1,1-Bis(3′-indolyl)-1-(p-substitutedphenyl)methanes containing p-trifluoromethyl (DIM-C-pPhCF3), p-t-butyl (DIM-C-pPhtBu), and phenyl (DIM-C-pPhC6H5) substituents have been identified as a new class of peroxisome proliferator-activated receptor γ (PPARγ) agonists that exhibit antitumorigenic activity. The PPARγ-active C-DIMs have not previously been studied against bladder cancer. We investigated the effects of the PPARγ-active C-DIMs on bladder cancer cells in vitro and bladder tumors in vivo. In this study, the PPARγ-active compounds inhibited the proliferation of KU7 and 253J-BV bladder cancer cells, and the corresponding IC50 values were 5 to 10 and 1 to 5 μmol/L, respectively. In the less responsive KU7 cells, the PPARγ agonists induced caveolin-1 and p21 expression but no changes in cyclin D1 or p27; in 253J-BV cells, the PPARγ agonists did not affect caveolin-1, cyclin D1, or p27 expression but induced p21 protein. In KU7 cells, induction of caveolin-1 by each of the PPARγ agonists was significantly down-regulated after cotreatment with the PPARγ antagonist GW9662. DIM-C-pPhCF3 (60 mg/kg thrice a week for 4 weeks) inhibited the growth of implanted KU7 orthotopic and s.c. tumors by 32% and 60%, respectively, and produced a corresponding decrease in proliferation index. Treatment of KU7 cells with DIM-C-pPhCF3 also elevated caveolin-1 expression by 25% to 30%, suggesting a role for this protein in mediating the antitumorigenic activity of DIM-C-pPhCF3 in bladder cancer. (Cancer Res 2006; 66(1): 412-8)

Metastatic urothelial carcinoma of the bladder is typically treated with various combinations of systemic chemotherapy (14). However, almost all patients with distant metastatic bladder cancer die within 1 to 2 years despite treatment with the most effective regimens available (2, 3). Since the mid-1980s, the standard treatment for metastatic urothelial carcinoma has been a combination of methotrexate, vinblastine, doxorubicin, and cisplatin. Recently, several novel combination regimens have been reported; however, there is no compelling evidence of enhanced patient survival with these treatments (57). As understanding of the biology of urothelial carcinoma improves, novel therapeutic approaches are being studied, including targeted therapies.

The peroxisome proliferator-activated receptor γ (PPARγ) is a promising target for targeted anticancer therapy. PPARγ is a ligand-activated receptor and a member of the nuclear receptor superfamily of transcription factors. PPARγ is differentially expressed in normal tissues and organs and play diverse roles in metabolism, anti-inflammatory responses, and differentiation (812). PPARγ is highly expressed in tumor samples from different sites, and, in one study, wild-type PPARγ mRNA was observed in multiple tumors and in 34 hematopoietic cancer cell lines as well as several lung (10), colon (10), duodenal (1), prostate (5), breast (4), and glioblastoma (2) cell lines (13). PPARγ has also been detected in bladder tumors and bladder cancer cell lines (1417) and is expressed in the epithelium of the bladder but not surrounding smooth muscle or interstitium (15). PPARγ is an excellent target for cancer chemotherapy not only because of its elevated expression in tumors but also because PPARγ activation results in decreased cell proliferation, decreased G0-G1 to S phase progression, increased differentiation, and apoptosis (1828).

We have identified a new class of PPARγ agonists from a series of 1,1-bis(3′-indolyl)-1-(p-substitutedphenyl)methanes. The PPARγ-active compounds we have identified contain p-trifluromethyl (DIM-C-pPhCF3), p-t-butyl (DIM-C-pPhtBu), and p-phenyl (DIM-C-pPhC6H5) substituents (2732). These PPARγ-active methylene-substituted diindolylmethanes (C-DIM) inhibit growth and/or induce apoptosis in breast, leukemia, pancreatic, and colon cancer cells, but have not previously been studied against bladder cancer. Herein, we investigated their effects on bladder cancer cells in vitro and bladder tumors in vivo. Our findings indicate that the PPARγ-active C-DIMs exhibit antitumorigenic activity against bladder cancer. We are currently evaluating these compounds for future clinical applications.

Cell lines, antibodies, plasmids, and reagents. The bladder cancer cell lines KU7 and 253J-BV were obtained from American Type Culture Collection (Manassas, VA) and generated in our laboratory, respectively. Cells were maintained in DMEM with Ham's F-12 medium (Sigma-Aldrich, St. Louis, MO) supplemented with 0.22% sodium bicarbonate, 0.011% sodium pyruvate, 5% fetal bovine serum (FBS), and 10 mL/L of 100× antibiotic antimycotic solution (Sigma-Aldrich). Cells were maintained at 37°C in the presence of 5% CO2. Antibodies for cyclin D1, p27, phospho-Akt, Akt, Sp1, and caveolin-1 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA) and anti-proliferating cell nuclear antigen (PCNA) antibody from DAKO (Carpinteria, CA). Monoclonal β-actin was purchased from Sigma-Aldrich. Reporter lysis buffer and luciferase reagent for luciferase studies were purchased from Promega (Madison, WI). β-Galactosidase reagent was purchased from Tropix (Bedford, MA) and Lipofectamine reagent was purchased from Invitrogen (Carlsbad, CA). Western Lightning chemiluminescence reagent was purchased from Perkin-Elmer Life Sciences (Boston, MA). Rosiglitazone was purchased from LKT Laboratories, Inc. (St. Paul, MN). The C-DIMs were prepared in this laboratory as previously described (2932). The Gal4 reporter containing 5× Gal4 response elements (pGal4) was kindly provided by Dr. Marty Mayo (University of North Carolina, Chapel Hill, NC). Gal4DBD-PPARγ construct was a gift of Dr. Jennifer L. Oberfield (GlaxoSmithKline Research and Development, Research Triangle Park, NC).

Cell proliferation assay. Bladder cancer cells (2 × 104 per well) were plated in 12-well plates and allowed to attach for 24 hours. The medium was then changed to DMEM:Ham's F-12 medium containing 2.5% charcoal-stripped FBS, and either vehicle (DMSO) or the indicated C-DIMs were added. Fresh medium and C-DIMs were added every 48 hours and cells were then trypsinized and counted after 48, 96, and 144 hours using a Coulter Z1 cell counter. Each experiment was done in triplicate and results are expressed as means ± SE for each set of three experiments.

Transfection and luciferase assay. KU7 or 253J-BV cells (1 × 105 per well) were plated in 12-well plates in DMEM:Ham's F-12 medium supplemented with 2.5% charcoal-stripped FBS. After 16 hours, various amounts of DNA (i.e., 0.4 μg pGal4, 0.04 μg β-galactosidase, and 0.04 μg PPARγ-GAL4) were transfected by Lipofectamine according to the protocol of the manufacturer. After 5 hours, the transfection mix was replaced with complete medium containing either vehicle (DMSO) or the indicated ligand. Twenty to 22 hours later, cells were lysed with 100 μL of 1× reporter lysis buffer and cell extracts (30 μL) were used for luciferase and β-galactosidase assays. A Lumicount luminometer (Perkin-Elmer Life and Analytical Sciences, Boston, MA) was used to quantitate luciferase and β-galactosidase activities, and the luciferase activities were normalized to β-galactosidase activity.

Western blot analysis. Bladder cancer cells were seeded in DMEM:Ham's F-12 medium containing 2.5% charcoal-stripped FBS. Twenty-four hours later, cells were treated with either vehicle (DMSO) or the indicated C-DIMs for different times as indicated. Cells were collected by scraping in 150 μL of high-salt lysis buffer [50 mmol/L HEPES, 0.5 mol/L NaCl, 1.5 mmol/L MgCl2, 1 mmol/L EGTA, 10% (v/v) glycerol, and 1% (v/v) Triton-X-100] and 5 μL/mL of Protease Inhibitor Cocktail (Sigma-Aldrich). The lysates were incubated on ice for 1 hour with intermittent vortexing followed by centrifugation at 40,000 × g for 10 minutes at 4°C. Samples were boiled for 3 minutes at 100°C before electrophoresis, protein concentrations were determined, and 60 μg of protein were applied per lane. Samples were subjected to SDS-PAGE on 10% gel at 120 V for 3 to 4 hours. Proteins were transferred onto polyvinylidene difluoride (PVDF) membranes (Bio-Rad, Hercules, CA) by semidry electroblotting in a buffer containing 25 mmol/L Tris, 192 mmol/L glycine, and 20% methanol for 1.5 hours at 180 mA. The membranes were blocked for 30 minutes with 5% TBST-Blotto [10 mmol/L Tris-HCl, 150 mmol/L NaCl (pH 8.0), 0.05% Triton X-100, and 5% nonfat dry milk] and incubated in fresh 5% TBST-Blotto with 1:1,000 primary antibody overnight with gentle shaking at 4°C. After washing with TBST for 10 minutes, the PVDF membrane was incubated with secondary antibody (1:5,000) in 5% TBST-Blotto for 90 minutes. The membrane was washed with TBST for 10 minutes, incubated with 10 mL of chemiluminescence substrate (Perkin-Elmer Life Sciences) for 1 minute, and exposed to Kodak X-OMAT AR autoradiography film (Eastman Kodak, Rochester, NY).

Animal experiments. Male athymic BALB/c nude mice, 4 to 6 weeks old, were purchased from the Animal Production Area of the National Cancer Institute, Frederick Cancer Research Facility (Frederick, MD). The mice were housed in laminar flow cabinets under specific pathogen-free conditions in facilities approved by the American Association for Accreditation of Laboratory Animal Care and in accordance with current regulations and standards of the U.S. Department of Agriculture, Department of Health and Human Services, and NIH. The use of mice in these experiments was approved by the M.D. Anderson Cancer Center Institutional Animal Care and Use Committee.

We implanted highly tumorigenic human bladder carcinoma KU7 cells (1 × 106 per injection) into the subcutis of athymic nude mice for the heterotopic xenograft experiment and into the bladder wall for the orthotopic xenograft experiment. One week later, mice were treated (10 per treatment group) with placebo or DIM-C-pPhCF3 (60 mg/kg/dose on days 1, 3, and 5). Treatment was continued for 4 weeks, after which the animals were sacrificed. Tumor volumes were measured and tumor kinetics were established in the various groups. Apoptosis and cell proliferation were subsequently evaluated in tumor sections using immunohistochemical techniques (see Results).

Quantification of PCNA and caveolin-1 in tissue samples. For immunohistochemical analysis, frozen tissue sections (8 μm thick) were fixed with cold acetone, chloroform/acetone, and acetone. Tissue sections (5 μm thick) of formalin-fixed, paraffin-embedded specimens were deparaffinized in xylene and then treated with a graded series of alcohol [100%, 95%, and 80% ethanol/double-distilled H2O (v/v)] and rehydrated in PBS (pH 7.5). Antigen retrieval for paraffin-embedded tissues was done with pepsin (Biomeda, Foster City, CA) for 15 minutes at 37°C. Endogenous peroxidase was blocked by the use of 3% hydrogen peroxide in PBS for 10 minutes. The samples were washed thrice with PBS and incubated for 20 minutes at room temperature with a protein blocking solution containing 5% normal horse serum and 1% normal goat serum in PBS (pH 7.5). Excess blocking solution was drained and the samples were incubated for 18 hours at 4°C with either a 1:100 dilution of mouse monoclonal anti-PCNA antibody or a 1:100 dilution of caveolin-1 antibody. The samples were then rinsed four times with PBS and incubated for 60 minutes at room temperature with the appropriate dilution of the secondary antimouse IgG (Jackson ImmunoResearch Laboratory, West Grove, PA). The slides were rinsed with PBS and incubated for 5 minutes with diaminobenzidine. The sections were then washed thrice with PBS, counterstained with Gill's hematoxylin (Biogenex Laboratories, San Ramon, CA), and again washed thrice with PBS. The slides were mounted using a water- and alcohol-based mounting medium (Universal Mount, Research Genetics). Cell proliferation was determined by immunohistochemical analysis of tissue sections with anti-PCNA antibodies. The tissue was photographed using a cooled charge coupled device Optotronics Tec 470 camera linked to a computer and digital printer. The intensity of the immunostaining was quantified in multiple points in five different areas of each sample by an image analyzer using Optimas image analysis software (Media Cybernetics, San Diego, CA) to obtain an average measurement. The density of proliferative cells and apoptotic cells was expressed as an average of the five highest densities identified within a single ×200 field.

Quantification of apoptosis in tissue samples. Frozen tissue sections fixed and treated as described in the preceding section were washed with PBS containing 0.1% Brij (v/v). Terminal deoxynucleotidyl transferase (TdT)–mediated nick end labeling (TUNEL) was done using a commercial kit (Promega) according to the instructions of the manufacturer with the following modifications. Samples were fixed with 4% paraformaldehyde (methanol free) for 10 minutes at room temperature, washed with PBS, and permeabilized by incubation with 0.2% Triton X-100 in PBS (v/v) for 15 minutes. The samples were incubated with equilibration buffer (from the kit). Reaction buffer containing equilibration buffer (45 μL), nucleotide mix (5 μL), and TdT (1 μL) was added to the sections, and sections were incubated in a humidified chamber for 1 hour at 37°C protected from light. The reaction was terminated by immersing the samples in 2× SSC [30 mmol/L NaCl and 3 mmol/L sodium citrate (pH 7.2)] for 15 minutes and then washing them thrice to remove unincorporated fluorescein-dUTP. Background reactivity was determined by processing the slides in the absence of TdT (negative control). Nuclei were stained with propidium iodide (1 μg/mL) for 10 minutes. Fluorescent bleaching was minimized with an enhancing reagent (Prolong, Molecular Probes, Eugene, OR). Immunofluorescence microscopy was done using a Zeiss Plan-Neofluar lens on an epifluorescence microscope equipped with narrow bandpass excitation filters mounted in a filter wheel (Ludl Electronic Products, Hawthorne, NY) to individually select for green, red, and blue fluorescence. Images were captured using a cooled charge coupled device camera (Photometrics, Tucson, AZ). DNA fragmentation was detected by localized green fluorescence within the nucleus of apoptotic cells.

Statistical analyses. Statistical analysis was done using SPSS software (SPSS, Inc., Chicago, IL) and Instat3 software (San Diego, CA). For in vitro data, ANOVA was done, and t tests with the Bonferroni correction were used to evaluate for significant differences between the treated cells at each drug concentration and untreated control cells. Tumor weights and expression intensities of TUNEL and PCNA counts were compared by unpaired Student's t test. For the primary end point of tumor size, the sample size of 10 mice per treatment group was expected to have >90% power to detect a minimum difference of 24 mm3 in tumor size at a statistical significance level of 0.05%. Statistical significance for this study was set at two-sided P < 0.05.

Effects of rosiglitazone and C-DIMs on cell proliferation and PPARγ activation. We investigated the effects of rosiglitazone, a widely used thiazolidinedione, and PPARγ-active C-DIMs on the proliferation of KU7 and 253J-BV bladder cancer cells.

Figure 1 shows the results in KU7 cells treated for 48 hours. Rosiglitazone did not decrease cell proliferation even at the highest concentration, 10 μmol/L (Fig. 1A). In contrast, DIM-C-pPhtBu and DIM-C-pPhC6H5 significantly inhibited proliferation starting at 1 μmol/L concentration and all PPARγ-active C-DIMs significantly decreased cell proliferation at 5 and 10 μmol/L concentrations (Fig. 1B-D). Moreover, following treatment with 10 μmol/L concentrations of the C-DIMs, the number of cells remaining was lower than the initial number of seeded cells, indicating that cell death was induced. We also examined the effects of treatment for 96 and 144 hours. At these longer treatment durations, results were similar to those observed after 48 hours with the exception that rosiglitazone at a concentration of 10 μmol/L also decreased (10-25%) cell proliferation (data not shown).

Figure 1.

KU7 cell proliferation inhibition assay. KU7 cells were treated with DMSO or 1, 5, or 10 μmol/L of rosiglitazone (A) or a C-DIM (B-D) for 48 hours, and the percentage inhibition of proliferation was determined as described in Materials and Methods. The number of cells in the DMSO treatment group was set at 100%. Columns, means for three replicate experiments for each concentration; bars, SE. *, P < 0.05, significantly decreased cell proliferation. Ten-micromolar concentrations of the C-DIMs significantly (P < 0.05) induced cell death (i.e., the numbers of cells in the treatment groups were lower than the original numbers of seeded cells).

Figure 1.

KU7 cell proliferation inhibition assay. KU7 cells were treated with DMSO or 1, 5, or 10 μmol/L of rosiglitazone (A) or a C-DIM (B-D) for 48 hours, and the percentage inhibition of proliferation was determined as described in Materials and Methods. The number of cells in the DMSO treatment group was set at 100%. Columns, means for three replicate experiments for each concentration; bars, SE. *, P < 0.05, significantly decreased cell proliferation. Ten-micromolar concentrations of the C-DIMs significantly (P < 0.05) induced cell death (i.e., the numbers of cells in the treatment groups were lower than the original numbers of seeded cells).

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Figure 2 shows the results in 253J-BV cells treated for 48 hours. Rosiglitazone exhibited minimal effects on cell survival even at the highest concentration, 10 μmol/L (Fig. 2A). In contrast, the 5 and 10 μmol/L concentrations of the C-DIMs significantly decreased cell proliferation (Fig. 2B-D). Moreover, treatment with 5 or 10 μmol/L concentrations of the C-DIMs for 96 or 144 hours induced cell death (data not shown).

Figure 2.

253J-BV cell proliferation inhibition assay. 253J-BV cells were treated with DMSO or 1, 5, or 10 μmol/L of rosiglitazone (A) or a C-DIM (B-D) for 48 hours, and percentage proliferation was determined as described in Materials and Methods. The number of cells in the DMSO treatment group was set at 100%. Columns, means for three replicate experiments for each concentration; bars, SE. *, P < 0.05, significantly decreased cell proliferation. Ten-micromolar concentrations of the C-DIMs significantly (P < 0.05) induced cell death. After treatment for 144 hours, 5 μmol/L concentrations also induced cell death.

Figure 2.

253J-BV cell proliferation inhibition assay. 253J-BV cells were treated with DMSO or 1, 5, or 10 μmol/L of rosiglitazone (A) or a C-DIM (B-D) for 48 hours, and percentage proliferation was determined as described in Materials and Methods. The number of cells in the DMSO treatment group was set at 100%. Columns, means for three replicate experiments for each concentration; bars, SE. *, P < 0.05, significantly decreased cell proliferation. Ten-micromolar concentrations of the C-DIMs significantly (P < 0.05) induced cell death. After treatment for 144 hours, 5 μmol/L concentrations also induced cell death.

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On the basis of the cell growth curves, the IC50 values for proliferation inhibition by the PPARγ-active C-DIMs ranged from 5 to 10 μmol/L for KU7 cells and from 1 to 5 μmol/L for 253J-BV cells; the IC50 for rosiglitazone was >10 μmol/L in both cell lines.

We also examined PPARγ activation in KU7 and 253J-BV cells treated with rosiglitazone or C-DIMs and transfected with the GAL4-PPARγ/GAL4-luc constructs. In KU7 cells (Fig. 3A), rosiglitazone and the C-DIMs caused a concentration-dependent increase in transactivation. In 253J-BV cells (Fig. 3B), rosiglitazone and the C-DIMs also induced transactivation and this effect was greater for rosiglitazone than for the C-DIMs. However, in contrast to the findings in KU7 cells, the increase in transactivation caused by the C-DIMs was not concentration dependent; this may be due, in part, to their potent growth inhibitory effects at the 5 and 10 μmol/L concentrations (Fig. 2B-D).

Figure 3.

Activation of GAL4/pGAL4 by C-DIM compounds and rosiglitazone. KU7 (A) and 253J-BV (B) bladder cancer cells were transfected with PPARγ-GAL4/pGAL4 and treated with DMSO or different concentrations of the C-DIMs, and luciferase was determined as described in Materials and Methods. Columns, means for at least three replicate experiments for each treatment group; bars, SE. *, P < 0.05, significant induction.

Figure 3.

Activation of GAL4/pGAL4 by C-DIM compounds and rosiglitazone. KU7 (A) and 253J-BV (B) bladder cancer cells were transfected with PPARγ-GAL4/pGAL4 and treated with DMSO or different concentrations of the C-DIMs, and luciferase was determined as described in Materials and Methods. Columns, means for at least three replicate experiments for each treatment group; bars, SE. *, P < 0.05, significant induction.

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Effects of rosiglitazone and C-DIMs on cell cycle proteins and caveolin-1. In some cancer cell lines, PPARγ agonists modulate genes and proteins involved in the G0-G1-S cell cycle progression. We investigated the effects of rosiglitazone and the C-DIMs on cell cycle proteins in KU7 and 253J-BV cells. In KU7 cells (Fig. 4A), cyclin D1 and p27 levels were unaffected by treatment, and both rosiglitazone and the C-DIMs increased p21 expression in a concentration-independent manner. The reasons for the differences in p21 protein levels in KU7 cells after treatment with 5.0 and 7.5 μmol/L DIM-C-pPhtBu and DIM-C-pPhC6H5 are unknown; however, because this response was not observed for DIM-C-pPhCF3, it is likely that these effects are compound-specific and PPARγ independent. In 253J-BV cells (Fig. 4B), cyclin D1 and p27 levels were unaffected by treatment and the C-DIMs but not rosiglitazone induced p21 expression in a concentration-independent manner. The lack of induction of p21 by rosiglitazone may be due to the concentrations used (5.0 and 7.5 μmol/L) because at these concentrations it did not inhibit cell growth (Figs. 1 and 2) either. The up-regulation of p21 expression by C-DIMs has previously been reported in Panc-28 cells (31) and is consistent with their growth-inhibitory effects (Fig. 2).

Figure 4.

Effects of C-DIMs on cell cycle protein expression. KU7 (A) and 253J-BV (B) cells were treated with DMSO or 5.0 or 7.5 μmol/L of the PPARγ-active C-DIMs and rosiglitazone for 24 hours. Whole cell lysates were analyzed in a Western blot assay as described in Materials and Methods. Sp1 protein served as a loading control.

Figure 4.

Effects of C-DIMs on cell cycle protein expression. KU7 (A) and 253J-BV (B) cells were treated with DMSO or 5.0 or 7.5 μmol/L of the PPARγ-active C-DIMs and rosiglitazone for 24 hours. Whole cell lysates were analyzed in a Western blot assay as described in Materials and Methods. Sp1 protein served as a loading control.

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Previous studies in several colon cancer cell lines related the growth inhibitory effects of C-DIMs to the induction of caveolin-1 (30), which has been shown to inhibit the proliferation of some colon cancer cells. We investigated the effects of rosiglitazone and the C-DIMs on caveolin-1 expression in KU7 and 253J-BV cells. In KU7 cells (Fig. 5A), the C-DIMs, but not rosiglitazone, induced an ∼3- to 4-fold increase in caveolin-1 protein expression similar to the response previously reported in colon cancer cells (30). In 253J-BV cells (Fig. 5B), constitutive caveolin-1 protein expression was relatively high and it was difficult to show further significant enhancement in levels of this protein. To further investigate the role of PPARγ agonists in mediating induction of caveolin-1, KU7 cells were treated with 5 or 7.5 μmol/L DIM-C-pPhCF3 or DIM-C-pPhC6H5 alone or in combination with the PPARγ antagonist GW9662 (Fig. 5C). Cotreatment with GW9662 inhibited induction of caveolin-1 by PPARγ-active C-DIMs.

Figure 5.

Induction of caveolin-1 by C-DIMs. KU7 (A) and 253J-BV (B) cells were treated with DMSO or 5.0 or 7.5 μmol/L PPARγ-active C-DIMs or rosiglitazone for 72 hours. Whole cell lysates were analyzed for caveolin-1 in a Western blot assay as described in Materials and Methods. Sp1 protein served as loading control. C, effects of the PPARγ agonist GW9662. KU7 cells were treated with DIM-C-pPhCF3 or DIM-C-pPhC6H5 alone or in combination with 10 μmol/L GW9662 and analyzed in a Western blot assay for caveolin-1, Akt, phospho-Akt (pAkt), and β-actin (loading control) as described in Materials and Methods.

Figure 5.

Induction of caveolin-1 by C-DIMs. KU7 (A) and 253J-BV (B) cells were treated with DMSO or 5.0 or 7.5 μmol/L PPARγ-active C-DIMs or rosiglitazone for 72 hours. Whole cell lysates were analyzed for caveolin-1 in a Western blot assay as described in Materials and Methods. Sp1 protein served as loading control. C, effects of the PPARγ agonist GW9662. KU7 cells were treated with DIM-C-pPhCF3 or DIM-C-pPhC6H5 alone or in combination with 10 μmol/L GW9662 and analyzed in a Western blot assay for caveolin-1, Akt, phospho-Akt (pAkt), and β-actin (loading control) as described in Materials and Methods.

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There is evidence that PPARγ-active compounds, including C-DIMs, enhance phosphatidylinositol-3-kinase (PI3K) activity in colon cancer cell lines (33),5

5

S. Safe, unpublished observations.

so we investigated the effect of the C-DIMs on PI3K in KU7 cells. As shown in Fig. 5C, both DIM-C-pPhCF3 and DIM-C-pPhC6H5 induced PI3K-dependent phosphorylation of Akt. The compounds did not affect Akt protein.

Thus, PPARγ-active C-DIMs decreased proliferation of KU7 and 253J-BV cells, and this correlated with increased expression of caveolin-1 and p21 in KU7 cells and p21 in 253J-BV cells. These results are another example of the cancer cell context–dependent differences in the growth inhibitory pathways induced by C-DIMs and other PPARγ agonists.

Effects of PPARγ-active C-DIM on bladder tumor growth in vivo. To confirm whether our in vitro findings would extend to an in vivo model of bladder cancer, we implanted KU7 cells orthotopically into the bladders of athymic nude mice. To establish tumor kinetics, another in vivo experiment was done using s.c. implanted tumors. Mice were then treated with DIM-C-pPhCF3 as described in Materials and Methods. No toxicity was seen in mice treated with DIM-C-pPhCF3, as shown by weighing the mice.

Compared with control, treatment with DIM-C-pPhCF3 substantially inhibited tumor growth in both orthotopic tumors (32% growth inhibition) and s.c. tumors (60% growth inhibition; Fig. 6A). The mean tumor weights of mice in the control and DIM-C-pPhCF3–treated groups were 265 and 191 mg, respectively, for the orthotopic tumors and 997 and 398 mg, respectively, for the s.c. tumors. DIM-C-pPhCF3 significantly inhibited cell proliferation in orthotopic and s.c. bladder tumors by 26% and 55%, respectively (PCNA staining; Fig. 6C). However, tumors obtained from mice in the treatment group displayed low levels of apoptosis that were not appreciably different from those in controls (data not shown). Immunohistochemical analysis showed that DIM-pPhCF3 significantly increased expression of caveolin in both orthotopic and s.c. bladder tumors, by 30% and 25%, respectively (Fig. 6D). These in vivo results complemented the results of in vitro studies (Fig. 5), which showed up-regulation of caveolin-1 protein expression.

Figure 6.

Inhibition of bladder tumor growth by DIM-C-pPhCF3. KU7 cells (1 × 106 per injection) were administered to athymic nude mice via s.c. injection or by direct injection into the bladder wall as described in Materials and Methods. Mice were administered DIM-C-pPhCF3 (60 mg/kg) thrice a week (days 1, 3, and 5) for 4 weeks. A, tumor weights at the end of the treatment; B, tumor volumes in the s.c. group. C, proliferation index and apoptosis (not shown) were determined as described in Materials and Methods. D, caveolin-1 index and immunostaining of bladder tumors for caveolin-1 expression were determined as described in Materials and Methods.

Figure 6.

Inhibition of bladder tumor growth by DIM-C-pPhCF3. KU7 cells (1 × 106 per injection) were administered to athymic nude mice via s.c. injection or by direct injection into the bladder wall as described in Materials and Methods. Mice were administered DIM-C-pPhCF3 (60 mg/kg) thrice a week (days 1, 3, and 5) for 4 weeks. A, tumor weights at the end of the treatment; B, tumor volumes in the s.c. group. C, proliferation index and apoptosis (not shown) were determined as described in Materials and Methods. D, caveolin-1 index and immunostaining of bladder tumors for caveolin-1 expression were determined as described in Materials and Methods.

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Our findings indicate that a new class of PPARγ agonists, PPARγ-active C-DIMs, have antitumorigenic activity against bladder cancer cells in vitro and against s.c. and orthotopic bladder tumors in mice. The PPARγ-active C-DIMs were more potent inhibitors of bladder cancer cell growth than was rosiglitazone, a widely used synthetic thiazolidinedione. In addition, we observed cell line–specific differences in the PPARγ-dependent growth inhibitory genes activated by the C-DIMs. Finally, our studies provide evidence that induction of caveolin-1 and p21 expression may mediate the antitumorigenic activity of PPARγ-active C-DIMs and that activation of PI3K might be important as well.

The potential anticarcinogenic activities of different structural classes of PPARγ agonists have been investigated in various cancer cell lines in vitro and in vivo (1832). PPARγ agonists induce genes and activities linked to differentiation, growth inhibition, and apoptosis; however, the induced responses are highly variable and dependent on ligand structure and cell context. Moreover, there is evidence that the effects of these compounds may be either PPARγ dependent or PPARγ independent. For example, the PPARγ agonists troglitazone and 15-deoxyΔ12,14-prostaglandin J2 induce the NSAID-activated gene (NAG-1) in HCT-116 colon cancer cells by receptor-independent and receptor-dependent pathways, respectively (34). PPARγ-active C-DIMs induce p21 expression in Panc-28 but not other pancreatic cell lines and this response is PPARγ dependent. Rosiglitazone induces caveolin-1 in HT-29 colon cancer cells expressing wild-type PPARγ but not in HCT-15 cells, which express a mutant (K422Q) form of this receptor (30). In contrast, PPARγ-active C-DIMs induce caveolin and other genes associated with differentiation in colon cancer cells expressing either wild-type or mutant PPARγ (30). A recent study showed that 2-cyano-3,12-diooxoolean-1,9-dien-oic acid (CDDO) and related PPARγ agonists also induce caveolin-1 in colon cancer cells, and this effect was inhibited by PPARγ antagonists, whereas higher concentrations of CDDO induce apoptosis in a receptor-independent fashion (33).

Bladder cancer cells also overexpress PPARγ (1417). One study showed that although prostaglandin J2 and thiazolidinediones both decreased cell survival, there were significant differences in the responsiveness of bladder cancer cells to these compounds (17). For example, the IC50 value for growth inhibition by troglitazone and pioglitazone was >50 μmol/L in 253J-BV bladder cancer cells; IC50 values in T24, RT4, and IT-1 cells were ≤30, 15, and 10 μmol/L, respectively (17). In our study, both PPARγ-active C-DIMs and rosiglitazone-activated PPARγ (Fig. 3) and the C-DIMs decreased cell survival in KU7 and 253J-BV cells (Figs. 1 and 2). KU7 cells were less responsive than 253J-BV cells to the growth inhibitory effects of C-DIMs at concentrations ≤10 μmol/L, whereas rosiglitazone at concentrations ≤10 μmol/L was relatively ineffective in both cell lines. Thus, PPARγ-active C-DIMs were more potent than rosiglitazone as inhibitors of bladder cancer cell growth.

The modulation of cell cycle genes (p21) associated with G0-G1 to S-phase progression and the induction of caveolin-1 have been linked to PPARγ-dependent inhibition of Panc-28 pancreatic and HT-29 and HCT-15 colon cancer cell growth after treatment with C-DIMs (30, 31). In our study, we observed differences in activation of specific PPARγ-dependent “growth inhibitory” genes by C-DIMs-C-DIMs induced p21 in 253J-BV cells and caveolin-1 and p21 in KU7 cells (Figs. 4 and 5). The reason for cell context–dependent regulation of p21 and caveolin-1 by C-DIMs is unknown; however, results of ongoing studies with colon cancer cells in our laboratory indicate that the degree of induction of caveolin-1 is inversely related to constitutive levels of this protein (data not shown). Thus, high basal expression of caveolin-1 in 253J-BV cells (Fig. 5) may preclude further induction of this protein.

We further investigated the in vivo anticarcinogenic activity of DIM-C-pPhC6H5 and DIM-C-pPhCF3 by implanting the highly tumorigenic KU7 cell line into the bladders and subcutis of athymic nude mice. Both PPARγ agonists inhibited tumor growth in the orthotopic and s.c. implant models in athymic nude mice. DIM-C-pPhCF3 significantly inhibited tumor growth in both models. In vivo studies with DIM-C-pPhC6H5 were not completed because of formulation problems (data not shown). We also showed that for DIM-C-pPhCF3, there was a significantly increased expression of caveolin-1 in the orthotopic (30%) and s.c. (25%) bladder cancer models, and these in vivo responses (Fig. 6D) correlated with induced expression of caveolin-1 in KU7 cancer cells (Fig. 5A). Thus, PPARγ-active C-DIMs inhibit growth of bladder cancer cells in vitro and bladder tumors in both an orthotopic and s.c. in vivo mouse models. This is the first example of the antitumorigenic activity of PPARγ agonists in an in vivo model of bladder cancer.

We also observed the somewhat paradoxical induction of PI3K activity by PPARγ-active C-DIMs in KU7 cells (Fig. 5). A previous study also showed that the triterpenoid PPARγ agonist CDDO also induced this kinase pathway in SW-480 colon cancer cells (33). Although PI3K has been linked to cell survival pathways, increased PI3K-dependent activity and caveolin-1 expression sensitizes HeLa and 293 cells to the cytotoxicity of arsenite and hydrogen peroxide (35) and sensitizes L929 cells to tumor necrosis factor α–induced cell death (36). Thus, activation of PI3K and induction of caveolin-1 expression may be important mediators of the antitumorigenic activity of PPARγ-active C-DIMs. We are currently investigating other receptor-dependent and receptor-independent pathways responsible for the anticancer activities of these compounds in the treatment of bladder cancer.

Grant support: M.D. Anderson Bladder Specialized Programs of Research Excellence Developmental Research grant 5P50CA091846-03 and NIH grants ES09106 and CA112337.

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.

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