Pancreatic cancer is the fourth leading cause of cancer mortality in the United States.1 It is a devastating disease with a 5-year survival of less than 5%.2 Chemotherapy is often either ineffective or effective for only a short duration. Further therapeutic agents are required in addition to the conventional chemotherapy for pancreatic cancer.
Histones are small basic proteins that, by complexing with DNA, form the nucleosome core.3 Repetitive units of these nucleosomes constitute the chromatin that package the human genome. Histone acetylation is a posttranslational modification of the core nucleosomal histones that affects chromatin structure and gene expression. The acetylation status of histones is regulated by the opposing activities of the corresponding enzymes, histone acetylases (HATs) and histone deacetylases (HDACs).4 Acetylation correlates with remodeling of nucleosomes, resulting in the relaxation of chromatin structure which facilitates the accessibility of a variety of factors to DNA causing transcriptional activation. In contrast, deacetylation of the histone tails induces transcriptional repression through chromatin condensation.5
Aberrant gene expression that results in either functional inactivation of HAT activity or overexpression of HDACs can enhance tumor cell proliferation.6 For example, in acute promyelocytic leukemia, the fusion protein PML-RARα recruits corepressors and HDACs.7 Inappropriate transcriptional repression of tumor suppressor genes mediated by HDACs may be a common molecular mechanism associated with tumorigenesis.8 In addition, HAT and HDACs can directly modify proteins, including transcriptional factors; these changes can either enhance or dampen the activity of the protein.
The HDAC inhibitor, depsipeptide, was demonstrated to have potent antiproliferative activities against a variety of cell lines including those from leukemias, melanomas, prostate and breast cancers that composed the NCI 60 cell lines (http://dtp.nci.nih.gov/docs/cancer/cancer_data.html). Our preliminary data showed that another HDAC inhibitor, SAHA, also had inhibitory activity against a variety of cancer cell lines (data not shown). Inhibitors of HDAC activity induce differentiation, growth arrest, and/or apoptosis of transformed cells in culture and inhibit tumor growth in animals.4 Linear hydroxamic acids such as suberoylanilide hydroxamic acid (Vorinostat, SAHA) are inhibitors of HDAC activities, with SAHA inhibiting HDAC1, 3 and 4.9 SAHA inhibits the in vitro and in vivo growth of transformed human cells, including erythroleukemia,9 lymphoma,10 as well as, cancers of the prostate,11 bladder,9 ovaries12 and endometrium.13 Tumor cells are often much more sensitive to the drug than are normal cells from the same tissue.14 SAHA is in phase I and II clinical trials for the treatment of various malignancies and has shown anticancer effects at doses that are tolerated by the patients.15, 16
To date, no effective drug therapy exists for pancreatic cancer; therefore in this study, we explored the activity of SAHA against pancreatic cancer cells and found that SAHA is a novel, promising therapeutic agent for human pancreatic cancer.
Material and methods
Cells and reagents
Cell lines used in this study were obtained from American Type Culture Collection (Rockville, MD) and were maintained according to their recommendations. Pancreatic cancer cells (PANC-1, AsPC-1, BxPC-3, Capan-2, HPAC, HPAF-II) were grown in Dulbecco's modified Eagle medium with 10% FCS at 37°C in 5% CO2. SAHA was generously provided by Dr. V.M. Richon (Merck Pharmaceuticals, USA). 5-Aza-dC was purchased from Sigma-Aldrich (St. Louis, MO).
3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide assay for cell proliferation and viability
3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT; Sigma) was dissolved in phosphate-buffered saline (PBS) at 5 mg/mL and used to measure either cell proliferation or viability. About 104 cells per well were incubated in culture medium for 96 hr in 96-well plates, followed by the addition of 10 μL of the MTT solution. After a 4-hr incubation, 100 μL of solubilization solution [20% sodium dodecyl sulfate (SDS)] was added, and the mixture was incubated at 37°C for 16 hr. In this assay, MTT is cleaved to an orange formazan dye by metabolically active cells. The absorbance of the formazan product is measured with an enzyme-linked immunosorbent assay reader at 540 nm.
Soft agar colony assay
The 2-layer soft agar system was used as previously described.17 Briefly, cells were removed from culture plates with trypsin and washed. Single-cell suspensions were counted, and 1 × 103 cells were plated per well containing a soft agar mixture (400 μL) into 24-well flat bottom plates and incubated for 14 days at 37°C in a humidified atmosphere containing 5% CO2/95% air; and then, the number of colonies was counted with an inverted light microscope.
Cell cycle analysis
Cultured cells were treated with 5 × 10−6 M SAHA, 1 × 10−6 M 5-Aza-dC, or a vehicle control (10% FCS in RPMI 1640 medium containing 0.01% ethanol) for 3 days. All cells (those in suspension and attached to the culture dish) were collected, washed, suspended in ice-cold PBS, fixed in 75% chilled methanol at 4°C, and stained with propidium iodine. Cell cycle status was analyzed on a Becton Dickinson Flow Cytometer. Analysis was performed immediately after staining using the CELLFit program (Becton Dickinson & Co., Mountain View, CA) whereby the S-phase was calculated using a RFit model.
Western blot analysis
Cells were washed twice in PBS, suspended in lysis buffer [50 mM Tris at pH 8.0, 150 mM NaCl, 0.1% SDS, 0.5% sodium deoxycholate, 1% Nonidet P-40, phenylmethylsulfonyl fluoride (PMSF) at 100 μg/mL, aprotinin at 2 μg/mL, pepstatin at 1 μg/mL and leupetin at 10 μg/mL] and placed on ice for 30 min. After centrifugation at 15,000g for 15 min at 4°C, the suspension was collected. Protein concentrations were quantitated using the Bio-Rad Protein Assay Dye Reagent Concentrate (Bio-Rad Laboratories, Hercules, CA), according to the manufacturer's recommendation. Proteins in whole lysates (40 μg) were resolved by SDS-polyacrylamide gel electrophoresis in a 4–15% gel, transferred to a polyvinylidene difuride membrane (Immobilon, Amersham, Arlington Heights, IL), and probed sequentially with antibodies against the following proteins: p21WAF1, p27KIP1, E-cadherin, RARα, Cyclin D1, Cyclin A, Cyclin B1, c-MYC and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (Santa Cruz Biotechnology, Santa Cruz, CA). The blots were developed with the Supersignal West Pico Chemiluminescent Substrate Kit (Pierce, Rockford, IL).
Preparation of nuclear extracts
For preparation of nuclear extracts, 5 × 106 cells were washed 3 times with ice-cold PBS. After the last wash, adherent cells were scraped off the dish with a rubber policeman and resuspended in 500-μL extraction buffer B [20 mmol/L HEPES, pH 7.9, 20% glycerol, 10 mmol/L NaCl, 0.2 mmol/L EDTA, 1.5 mmol/L MgCl2, 0.1% Triton X, 1 mmol/L dithiothreitol, 1 mmol/L PMSF, 40 μL/mL Complete (Boehringer, Indianapolis, IN)]. After 15 min of incubation on ice, the nuclei were pelleted at 250g for 10 min. Nuclei were resuspended in extraction buffer B, and NaCl was added dropwise with mixing, to a final concentration of 300 mmol/L NaCl. Nuclei were rocked for 60 min at 4°C. Samples were microcentrifuged at 12,000 rpm, and supernatants were frozen at −80°C.
Measurement of apoptosis
Cells in the sub-G1 population, an indication of the number of apoptotic cells, were detected by cell cycle analysis with flow cytometry. The terminal deoxynucleotidyltransferase-mediated uridine 5′-triphosphate end labeling (TUNEL) assay was also used to detect and quantify apoptosis (In Situ Cell Death Detection, POD, Roche, Indianapolis, IN), as described earlier.18 Early apoptosis was also detected by measuring Annexin V protein in the cell membrane using Annexin V-FITC Kit (Clontech, Palo Alto, CA) followed by flow cytometric analysis according to the manufacturer's recommendation.
RNA extraction and reverse transcription (RT) were done with TRIzole (Invitrogen, Carlsbad, CA) and reverse transcriptase (Promega, Madison, WI); 20 μL of cDNA was prepared from 1 μg of RNA. cDNAs were amplified by polymerase chain reaction (PCR) with specific primers for 24-hydroxylase and 18S rRNA, with 25 PCR cycles for 18S rRNA and 32 cycles for p57Kip2 and C/EBPα genes. The PCR primers used to amplify p57Kip2 and C/EBPα genes were 5′-CTGATCTCCGATTTCTTCGC-3′ (forward) and 5′-TCTTTGGGCTCTAAATTGG-3′ (reverse), and 5′-TGGACAAGAACAGCAACGAG-3′ (forward) and 5′-TTGTCA-CTGGTCAGCTCC AG-3′ (reverse), respectively; and those used to amplify 18S rRNA were 5′-AAACGGCTACCACATCCAAG-3′ (forward) and 5′-CCTCCAATGGATCCTCGTTA-3′ (reverse). Annealing temperatures for the PCRs were 56°C for p57Kip2 and C/EBPα genes and 58°C for 18S rRNA. PCR products were separated on a 2% agarose gel, stained with ethidium bromide and photographed.
Real-time quantitative PCR
RNA was prepared using RNeasy kits obtained from Qiagen (Valencia, CA). Gene expression was quantified using real-time quantitative PCR (TaqMan, iCycler, Bio-Rad) technique. We used TaqMan and SybrGreen methods. The sequences of the primers used for the analysis of CK7 and β-actin expression were as follows: CK7 forward 5′-AAGAACCAGCGTGCCAAGTT-3′, cytokeratin 7 (CK7) reverse 5′-CACGCTCATGAGTTCCTGGT-3′, β-actin forward 5′-AAGAGAGGCATCCTCACCCT-3′, β-actin reverse 5′-AAGAGAGGCATCCTCACCCT-3′. Primers were synthesized by Invitrogen. The real time RT-PCR conditions were 3 min at 95°C, followed by 50 cycles: 20 sec 95°C, 20 sec 60°C, 20 sec 72°C and 20 sec 84°C. To determine the relative expression level of each sample, additional reactions with 4 serial 5-fold dilutions of cDNA from PANC-1 cells treated with 5-Aza-dC and SAHA were performed to generate a standard curve, which related the threshold cycle to the log input amount of template.
A melting curve analysis was performed following PCR to identify the correct product by its specific melting temperature, because SybrGreen intercalates nonspecifically into PCR products. Melting curve analysis included 95°C for 5 sec, 65°C for 15 sec, and heating to 95°C at a rate of 0.1°C/2 sec with continuous reading of fluorescence. The signal in this study was generated by the binding of the fluorophore SybrGreen™ (SG, Biozym, Hess. Oldendorf, Germany) to double-stranded DNA. Baseline and threshold calculations were done with the I-Cycler software. The chemical composition of the PCR assays was according to the descriptions of the Taq polymerases. The amplification followed a 3-step PCR with 20-sec denaturation (94°C), 10-sec annealing (60°C), elongation at 65°C for 25 sec and 20 sec at melting temperature.
Chromatin immunoprecipitation assay
Cells were plated at a density of 1 × 106 cells/10-cm dish and incubated overnight at 37°C with 5% CO2. After 12 hr, cells were cultured either in 5 × 10−6 M SAHA or control cultures for 24 hr. Formaldehyde was then added to the cells to a final concentration of 1%, and the cells were incubated at 37°C for 10 min. The medium was removed, and the cells were suspended in 1 mL of ice-cold PBS containing protease inhibitors (Complete). Cells were pelleted, resuspended in 0.5 mL of SDS lysis buffer [1% SDS/10 mM EDTA/50 mM Tris·HCl (pH 8.1)], and incubated on ice for 10 min. Lysates were sonicated with three 10-sec bursts. Debris was removed from samples by centrifugation for 10 min at 15,000g at 4°C. An aliquot of the chromatin preparation (200 μL) was set aside and designated as the input fraction. Supernatants were diluted 5-fold in immunoprecipitation buffer [0.01% SDS/1.1% Triton X-100/1.2 mM EDTA/16.7 mM Tris·HCl (pH 8.1)/16.7 mM NaCl], and 80 μL of a 50% Protein A-Sepharose slurry containing 20 μg of sonicated salmon sperm DNA and 1 mg/mL BSA in TE buffer [10 mM Tris·HCl (pH 8.0)/1 mM EDTA] were added and incubated, rocking for 30 min at 4°C. Beads were pelleted by centrifugation, and supernatants were placed in fresh tubes with 5 μg of antiacetylated histone H3 antibody (Upstate Biotechnology) or normal rabbit serum and incubated overnight at 4°C. Protein A-Sepharose slurry (60 μL) was added, and samples were rocked for 1 hr at 4°C. Protein A complexes were centrifuged and washed 5 times for 5 min each, according to the manufacturer's protocol. Immune complexes were eluted twice with 250 μL of elution buffer (1% SDS/0.1 M NaHCO3) for 15 min at room temperature. A total of 20 μL of 5 M NaCl was added to the combined eluates, and the samples were incubated at 65°C for 4 hr. EDTA, Tris·HCl (pH 6.5) and proteinase K were then added to the samples at a final concentration of 10 mM, 40 mM and 0.04 μg/μL, respectively; and the samples were incubated at 45°C for 1 hr. Immunoprecipitated DNA (both immunoprecipitation samples and input) was recovered by phenol/chloroform extraction and ethanol precipitation, and analyzed by PCR. p21WAF1-specific primers were used to carry out PCR from DNA isolated from ChIP experiments and input samples. The optimal reaction conditions for PCR were determined for each primer pair. Primers were denatured at 95°C for 1 min and annealed at 66°C for 1 min, followed by elongation at 72°C for 1 min. PCR products were analyzed by 2.5% agarose/ethidium bromide gel electrophoresis. The primer pairs used for p21WAF1 ChIP analysis were as follows: 5′-GGT GTC TAG GTG CTC CAG GT-3′ (forward), 5′-GCA CTC TCC AGG AGG ACA CA-3′ (reverse).
Cells were plated at a subconfluent density and cotransfected with 1 μg of the reporter plasmid and the appropriate expression vectors. TOPFLASH construct contains 4 copies of T cell factor (TCF) binding sites placed upstream of thymidine kinase minimal promoter in front of luciferase cDNA (Upstate Biotechnologies). Negative control FOPFLASH contains mutated TCF binding sites. Fifty nanograms of Renilla luciferase pRL-TK were cotransfected as an internal control for transfection efficiency. Lipofectamine 2000 and Opti-MEM were used for transfections, following the manufacturer's instructions. Cell lysates were prepared 48 hr after transfection, and the reporter activity was measured using the Dual-Luciferase Reporter Assay System (Promega).
Transfection and colony formation assay
The entire coding region (nucleotides 132–1,230) of the wild-type C/EBPα was amplified as described previously.19 The fragments were gel-purified and ligated into the pcDNA3 expression vector (TM1, Invitrogen). Cells were plated at 2 × 104 per well using 6-well plates, and transfected with either pcDNA3-C/EBP-α, or vector control (1 μg) using Effectene Transfection Reagent (Qiagen) according to the manufacturer's protocol. The cells were then detached and plated on 100-mm tissue culture dishes at 24–48-hr posttransfection, and simultaneously harvested at 48 hr after transfection to examine their expression of protein using Western blotting.
The differences between 2 groups were analyzed by Student's t test. These data met the assumptions for the Student's t test. All statistical tests were 2-sided.
IC50 is the drug concentration producing the median effect of 50%, and it was calculated by the linear regression method.20 The qualitative information on the nature of drug interaction was calculated by combination index (CI) as described in equation (A).
CA,x and CB,x are the concentrations of drug A and drug B used in combination to achieve x% drug effect. ICx,A and ICx,B are the concentrations for single agents to achieve the same effect. A CI of less than, equal to and more than 1 indicates synergy, additivity and antagonism, respectively.20
SAHA or SAHA with 5-Aza-dC have prominent antitumor activities against human pancreatic cancer cells
The antiproliferative effects of SAHA were examined against various pancreatic cancer cell lines in vitro. Antiproliferative studies used an extremely sensitive soft agar colony assay (Fig. 1a). The concentration of SAHA that caused 50% inhibition (ED50) of clonal growth was as follows: PANC-1, 1.0 × 10−7 μM; AsPC-1, 2.3 × 10−7 μM; BxPC-3, 2.0 × 10−7 μM; HPAC, 1.0 × 10−6 μM; HPAF-II, 3.4 × 10−7 μM; Capan-2, 4.5 × 10−7 μM. In addition, SAHA suppressed the growth of PANC-1 cells in a dose-dependent manner as measured by MTT assay (Fig. 1b). Furthermore, SAHA (2.5 × 10−6 M), in combination with 5-Aza-dC (10−6 M), prominently decreased the growth of PANC-1 (31% of the control) and AsPC-1 cell lines (16% of the control) compared with either agent alone (Fig. 1c). The concentrations for SAHA alone to achieve the same effect (31% of the control) for PANC-1 was 7.2 × 10−6 M, which was calculated from MTT assay (Fig. 1b). Also, the concentration for 5-Aza-dC alone to achieve the same effect calculated by MTT assay (data not shown) was 5.1 × 10−6 M. The CI for PANC-1 was calculated as 0.54 (<1.0). Similarly, the CI for AsPC-1 based on MTT assays (data not shown) was 0.48 (<1.0). These results suggested that the combination of SAHA and 5-Aza-dC had synergistic antiproliferative effects on these pancreatic cancer cells.
Cell cycle analysis of the pancreatic cancer cell lines treated with SAHA
Cell cycle analysis was performed on various pancreatic cancer cells (PANC-1, AsPC-1, BxPC-3, HPAC, HPAF-II, Capan-2) treated with either 5-Aza-dC (10−6 M), SAHA (5 × 10−6 M) or both, as well as vehicle alone (control) for 72 hr (Fig. 2a). The S phase decreased in PANC-1, AsPC-1, HPAC, HPAF-II and Capan-2 treated with SAHA (5 × 10−6 M, 72 hr) compared with control, diluent treated cells. G2/M cell cycle arrest was also induced by SAHA in PANC-1, AsPC-1, HPAC, HPAF-II and Capan-2. Likewise, 5-Aza-dC (10−6 M, 72 hr) produced G2/M arrest in ASPC-1 and HPAC. In ASPC-1, G2/M cell cycle arrest was enhanced when treated with SAHA and 5-Aza-dC, compared with either drug alone (Fig. 2a). The number of cells in the sub-G1 population (late apoptotic cells) increased in all the cell lines (PANC-1, AsPC-1, BxPC3, HPAC, HPAF-II and Capan-2) cultured with SAHA (21, 10, 34, 37, 26 and 27%, respectively) compared with controls (3, 2, 4, 2, 5 and 3%, respectively), suggesting that SAHA induced apoptosis. The sub-G1 population was slightly increased in the cells cultured with 5-Aza-dC (5, 19, 5, 5, 10 and 7%, respectively) compared with controls, but the combination of both drugs had the greatest number of cells in the sub-G1 phase in all the cell lines (28, 22, 43, 55, 40 and 49%, respectively), suggesting that 5-Aza-dC enhanced the apoptosis induced by SAHA (Fig. 2a). These results indicated that SAHA induces G2/M cell cycle arrest and apoptosis in pancreatic cancer; and the apoptosis was enhanced by 5-Aza-dC.
Apoptosis was next examined with TUNEL assay. PANC-1 and ASPC-1 cells were treated with SAHA (5 × 10−6 M), either with or without 5-Aza-dC (10−6 M) for 72 hr, and subjected to TUNEL assay (Fig. 2b). Twelve percent of cells in SAHA-treated cultures, and 3% of those in the 5-Aza-dC-treated cultures were apoptotic in PANC-1 cells, while 2% of vehicle-treated control cells were apoptotic. The percent of apoptotic cells was 17% when both reagents were combined. In ASPC-1 cells, SAHA, 5-Aza-dC or both induced 5–7% apoptosis.
Annexin V measurements were done to detect early apoptosis of the PANC-1 cells exposed to SAHA (5 × 10−6 M), either with or without 5-Aza-dC (10−6 M) for 48 hs. Fourteen percent of SAHA-treated PANC-1 (Annexin V+/PI-) underwent early apoptosis, while control cells exposed to vehicle alone showed 1% early apoptosis (Fig. 2c). 5-Aza-dC induced early apoptosis in only 4% of the cells. The combination of both drugs induced apoptosis in 22% of cells, suggesting that 5-Aza-dC enhanced the apoptosis induced by SAHA in PANC-1 cells (Fig. 2c).
SAHA stimulated differentiation of PANC-1 cells
We initially examined whether addition of SAHA to PANC-1 cells produced an effect on cellular morphology. A photomicrograph composite of PANC-1 cells incubated either in the absence (control) or presence (5 × 10−6 M) of SAHA for 48 hr is shown in Figure 3a. Control cells (left panel) appear as undifferentiated, small round clumps of cells with indistinct borders. When the cells were cultured with SAHA (right panel), they spread out in a more uniform monolayer with well-defined cell borders, with increased cytoplasm and cytoplasmic projections, suggesting a more differentiated state. Thus, SAHA appears to induce morphological differentiation of PANC-1 cells.
Cytokeratin 7 (CK7) has been reported to be associated with pancreatic cancer cell differentiation.21 To examine further if SAHA might induced differentiation of PANC-1 cells, we measured the expression level of CK7 gene in these cells after exposure to SAHA (5 × 10−6 M, 24, 48, 72 hr) using real-time PCR assay. Transcriptional level of CK7 increased in a time-dependent manner, and was significantly upregulated after a 72-hr incubation with SAHA (Fig. 3b).
Effect of SAHA on acetylation of histone
To examine whether SAHA modulates the acetylation of histones in pancreatic cancer cells, we initially first examined the global acetylation of the cells by Western blot. Expression of acetylated H3 was very low in control cells and 5-Aza-dC-treated (10−6 M, 72 hr) cells, but increased in those cultured with SAHA (5 × 10−6 M, 72 hr). Interestingly, the cells incubated with both SAHA and 5-Aza-dC had higher levels of acetylated-H3 than those treated with SAHA alone (Fig. 4a).
Next, we examined the acetylation of the promoter of the p21WAF gene using the chromatin immunoprecipitation (ChIP) assay. Antiacetyl histone H3 antibody was used to immunoprecipitate soluble chromatin from PANC-1 cells cultured with either SAHA or vehicle (control) for 48 hr. The immunoprecipitated DNAs were subjected to PCR using primers for the p21WAF1. Input DNA, before immunoprecipitation, was also subjected to PCR as a control. Treatment with SAHA markedly enhanced the acetylation of histone H3 in the promoter region of the p21WAF1 gene (Fig. 4b).
Changes in expression of cell-cycle and apoptosis-associated genes in PANC-1 cells treated with SAHA either with or without 5-Aza-dC
To explore the mechanism of the antiproliferative effects of SAHA, we further examined the expression of genes associated with cell proliferation. Because SAHA modulated the cell cycle in pancreatic cancer cells (Fig. 2), we examined the expression of cyclin-dependent kinase inhibitors (CDKIs): p21WAF1, p27KIP1 and p57KIP2. Protein expression of p21WAF1 increased to the same levels after exposure to either 5-Aza-dC (10−6 M, 72 hr), SAHA (5 × 10−6 M, 72 hr) or both, but levels of p27KIP1 remained unchanged after exposure to these agents as measured by Western blot (Fig. 5a). Expressions of the apoptosis-related genes, Bax and Bcl-2, in PANC-1 cells treated with SAHA were unchanged (data not shown). Levels of p57KIP2 were examined by RT-PCR. It increased in the presence of SAHA, either alone or with 5-Aza-dC, but not after exposure to 5-Aza-dC alone (Fig. 5b).
Tumor-suppressor genes are often silenced in cancer cells, and SAHA may increase their expression associated with its antiproliferative effects. We found that levels of both E-cadherin and RARα increased after PANC-1 pancreatic cells were cultured with SAHA but not with 5-Aza-dC, and their expression further increased by exposure to both SAHA and 5-Aza-dC (Fig. 5a).
SAHA decreased expression of cyclin D1, B1 and c-myc independently of the β-catenin pathway and increased C/EBPα
To explore further the mechanism of the antiproliferative activities of SAHA, we also studied the expression of progrowth associated genes. Levels of cyclin B1 decreased by 52% in PANC-1 cells cultured with SAHA (5 × 10−6 M, 72 hr), but not after these cells were treated with 5-Aza-dC (10−6 M, 72 hr). Levels of cyclin A were unchanged by SAHA and/or 5-Aza-dC (Fig. 6a). The decreased expression of cyclin B1 may be associated with the G2/M cell cycle arrest induced by SAHA.
Also, SAHA or SAHA in combination with 5-Aza-dC markedly decreased the expression of c-myc and cyclin D1 (Fig. 6b). 5-Aza-dC alone slightly decreased the level of c-myc, but did not change the expression of cyclin D1.
β-catenin, when it migrates to the nucleus, acts as a costimulatory protein for the TCF family of transcription factors. Activation of these transcription factors increases the expression of a number of progrowth genes, including cyclin D1 and c-myc.22, 23 Therefore, we examined whether the nuclear levels of β-catenin decreased after culturing with SAHA (Figs. 6b and 6c). To our surprise, nuclear β-catenin was higher after treatment with SAHA, either with or without 5-Aza-dC, suggesting that SAHA increased the activity of β-catenin rather than decreasing it (Fig. 6b). Furthermore, we examined the ability of SAHA to affect the TCF-dependent transcriptional activity as measured by reporter assays. TOP- or FOP-FLASH was transfected into PANC-1 cells, and the cells were incubated with either diluent, SAHA or 5-Aza-dC. Stimulation of TCF reporter activity was found when the cells were treated with SAHA, either with or without 5-Aza-dC, but not with either vehicle or 5-Aza-dC alone, suggesting that SAHA activated the TCF-dependent transcriptional pathway (Fig. 6c). These result indicated that the decreased expression of c-myc and cyclin D1 by SAHA occurred independently of β-catenin pathway.
Recently, c-myc was identified as a C/EBPα negatively regulated gene in myeloid precursor cells stimulated to enter the differentiation pathway.24 Furthermore, we found that forced expression of C/EBPα in breast cancer cells downregulated their expression of c-myc and induced expression of genes associated with ductal epithelial cells.25 The promoter region of c-myc has an E2F binding site, which is critical for its expression. C/EBPα binds E2F and blocks its transcriptional activity.24 Therefore, we examined the expression of C/EBPα in PANC-1 cultured with SAHA. By RT-PCR, we found that C/EBPα was upregulated by SAHA in PANC-1 cells (Fig. 7a).
In further studies, C/EBPα was transfected into PANC-1 cells, and the cells were examined by Western blot 2 days later. As expected, these cells had abundant levels of C/EBPα compared with vector transfected control cells (Fig. 7b). Expression levels of cyclin B1 and phospholylated Rb in the C/EBPα-transfected PANC-1 cells were decreased compared with the control cells (Fig. 7b). These results together with the results as shown in Figures 2a and 6a suggest that the decreased expression of cylin B1 and phosphorylated Rb, as well as increased number of cells in G2/M in the SAHA treated cells may be mediated by the induction of C/EBPα expression.
In this study, we have found that the HDACI, SAHA at submicromolar concentrations, inhibited growth of pancreatic cancer cells by inducing apoptosis, (G2/M) cell cycle arrest and differentiation. We and other groups have previously reported the in vitro and in vivo ability of SAHA to mediate similar antiproliferative events in other malignancies.9, 10, 11, 12, 13 The morphological differentiation as we have shown in pancreatic cancer cells has also been observed in ovarian cancer cells treated with the HDACI, Tricostatin A.26
We further studied the mechanism of the antitumor activity of SAHA against pancreatic cancer cells. The drug induced p21WAF1 expression in PANC1 cells. This increased expression was independent of p53 expression, because PANC1 cells have a mutant p53 gene.27 A similar p53-independent induction of p21WAF1 by SAHA in cancer cells has been noted, and was thought to be mediated through Sp1- and Sp3-mediated transcriptional modulation.28, 29
We also showed that the elevated level of p21WAF1 was associated with increased acetylation of histones surrounding the p21WAF1 promoter, as demonstrated by ChIP assay. We have noted a similar effect of SAHA in cells from endometrial cancer and lymphoma.10, 12, 13 SAHA may induce p21WAF1 expression not only by acetylation of histones surrounding its promoter, but also by changing the expression of genes associated with transcriptional control of p21WAF1. Also we found that SAHA decreased protein expression of c-myc. c-myc has been shown to be recruited to the p21WAF1 promoter by MIZ-1; and this blocked the transcription of the p21WAF1 gene by various activators.30 Therefore, the decreased expression of c-myc is possibly a factor leading to the increased expression of the p21WAF1 gene in SAHA-treated pancreatic cancer cells.
Most pancreatic cancer cell lines in this study underwent G2/M cell cycle arrest after treatment with SAHA. This may be associated with a deceased level of cyclin B1 as well as an increased p21WAF1 expression. Although p21WAF1 is commonly associated with the G1 check point, its association with G2/M cell cycle arrest has been also demonstrated.31 We also found that SAHA decreased the level of cyclin B1, which may be associated with G2/M cell cycle arrest induced by SAHA.
In addition, we also examined the expression of other members of the kinase inhibitor proteins (KIPs) which potently inhibit several cyclin/CDK complexes, including p27KIP1 and p57KIP232, 33, 34, 35; p57KIP2 was induced by SAHA. This CDKI may also play an important role in the antipancreatic tumor activity of SAHA.
SAHA (10−6 M, 72 hr) did not decrease the percent of BXPC3 cells in S phase, but did potently decrease clonal proliferation and induce apoptosis of these cells. Why these cells had a different cell cycle response to SAHA compared with the other pancreatic cells is unclear.
Another paradox concerns cyclin D1 and c-myc. We have also found that SAHA decreased the expression of both of these proteins in pancreatic cancer cells (PANC-1). In colon cancer cells having a mutation of the APC gene, β-catenin accumulates in their nuclei, acting as a costimulatory protein for the T cell factor (TCF) family of transcription factors, which then stimulate a number of progrowth genes, including cyclin D1 and c-myc.22, 23, 36, 37, 38 We found that SAHA caused the nuclear accumulation of β-catenin in PANC-1 cells associated with enhanced TCF-dependent transcriptional activity as measured by reporter gene assays, suggesting that the β-catenin pathway was activated by SAHA. A recent study showed that the LEF/TCF proteins, in the absence of Wnt signals, functions as transcription repressors rather than activators.39 Furthermore, the repression mediated by LEF1 required HDAC activity.40 At this time, we do not understand how SAHA caused both β-catenin accumulation in the nucleus associated with enhanced TCF-mediated transcriptional activities and decreased levels of c-myc and cyclin D1.
A previous study showed that the transcription factor C/EBPα is integral to normal granulocytic development.41 We found that SAHA induced the expression of C/EBPα in PANC-1 cells; and the forced expression of C/EBPα in these cells increased their levels of cyclin B1 and decreased expression of phospholylated Rb. The decreased cyclin B1 levels associated with G2/M cell cycle arrest may result from SAHA induction of C/EBPα.
SAHA increased the expressions of 2 known tumor suppressor genes, E-cadherin and RARα. E-cadherin is a transmembrane linker protein located in the intercellular adherent junctions, which maintain the adhesive and polarized phenotype of epithelial cells.42, 43 Loss of E-cadherin expression occurs as cells acquire the capacity to invade during the transition from adenoma to carcinoma,44, 45 and its loss is associated with a poor prognosis.46, 47, 48 Retinoids bind to RARs including RARα, which heterodimerizes with retinoid X receptors, and transactivates retinoid-responsive genes.49 Many of these expressed proteins are important in suppressing growth and inducing cellular differentiation.49, 50 Taken together, SAHA enhanced expression of 2 proteins, E-cadherin and RARα, and these proteins are associated with slow growth and induced differentiation.
A number of observations indicate that acetylation and methylation act jointly to regulate gene expression.51, 52, 53, 54, 55 Densely methylated DNA associated with transcriptionally repressive chromatin usually has hypoacetylated histones.56, 57 These 2 epigenetic processes are linked. The methyl-CpG-binding protein MeCP2 resides in a complex with HDAC activity.51, 52 MeCP2 can mediate formation of transcriptionally repressive chromatin on methylated promoter templates in vitro, and this process can be reversed by a HDACI.51, 52, 58 Preclinical studies have shown that inhibition of DNA methylation and histone deacetylation strongly suppresses cancer growth.59, 60, 61 These studies prompted us to examine the combination of an inhibitor of DNA methylation, (5-aza-2′-deoxycytidine), and a HDACI (SAHA) against pancreatic cancer. 5-Aza-dC enhanced the antiproliferative activity of SAHA against the PANC-1 and AsPC-1 pancreatic cancer cells associated with enhanced levels of acetylated histone H3, E-cadherin and RARα in PANC-1 cells.
In conclusion, our studies showed that SAHA inhibited the growth of pancreatic cancer cells by inducing apoptosis, cell cycle arrest and differentiation associated with reactivation of several silenced genes, whose expression is associated with growth suppression. SAHA is a novel, promising therapeutic agent that might be combined with other agents for treatment of human pancreatic cancer.
We particularly thank Dr. Victoria M. Richon of Merck & Co. for generously providing SAHA and for helpful discussions.