (GIBCO BRL) in a humidified incubator with 5% CO2 and 95% air. Stock solution of BITC was prepared in 100% dimethyl sulfoxide (DMSO) and subsequently diluted in medium so that the final concentration of DMSO was <0.2% in the medium. The cells were treated with BITC for 12, 24, 48, or 72 h. Effect of BITC on proliferation of Capan-2 or acinar and ductal cells was determined by Sulforhodamine B assay as described previously (22). The plates were read at 570 nm with a Bio Kinetics plate reader.

    Cell cycle analysis. The effect of BITC on cell cycle distribution was assessed by flow cytometry after staining the cells with propidium iodide. Briefly, 0.5 x 106 cells were plated and allowed to attach overnight. The medium was replaced with fresh complete medium containing the desired concentration of BITC and equal volume of DMSO was added to controls so that final concentration of DMSO was <0.2%. After incubation of cells at 37°C for specified time, floating and adherent cells were collected by using 0.1% trypsin, washed twice with cold PBS, and fixed with ice-cold 70% ethanol overnight at 4°C. The cells were then treated with 80 mg/L RNase A and 50 mg/L propidium iodide for 30 min. The stained cells were analyzed using a Coulter Epics XL Flow Cytometer. In another experiment, to establish whether the cell cycle arrest induced by BITC is permanent, cells were treated with 5 or 10 µmol/L BITC for 24 h and then further cultured in fresh BITC-free medium for an additional 48 h and processed for cell cycle distribution. Control cells were treated with DMSO and cultured in the similar fashion. The cell cycle data were reanalyzed using MODFIT software.

    Apoptosis determination. Apoptosis induction in control and BITC-treated Capan-2 cells was determined by flow cytometry by quantitating: 1) the cells after staining with annexinV-fluorescein isothiocyanate and propidium iodide, or 2) the cells with sub G0/G1 DNA content following staining with propidium iodide, as described above for cell cycle analysis. Briefly, 0.5 x 106 cells were plated and allowed to attach overnight. After treatment of cells with BITC at 37°C for specified time, floating and adherent cells were collected by using 0.1% trypsin, washed twice with cold PBS, and suspended in 500 µL binding buffer (10 mmol/L HEPES buffer, pH 7.4, 140 mmol/L NaCl, and 2.5 mmol/L CaCl2). The cells were then treated with 5 µL of Annexin V-fluorescein isothiocyanate and 10 µL propidium iodide (50 mg/L) and incubated in dark for 15 min. The stained cells were analyzed using a Coulter Epics XL Flow Cytometer.

    Lactacystin treatment. Cells were treated with 5 µmol/L lactacystin for 2 h at 37°C and then subsequently exposed to 10 µmol/L BITC for 24 h without removing lactacystin. Control cells received DMSO only. Subsequently, cells were collected, washed with PBS, and processed for determination of cell cycle distribution or apoptosis as described above. In a separate experiment, control and treated cells were collected, lysed, and subjected to western blotting for various G2/M regulatory proteins.

    Western-blot analysis. Cells were exposed to varying concentrations of BITC for the indicated time periods as described above. The cells were washed twice with ice-cold PBS, lysed on ice with a solution containing 50 mmol/L Tris, 1% Triton X-100, 0.1% SDS, 150 mmol/L NaCl, 2 mmol/L Na3VO4, 2 mmol/L EGTA, 12 mmol/L ß-glycerol phosphate, 10 mmol/L NaF, 16 mg/L benzamidine hydrochloride, 10 mg/L phenanthroline, 10 mg/L aprotinin, 10 mg/L leupeptin, 10 mg/L pepstatin, and 1 nmol/L phenyl methyl sulfonyl fluoride. The cell lysate was cleared by centrifugation at 14,000 x g; 15 min. Protein content in the supernatant fraction was determined by the method of Bradford (31). Lysate containing 20 to 80 µg protein was subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis according to the method of Laemmli (32) and the proteins were transferred onto polyvinylidene fluoride membrane (33). After blocking with 10% nonfat dry milk in Tris buffered saline, the membrane was incubated overnight with the desired primary antibody. Subsequently, the membrane was incubated with appropriate secondary antibody, and the immunoreactive bands were visualized using enhanced chemiluminescence kit (NEN Life Science Products) according to the manufacturer's instructions. The same membrane was reprobed with the antibody against actin (1:50000 dilution) that was used as an internal control for equal protein loading.

    Cdc2 kinase activity. Control and BITC-treated Capan-2 cells were lysed on ice by lysis buffer containing 50 mmol/L Tris, pH 7.4, 1% Triton X-100, 150 mmol/L NaCl, 2 mmol/L Na3VO4, 2 mmol/L EDTA, 12 mmol/L glycerol phosphate, 10 mmol/L NaF, 10 mg/L aprotinin, 10 mg/L leupeptin, 10 mg/L pepstatin, and 1 nmol/L phenyl methyl sulfonyl fluoride. Approximately 500 µg protein lysate was incubated with 3 µg Cdc2 antibody for 2 h at 4°C followed by the addition of 35 µL protein A agarose and the complex was rocked gently overnight at 4°C. Cdc2 kinase activity was essentially measured using a Cdc2 kinase assay kit (Upstate) according to the manufactures instructions.

    Densitometric scanning and statistical analysis. The intensity of immunoreactive bands was determined using a densitometer (Molecular Dynamics) equipped with Image QuaNT software. Results are expressed as means ± SEM of at least 2 independent experiments, each conducted in triplicate. Data were analyzed by ANOVA followed by Bonferroni's post hoc analysis for multiple comparisons. All statistical calculations were performed using InStat software and GraphPad Prizm 4.0. Differences between control and BITC treatment were analyzed by 1-way ANOVA. The effect of BITC treatment when compared with cells cultured in BITC-free medium after single BITC exposure, or in combination with/without lactacystin, was analyzed by 2-way ANOVA. Differences were considered significant at P < 0.05.


   Results
TOP
 ABSTRACT
 Introduction
 Materials and Methods
 Results
Discussion
 LITERATURE CITED
 
    Antiproliferative effect of BITC. Treatment of Capan-2 cells with increasing concentrations of BITC for 24 h significantly reduced the survival of cells with an IC50 of 10 ± 0.5 µmol/L (Fig. 1). However, we observed a differential response of BITC in Capan-2 cells (Fig. 1). Treatment of cells with 0–5 µmol/L BITC revealed a modest but significant 25% growth inhibition compared with the control. The maximum inhibition (~60%) was observed in the cells treated with 5–10 µmol/L BITC, whereas a modest growth inhibition was observed at the higher 10–40 µmol/L BITC concentration. Nonetheless, a small percentage of cells escaped the growth inhibitory effects of BITC. We obtained similar results by Trypan blue dye exclusion assay (data not shown). On the other hand, survival of acinar cells was not affected following exposure to BITC up to 40 µmol/L, concentrations which were very toxic to Capan-2 cells (Fig. 1).


Figure 1
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  		Figure 1  Effect of BITC on the proliferation of Capan-2 and acinar cells. Values are means ± SEM of 3 independent experiments (each conducted in triplicate). Means without a common letter differ, P < 0.05.

 
    BITC induces apoptosis in Capan-2 cells. A 24-h treatment of cells with 10 or 20 µmol/L BITC resulted in the significant increase in cells undergoing apoptosis compared with the controls (Fig. 2A). The time course treatment of Capan-2 cells also revealed that in comparison to control cells, the percentage of sub G0/G1 cells increased significantly in BITC-treated cells in a time-dependent manner (Fig. 2B). Compared with DMSO-treated controls, the number of apoptotic cells was ~3.7-fold greater in BITC-treated cells (10 µmol/L, 24 h) (Fig. 2B). To confirm the induction of apoptosis by BITC, we determined the activation of caspase-3 and PARP in the control and BITC-treated cells by western blotting. Treatment of Capan-2 cells with BITC for 24 h resulted in the activation of caspase-3 and PARP, as is apparent by the appearance of its cleaved products at 19 and 17 kDa (caspase-3) and 89 kDa (PARP) (Fig. 2C), suggesting that apoptosis induced by BITC in these cells is mediated by caspase-3 cascade.


Figure 2
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  		Figure 2  Effects of dose of BITC (A) and time of incubation (B) on the induction of apoptosis in Capan-2 cells. To validate BITC-induced apoptosis, cleavage of caspase-3 and PARP was determined by western-blot analysis (C). Blots were stripped and reprobed with anti-actin antibody to ensure equal protein loading. Values are means ± SEM of 2 independent experiments (each conducted in triplicate). Means without a common letter differ, P < 0.05.

 
    BITC causes G2/M cell cycle arrest. To gain further insight into the mechanism of growth inhibitory effects of BITC, we assessed cell cycle distribution of Capan-2 cells by flow cytometry (Table 1). Treatment of Capan-2 cells with varying concentrations of BITC for 24 h resulted in the increased accumulation of the cells in G2/M phase (Fig. 3A). Incubation of cells with 10 µmol/L BITC for 24 h caused ~42% enrichment of cells in G2/M phase compared with the controls. In a time-dependent experiment, we observed maximum accumulation of cells in the G2/M phase after treatment of cells with 10 µmol/L BITC for 24 h (Fig. 3B), which was reduced after 48 and 72 h of exposure time (Fig. 3B). The DMSO-treated control cells remained unchanged in G2/M phase (Fig. 3B). Interestingly, the decrease in the percentage of cells in G2/M phase after a 48- or 72-h treatment with 10 µmol/L BITC was associated with a concomitant increase in apoptosis (Fig. 2B).


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  		TABLE 1 Effect of BITC on the cell cycle distribution of Capan-2 cells1

 

Figure 3
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  		Figure 3  Effect of BITC on the cell cycle distribution in Capan-2 cells. The cells were treated with BITC at indicated dose (A) or time period (B). Values are means ± SEM of 2 independent experiments (each conducted in triplicate). Means without a common letter differ, P < 0.05.

 
    BITC treatment causes permanent damage to the cells. A significant 26% increase in the number of apoptotic cells (sub G0/G1) was associated with a 22% decline of cells in G2/M phase in 10 µmol/L BITC-treated cells cultured for an additional 48 h in fresh BITC-free medium compared with 24-h BITC-treated cells (Fig. 4A,B). The percentage of DMSO-treated control cells in G2/M phase did not change following culture in fresh medium and no difference was observed in the population of sub G0/G1 cells either. We further confirmed these observations by performing cell viability assay in both treatment regimens as described above. Consistent with these observations, we found a significant 25% reduced viability of BITC-treated cells. We then cultured these cells in fresh medium for an additional 48 h and compared them with cells treated with 10 µmol/L BITC for 24 h (Fig. 4C). The difference between the 2 treatments became greater between 10–20 µmol/L BITC (Fig. 4C). These results indicate that BITC-mediated permanent damage leads to increased apoptosis and reduced survival of Capan-2 cells. Therefore, even though cells were allowed to grow in fresh medium for 48 h after a single exposure to BITC, the cells did not recover from the damage incurred by BITC and eventually died.


Figure 4
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  		Figure 4  BITC induces permanent damage in Capan-2 cells. The cells were treated with DMSO or 10 µmol/L BITC for 24 h. Subsequently, the cells were analyzed or cultured in BITC-free medium for additional 48 h before being subjected to cell cycle analysis for the quantitation of cells in G2/M phase (A) and with sub-G0/G1 DNA content (B). These results were further confirmed by cell viability assay (C). Means without a common letter differ, P < 0.05.

 
    BITC treatment causes modulation of G2/M regulatory proteins. First, we sought to determine whether BITC treatment causes DNA damage by examining the phosphorylation of H2A.x at Ser-139, which is considered to be an important marker for the presence of DNA double-strand breaks. BITC treatment resulted in the significant phosphorylation of H2A.x (Fig. 5A). Compared with DMSO-treated cells, 10 µmol/L BITC treatment for 24 h resulted in ~2-fold greater phosphorylation of H2A.x at Ser-139 as shown by western blotting (Fig. 5B), indicating that BITC treatment causes DNA double-strand breaks in Capan-2 cells.


Figure 5
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  		Figure 5  Effect of BITC on G2/M regulatory proteins in Capan-2 cells. Representative immunoblots show the effect of BITC treatment on the phosphorylation of H2A.x (Ser-139), Chk2 (Thr-168), Cdc25C (Ser-216) and Cdc2 (Tyr-15) and protein expression of Chk2, Cdc25C, Cdc2, Cyclin B1 and p21Waf1/Cip1. Each blot was stripped and reprobed with anti-actin antibody to ensure equal protein loading. Intensities of the immunoreactive bands were quantified by densitometric scanning. Values are means ± SEM of 3 independent experiments. *Different from the control, P < 0.05.

 
The next logical step was to determine whether BITC-mediated DNA damage activates the Chk2 pathway. Indeed, treatment of cells with either 10 or 20 µmol/L BITC increased phosphorylation of Chk2 at Thr-68 by ~60% relative to controls. The Chk2 protein expression was also up-regulated in response to BITC treatment (Fig. 5A,B).

Further, we determined the effect of BITC treatment on the protein expression and phosphorylation of Cdc25C at Ser-216. The phosphorylation and protein expression of Cdc25C was drastically reduced following treatment of cells with BITC for 24 h (~70% reduction relative to the control), indicating its role in BITC-mediated G2/M arrest (Fig. 5A,B).

Similarly, treatment of cells with 10 µmol/L BITC resulted in ~30% reduced phosphorylation of Cdc2 at Tyr-15 compared with control (Fig. 5A,C). Protein expression of Cdc2 was 20–75% lower in BITC-treated cells than in control cells (Fig. 5A,C). However, the expression of Wee-1 did not differ between cells treated with BITC and controls (data not shown).

The activation of Cdc2/CyclinB1 complex is the rate-limiting factor for cells to enter into mitosis, whereas its inactivation leads to G2/M arrest. Treatment of Capan-2 cells with 10 µmol/L BITC for 24 h resulted in the inhibition of ~55% of Cdc2 kinase activity compared with DMSO-treated control cells (data not shown).

Several recent studies suggest that p21Waf1/Cip1 regulate the entry of cells at DNA damage-induced G2/M checkpoint and induce apoptosis (34–37). Western-blot analysis revealed that BITC treatment of the cells for 24 h resulted in a marked induction of the protein expression of p21Waf1/Cip1 (up to 6-fold higher) in a dose-dependent manner compared with DMSO-treated control cells (Fig. 5A,C), indicating its involvement in BITC-mediated G2/M arrest.

    Effect of lactacystin on BITC-induced G2/M arrest and apoptosis. The drastic reduction in the protein expression of Cdc25C mediated by BITC treatment was completely prevented in the cells, which were pretreated with 5 µmol/L lactacystin, a specific proteasome inhibitor, for 2 h prior to treatment with 10 µmol/L BITC for 24 h (Fig. 6A). In addition, high molecular weight conjugates were observed in the blot that was reprobed against ubiquitin antibody (Fig. 6A). These results indicate that BITC-mediated degradation of Cdc25C involves the ubiquitin/proteasomal pathway.


Figure 6
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  		Figure 6  Effect of lactacystin on BITC induced degradation of Cdc25C. Western-blot analysis for the detection of Cdc25C expression and presence of high molecular weight polyubiquitin conjugates (A). Blots were stripped and reprobed with anti-actin antibody to ensure equal protein loading. Cells were also analyzed by flow cytometry for cell cycle distribution (B) and apoptosis (C). Values are means ± SEM of 2 independent experiments (each conducted in triplicate). Means without a common letter differ, P < 0.05.

 
The next step was to determine whether blocking Cdc25C degradation could prevent BITC-mediated G2/M arrest and apoptosis. Interestingly, lactacystin treatment offered only modest (10% reduction compared with BITC treatment) protection against BITC-mediated G2/M arrest (Fig. 6B). Similarly, lactacystin offered ~15% protection against BITC-induced apoptosis (Fig. 6C).


   Discussion
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 Introduction
 Materials and Methods
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 Discussion
LITERATURE CITED
 
To ensure proper progression through the cell cycle, cells have evolved a series of checkpoints that prevent them from entering into a new phase until they have successfully completed the previous one. These checkpoints allow progression through the cell cycle or arrest the cells in G2/M phase in response to DNA damage for DNA repair (35,36). The cell cycle arrest may lead to apoptotic cell death in case of severe DNA damage. Our study demonstrates that BITC exposure led to G2/M cell cycle arrest and apoptosis in Capan-2 cells. Moreover, cells cultured for 48 h in BITC-free medium after 24-h exposure to BITC exhibited increased apoptosis compared with 24-h treatment with BITC. These results suggest that DNA damage in the cells caused by BITC may be irreparable and therefore force the arrested cells into apoptosis. Further, we observed that BITC treatments resulted in the increased phosphorylation of H2A.x at Ser-139 in Capan-2 cells, suggesting the presence of DNA double-strand breaks. Previous studies have suggested that reactive oxygen species can directly cause DNA damage as well as oxidize nucleotides, which can be converted to double-strand breaks during replication (38,39). In our unpublished studies, treatment of Capan-2 cells with 10 µmol/L BITC resulted in the generation of reactive oxygen species, which may in part be one plausible mechanism for inducing double-strand breaks.

Our data showing the differential dose response of BITC in cell proliferation and apoptosis is quite intriguing. Cells treated with 5 µmol/L BITC were significantly retained in the G2/M phase (Fig. 3A) and thus prevented further division and left fewer cells compared with the control (Fig. 1). This might be one possible explanation why a 5 µmol/L BITC treatment reduces cell survival but does not induce apoptosis. On the other hand, at higher BITC concentration, both cell cycle arrest and apoptosis were significantly higher compared with control cells, resulting in sharp decline in the cell growth.

DNA damage checkpoint is associated with activation of Chk2 kinase, which in turn phosphorylates and inactivates Cdc25C, further allowing the inactivation of Cdc2-CyclinB1 complex leading to G2/M arrest (34,35). The results of this study indicate that treatment of Capan-2 cells with BITC results in the activation of Chk2 kinase and significantly decreased the protein expression of Cdc25C, Cdc-2, and CyclinB1, as reported previously (40). However, we did not observe increased phosphorylation of Cdc25C at Ser-216 and Cdc2 at Tyr-15 following activation of Chk2 as shown in other studies (26). This raises the possibility that BITC-mediated G2/M arrest in our model may be due to reduced interaction between Cdc2 and CyclinB1, which leads to the inhibition of Cdc2 kinase activity.

The sharp decline in the protein level of Cdc25C in Capan-2 cells was proteasome mediated, which was blocked once the cells were pretreated with lactacystin (a specific proteasome inhibitor). Surprisingly, lactacystin treatment did not significantly protect the cells from BITC-induced G2/M arrest and apoptosis, suggesting the presence of other pathways contributing to strong growth inhibitory effects of BITC in this cell line. The inactivation of Cdc2-CyclinB1 complex by its subsequent binding with cyclin-dependent kinase inhibitor p21Waf1/Cip1 is one of the probable mechanisms of G2/M arrest (35–37). Our data demonstrate a significant up-regulation of p21Waf1/Cip1, indicating its role in BITC-mediated G2/M arrest in Capan-2 cells. The p21Waf1/Cip1 is known to regulate G1 phase of the cell cycle, but recent evidence suggests that it negatively regulates G2/M phase as well (35–37). Based on the results of our study, a possible mechanism by which BITC induces G2/M arrest and apoptosis in Capan-2 cells is summarized in Figure 7.


Figure 7
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  		Figure 7  Proposed mechanism of BITC-induced G2/M arrest and apoptosis in Capan-2 cells. BITC-induced DNA damage results in the activation of Chk2, up-regulation of p21, and decline in the expression of Cdc25C and Cdc-2, which leads to G2/M arrest and eventually apoptosis.

 
Our data are consistent with previous cellular studies using other isothiocyanate analogs such as AITC, PEITC, and sulforaphane, where cell growth arrest and induction of apoptosis were observed at isothiocyanate concentrations of 50 µmol/L or lower (17–19,26). Because the pharmacokinetics of BITC in humans has not been determined, it is difficult to predict how much Brassica vegetable would need to be consumed to achieve a circulating concentration of 10 µmol/L BITC. However, a very recent study suggested that orally feeding male Sprague-Dawley rats with 10 or 100 µmol PEITC/kg (an analogue of BITC) resulted in rapid absorption and reached a peak concentration of 9.2 ± 0.6 and 42.1 ± 11.4 µmol/L PEITC, respectively, in the plasma after 0.44 ± 0.1 and 2.0 ± 1 h of PEITC feeding, respectively, suggesting that micromolar concentrations may be achieved in vivo (41). In another pharmacokinetics study, 4 human volunteers were fed with a single dose of myrosinase-hydrolyzed extract of 3-d-old broccoli sprouts (containing ~200 µmol of total isothiocyanates), and a peak concentration of 0.94–2.27 µmol/L isothicyanates was reached in the plasma, serum, and erythrocytes 1 h after broccoli extract ingestion (42). More pharmacokinetic studies of BITC are needed before conducting clinical testing of BITC as a cancer chemopreventive agent.

Our study reveals the chemotherapeutic effects of BITC against human pancreatic cancer cells. Our data demonstrate that G2/M arrest and apoptosis induced by BITC in these cells may be mediated by the following interrelated mechanisms: 1) DNA damage; 2) activation of Chk2 and down-regulation of key G2/M regulators such as Cdc25C, Cdc-2, and CyclinB1; and 3) induction of cyclin-dependent kinase inhibitor p21Waf1/Cip1. These observations are in agreement with the overall effectiveness of the growth suppressive effects of BITC in Capan-2 cells. Nevertheless, further studies are needed to determine the mechanism of DNA damage and pinpoint the pivotal regulator(s) of the pathway mediated by BITC in Capan-2 cells.


   ACKNOWLEDGMENTS
 
The authors thank Dr. Massimo Trucco, University of Pittsburgh, for providing acinar cells isolated from normal human pancreas and Jeffrey Richards for technical assistance.


   FOOTNOTES
 
1 Supported by USPHS RO1 grant CA106953 (to S.K.S.) awarded by the National Cancer Institute. The startup fund from University of Pittsburgh Cancer Institute (to S.K.S.) is also acknowledged. Back

2 Abbreviations used: BITC, benzyl isothiocyanate; Cdc25C, cell division cycle 25C; Chk2, checkpoint kinase 2; DMSO, dimethyl sulfoxide; PARP, poly(ADP-ribose)polymerase; PEITC, phenethyl isothiocyanate. Back

Manuscript received 6 June 2006. Initial review completed 10 July 2006. Revision accepted 30 August 2006.


   LITERATURE CITED
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 ABSTRACT
 Introduction
 Materials and Methods
 Results
 Discussion
 LITERATURE CITED
 

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