3,3′-Diindolylmethane Enhances the Efficacy of Butyrate in Colon Cancer Prevention through Down-Regulation of Survivin Free

Colon cancer is one of the leading causes of cancer death in developed countries. A large number of preclinical studies have examined various agents for the prevention of colon cancer. However, significant tumor suppression activity was rarely observed and tumor occurrence even increased after treatment in certain cases (1, 2). Butyrate is produced naturally in the colon by bacterial fermentation of dietary fibers and has been extensively studied in colon cancer prevention. Butyrate inhibits tumor growth and induces apoptosis through the inhibition of histone deacetylases (HDAC; refs. 3, 4). HDACs together with histone acetyltransferases regulate the acetylation of core nucleosomal histones, which is important for the transcription activity of the target genes (5). Abnormal HDAC activity has been associated with the development of many types of cancer (6). HDAC inhibitors induce differentiation, growth arrest, and apoptosis in cancer cells, whereas they are relatively nontoxic to normal cells (6–8). As a HDAC inhibitor, butyrate is a promising prevention agent for colon cancer (9, 10). However, the activity of butyrate has been moderate in chemoprevention studies using animal models (11–13). For a potential explanation of the ineffectiveness of butyrate, we recently showed that mutations in the adenomatous polyposis coli (APC) gene in colon cancers can confer resistance to HDAC inhibitor–induced apoptosis (14).

Colon cancer is caused by the accumulation of mutations in a number of oncogenes and tumor suppressor genes. APC, deleted in colon cancer (DCC), K-Ras, and p53 are among the most frequently mutated genes in colon cancer (15–18). APC is a key component of the β-catenin destruction complex (consisting GSK-3β, axin, and APC) and involved in the Wnt signaling pathway (19). Axin and APC form a structural scaffold that allows GSK-3β to phosphorylate β-catenin. Phosphorylated β-catenin is subsequently degraded by the proteasome (19). Loss of wild-type APC expression results in the nuclear accumulation of β-catenin, which interacts with Tcf-4/Lef1 transcription factors to cause aberrant gene transcription and formation of cancer (20, 21). The role of APC as a tumor suppressor has been further shown in mice with APC mutations that develop multiple intestinal neoplasia (APC^min/+ mice; refs. 22, 23).

Previous studies in our laboratory have shown that resistance to HDAC inhibitors by colon cancers expressing mutant APC was due to a failure to down-regulate survivin (14). Survivin is an antiapoptotic protein of the inhibitor of apoptosis family. Survivin is expressed at a high level in many types of cancers, but not in normal tissues from the same organs (24). Survivin blocks apoptosis by inhibiting caspases and antagonizing mitochondrial-dependent apoptosis (24). Survivin also regulates cell division through interaction with INCENP and Aurora B kinase (25).

3,3′-Diindolylmethane (DIM) is a natural compound formed during the autolytic breakdown of glucobrassicin present in food plants of the Brassica genus, including broccoli, cabbage, Brussels sprouts, cauliflower, and kale (26). The anticancer activity of DIM was linked with inhibition of mitochondrial H(+)-ATP synthase and induction of p21(Cip1/Waf1) expression in breast cancer cells (27); down-regulation of androgen receptor (28); and inhibition of mammalian target of rapamycin (29) in prostate cancer cells, inhibition of AKT signaling and FLICE-like inhibitory protein in cholangiocarcinoma cells (30), activation of caspase-8 in colon cancer cells (31), as well as inactivation of NF-κB (32), and down-regulation of survivin in breast cancer cells (33). DIM was also shown to inhibit angiogenesis and invasion by repressing the expression of matrix metalloproteinase MMP-9 and urokinase-type plasminogen activator (34).

In the present study, we found that colon cancer cells expressing mutant APC are resistant to butyrate-induced apoptosis due to the failure to down-regulate survivin. Importantly, we showed that DIM was able to down-regulate survivin and potentiate butyrate-induced apoptosis in these resistant cells. DIM can be used to enhance the efficacy of butyrate in colon cancer prevention.

Human colon cancer cell line HT-29 was purchased from the American Type Culture Collection. The cells were cultured in DMEM containing 10% fetal bovine serum. Human colon cancer HT-29/APC and HT-29/β-Gal cells were generously provided by Dr. Vogelstein and cultured as described (35). IMCE cells were kindly provided by Dr. Robert H. Whitehead and cultured at 33°C, in RPMI 1640 containing 5% FCS, 1 μg/mL insulin, 10⁻⁵ mol/L α-thioglycerol, and 5 units/mL of mouse γ-IFN. For transient transfection, plasmids were transfected into cells using Lipofectamine Plus Reagent (Invitrogen) following the manufacturer's protocol.

Butyrate and cycloheximide were purchased from Sigma. DIM was purchased from LKT Laboratories. Proteasome inhibitor MG-132 was purchased from Calbiochem. Disuccinimidyl suberate was purchased from Pierce.

Human cDNA encoding the full-length survivin gene was obtained by PCR amplification using an EST clone (I.M.A.G.E. clone ID 4477581) as template. Survivin cDNA was subcloned into phrGFP-C and pCMV-Tag2A expression vectors (Stratagene).

In Terminal Deoxynucleotidyl Transferase Biotin-dUTP Nick End Labeling (TUNEL) assay, cells were fixed with 1% paraformaldehyde on ice for 1 h and then washed with PBS and permeabilized with 70% ethanol at −20°C for 12 h. Cells were then labeled with Guava TUNEL Assay reagents and analyzed on the Guava PCA microcytometer following the manufacturer's protocol. In 4′,6-diamidine-29-phenylindole dihydrochloride staining, Green Flourescent Protein (GFP) or GFP-survivin–expressing cells were fixed with PBS containing 3.7% formaldehyde and stained with 0.5 μg/mL 4′,6-diamidine-29-phenylindole dihydrochloride in PBS. The percentages of apoptotic cells were determined by confocal microscopy, counting GFP-positive cells having nuclear fragmentation and/or chromatin condensation. In the ELISA assay, the Cell Death Detection Elisa^PLUS kit (Roche) was used following the manufacturer's protocol. This assay determines apoptosis by measuring mononucleosomes and oligonucleosomes in the lysates of apoptotic cells. The cell lysates were placed into a streptavidin-coated microplate and incubated with a mixture of anti–histone-biotin and anti–DNA-peroxidase. The amount of peroxidase retained in the immunocomplex was photometrically determined with ABTS as the substrate. Absorbance was measured at 405 nm.

Western blot was done as described previously (14). Briefly, cells were lysed in radioimmunoprecipitation assay buffer (1% NP40, 0.5% sodium deoxycholate, 0.1% SDS in PBS). Protein samples were subjected to SDS-PAGE and transferred to nitrocellulose membrane. The membrane was blocked in 5% nonfat milk in PBS overnight and incubated with primary antibody and subsequently with appropriate horseradish peroxidase–conjugated secondary antibody. Signals were developed with ECL reagents (Amersham) and exposed to X-ray films.

Image digitization and quantification were done with UN-SCAN-IT software from Silk Scientific. Anti-survivin, anti-Cdc2, anti-cyclin B1, anti-Bax, and anti–β-tubulin monoclonal antibodies and anti-Bak polyclonal antibodies were purchased from Santa Cruz Biotechnology. Anti-Smac monoclonal antibody and anti–cleaved caspase-3, anti–cleaved caspase-7, anti–cleaved caspase-9, anti-HDAC1, anti-HDAC2, anti-HDAC3, anti-HDAC4, anti-HDAC5, anti-HDAC7, and anti–cleaved PARP polyclonal antibodies were purchased from Cell Signaling Technology. Anti–phospho-survivin (Thr³⁴) polyclonal antibody was purchased from Novus. Anti-Hsp60 monoclonal antibody was purchased from BD Biosciences. APC protein was detected by Western blot as described previously (14). Anti-APC monoclonal antibody was purchased from Calbiochem.

Disuccinimidyl suberate was dissolved in DMSO at 25 mmol/L concentration. Before protein isolation for Western blot, cells were treated with 1 mmol/L disuccinimidyl suberate in PBS for 30 min at 37°C. The stop solution [1 mol/L Tris (pH 7.5)] was then added to a final concentration of 10 mmol/L and incubated for 15 min. Total protein was isolated and Western blot was done as described above.

Cells (20 × 10⁶) were harvested by centrifugation at 600 × g for 10 min at 4°C. The cell pellets were washed once with ice-cold PBS and resuspended with 5 volumes of buffer A [20 mmol/L HEPES-KOH (pH 7.5), 10 mmol/L KCI, 1.5 mmol/L MgCI₂, 1 mmol/L sodium EDTA, 1 mmol/L sodium EGTA, 1 mmol/L DTT, and 0.1 mmol/L phenylmethylsulfonylfluoride] containing 250 mmol/L sucrose. Cells were homogenized with 10 strokes of a Teflon homogenizer and centrifuged twice at 750 × g for 10 min at 4°C. Supernatants were centrifuged at 10,000 × g for 20 min at 4°C. The resulting mitochondrial pellets were resuspended in buffer A and frozen at −80°C for future use. The supernatants of the 10,000 × g spin were further centrifuged at 100,000 × g for 1 h at 4°C, and the resulting supernatants were designated cytosol fraction and stored at −80°C.

Silencer validated small interfering RNAs (siRNA) and negative control siRNA were purchased from Ambion. The sequence for survivin siRNA is sense (5′-GGAACAUAAAAAGCAUUCGtt-3′) and antisense (5′-CGAAUGCUUUUUAUGUUCCTC-3′). Survivin siRNA was transfected into HT-29 cells using X-tremeGENE siRNA transfection reagent (Roche) following the manufacturer's protocol. Cells were cultured and transfected in six-well plates (1 × 10⁵ per well) and the final siRNA concentration was 100 nmol/L. Protein samples were collected at 48 h after transfection for Western blot analysis. For apoptosis assay, drugs were added at 24 h after siRNA transfection and cells were collected at 72 h posttransfection.

The mRNA level of survivin was measured by real-time PCR using the TaqMan Gene Expression assay from Applied Biosystems. Total RNA was isolated from HT-29 cells using the RNeasy kit (Qiagen). Five micrograms of total RNA were used in reverse transcription reaction. The cDNAs were used as templates to perform PCR on an Applied Biosystems 7500 Real-time PCR system following the manufacturer's protocol.

APC^min/+ mice were purchased from The Jackson Laboratory. Mice were housed in an animal-holding room under controlled light, temperature, and humidity and were fed Purina lab rodent diet (LabDiet 5K20) and water ad libitum. All procedures were approved by the Institutional Animal Care and Use Committee of North Dakota State University. For treatment, APC^min/+ mice were separated into four groups (five each): control group, butyrate group, DIM group, and DIM/butyrate combination group. Treatment began at 6 wk of age. For DIM and DIM/butyrate group, DIM was administered in drinking water at a calculated dose of 10 mg/kg/d on days 1, 3, and 5 during week 6. Butyrate was administered in drinking water for 1 wk, at a calculated dose of 24 mg daily, starting on day 1 of week 6 for the butyrate group and on day 7 of week 6 for the DIM/butyrate group. Calculation of the doses was done as shown in the following example: a 20-g mouse drinks ∼3 mL water per day (36); 10 mg/kg equals to a dose of 0.067 mg/mL of DIM in water. Mice in the control group were treated with regular water. The mice were weighed twice weekly and monitored daily for signs of weight loss or lethargy that might indicate intestinal obstruction or anemia. At 16 wk of age, Mice were euthanized by pentobarbital injection (200 mg/kg) and cervical dislocation. After necropsy, the intestinal tracts were dissected from the esophagus to the distal rectum, spread onto filter paper, opened longitudinally with fine scissors, and cleaned with sterile saline. Tumor counts were done under a dissection microscope with ×5 magnification. For in vivo apoptosis detection, separate groups of mice were treated with doses and schedule as above and sacrificed at 24 h after the last dose of treatments and the tumor specimens were fixed in formalin.

The apoptotic cells in tumors were detected using an ApopTag In situ Apoptosis Detection kit (Chemicon). The assay was done according to the manufacturer's protocol. Tumor were fixed in 10% neutral-buffered formalin, dehydrated, and embedded in paraffin. After deparaffinization, the tissue sections were pretreated with proteinase K and then incubated with terminal deoxynucleotidyl transferase enzyme at 37°C for 1 h, washed thrice with PBS, and incubated with antidigoxigenin conjugate in a humidified chamber at room temperature for 30 min. The color was developed by incubating the sections with peroxidase substrate. Apoptosis indices were calculated as the percentage of apoptotic cells among 1,000 tumor cells in randomly selected fields (>5 fields per slide) of nonnecrotic tumor portions from five to six slides for each treatment group.

Differences between the mean values were analyzed for significance using the unpaired two-tailed Student's test for independent samples; P ≤ 0.05 was considered to be statistically significant.

Butyrate has been extensively studied as a cancer prevention agent for colon cancers but with only limited activity observed (11–13). We have previously shown that mutations in the APC gene (which occur in >85% of sporadic colon cancers) render colon cancer cells resistant to HDAC inhibitors (14). Because butyrate acts as a HDAC inhibitor, we hypothesize that APC mutations may also cause resistance to butyrate-induced apoptosis. To determine whether APC plays a role in colon cancer cell apoptosis in response to butyrate, we compared butyrate-induced apoptosis in HT-29/β-Gal and HT-29/APC cells. HT-29 colon cancer cells express two COOH-terminal–truncated mutant APC proteins. HT-29/APC are genetically engineered HT-29 cells in which wild-type APC is expressed from a Zn²⁺-inducible transgene (35). Expression of APC induces apoptosis in HT-29 cells (35). To avoid apoptosis induced by APC expression alone, we used 50 μmol/L zinc to induce APC expression (14). After induction of wild-type APC, apoptosis was observed in HT-29/APC cells when treated with butyrate (Fig. 1A). In contrast, the HT-29/β-Gal cells were resistant. When Zn²⁺ was not added to the culture medium to induce APC expression, HT-29/APC cells showed comparable resistance to butyrate-induced apoptosis (data not shown). We have previously shown that a failure to down-regulate survivin is the key mechanism of APC mutation–induced resistance to HDAC inhibitors (14). To further understand the mechanism of APC-mediated apoptosis after butyrate treatment, we examined the expression of survivin. Down-regulation of survivin was observed in HT-29/APC cells after induction of APC expression and treatment with butyrate, but not in HT-29/β-Gal cells (Fig. 1B). Because HT-29 cell lines express mutant p53 proteins, the down-regulation of survivin seems to be p53 independent.

Because APC is frequently mutated in colon cancer patients, the data above predict the ineffectiveness of butyrate in preventing colon cancers. To overcome resistance to butyrate-induced apoptosis in APC mutant tumors, we tested various agents (including genistein, selenium, DIM, and others) to identify a nontoxic agent that can down-regulate survivin. We found that DIM, a cancer prevention agent from food plants including cabbage and broccoli, was able to down-regulate survivin in HT-29 cells. Treatment with DIM down-regulated survivin in a dose-dependent manner (Fig. 2A). We determined whether down-regulation of survivin by DIM occurred at the transcription level. Using real-time PCR, we found that treatment with 40 μmol/L DIM for 24 hours decreased survivin mRNA level by 53% in HT-29 cells compared with untreated cells (Fig. 2B). Next, we determined whether proteasome-dependent degradation is also involved in the down-regulation of survivin in response to DIM. As shown in Fig. 2C, cotreatment with the proteasome inhibitor MG-132 (10 μmol/L) completely blocked the DIM-induced down-regulation of survivin protein in HT-29 cells. To determine if DIM promotes the degradation of survivin protein, HT-29 cells were treated with 20 μmol/L cycloheximide or 20 μmol/L cycloheximide plus 40 μmol/L DIM, degradation of survivin was determined by Western blotting. DIM exposure promoted survivin degradation in HT-29 cells (Fig. 2D). The stability of survivin protein is maintained by p34^cdc2-cyclin B1 phosphorylation on Thr³⁴ of the protein, and Thr³⁴ dephosphorylation causes survivin degradation (37, 38). To further determine the mechanism of DIM-promoted survivin degradation, we examined the effects of DIM on p34^cdc2 and cyclin B1. As shown in Fig. 2E, DIM treatment caused a significant decrease of cyclin B1 protein level in a dose-dependent manner, whereas the p34^cdc2 level remained unchanged. As a result, survivin Thr³⁴ phosphorylation was decreased after DIM treatment (Fig. 2F), contributing to the decreased stability of survivin. This result indicates that DIM causes survivin degradation independent of wild-type APC, through a mechanism involving decreased survivin Thr³⁴ phosphorylation by p34^cdc2-cyclin B1.

Because DIM is able to down-regulate survivin in HT-29 cells, we tested whether DIM can help overcome APC mutation–mediated resistance to butyrate-induced apoptosis. Pretreatment with 40 μmol/L DIM significantly enhanced butyrate-induced apoptosis in HT-29 cells (Fig. 3A). DIM alone at this dose did not induce a significant level of apoptosis. To identify the key mediators of DIM/butyrate–induced apoptosis, HT-29 cells were treated with DIM, butyrate, or DIM/butyrate combination. Western blot analysis with various antibodies identified significantly higher levels of cleaved (activated) caspase-7 and cleaved (activated) caspase-9 in DIM/butyrate–treated cells (Fig. 3B), indicating that these two caspases potentially mediate apoptosis in the combination treatment. We did not observe caspase-3 activation in these treatments (data not shown). Interestingly, the combination of DIM and butyrate was synergistic in causing survivin down-regulation (Fig. 3B). We also determined the effects of DIM on the expression level of HDAC proteins in HT-29 cells. As shown in Fig. 3C, DIM treatment induced significant decreases in the levels of class I HDACs (HDAC1, HDAC2, HDAC3) in a dose-dependent manner. However, DIM had no effects on the levels of class II HDACs (HDAC4, HDAC5, and HDAC7; Fig. 3C and data not shown). By down-regulating the HDAC proteins, DIM should enhance the effects of butyrate on HDAC inhibition. To further understand the mechanisms of how DIM increases butyrate-induced apoptosis in HT-29 cells, we examined the effects of DIM/butyrate combination on the key apoptosis regulators Bax and Bak. Bax and Bak double-knockout mice displayed developmental defects in multiple tissues, indicating that Bax and Bak are critical apoptosis regulators (39). Deletion of both Bax and Bak also results in resistance to various apoptotic stimuli known to trigger mitochondria-dependent apoptosis (40). In contrast, the BH3-only proteins of the Bcl-2 family (Bid, Bad, etc.) are signal transducers and they require Bax or Bak to induce apoptosis (40, 41). We found that the DIM/butyrate combination treatment caused significant increases of both Bax and Bak proteins, as well as the dimerization of the 21-kDa monomer Bax into 42-kDa dimers (representing the activated Bax; refs. 42, 43; Fig. 3D). Activation of Bax and Bak may facilitate or directly cause the release of proapoptotic mitochondrial proteins into the cytosol. To test this hypothesis, the localization of cytochrome c and Smac proteins was determined by subcellular fractionation experiments. As shown in Fig. 3E, the DIM/butyrate combination caused significant release of cytochrome c and Smac proteins from the mitochondria of HT-29 cells. Cytochrome c together with Apaf-1 promote apoptosome formation and activation of pro-caspase-9 (44, 45) whereas Smac antagonizes the activities of inhibitor of apoptosis proteins and activates caspases (46), which explain the observed activation of caspases and apoptosis in HT-29 cells (Fig. 3A and B).

We understand that survivin is not the only target of DIM. However, survivin may play a key role in the apoptotic effects observed during DIM/butyrate combination treatment. To determine the role of survivin, we overexpressed survivin protein in HT-29 cells by transient transfection. As shown in Fig. 4A, overexpression of survivin blocked DIM/butyrate–induced apoptosis in HT-29 cells. Furthermore, knocking down survivin by siRNA sensitized HT-29 cells to butyrate-induced apoptosis (Fig. 4B). These data suggest that although survivin is not the only target of DIM, it plays a critical role in the activity of DIM when in combination with butyrate. Down-regulation of survivin by DIM lowers the threshold of apoptosis induction and allows the activation of downstream mechanisms of apoptosis (such as Bax activation, cytochrome c/Smac release, and activation of caspases).

To determine whether DIM can be effective when used in combination with butyrate for in vivo cancer prevention using APC^min/+ mice, we examined the effects of DIM/butyrate combination in IMCE cells. IMCE cells are colon epithelial cells derived from APC^min/+ mice, a mouse model widely used in colon cancer prevention research (47). As shown in Fig. 5A and B, DIM was able to down-regulate survivin in IMCE cells and significantly enhance butyrate-induced apoptosis in these cells. Comparing with HT-29 cells, caspase-3 and caspase-7 were activated by DIM/butyrate combination treatment in IMCE cells (Fig. 5C) and potentially mediated apoptosis in these cells. We did not observe caspase-9 activation in these cells (data not shown). The different profiles of caspase activation between IMCE and HT-29 cells could be due to the different genetics of human and mouse cells. Thus, different caspases may mediate apoptosis between HT-29 and IMCE cells. Comparing with DIM-only treatment, the combination of DIM/butyrate did not further reduce the level of survivin protein (Fig. 5C).

We tested whether the DIM/butyrate combination strategy can be applied in vivo for cancer prevention in APC^min/+ mice. First, we tested whether DIM can down-regulate survivin in vivo in APC^min/+ mice and determined the optimal dose of DIM. As shown in Fig. 6A, oral administration of various doses of DIM was effective in down-regulating survivin protein in the intestinal tumors from APC^min/+ mice. The optimal dose was determined to be 10 mg/kg/d due to the solubility of DIM. No toxicities (body weight loss, etc.) were observed at these doses. APC^min/+ mice were then randomly assigned to four groups and treated with DIM or butyrate alone, or DIM/butyrate combination. As shown in Fig. 6B, treatment with DIM/butyrate combination significantly decreased tumor numbers in APC^min/+ mice. Whereas the butyrate-treated group has slightly decreased the number of tumors, DIM alone has no effects on the number of tumors in these mice. To examine apoptosis in tumors, separate groups of mice were treated with doses and schedule as above and sacrificed immediately after drug treatment because apoptotic cells are cleared quickly in vivo. We performed in vivo TUNEL assay to determine the level of apoptosis in the tumors isolated from APC^min/+ mice. As shown in Fig. 6C, significant higher levels of apoptosis were observed in the mice receiving DIM/butyrate combination treatment, comparing with the low levels of apoptosis in control group or mice receiving single-drug treatment.

This study has provided strong support for a novel combination strategy of using DIM and butyrate for colon cancer prevention. Short-chain fatty acids, mainly acetate, propionate, and butyrate, are produced during bacterial fermentation of resistant starch and nonstarch polysaccharides (major components of dietary fiber) in the human colon (48). Butyrate is the major energy source for intestinal epithelial cells (49). By promoting cell differentiation, cell cycle arrest, and apoptosis of transformed colonocytes (50–52), butyrate has been regarded as a cancer-preventive agent and extensively evaluated for colon cancer prevention (53, 54). However, the effects of butyrate in prevention of animal models of colon cancer are often limited and in some cases butyrate even promotes tumor occurrence (11–13, 55). Butyrate is an inhibitor of HDAC enzymes (3, 56). We have recently shown that APC mutations in colon cancer cells cause resistance to HDAC inhibitors in general due to a failure of survivin down-regulation (14). In the present study, we found that APC mutations also caused resistance to apoptosis in colon cancer cells treated with butyrate, as survivin was not down-regulated by butyrate in these cells. Thus, down-regulation of survivin is the key target to overcome colon cancer resistance to butyrate in cancer prevention.

We found that DIM is very effective in down-regulating survivin and that DIM is able to significantly potentiate butyrate-induced apoptosis in colon cancer cells expressing mutant APC. Although survivin is not the only target for DIM, our data indicate that down-regulation of survivin plays a key role in DIM-mediated effects on apoptosis. Whereas overexpression of survivin blocked DIM/butyrate–induced apoptosis in HT-29 cells, knocking down survivin protein by siRNA successfully bypassed the requirement of DIM and induced apoptosis after butyrate treatment (Fig. 4). Other efforts have been developed to inhibit survivin by antisense and ribozymes (57–59) and by using survivin mutants to suppress wild-type survivin (60). However, as a nontoxic small-molecule agent, DIM may prove to be more effective and safer for the cancer prevention purpose.

We have previously shown that HDAC inhibitors down-regulate survivin through a GSK-3β/β-catenin/Tcf-4 pathway, which is dependent on the expression of wild-type APC (14). Here, we show that DIM is able to suppress survivin mRNA expression independent of wild-type APC (Fig. 2B). Survivin expression is regulated by AKT and nuclear factor-κB signaling molecules (61). Inhibition of survivin expression by DIM may be mediated by the effects of DIM on these signaling pathways, which has been documented previously (30, 32). We also show that by down-regulating cyclin B1, DIM is able to cause survivin degradation through an alternative mechanism independent of the status of APC. This mechanism involves a decrease of survivin Thr³⁴ phosphorylation and results in destabilization of survivin, successfully bypassing the requirement of wild-type APC. Through these mechanisms, DIM is able to promote survivin down-regulation in HT-29 cells.

Down-regulation of survivin in HT-29 cells by transfection of survivin-targeting siRNA is not enough to cause apoptosis in our experiments. Apparently, other changes that occurred in the colon cancer cells (inhibition of HDACs, activation of Bax and Bak, and release of Smac and cytochrome c) in response to DIM/butyrate treatment contribute to the increased induction of apoptosis (Fig. 3). The decrease of survivin lowers the threshold for apoptosis induction. We found that activation of caspase-3, caspase-7, and caspase-9 is involved in DIM/butyrate–induced apoptosis in HT-29 and IMCE cells, further indicating a role for the mitochondria-mediated intrinsic pathway of apoptosis (62).

In this report, we showed that DIM is not only effective in in vitro studies but, more importantly, also in enhancing butyrate-induced apoptosis and improving the efficacy of cancer prevention in APC^min/+ mice. We used a three-dose schedule for DIM (one dose every other day and 1-week treatment of butyrate in drinking water). By fine-tuning the doses of DIM and butyrate as well as the scheduling of treatment, further improvement on the efficacy of cancer prevention can be potentially achieved. Taken together, the present findings have important implications for the clinical use of butyrate in colon cancer prevention. Using DIM in combination with butyrate may prove to be an effective strategy for the prevention of colorectal cancers in patients.

No potential conflicts of interest were disclosed.

We thank Jodie Haring (Center for Protease Research, North Dakota State University) for providing help with real-time PCR, Dr. Bert Vogelstein (Johns Hopkins University) for providing the HT-29/APC and HT-29/β-Gal cells, and Dr. Robert H. Whitehead (Vanderbilt University) for providing the IMCE cells.