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The Histone Deacetylase Inhibitor Suberoylanilide Hydroxamic Acid Induces Growth Inhibition and Enhances Gemcitabine-Induced Cell Death in Pancreatic Cancer

Authors' Affiliation: Departments of Medicine, and Pharmacology and Toxicology, Dartmouth-Hitchcock Medical Center, Dartmouth Medical School, Hanover, New Hampshire

Requests for reprints: Murray Korc, Department of Medicine, Dartmouth-Hitchcock Medical Center, Lebanon, NH 03756. Phone: 603-650-7936; Fax: 603-650-6122; E-mail: Murray.Korc{at}Dartmouth.edu.

	Abstract

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Purpose: Pancreatic cancer is an aggressive human malignancy that is generally refractory to chemotherapy. Histone deacetylase inhibitors are novel agents that modulate cell growth and survival. In this study, we sought to determine whether a relatively new histone deacetylase inhibitor, suberoylanilide hydroxamic acid (SAHA), inhibits pancreatic cancer cell growth.

Experimental Design: The effects of SAHA on the growth of three pancreatic cancer cell lines (BxPC3, COLO-357, and PANC-1) were examined with respect to cell cycle progression, p21 induction and localization, and interactions with the nucleoside analogue gemcitabine.

Results: SAHA induced a G₁ cell cycle arrest in BxPC-3 cells and COLO-357 cells but not in PANC-1 cells. This arrest was dependent, in part, on induction of p21 by SAHA, as p21 was not induced in PANC-1 cells, and knockdown of p21 using small interfering RNA oligonucleotides nearly completely suppressed the effects of SAHA on cell cycle arrest in COLO-357 and partly attenuated the effects of SAHA in BxPC-3. COLO-357 and BxPC-3 cells, but not PANC-1 cells, were also sensitive to gemcitabine. In the gemcitabine-resistant PANC-1 cells, a 48-h cotreatment with SAHA rendered the cells sensitive to the inhibitory and proapoptotic effects of gemcitabine. An additive effect on growth inhibition by SAHA and gemcitabine was observed in COLO-357 and BxPC-3 cells. Moreover, analysis of p21 distribution in COLO-357 cells revealed that SAHA induced the cytoplasmic localization of both p21 and phospho-p21.

Conclusions: These data indicate that SAHA exerts proapoptotic effects in pancreatic cancer cells, in part, by up-regulating p21 and sequestering it in the cytoplasm, raising the possibility that SAHA may have therapeutic potential in the treatment of pancreatic cancer.

Pancreatic ductal adenocarcinoma is characterized by mutations and/or silencing of tumor suppressor genes, such as and p53 and Smad4, the overexpression of mitogenic growth factors and their cognate high affinity tyrosine kinase receptors, and mutation of K-ras (6). There are also defects in cell cycle–regulating genes, such as the increased expression of cyclin D1 and mutation/silencing of p16, which contribute to the inactivation of the retinoblastoma protein (7). These alterations contribute to the excessive growth of pancreatic tumors and to the resistance of pancreatic cancers to chemotherapeutic agents (8).

Histone acetylation is a posttranslational modification of the nucleosomal histones that affects chromatin structure and modulates gene expression. The acetylation status of histones is modulated by histone acetyl transferases and histone deacetylases (HDAC). Histone acetyl transferases are generally considered to be transcriptional activators because histone acetylation is associated with transcriptionally active chromatin, whereas HDACs are considered as transcriptional inhibitors because histone deacetylation is associated with transcription repression (9, 10). Moreover, HDAC activity is increased in cancer cells and this increase has been shown to induce oncogenic transformation by modulating the function of transcription factors, such as p53 and nuclear factor-B (11), resulting in altered gene transcription and increased proliferation (12). HDAC inhibitors (HDACI) represent a new class of targeted anticancer agents, and several structural classes of HDACI have been developed and are in clinical trials. Second-generation, novel compounds that inhibit class I/II HDAC are promising new anticancer agents due to their low toxicity and high tolerance in cancer treatment (13). These compounds have been shown to inhibit growth, induce differentiation, and decrease the survival of transformed/cancer cells but not normal cells (14, 15). One such compound, suberoylanilide hydroxamic acid (SAHA), induces differentiation, growth arrest, or apoptosis of transformed human cells in culture at micromolar concentrations (16). SAHA was originally identified based on its ability to induce differentiation of murine erythroleukemia cells (17). Subsequently, it was found to induce differentiation of human breast adenocarcinoma cells (18) and growth arrest in human prostate carcinoma (19, 20) and bladder transitional cell carcinoma cells (21). SAHA induces apoptosis in transformed hematopoietic cells, including Jurkat, CEM, and ARP-1 cells (22–24).

In the present study, we examined the effects of SAHA on three human pancreatic cancer cell lines. We report that BxPC3 and COLO-357 cells are sensitive to the growth-suppressive effects of SAHA. We further show that SAHA can be used to sensitize pancreatic cancer cells to the apoptosis-inducing effects of gemcitabine. These results indicate that SAHA may have a therapeutic potential in pancreatic cancer.

	Materials and Methods

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Cell culture. The human pancreatic cancer cell lines PANC-1 and BxPC-3 were purchased from American Type Culture Collection (Manassas, VA). BxPC-3 was maintained in RPMI 1640 supplemented with 10% fetal bovine serum, 100 units/mL penicillin, and 100 µg/mL streptomycin (Irvine Scientific, Irvine, CA) in 5% CO₂. COLO-357 cells (a gift from R.S. Metzger, Duke University, Durham, NC) and PANC-1 cells were maintained in DMEM supplemented with 10% fetal bovine serum, 100 units/mL penicillin, and 100 µg/mL streptomycin in 5% CO₂. For all experiments, the cell lines were treated in serum-free medium (0.1% bovine serum albumin, 5 µg/mL transferrin, and 5 ng/mL sodium selenite and antibiotics). SAHA was purchased from BioMol (Plymouth Meeting, PA), and gemcitabine was purchased from Ely Lilly.

Immunoblotting. Cells were washed and lysed in lysis buffer in the presence of a protease inhibitor solution consisting of 1 mmol/L phenylmethylsulfonyl fluoride, 1 µg/mL aprotinin, 1 µg/mL leupeptin, and 1 µg/mL pepstatin. Lysates were then subjected to SDS-PAGE, transferred to nitrocellulose membranes (25 µg/lane), and incubated with the indicated antibodies. The anti–poly(ADP-ribose) polymerase (PARP) antibody was purchased from Cell Signaling Technologies (Beverly, MA). Anti-p21 and anti–acetylated lysine-specific histone H3 antibodies were purchased from Upstate Biotechnologies (Waltham MA). Membranes were then stripped and probed with an anti–extracellular signal-regulated kinase 2 (ERK2) antibody from Santa Cruz Biotechnology (Santa Cruz, CA) to normalize for protein loading (25).

Flow cytometry. Cells were plated in six-well plates at a density of 100,000 per well and allowed to adhere overnight. Cells were treated with 10 µmol/L SAHA in serum-free medium for 48 h. Cells were then trypsinized and fixed in ice cold 70% ethanol and stained with propidium iodide in PBS. Flow cytometric data were acquired using a FACSCan analysis system equipped with a FACStation, MAC PowerPC computer and CellQuest Acquisition software from Becton Dickinson (San Jose, CA). Data were analyzed using ModFit LT 2.0 software from Verity Software House (Topsham, ME) as reported previously (26).

Growth assay. The 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide growth assay (27), which in pancreatic cancer cells correlates well with results obtained by cell counting with a hemocytometer and by monitoring [³H]thymidine incorporation (28, 29), was used to monitor cell growth. Cells were plated at a density of 10,000 per well, using 96-well microtiter plates (Corning, Acton, MA), and allowed to adhere overnight before SAHA addition for 48 h. Colorimetric changes were measured using a microtiter plate reader with a 570 nm filter.

p21 Small interfering RNA. BxPC-3 and COLO-357 cells were transfected with 20 mmol/L SMARTpool p21small interfering RNA (siRNA; Dharmacon, Lafayette, CO) and jetPEI using the recommended fast protocol (Qbiogene, Solon, OH). Cells were plated to 60-mm dishes and allowed to adhere overnight. The following day, the cells were split to 96-well plates for a 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay or 60-mm dishes for Western blot analysis. The cells were then treated with 10 µmol/L SAHA for 48 h. Cell proliferation was determined using a 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay (described above), and p21 expression was determined by Western blotting.

Measurement of caspase-3 activity. Caspase-3 activity was measured using Colorimetric Assay kits from R&D Systems (Minneapolis, MN). In brief, cells were scraped into PBS, pelleted at low speed, and resuspended in lysis buffer for 10 min at 4°C. Cell lysates were cleared by centrifugation and assayed for caspase-3 activity using a DEVDpNA peptide substrate and incubated for 6 h at 37°C. The activities were quantified spectrophotometrically at a wavelength of 405 nm. Caspase activity was calculated as the change in absorbance at 405 nm and divided by total protein concentration.

Confocal microscopy. Cells were seeded overnight on Lab-Tek chamber slides (Nalge Nunc, Rochester, NY) followed by a 48-h incubation in the absence or presence of SAHA. Cells were then washed twice with PBS and fixed for 10 min with 2% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA) in PBS, washed with PBS, and incubated with 50 nmol/L NH₄Cl in PBS for 10 min. After washing with PBS, cells were permeabilized for 15 min with 0.1% Triton X-100/2% bovine serum albumin in PBS and incubated with anti-p21 or anti–phospho-p21 antibody (1:500 dilution) for 1 h at 23°C. Cells were then washed twice with PBS and incubated for 30 min at 23°C with 5 µg/mL Alexa Fluor 555 goat anti-rabbit IgG (H+L). Nuclei were stained with 0.2 µg/mL Hoechst 33258 pentahydrate (Molecular Probes, Eugene, OR). After three washes with PBS and one with water, slides were mounted with PermaFluor (Thermo Electron Corp., Pittsburgh, PA). To block phosphatase activity, slides were incubated for 15 min with 20 mmol/L NaF/2 mmol/L sodium orthovanadate/PBS before fixation and fixed with paraformaldehyde containing the same phosphatase inhibitors. All images were captured using Zeiss LSM 510 META confocal system and LSM release 3.2 software (Carl Zeiss, Inc., Thornwood, NY) as reported previously (26).

	Results

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SAHA inhibits the growth of pancreatic cancer cell lines. HDACIs have been shown to induce cell cycle arrest, differentiation, and apoptosis of tumor-derived cell lines (30, 31). To elucidate the effects of the HDACI SAHA in pancreatic cancer cells, three pancreatic cancer cell lines, BxPC-3, COLO-357, and PANC-1, were treated with increasing concentrations of SAHA and cell proliferation was assessed by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay. Treatment with SAHA caused a dose-dependent decrease in the proliferation of BxPC-3 and COLO-357 cells (Fig. 1A ). At its maximally effective concentration (25 µmol/L), SAHA inhibited the growth of BxPC-3 and COLO-357 cells by 42% and 50%, respectively. This decrease in cell proliferation was not due to SAHA-induced apoptosis as SAHA did not induce cleavage of PARP or increase Annexin 5 staining (data not shown). By contrast, the growth of PANC-1 cells was not inhibited by SAHA (Fig. 1A).

View larger version (37K): [in this window] [in a new window]   Fig. 1. Effects of SAHA on cell proliferation and histone H3 acetylation. A, proliferation assay. BxPC-3, COLO-357, and PANC-1 cells were plated at a density of 8,000 per well in 96-well microtiter plates, allowed to adhere overnight, and incubated for 48 h in the absence (white columns) or presence of 5 µmol/L (black columns), 10 µmol/L (gray columns), or 25 µmol/L (stipled columns) SAHA. Growth was assessed as described in Materials and Methods. Columns, mean of three experiments; bars, SE. *, P = 0.013; **, P = 0.001; , P = 0.034; and , P = 0.0001, when compared with respective controls. B, histone H3 acetylation. BxPC-3, COLO-357, and PANC-1 cells were incubated in the absence or presence of 10 µmol/L SAHA for 48 h, lysed, and subjected to immunoblotting using an antibody directed at acetylated lysine-specific histone H3 (H3-AcK). The membrane was then probed for ERK2 to confirm equal loading of lanes.

SAHA induces a G₁ cell cycle arrest. SAHA treatment of cancer cell lines has been shown to induce a G₁ cell cycle block by up-regulating the cyclin-dependent kinase inhibitor p21 (21, 32). Because SAHA decreased the proliferation of BxPC-3 and COLO-357 cells, the effects of SAHA on cell cycle arrest and p21 regulation were examined next by fluorescence-activated cell sorting analysis and p21 immunoblotting. After the treatment with 10 µmol/L SAHA for 48 h, BxPC-3 and COLO-357 cells exhibited a shift into G₁ arrest by 15% to 30%, whereas PANC-1 cells were unaffected (Fig. 2 ). Analysis of p21 protein levels showed that SAHA treatment induced an increase in p21 expression in BxPC-3 and COLO-357, whereas PANC-1 cells exhibited minimal p21 induction (Fig. 3A ). To further determine whether p21 up-regulation was necessary for SAHA-induced growth arrest, knockdown of p21 protein levels using a siRNA strategy was done next. BxPC-3 and COLO-357 cells were transfected with p21 double-stranded siRNA oligonucleotides or nontargeting double-stranded siRNA oligonucleotides. After 48 h of SAHA treatment, the presence of p21 siRNA blocked the SAHA-mediated up-regulation of p21 protein in both cell lines and this effect was especially dramatic in COLO-357 cells (Fig. 3B). Moreover, in COLO-357 cells, suppression of p21 nearly completely reversed SAHA-mediated growth inhibition (Fig. 3C). In contrast, p21 suppression only partially decreased SAHA-induced growth inhibition in BxPC-3 cells. These findings indicate that up-regulation of p21 is necessary for SAHA-induced cell cycle arrest in COLO-357 cells but not in BxPC-3 cells.

View larger version (28K): [in this window] [in a new window]   Fig. 2. Effects of SAHA on the cell cycle. Fluorescence-activated cell sorting analysis. BxPC-3, COLO-357, and PANC-1 cells were plated onto six-well plates and treated with 10 µmol/L SAHA for 48 h. The cells were trypsinized, stained with propidium iodide, and subjected to fluorescence-activated cell sorting analysis as described in Materials and Methods.

View larger version (31K): [in this window] [in a new window]   Fig. 3. Effects of p21 suppression on SAHA-induced growth arrest. A, effects of SAHA on p21 levels. Total cell lysates (25 µg/lane) from untreated (control) or SAHA-treated (48 h) cells were subjected to 13% SDS-PAGE, transferred to nitrocellulose membranes, and probed with an anti-p21 antibody. ERK2 immunoblotting confirmed equal loading of lanes. B, p21 knockdown. BxPC-3 and COLO-357 cells were transfected with nontargeting (NT) or double-stranded p21 siRNA (p21si) oligonucleotides, plated (60-mm plates) at a density of 400,000 per plate, allowed to adhere overnight, and incubated for 48 h in the absence or presence of 10 µmol/L SAHA. Cell lysates (25 µg/lane) were probed with an anti-p21 antibody and an anti-ERK2 antibody as described above. C, proliferation assay. BxPC-3 and COLO-357 cells were transfected with nontargeting or double-stranded p21 siRNA oligonucleotides, plated at a density of 8,000 per well in 96-well plates, allowed to adhere overnight, and incubated for 48 h in the absence or presence of 10 µmol/L SAHA. Growth was assessed as described in Material and Methods. Columns, mean of three experiments; bars, SE. *, P = 0.002; **, P = 0.021; , P = 0.012; and , P = 0.041, when compared with respective controls.

View larger version (23K): [in this window] [in a new window]   Fig. 4. Effects of gemcitabine on proliferation and apoptosis. A, proliferation. BxPC-3, COLO-357, and PANC-1 cells were plated at a density of 8,000 per well in 96-well microtiter plates, allowed to adhere overnight, and incubated for 48 h in the absence (white columns) or presence of 1 µmol/L (black columns), 10 µmol/L (gray columns), or 25 µmol/L (stipled columns) gemcitabine. Growth was assessed as described in Materials and Methods. Columns, mean of three experiments; bars, SE. B, immunoblotting. Cells were incubated in the absence or presence of 10 µmol/L gemcitabine (Gem) for 24 h, lysed, and subjected to immunoblotting with an antibody that recognizes the cleaved form of PARP (89 kDa). ERK2 immunoblotting confirmed equal loading of lanes. C, caspase activation. Cells were incubated for 24 h in the absence (white columns) or presence (black columns) of 1 µmol/L gemcitabine, and cell lysates were assayed for caspase-3 activity as described in Materials and Methods. Columns, mean of three experiments; bars, SE. *, P = 0.02 and **, P = 0.03, when compared with respective controls.

View larger version (27K): [in this window] [in a new window]   Fig. 5. SAHA-mediated potentiation of gemcitabine-induced apoptosis. A, proliferation assay. BxPC-3, COLO-357, and PANC-1 cells were plated at a density of 8,000 per well in 96-well microtiter plates, allowed to adhere overnight, and incubated for 24 h in the absence (white columns) or presence of 10 µmol/L SAHA (black columns), 1 µmol/L gemcitabine (gray columns), the combination of SAHA and 1 µmol/L gemcitabine (hatched columns), 10 µmol/L gemcitabine (stipled columns), and the combination of SAHA and 10 µmol/L gemcitabine (dark stipled columns). Growth was assessed as described in Materials and Methods. Columns, mean of three experiments; bars, SE. B, immunoblotting. Cells were incubated in the absence or presence of 10 µmol/L gemcitabine or 10 µmol/L SAHA for 24 h. Cell lysates were subjected to immunoblotting with an antibody that recognizes the cleaved form of PARP (89 kDa). ERK2 immunoblotting confirmed equal loading of lanes. C, caspase activation. Cells were incubated in the absence (white columns) or presence (black columns) of 10 µmol/L SAHA for 6 h and then for 24 h in the absence (gray columns) or presence (stipled columns) of 10 µmol/L gemcitabine. Cell lysates were assayed for caspase-3 activity as described in Materials and Methods. Columns, mean of three experiments; bars, SE. *, P 0.005, by comparison with respective controls; , P 0.05, by comparison with SAHA alone.

View larger version (24K): [in this window] [in a new window]   Fig. 6. Induction of apoptosis by SAHA and gemcitabine in PANC-1 cells. A, proliferation. PANC-1 cells were plated at a density of 8,000 per well to 96-well microtiter plates, allowed to adhere overnight, and incubated for 48 h in the absence (white column) or presence of 10 µmol/L SAHA (black column), 10 µmol/L gemcitabine (gray column), both agents at these concentrations (stipled column), 25 µmol/L gemcitabine (column with horizontal lines), or SAHA with 25 µmol/L gemcitabine (hatched column). Growth was assessed as described in Materials and Methods. Columns, mean of three experiments; bars, SE. *, P = 0.02 and ** P = 0.004, by comparison with control. B, PARP cleavage. Cells were incubated for 48 h in the absence or presence of 25 µmol/L gemcitabine and 25 µmol/L SAHA. Cell lysates were subjected to immunoblotting with an antibody that recognizes the cleaved form of PARP (89 kDa). ERK2 immunoblotting confirmed equal loading of lanes. C, caspase activation. Cells were incubated for 48 h in the absence (white column) or presence of 10 µmol/L SAHA (black column), 25 µmol/L gemcitabine (column with horizontal lines), or both SAHA and gemcitabine (hatched column). Cell lysates were assayed for caspase-3 activity as described in Materials and Methods. Columns, mean of three experiments; bars, SE. *, P = 0.012, by comparison with control.

View larger version (13K): [in this window] [in a new window]   Fig. 7. Additive effects of SAHA and gemcitabine on proliferation. A, BxPC3 cells were plated at a density of 10,000 per well in 96-well microtiter plates, allowed to adhere overnight, and incubated for 48 h in the absence (white column) or presence of 10 µmol/L SAHA (black column), 500 nmol/L gemcitabine (column with horizontal lines), and the combination of SAHA and 500 nmol/L gemcitabine (stipled column). *, P < 0.004 and **, P < 0.0006, when compared with control; , P < 0.002, when compared with either SAHA or gemcitabine alone. B, COLO-357 cells were plated at a density of 8,000 per well in 96-well microtiter plates, allowed to adhere overnight, and incubated for 48 h in the absence (white column) or presence of 10 µmol/L SAHA (black column), 10 nmol/L gemcitabine (column with horizontal lines), and the combination of SAHA and 10 nmol/L gemcitabine (stipled column). *, P < 0.004 and **, P < 0.0001, when compared with control; , P < 0.015, when compared with either SAHA or gemcitabine alone.

View larger version (11K): [in this window] [in a new window]   Fig. 8. Immunofluorescence analysis of p21. COLO 357 cells were grown on glass slides and incubated for 48 h at 37°C in the absence (A) or presence (B) of 10 µmol/L SAHA. Cells were then subjected to immunofluorescent staining, using an anti-p21 antibody (1). Nuclei were stained with Hoechst 33258 dye (2), and regions were merged to assess signal colocalization (3). Original magnification, x400.

View larger version (16K): [in this window] [in a new window]   Fig. 9. Immunofluorescence analysis of phospho-p21. COLO 357 cells were grown on glass slides and incubated for 48 h at 37°C in the absence (A) or presence (B) of 10 µmol/L SAHA. Cells were then subjected to immunofluorescent staining, using an anti–phospho-p21 antibody (1). Nuclei were stained with Hoechst 33258 dye (2), and regions were merged to assess signal colocalization (3). Original magnification, x400.

	Discussion

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In this study, we tested the effects of SAHA on three distinct pancreatic cancer cell lines BxPC-3, COLO-357, and PANC-1, which harbor mutations in K-ras, p53, and Smad4 genes, alterations that are commonly found in pancreatic cancers. SAHA (10 µmol/L) significantly decreased the proliferation of BxPC3 and COLO-357, which, by fluorescence-activated cell sorting analysis, was associated with a G₁ cell cycle arrest. By contrast, PANC-1 cells were resistant to the growth-inhibitory effects of SAHA at all concentrations tested. PANC-1 cells also exhibited a weak increase in the acetylation of histone H3, indicating that the histone deactylase activity was not decreased by SAHA in this cell line.

It has been shown previously that SAHA induces the up-regulation of the cyclin-dependent kinase inhibitor p21^WAF1 (44) by changing the association of proteins on the p21 promoter (22, 45), indicating that p21 is a direct target of HDACI. In BxPC3 and COLO-357 cells, but not in PANC-1 cells, there was marked up-regulation of p21 by SAHA. When the induction of p21 was prevented by using a p21 siRNA knockdown strategy, COLO-357 cells were no longer sensitive to SAHA-mediated growth inhibition. By contrast, in BxPC3 cells, SAHA-mediated growth inhibition was only partly attenuated. These observations suggest that a major mechanism in SAHA-induced growth arrest in some pancreatic cancer cells is dependent on p21 up-regulation. In support of this conclusion, trichostatin A induces the expression of p21 in pancreatic cancer cell lines (46). Furthermore, in the present study, SAHA did not induce the up-regulation of p21 in PANC-1 cells, which were resistant to its growth-inhibitory effects.

BxPC-3 and COLO-357 exhibited a slight increase in apoptosis when incubated with both SAHA and gemcitabine, by comparison with their response to gemcitabine alone. This finding was not surprising because both cell lines were very sensitive to the effects of gemcitabine alone. However, when the cells were incubated for 48 h with submaximal concentrations of gemcitabine, the addition of SAHA resulted in a significant increase in growth inhibition that was greater than that observed with either agent alone. It is possible, therefore, that the concentration of gemcitabine could be decreased in clinical protocols with SAHA cotreatment, as suggested previously for multiple myeloma (47). Similarly, PANC-1 cells, which were resistant to gemcitabine alone, exhibited a marked increase in sensitivity when treated with both gemcitabine and SAHA for 48 h. Moreover, this effect was associated with a proapoptotic action, as evidenced by PARP cleavage and caspase-3 activation. Thus, a pancreatic cancer cell line that is resistant to the growth-inhibitory and proapoptotic effects of gemcitabine can be rendered responsive to this nucleoside analogue by treatment with SAHA.

A surprising finding in the present study was the SAHA-mediated cytoplasmic localization of p21 and phospho-p21. It is generally accepted that p21 is located in the nucleus, where it interacts with cyclin-dependent kinase 4/6 complexes, thereby inhibiting their activity. However, several studies have reported on the cytoplasmic localization of p21 and indicated that when found in the cytoplasm p21 may promote cell proliferation (48, 49) and may contribute to p21-mediated resistance to apoptosis (50, 51). Other studies have suggested that the cytoplasmic accumulation of p21 abrogates the inhibitory effect of p21 (52). By contrast, in the present study, we determined that SAHA caused the cytoplasmic localization of both p21 and phospho-p21 in COLO-357 cells and that suppression of p21 levels in these cells completely eliminated the growth-inhibitory actions of SAHA. These observations suggest that in this cell line, the SAHA-induced cytoplasmic localization of p21 and phospho-p21 impedes cell cycle progression and facilitates apoptosis. In support of this conclusion, p21 has been reported to enhance cell cycle progression by acting as a nuclear import factor for cyclin D1 and its target kinases, cyclin-dependent kinase 4/6 (53), and to promote apoptosis (54–57).

It was shown recently that HDACIs can enhance the proapoptotic actions of bortezomib, a proteosome inhibitor, in pancreatic cancer cells (58). Given the low toxicity of SAHA, the recent findings with bortezomib, and our current findings, it is possible that SAHA and related compounds may ultimately be useful in various combination treatment strategies for pancreatic cancer.

	Footnotes

 
Grant support: U.S. Public Health Service grant CA-102687 (M. Korc).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 4/17/06; revised 8/27/06; accepted 10/ 9/06.

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