PT was obtained from Calbiochem (San Diego, CA, USA). Propidium iodide (PI) was purchased from Sigma-Aldrich (St. Louis, MO, USA). Growth factor-reduced Matrigel was purchased from BD Biosciences (San Diego, CA, USA). Mouse recombinant VEGF was purchased from R&D Systems Inc., (Minneapolis, MN, USA). Anti-VEGF, anti-VEGF receptor (VEGFR)1, anti-VEGFR2 and anti-actin antibodies were obtained from Cell Signaling Technology (Danvers, MA, USA). PT was dissolved in dimethylsulfoxide (DMSO; Sigma-Aldrich) to a concentration of 100 μM and stored in the dark at −20°C. An aliquot of DMSO was stored under the same conditions and used as a control treatment.

Human umbilical vein endothelial cells (HUVECs) were purchased from the American Type Culture Collection (ATCC; Rockville, MD, USA) and cultured in endothelial cell growth medium (EGM^®-2 MV Bullet kit; Lonza Inc., Walkersville, MD, USA) containing 2% fetal bovine serum (FBS). The HUVECs were used at passages 3–4.

The human CRC cell lines, HT-29, SW620 and HCT116, were obtained from ATCC and cultured in RPMI-1640 medium supplemented with 10% FBS, 100 units penicillin, and 100 units streptomycin. For the treatment of the cells with PT, the cells were subcultured in RPMI-1640 medium without FBS for 12 h. PT dissolved in DMSO was diluted with FBS-free medium to achieve the indicated concentrations. The same concentrations of DMSO were applied to the cells as a control.

The HUVECs and CRC cells were plated at a density of 1×10⁴ cells/well in 96-well plates. The cells were treated with PT for 24 h, the medium was removed, and 200 μl of fresh medium plus 20 μl of MTT, 2.5 mg dissolved in 50 μl of DMSO were added to each well. Following incubation for 4 h at 37°C, the culture medium containing MTT was withdrawn and 200 μl of DMSO were added, followed by shaking until the crystals were dissolved. Viable cells were determined by measuring the absorbance at 570 nm using a microplate reader (Molecular Devices, Sunnyvale, CA, USA).

Cell cycle and sub-G1 distribution were determined by the staining of DNA with PI (Sigma-Aldrich) [excitation (Ex)/emission (Em), 488/617 nm]. In brief, 1×10⁶ cells were incubated with PT for 24 h. The cells were then washed with phosphate-buffered saline (PBS) and fixed in 70% ethanol overnight. The cells were washed again with PBS and then incubated with PI (10 μg/ml) and simultaneously treated with RNase at 37°C for 1 h. The percentages of cells in the different phases of the cell cycle or the sub-G1 DNA content were measured using a BD LSR flow cytometer and analyzed using CellQuest software (Becton Dickinson, New York, NY, USA).

Cell migration was evaluated by a wound healing assay, according to the method described in the study by Reischer et al (17). HUVECs (1×10⁶/well) were seeded in 6-cm culture plates and allowed to form a confluent monolayer. The monolayer was then scraped with a P200 pipette tip to generate a wound of approximately 1 mm in diameter. Subsequently, the medium was changed to fresh endothelial basal medium (EBM-2; supplemented with 20 ng/ml of VEGF) in the absence or presence of PT. Images of the wounds were captured after 24 h of incubation at ×40 magnification, and the wound area was determined using an inverted microscope (IX71; Olympus, Center Valley, PA, USA). The ability of the cells to close the wound, that is their motility, was evaluated by determining the healed area. The percentage of the healed area was calculated as follows:

Matrigel (200 μl/well) was added to a 24-well plate and polymerized for 30 min at 37°C. The HUVECs (1×10⁴ cells/well) were seeded onto each well of the Matrigel-coated 24-well plate and then incubated in EGM-2 containing 2% FBS. Following overnight incubation, the medium was changed to fresh EBM-2 (supplemented with 20 ng/ml of VEGF) in the absence or presence of PT. The formation of endothelial cell tubular structure was visualized under an inverted microscope and photographed at ×40 magnification. Furthermore, the tube formation was quantified by calculating the number of polygons and was expressed as a percentage by normalization with PT-untreated control cells.

The cells were collected, washed twice with PBS, and then lysed for 30 min on ice in a lysis buffer (50 mM Tris-HCl pH 8.0, 150 mM EDTA, 1% Triton X-100, 0.5% SDS and protease inhibitor cocktail). The protein concentration in the cell lysates was measured using a Protein Quantification kit (Bio-Rad, Hercules, CA, USA). Samples (50 μg protein/well) were loaded onto a SDS-PAGE gel. After transferring and blocking, the membrane was probed with anti-VEGF, anti-VEGFR1, anti-VEGFR2 and anti-actin antibodies. The signal was detected using WEST-one™ enhanced chemiluminescence solution (Intron Biotechnology Inc., Daejeon, Korea), captured and analyzed by a Luminescent Image Analyzer (LAS-3000; Fujifilm, Tokyo, Japan).

All animal experiments were approved by the Chonbuk National University Animal Care and Use Committee (Approval no: CBU 2012-0035). The HT-29 (6×10⁶) cells were injected into nude mice. The mice were randomly assigned to the control and treatment group and intraperitoneally injected 3 times a week with the vehicle (DMSO) or 4 mg/kg PT. Treatment with PT or the vehicle commenced 7 days after tumor cell implantation (25 mm³ tumor volume). The experiment was terminated at 28 days, and the tumors were harvested for immunohistochemistry. Immunohistochemistry was carried out in paraffin-embedded 5-μm-thick tissue sections. The sections were immunostained with an antibody against mouse CD31 (sc-507; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA), visualized by appropriate biotin-conjugated secondary antibodies followed by immunoperoxidase detection with the Vectastain ABC Elite kit (Linaris, Dossenheim, Germany) and diaminobenzidine (DAB) substrate (Vector Laboratories, Peterborough, UK). Five equal-sized fields were randomly selected. Four fields at ×40 magnification were selected and the CD31-positive cells were counted.

The data are presented as the means ± standard error (SE) of at least 3 independent experiments performed in duplicate. Representative blots are shown. All the data were entered into Microsoft Excel 5.0 and SPSS software was used to perform the two-tailed t-tests or the analysis of the variance, where appropriate. P-values <0.05 were considered to indicate statistically significant differences.

To determine the anti-angiogenic effects of PT in vitro, we used MTT assay to examine HUVEC viability and survival. Treatment of the HUVECs with PT at a dose of up to 2.5 μM did not decrease cell viability. However, at a dose of >5 μM PT, the viability of the HUVECs was significantly reduced in a dose-dependent manner (P<0.05), showing 2.94±1.31% viability at 20 μM (Fig. 1A). We also evaluated the effects of PT on cell cycle modifications. After 24 h of incubation of the HUVECs with or without PT, the cells were analyzed by PI staining and flow cytometric analysis. Treatment with PT resulted in the presence of a sub-G1 cell population, suggestive of cell death. The sub-G1 cell population was also increased in a dose-dependent manner (Fig 1B).

Cell migration is a key step in angiogenesis (18). We therefore investigated the effects of PT on the motility of the endothelial cells by wound healing assay. The results revealed that PT significantly reduced the migration ability of the HUVECs at 2.5 μM (P<0.05; Fig. 2A).

To further investigate the effects of PT on endothelial cells, we examined the VEGF-stimulated tube formation of HUVECs using Matrigel. It is well known that endothelial cells are able to spontaneously form a capillary-like network on Matrigel in vitro, an important feature in the process of angiogenesis (19). HUVECs spontaneously formed a capillary-like tube structure following 24 h of incubation on Matrigel (Fig. 2B). However, treatment with PT (2.5 μM) markedly inhibited the VEGF-stimulated tube formation of the HUVECs, resulting in less elongated, broken and foreshortened tubes.

VEGF induces endothelial cell proliferation, migration and survival through its cell surface receptors, VEGFR1/VEGFR2 (20). To elucidate the underlying mechanisms responsible for the anti-angiogenic activity of PT, the effects of PT on the expression of VEGF and its receptors in the HUVECs were investigated. Western blot analyses revealed that the treatment of the HUVECs with PT markedly reduced the protein levels of VEGF, VEGFR1 and VEGFR2 compared to the levels in control cells (Fig. 3).

To assess the anti-angiogenic effects of PT in human CRC cells, we investigated its effects on cell viability by MTT assay. The HT-29, SW620 and HCT116 cells were treated with various concentrations (0, 5, 10 and 20 μM) of PT for 24 h. The viability of these 3 cell lines was decreased in a dose-dependent manner, indicating that the IC₅₀ of PT is approximately 20 μM (Fig. 4A).

We also determined the expression levels of VEGF and its receptors in human CRC cells. In addition, western blot analyses revealed that the levels of VEGF and VEGFR2 were significantly decreased in a dose-dependent manner following treatment with PT (Fig. 4B); the results obtained for the CRC cells are in accord with the results obtained for the HUVECs.

To examine the effects of PT on angiogenesis in vivo, we used a xenograft nude mouse tumor model in which mice were subcutaneously implanted with HT-29 cells. Immunohistochemical analysis of CD31 revealed that the blood vessel network was well developed in the tumors from the control (vehicle-treated) mice, whereas the development of the blood vessel network was substantially inhibited by the administration of PT (Fig. 5). Moreover, the tumor tissues from the mice treated with PT displayed a significantly lower number of CD31-positive stained cells, as compared to those from control mice (P<0.01).