Human colon carcinoma HT-29 cells were obtained from the Cell Bank of Chinese Academy of Sciences (Shanghai, China) and grown in DMEM supplemented with 10% (v/v) FBS, 100 U/ml penicillin and 100 μg/ml streptomycin (Invitrogen, Carlsbad, CA, USA). Cells were cultured at 37°C, in 5% CO₂ humidified incubator.

Oleanolic αcid (OA, ≥90%) was purchased from Sigma-Aldrich (St. Louis, MO, USA). For cell-based experiments, stock solution of OA was prepared by dissolving OA powder in DMSO to a concentration of 20 mM. The working concentrations of OA were made by diluting the stock solution in the culture medium. The final concentrations of DMSO in the medium were 0.1%. For animal experiments, OA was dissolved in saline.

Six-week-old male BALB/c athymic (nude) mice (20–22 g) were obtained from Shanghai SLAC Laboratory Animal Co., Ltd. (Shanghai, China) and housed in the specific pathogen-free controlled conditions (22°C, a 12-h light/dark cycle). Food and water were given ad libitum. HT-29 CRC xenograft mice were produced as previously described (19). Briefly, 2.5×10⁶ of HT-29 cells were subcutaneously injected into the right flank of mice to initiate tumor growth. At day 3, following xenograft implantation, mice were randomly divided into two groups (n=10) and intraperitoneally injected with 12.5 mg/kg of OA or saline, daily, 6 days per week for 16 days. Tumor volume was measured and calculated by the formulation of ‘π/6 × L × W²’, where ‘L’ and ‘W’ refer to length and width, respectively. All animal experiments were approved by the Institutional Animal Care and Use Committee of Fujian University of Traditional Chinese Medicine.

Tumor tissues were analyzed by immunohistochemistry staining as described before (19). Briefly, 8 tumors were randomly selected from OA-treatment or control groups. Tumor tissues were fixed in 10% formaldehyde for 12 h, paraffin-embedded, sectioned and placed on slides. The slides were subjected to antigen retrieval and endogenous peroxidase activity was quenched with hydrogen peroxide. Non-specific binding was blocked with normal serum in PBS (0.1% Tween-20). Rabbit polyclonal antibody against PCNA at dilution of 1:200 (Santa Cruz Biotechnology, CA, USA) was used to detect the relevant proteins. The binding of the primary antibody was demonstrated with a biotinylated secondary antibody, horse-radish peroxidase (HRP)-conjugated streptavidin (Dako) and diaminobenzidine (DAB) as the chromogen. The tissues were counterstained with diluted Harris hematoxylin. After staining, five high-power fields (at magnification of ×400) were randomly selected in each slide and the proportion of positive cells in each field was determined using a true color multi-functional cell image analysis management system (Image-Pro Plus, Media Cybernetics, USA). PBS was used to replace the primary antibody as a non-specific control.

HT-29 cells were seeded into 96-well plates at 1.0×10⁴ cells per well and treated with various concentrations of OA for 24 h or 50 μM of OA for different time-point. After treatment, MTT assay was performed as previously described (20). Briefly, 10 μl MTT [5 mg/ml in phosphate-buffered saline (PBS)] was added to each well, and the samples were incubated for an additional 4 h at 37°C. The purple-blue MTT formazan precipitate was dissolved in 100 μl DMSO. The cell viability was measured at 570 nm with a Model ELX800 Microplate Reader (BioTek, VT, USA).

HT-29 cells were seeded into 6-well plates at a density of 2×10⁵ cells/well and treated with various concentrations of OA for 24 h. The cells were harvested, resuspended in medium and re-seeded into 6-well plates at a density of 1.0×10³ cells/well. After incubation for 10 days in a 37°C humidified incubator with 5% CO₂, colonies were counted by light microscopy.

HT-29 cells were seeded into 6-well plates at a density of 2×10⁵ cells/well and treated with indicated concentrations of OA for 24 h. The cells were harvested, resuspended and adjusted to a concentration of 1×10⁶ cells/ml and fixed in 70% ethanol at 4°C overnight. The fixed cells were washed twice with cold PBS and incubated for 30 min with RNase (8 μg/ml) and PI (10 μg/ml). The cell cycle phases were analyzed by flow cytometry (Caliber, Becton-Dickinson, CA, USA) as previously described (22). The fluorescent signal was detected through the FL2 channel and the proportion of DNA in different phases was analyzed using ModfitLT Version 3.0 (Verity Software House, Topsham).

Paraffin-embedded sections of tumors (4-μm thick) were analyzed by TUNEL staining with TumorTACS in situ kit (R&D Systems, Minneapolis, MN, USA) according to the manufacturer's recommended protocol. The proportion of apoptotic cells were recorded as DAB-positive cells (brown stained). Five high-power fields (x400) were randomly selected in each slide and the average proportion of positive cells in each field was counted.

HT-29 cells were seeded into 6-well plates at a density of 2×10⁵ cells/well and treated with various concentrations of OA for 24 h. The cells were harvested, resuspended and stained with Annexin V/PI (Becton-Dickinson, CA, USA) according to the manufacturer's instructions followed by flow cytometry analysis. The percentage of cells in early apoptosis was calculated by Annexin V-positivity and PI-negativity and the percentage of cells in late apoptosis was calculated by Annexin V-positivity and PI-positivity.

The protein expression was detected by western blot analysis. Protein of tumor tissues or HT-29 cells was extracted using RIPA protein extraction kit (Tiangen Biotech, Beijing, China), respectively. The protein were separated by SDS-PAGE and transferred onto PVDF membranes. The membranes were blocked for 1 h with 5% non-fat milk and incubated with primary antibody against Cyclin D1, CDK4, p21, Bcl-2, Bax or β-actin (all in 1:1,000 dilutions; Cell Signaling Technology, Beverly, MA, USA) overnight at 4°C. The HRP-conjugated secondary antibodies (1:2,000 dilution) were added. Then the levels of protein expression were detected with enhanced chemiluminescence detection.

The phosphorylation of multiple proteins was simultaneously detected by Bio-Plex phosphoprotein assay as described before (22). Briefly, 5×10⁵ of HT-29 cells were seeded into 25-cm² flasks with 5 ml medium and treated with 50 μM of OA for 24 h. Eight tumors were randomly selected from each group and homogenized. After lysis of treated cells or tumor tissues the expression of pAkt, pErk1/2, pJNK, pp38, pp70S6K and pp53 was examined using the Bio-Plex 200 suspension array system Bio-Plex Phosphoprotein assay (Bio-Rad, Hercules, CA, USA) according to the manufacturer's protocol.

The data were analyzed using SPSS for Windows (Version 17.0) and shown as average with SD. Statistical analysis was performed with Student's t-test or ANOVA.

Tumor growth in vivo was evaluated by measuring the tumor volume and tumor weight in CRC xenograft mice. As shown in Fig. 1A, administration with OA significantly decreased both tumor volume and tumor weight. At the end of experiment, the tumor volume and tumor weight per mouse in control group were 1.36±0.32 cm³ and 0.94±0.14 g, whereas those in OA-treated group were 0.79±0.14 cm³ and 0.51±0.08 g (P<0.05). The in vitro effect of OA on CRC growth was determined by MTT assay to compare the relative viability of HT-29 cells in OA-treated monolayers and untreated controls. As shown in Fig. 1B, treatment with 25–150 μM of OA for 24 h reduced HT-29 cell viability by 16–78% compared to untreated control cells; and 50 μM of OA gradually decreased cell viability with the increase of exposure time (P<0.05). Taken together, it is suggested that OA possesses inhibitory effects on CRC growth both in vivo and in vitro, in a dose- and time-dependent manner.

Cancers are typically characterized by an uncontrolled increase of cell proliferation (30); which therefore is a critical target of anticancer treatment. To determine the in vivo effect of OA on cancer cell proliferation, we performed immunohistochemical staining (IHS) to evaluate the expression level of a proliferation marker PCNA in CRC tumor tissues. As shown in Fig. 2A, OA treatment significantly decreased the protein expression of PCNA in CRC xenograft mice. The percentage of PCNA-positive cells in tumors from control and OA-treated CRC mice was 30.64±5.18 and 14.75±5.17%, respectively (P<0.05). The effect of OA on CRC cell proliferation in vitro was assessed by colony formation assay. As shown in Fig. 2B, OA significantly and dose-dependently decreased the survival rate of HT-29 cells (P<0.05). Eukaryotic cell proliferation is tightly regulated by the cell cycle, and G1/S transition is a main checkpoint critical for the control of the cell cycle progress. To investigate the mechanism of anti-proliferative effect of OA, we performed FACS analysis with PI staining to evaluate cell cycle. As shown in Fig. 2C, OA treatment significantly and dose-dependently reduced the S-phase proportion of HT-29 cells. Collectively, OA can inhibit CRC cell proliferation via G1/S cell cycle arrest.

Apoptosis eliminates abnormal cells and hence is essential for tissue homeostasis. Dysregulation of apoptosis represents a major causative factor of tumorigenesis (31). Therefore, promoting cell apoptosis has been a major focus for antitumor therapies. The in vivo apoptosis was evaluated by TUNEL assay. As shown in Fig. 3A, treatment with OA significantly enhanced the percentage of TUNEL-positive cells in tumors from CRC mice. HT-29 cell apoptosis in vitro was examined using Annexin V/PI staining followed by FACS analysis. As shown in Fig. 3B, OA treatment increased the percentage of cells undergoing apoptosis in a dose-dependent manner.

The mitochondrion-dependent pathway is the most common apoptotic pathway in vertebrate animal cells. Mitochondrial outer membrane permeabilization (MOMP) results in the release of numerous apoptogenic proteins and eventually induces apoptosis, which thereby is a key commitment step in the induction of cellular apoptosis. During the process of MOMP the electrochemical gradient across the mitochondrial membrane collapses. Therefore, the loss of mitochondrial membrane potential is a hallmark for apoptosis. Since the membrane-permeant JC-1 dye displays potential-dependent accumulation in mitochondria, indicated by a fluorescence emission shift from green (525 nm) to red (590 nm), collapse of mitochondrial potential during apoptosis can be represented by a decrease in the ratio of red/green fluorescence intensity of JC-1. As shown in Fig. 3C, OA treatment significantly and dose-dependently reduced JC-1 red/green fluorescent ratio or mitochondrial membrane potential in HT-29 cells (P<0.05). These data together suggest that OA induces CRC cell apoptosis in vivo and in vitro via the mitochondrion-dependent pathway.

G1/S progression is highly mediated by Cyclin D1 and its catalytic partner CDK4 (32,33). An unchecked or hyperactivated Cyclin D1/CDK4 complex often leads to uncontrolled increase in cell proliferation (34,35). As a proliferation inhibitor, p21 protein plays a role in G1 arrest by binding to and inhibiting the activity of Cyclin-CDK complexes (36). Bcl-2 family proteins are critical mediators of mitochondrion-dependent apoptosis, functioning as either promoters (e.g., Bax) or inhibitors (e.g., Bcl-2) (37). The ratio of Bcl-2 to Bax is critical for determining the fate of cells. Higher Bcl-2/Bax ratio is commonly found in various types of cancer, conferring a survival advantage to cancer cells (38–40).

To explore the mechanisms whereby OA exerts its anti-proliferative and pro-apoptotic activities, we performed western blot analysis to determine the protein expression of the above-mentioned factors. As shown in Fig. 4, OA significantly reduced the protein level of pro-proliferative Cyclin D1 and CKD4 as well as anti-apoptotic Bcl-2 in both CRC tumors and HT-29 cells, whereas that of anti-proliferative p21 and pro-apoptotic Bax was significantly increased after OA treatment. Taken together, these data demonstrate that OA exerts its antitumor activities through modulating the expression of critical genes involved in cell proliferation and apoptosis.

The development of malignant tumors including CRC is strongly associated with multiple intracellular signal transduction cascades such as MAPK, Akt, p70S6K and p53 pathways (3–11). After activation by PI3K, Akt phosphorylates mTOR which in turns regulates p70S6K phosphorylation and activation. The Akt-mTOR-p70S6K signaling pathway is considered as a central regulatory pathway of protein expression involved in regulating cell proliferation and survival. Mammals have three major subfamilies of MAPK, including ERK, JNK and p38. The MAPK signaling proceeds via a three-tiered kinase core consisting of MAPK kinase kinase, MAPK kinase, and ultimately a given MAPK. Signaling of both Akt and MAPK are major cell-survival and proliferation pathways and have been shown to be activated in several cancers including CRC. In addition, Akt and MAPK pathways can influence the tumor suppressor p53, a transcription factor involved in many cellular processes such as DNA repair, cell cycle arrest and apoptosis. p53 is one of the most frequently mutated genes in human cancers. To further elucidate the mechanisms of antitumor activity of OA, we used Bio-Plex Phosphoprotein assay to examine the activation/phosphorylation of multiple proteins in CRC xenograft tumor tissues and HT-29 cells. As shown in Fig. 5, OA treatment significantly reduced the phosphorylation levels of Akt, ERK, JNK, p38 and p70S6K both in vivo and in vitro, while p53 phosphorylation was remarkably increased after OA treatment, suggesting that OA profoundly modulates the activation of multiple CRC-related signaling pathways in vivo and in vitro.

In conclusion, OA exerts its antitumor effects via modulation of multiple cellular targets. Results from this study may provide a strong scientific foundation for the development of novel multi-targeted anticancer agents from the bioactive ingredients in medicinal herbs.