HeLa and KM12C cells were grown in DMEM with penicillin (100 U/ml)-streptomycin (100 U/ml) (Invitrogen, Carlsbad, CA, USA) and 10% heat-inactivated fetal bovine serum (Hyclone, Logan, UT, USA). HCT116 and SNU638 cells were maintained in RPMI-1640 medium supplemented as above. The cells were purchased from American Type Culture Collection (ATCC, Manassas, VA, USA). Cobalt chloride (CoCl₂), desferrioxamine (DFO), MG-132, and cycloheximide (CHX) from Sigma Chemical Co. (St. Louis, MO, USA). Antibody for HIF-1α was obtained from BD Biosciences (San Diego, CA, USA). CoCl₂ was reported as a widely used mimetic of hypoxia in a large range of cells, the molecule is known to inhibit prolyl hydroxylases leading to HIF-1α stabilization (14). Thus, in this study, CoCl₂ was used to induce hypoxia mimicking condition. Then, the cell culture was kept in a gas-controlled chamber (Thermo Electron Corp., Marietta, OH, USA) maintained at 5% CO₂ and 37°C. Kamebakaurin was isolated from Isodon excia (Maxin.) Hara and the structure is shown in Fig. 1A. The purity of kamebakaurin was >98% in HPLC analysis.

The ability of the compound to inhibit hypoxia inducible factor was determined by HRE-dependent reporter assay as previously described (15). In brief, at 50-80% confluence, HCT116 cells were cotransfected with the vectors for pGL3-HRE-Luciferase plasmid containing six copies of HREs derived from the human VEGF gene and with pRL-CMV (Promega, Madison, WI, USA) using Lipofectamine 2000 reagent (Invitrogen). Following 24 h of incubation, the cells were treated with various concentrations of kamebakaurin and incubated for 16 h in hypoxia. Luciferase assay was performed using Dual-luciferase reporter assay system according to the instructions of the manufacturer (Promega). Luciferase activity was determined in Microlumat plus luminometer (EG&G Berthold, Bad Wildbad, Germany) by injecting 100 µl of assay buffer containing luciferin and measuring light emission for 10 sec. The results were normalized to the activity of renilla expressed by cotransfected Rluc gene under the control of a constitutive promoter. Data were analyzed using ANOVA (analysis of variance).

HCT116 cells were seeded at 1×10⁵ cells/ml in 96-well plates containing 100 µl of RPMI-1640 with 10% FBS and incubated overnight. Kamebakaurin was dissolved in DMSO and DMSO was added to all plates to compensate the same volume of DMSO. After 24 h, the cells were pretreated with different concentrations of kamebakaurin for 24 h. Subsequently, cells were cultured with MTT solution (5 mg/ml) [3-(4,5-dimethyl-thiazol-2-yl)-2,5-diphenyltetrazolium bromide] (Sigma, St. Louis, MO, USA) for 4 h. The viable cells converted MTT to formazan, which generated a blue-purple color after dissolving in 100 µl of DMSO. The absorbance at 570 nm was measured by micro-plate reader (Molecular Devices, Sunnyvale, CA, USA).

Whole-cell extracts were obtained by lysing cells in ice-cold lysis buffer (50 mM Tris-HCl, pH 7.5, 1% Nonidet P-40, 1 mM EDTA, 1 mM phenylmeth-ylsulfonylfluoride) supplemented with the protease inhibitor cocktail (Sigma-Aldrich). HIF-1α protein was analyzed in nuclear extracts prepared from cells using NE-PER reagent (Pierce, Rockford, IL, USA), according to the instructions of manufacturer. An aliquot of protein extracts were used to determine protein concentration by the Bradford method. Fifty microgram of whole-cell extracts or 30 µg of nuclear extract protein per lane was separated by SDS-polyacrylamide gels and followed by transferring to a polyvinylidenedifluoride membrane (Millipore, Bedford, MA, USA). The membrane was blocked with 5% skim milk, and then incubated with the corresponding antibody. Antibody for HIF-1α was obtained from BD Biosciences. The primary antibodies for VEGF, Topo-I, GLUT1, c-Myc and α-tubulin were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). The primary antibody for cyclin D1 was purchased from Cell Signaling Technology (Beverly, MA, USA). After binding of an appropriate secondary antibody coupled to horseradish peroxidase, proteins were visualized by enhanced chemiluminescence according to the instructions of the manufacturer (Amersham Pharmacia Biotech, Buckinghamshire, UK).

HCT116 cells were plated in a 96-well plate at a density of 1×10⁵ cells per well and treated with various concentrations of kamebakaurin for 12 h under hypoxia conditions. The VEGF levels in the culture supernatant and the serum were determined by ELISA using the Duo-Set ELISA development kit (R&D Systems, Inc., Minneapolis, MN, USA) according to the manufacturer's instructions.

Total RNA from HCT116 cells was obtained using RNA Mini kit (Qiagen, Valencia, CA, USA). Total RNA (2 µg) was used to perform reverse transcription-PCR (RT-PCR) using RT-PCR kit (Invitrogen) according to the manufacturer's protocol. The PCR primers for VEGF were 5′-GCTCTACCTCCACCATGCCAA-3′ (sense) and 5′-TGGA AGATGTCCACCAGGGTC-3′ (antisense); for EPO were 5′-CACTTTCCGCAAACTCTTCCG-3′ (sense) and 5′-GTC ACAGCTTGCCACCTAAG-3′ (antisense); for HIF-1α were 5′-CTCAAAGTCCGACAGCCTCA-3′ (sense) and 5′-CCCT GCAGTAGGTTTCTGCT-3′ (antisense); for GAPDH were 5′-ACCACAGTCCATGCCATCAC-3′ (sense) and 5′-TCCA CCACCCTGTTGCTGTA-3′ (antisense). The oligonucleotide sequences of the reaction products were confirmed by sequencing.

Distribution of cells in different stages of cell cycle was analyzed by BD Accuri™ C6 flow cytometry (Becton-Dickinson, Franklin Lakes, NJ, USA). The kit utilizes propidium iodide (PI) staining to allow quantitative measurements of percentage of cells in the G0/G1, S and G2/M phases on the Cell Quest software (Becton-Dickinson). For all assays, 10,000 events were counted. The ModFit LT V4.0 software package (Verity Software, Topsham, ME, USA) was used to analyze the data. The cell cycle analysis was performed according to the manufacturer's protocol, and as previously described (16). Briefly, HCT116 cells (5×10⁵ cells/ml) were treated with different concentrations of kamebakaurin for 12 h and harvested from culture dishes. After washing with PBS, HCT116 cells were fixed with ice-cold 70% ethanol at −4°C for 12 h. The cells were then washed with PBS containing 0.1% Triton X-100, stained with PI/RNase reagent for 30 min and analyzed by BD Accuri™ C6 flow cytometry. The ModFit LT V4.0 software package (Verity Software) was used to analyze the data.

All surgical procedures and care applied to the animals were in accordance with IACUC guidelines. Six weeks old specific-pathogen-free Crj:BALB/c nu/nu female athymic nude mice (Vital River, China) were randomly assigned to three groups, each of which consists of five mice (n=5 per group), and then were subcutaneously inoculated with 0.2 ml of HCT116 cells (5×10⁷ cells/ml) in the left flank region. Kamebakaurin, dissolved in DMSO, was administered orally every other day for 40 days at a dose of 15 and 50 mg/kg body weight starting from day 10 post cell implantation to mice. Tumor weight was calculated every five days using the equation: [length × (width)²]/2. Tumors were harvested 4 h after the last treatment, followed by homogenising in RIPA for western blot analysis.

All values are expressed as mean ± SD. A comparison of the results was performed with one-way ANOVA and Tukey's multiple comparison tests (Graphpad Software, Inc, San Diego, CA, USA). Statistically significant differences between groups were defined as p-values <0.05.

To investigate whether kamebakaurin (Fig. 1A) inhibited HIF-1α transcriptional activation, we transfected HCT116 cells with a luciferase reporter gene driven by six specific HRE. A substantial increase of luciferase activity was observed in cells cultured in hypoxic conditions, whereas kamebakaurin dose-dependently inhibited hypoxia-induced luciferase activity (Fig. 1B). Given that the inhibition of HIF-1α transcriptional activation might be correlated with kamebakaurin-induced cytotoxicity, parallel studies of cell viability were performed (Fig. 1C). After the HCT116 cells were treated with different concentrations of kamebakaurin for 24 h, no significant alteration of cell viability was observed relative to the untreated control group.

To explore the underlying mechanism of kamebakaurin activity, we investigated its effect on HIF-1α protein levels. In HCT116 cells, HIF-1α protein is undetectable under normoxia, whereas it is stabilized under hypoxia or in the presence of CoCl₂ and becomes readily detectable by western blotting. Following 12 h of treatment, kamebakaurin exerted dose-dependent inhibition of HIF-1α protein levels induced by hypoxia or CoCl₂ in HCT116 cells, with complete abrogation at 30 µM (Fig. 2A). In contrast to the decrease of HIF-1α levels, kamebakaurin had almost no effect on the levels of Topo-I protein. Next, in order to address whether the inhibition of HIF-1α by kamebakaurin was cell line specific, we extended these studies to a diverse set of tumor cell lines with tissues of various origins, including the cervical cancer cell line HeLa, gastric cancer cell line SNU638 and colorectal-cancer cell line KM12C (Fig. 2A). In addition, induced-accumulation of HIF-1α by well-characterized hypoxia mimetic reagents, including CoCl₂ and DFO, could also be abrogated by kamebakaurin (Fig. 2B).

We next performed an immunofluorescence assay to evaluate the effect of kamebakaurin on HIF-1α expression in HCT116 cells. Following 12 h of treatment, kamebakaurin (30 µM) exerted almost complete inhibition of HIF-1α protein levels in cell nuclei induced by hypoxia in HCT116 cells (Fig. 2C). To further confirm the effects of kamebakaurin, we conducted both time-course experiments and dose-response experiments to determine the expression of HIF-1α protein in the presence of kamebakaurin. The addition of kamebakaurin to the culture medium remarkably inhibited HIF-1α accumulation, and this inhibition persisted as long as the drug was present, at least up to 12 h (Fig. 2D). Dose-response experiments indicated that kamebakaurin dose-dependently inhibited hypoxia-induced accumulation of HIF-1α in HCT116 cells, with complete abrogation at 30 µM (Fig. 2E).

Generally, the accumulation of HIF-1α is dependent on the balance between its protein synthesis and degradation. To further address the mechanism by which kamebakaurin inhibits HIF-1α protein level, we examined whether kamebakaurin modulates HIF-1α protein synthesis in the presence of a proteasome inhibitor MG-132 to prevent HIF-1α degradation. As expected, addition of proteasome inhibitor caused the increased accumulation of HIF-1α protein levels in the presence of CoCl₂ (Fig. 3A). Kamebakaurin inhibited the accumulation of HIF-1α protein induced by CoCl₂ despite of the presence of MG-132. No significant effects were observed on Topo-I levels.

To address the effect of kamebakaurin on HIF-1α protein stability, the protein translation inhibitor cycloheximide (CHX) was used to prevent de novo HIF-1α protein synthesis and then the cells were exposed to normoxia for increasing periods up to 60 min. The nuclear extracts were prepared to detect HIF-1α protein by western blotting. Under these conditions, HIF-1α protein levels mainly reflected the rate of HIF-1α degradation. As shown in Fig. 3B, the degradation of HIF-1α was similar in the kamebakaurin-treated and the control cells. Therefore, it is confirmed in our experiments that kamebakaurin does not promote the degradation of HIF-1α. To determine whether HIF-1α synthesis inhibition by kamebakaurin was a downstream effect from decreased HIF-1α gene transcription or HIF-1α mRNA stability, we analyzed HIF-1α mRNA levels by RT-PCR. Kamebakaurin did not reduce HIF-1α mRNA levels significantly (Fig. 4A). This suggests that kamebakaurin-mediated decrease of HIF-1α synthesis is likely due to downregulation of HIF-1α mRNA translation.

The expression of VEGF and EPO, which are involved in tumor cells proliferation, angiogenesis, invasion and metastasis, is known to be regulated by HIF-1α (5,11). We therefore examined whether kamebakaurin decreases the expression of these genes. VEGF and EPO mRNA levels were measured by RT-PCR analysis in HCT116 cells. Treatment of the cells with kamebakaurin resulted in a dose-dependent inhibition of VEGF and EPO mRNA expression (Fig. 4A). Then, we also examined the hypoxia induction of VEGF or GLUT1 (glucose transporter 1) protein expression. Consistently, they were dose-dependently inhibited by kamebakaurin (Fig. 4B). The concentrations to inhibit the expression of HIF-1α target genes were comparable with those of HIF-1α protein accumulation. This result led us to measure the VEGF protein concentration in the culture supernatant by ELISA. Consistently, the hypoxic induction of secreted VEGF protein was dose-dependently inhibited by kamebakaurin (Fig. 4C).

To evaluate the effect of kamebakaurin on cell proliferation, we tested whether the antiproliferative effect of kamebakaurin is associated with cell cycle arrest by measuring the DNA content of nuclei of HCT116 cells in flow cytometric analysis. As shown in Fig. 5B, treatment with 30 µM kamebakaurin markedly induced G1/S phase cell cycle arrest. FACS analysis revealed that 24 h exposure to kamebakaurin increased the population of G1/S phase cells in a dose-dependent manner. Cells at the G1/S phase increased from 69.74% in medium alone to 76.14, 80.00 and 82.49% in the presence of 3, 10 and 30 µM kamebakaurin, respectively (Fig. 5A). While kamebakaurin treatment retarded the progression of G1 to S/G2 phase.

Next, we determined the specific cell cycle regulators responsible for the cell cycle arrest induced by kamebakaurin by western blot analysis using antibodies specific to cyclin D1 and c-Myc. The result showed that cyclin D1 and c-Myc were decreased by kamebakaurin dose-dependently (Fig. 5B). Taken together, these results clearly suggest that antiproliferative effect of kamebakaurin is associated with its induction of cell cycle arrest at G1 phase, and further confirmed that kamebakaurin downregulates cyclin D1 and c-Myc protein levels on the cell cycle.

To further reveal the effect of kamebakaurin on the expression of HIF-1α and VEGF in vivo, we next determined whether these results could be translated into an in vivo xenograft model. HCT116 cells were subcutaneously implanted in athymic nude mice, and the experimental mice were treated with kamebakaurin (15 and 50 mg/kg) every other day until the end of the study. As expected, kamebakaurin (50 mg/kg) produced significant growth inhibition of HCT116 cells in a tumor xenograft model, compared to that of the vehicle-treated control group (Fig. 6A). Due to the key roles of HIF-1α in tumor angiogenesis, we studied its expression in the tumors by western blotting. Consistent with the finding in cultured cells, kamebakaurin significantly decreased the protein levels of HIF-1α in the tumors, whereas no significant difference was observed in Topo-I levels (Fig. 6B).

VEGF has key roles in tumor angiogenesis, therefore, we measured the concentration of secreted VEGF in the serum of xenograft mouse. Consistent with the findings in cultured cells, kamebakaurin significantly decreased the serum VEGF levels dose-dependently (Fig. 6C). Taken together, our study showed that the downregulation of HIF-1α by kamebakaurin could contribute to the inhibition of tumor growth and VEGF secretion in a tumor xenograft model with HCT116 cells.