Extraction and isolation of the two ETSs, 5α-hydroxycostic acid (Comp. 1) and hydroxyisocostic acid (Comp. 2), from Laggera alata was carried out as described previously (16). Their structures are shown in Fig. 1A. The two chemicals were dissolved in DMSO at a final concentration of 50 mM respectively, and diluted in culture medium to the indicated concentration for assigned assay. The vehicle group contained DMSO at 0.1% in its culture medium.

The human Ang2 was purchased from Adipogen (San Diego, CA, USA). VEGF (VEGFA-165) was obtained from Peprotech (Rockyhill, NJ, USA). The concentrations of Ang2 and VEGF used in the subsequent experiments were 400 and 20 ng/ml, respectively. Fetal bovine serum (FBS), low serum growth supplement (LSGS), Dulbecco's modified Eagle's medium (DMEM), phosphatase substrate kit, BCA protein assay kit, Medium 200, Matrigel matrix, M-MLV and TRIzol reagent were all purchased from Thermo Fisher Scientific (Waltham, MA, USA), and Taq DNA polymerase was from Takara (Tokyo, Japan). Evagreen dye was purchased from Biotium (Hayward, CA, USA). Reagents including semaxanib (SU5416, a specific VEGFR inhibitor), phalloidin, heparin, bull serum albumin (BSA), MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) and non-protein chemicals were obtained from Sigma Chemical Co. (St. Louis, MO, USA). The pexmetinib (ARRY614, a specific inhibitor of Tie2 and p38) was purchased from APExBIO (Houston, TX, USA).

The antibodies against VEGFR, p-VEGFR2_Tyr1175, p-Tie2Tyr992, AKT, p-AKTSer473, ERK1/2, p-ERK1/2Thr202/Tyr204, p38, p-p38_Thr180/Tyr182, p-PLCγSer1248, p-SrcTyr416, p-FAKTyr397, p-eNOSSer1177, anti-mouse and anti-rabbit horseradish perioxidase (HRP)-conjugated secondary antibodies were purchased from Cell Signaling Technology (CST, Danvers, MA, USA). The antibodies against Tie2, and p-Tie (Tie2Tyr992 + Tie1Tyr1007) were from Abcam (Cambridge, UK). The anti-β-actin antibody was obtained from Santa Cruz Biotechnology (Dallas, TX, USA). The antibody against E-cadherin was from BD Biosciences (BD, San Jose, CA, USA). Both DAPI and proteinase inhibitor cocktail tablets were from Roche (Basel, Switzerland).

Human umbilical vein endothelial cells (HUVECs) were purchased from Life Technologies Corp. (Carlsbad, CA, USA), and were cultured in the Medium 200 supplemented with LSGS, heparin (0.1 mg/ml), and heat-inactivated FBS (20%, v/v). The human breast cancer MCF-7 cells were obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA), and were cultured in DMEM containing 10% FBS. Both cell types were cultured in a 5% CO₂ humidified atmosphere at 37°C. For some of the in vitro experiments, HUVECs were starved in both serum- and LSGS-free Medium 200 (maintenance medium) containing 0.5% BSA for 4 h, and MCF-7 cells were serum-starved in DMEM medium (maintenance medium) overnight (~8 h). Then, the effects of ETSs were detected. Wild-type zebrafish (AB strain) were bought from Hong Kong goldfish market and maintained in flow-through aquaria at 28°C. The fish were fed with general tropical fish food and brine shrimp. Embryos were collected by natural pairwise mating, and transferred to beakers (25 fish/300 ml beaker) with clean embryo water (60 µg/ml instant ocean salt).

The cellular viability of the HUVECs in the presence of either ETS compound was assayed using the MTT method (17). In brief, HUVECs (5×10³ cells/well) were seeded in 96-well plates and allowed to attach overnight. Cells were then treated with ETSs (from 25 to 200 µM) for 48 h. The staining reagent of 30 µl MTT (5 mg/ml) was added to each well for 4 h. The medium was then removed and 200 µl DMSO was added. After complete dissolution of the precipitated formazan, optical density was measured at 570 nm using a UV-VIS spectrophotometer (Genesys 5, Spectronic Instruments, USA). The cell viability was expressed as percentage (%) of the vehicle group.

The effect of ETSs on VEGF-induced HUVECs proliferation was also evaluated by MTT assay. Starved HUVECs (5×10³ cells/well) were stimulated by VEGF in the presence or absence of the ETSs for 24 h, then the effect of ETSs on the proliferation of HUVECs was evaluated.

Total RNA in HUVECs or zebrafish embryos was isolated using TRIzol reagent and cDNA was conducted using M-MLV (18). Real-time PCR was performed on a Bio-Rad detection system CFX96 in a volume of 20 µl containing Evagreen dye 1 µl, Taq polymerase 0.1 µl, cDNA 4 µl (200 ng), 1 µl of each primer and H₂O 12.9 µl. The reaction protocol was 40 cycles at 95°C for 20 sec, 57°C for 20 sec, and 72°C for 20 sec. A melt curve analysis was performed at the end of the reaction to assess the specificity of amplification. The mRNA levels of the genes were normalized to the internal controls, namely, GAPDH and ELF1α, in HUVECs and zebrafish, respectively. The primer sequences for HUVECs are as follows: hGAPDH forward, 5′-CCCACTCCTCCACCT TTGAC-3′ and reverse, 5′-TCTTCCTCTTGTGCTCTTGC-3′; hVEGFR2 forward, 5′-GGAACCTCACTATCCGCAGAGT-3′ and reverse, 5′-CCAAGTTCGTCTTTTCCTGGGC-3′; hTie1 forward, 5′-GACCCACACTGACCAACACA-3′ and reverse, 5′-GTTGTGCACGTAGATGACGC-3′; hTie2 forward, 5′-CTGCAGTGCAATGAAGCATGC-3′ and reverse, 5′-CTGCAGACCCAAACTCCTGAG-3′. The primer sequences for zebrafish are as follows: zELF1α forward, 5′-GGCTGACTGTGCTGTGCTGATTG-3′ and reverse, 5′-CTTGTCGGTGGGACGGCT AGG-3′; zVEGFA-121 forward, 5′-CTACCCAGTGACCAACGC-3′ and reverse, 5′-GCTCTACAGCTCCTGACGAT-3′; zVEGFA-165 forward, 5′-TGCTCCTGCAAATTCACACAA-3′ and reverse, 5′-ATCTTGGCTTTTCACATCTGCAA-3′; zVEGFR2 forward, 5′-GCCTGATCCACAACTGCTTCC-3′ and reverse, 5′-CTCTCCTCACACGACTCAATGC-3′; zAng2 5′-AGGTGGAGGCTGGACTGTC-3′ and reverse, 5′-GTGGTGAGCAGGTGGATGAC-3′; zTie1 forward, 5′-CAAGAGGCACGGAAGGCTTA-3′ and reverse, 5′-AGTGACAGTAACGCAGAGCC-3′; zTie2 forward, 5′-CTACCCAGTGACCAACGC-3′ and reverse, 5′-GCTCTACAGCTCCTGACGAT-3′.

The effect of the ETSs on cell migration was evaluated by the wound-healing assay according to protocols reported previously (19) with some modifications. Briefly, HUVECs and MCF-7 cells were plated in 24-well plates at a density of 2×10⁵ and 2.5×10⁵ cells/well, respectively. When the cells reached 100% confluence, they were starved to inactivate cell proliferation and a scrape was created with sterilized pipette tip. After washings twice with PBS, the cells were overlaid with fresh medium containing various concentrations of ETSs or SU5416 (4 µM), co-treated with either VEGF or Ang2. Images were taken using an inverted microscope (Nikon TE300, Japan) at 0 h and after 20-h incubations. The extent of migration was analyzed using an Image-Pro Plus software (Media Cybernetics, USA). Wound size of the vehicle group was set as 100%.

Tube formation assay was carried out as described previously (20) with some modifications. In brief, HUVECs (6×10³ cells/well) in the medium were seeded onto the Matrigel matrix-coated 96-well plates along with VEGF. Various dilutions of ETSs or SU5416 (4 µM) were added to the wells and kept for 6 h. The tubular structures were captured and quantified using an Image-Pro Plus software.

HUVECs (1×10⁵ cells/ml) or MCF-7 cells (2×10⁵ cells/ml) were seeded to the clear cover glass-bottom 35-mm Petri-dishes for confocal imaging overnight at 37°C before starvation. During cell starvation, ETSs (100 µM) were added to the cells 4 h before the cells were treated with either VEGF or Ang2 for specified duration (20 min to 6 h) depending on the objective of the experiment. Collected cells were washed twice with ice-cold PBS, fixed with 4% paraformaldehyde for 15 min, and permeabilized with 0.1% Triton X-100 in PBS. After rinsing twice with PBS, the cells were blocked in the 3% BSA in PBS for 1 h and immunostained with corresponding primary antibody at 4°C overnight. Then, the cells were incubated with the secondary fluorescent antibody for 1 h at room temperature. Finally, the cell nucleus was visualized with DAPI, after washing with PBS, and images were captured using a confocal microscope (Olympus FV1000 IX81, Tokyo, Japan) (21).

HUVECs or MCF-7 cells (1×10⁶ cells/dish) were seeded in 100-mm culture dishes for 24 h before starvation. During cell starvation, ETSs were exposed to cells for 4 h and then stimulated with VEGF or Ang2 for specified time interval. Whole-cell extracts were collected using a lysis buffer supplemented with proteinase inhibitors. Protein concentration was measured using the BCA protein assay kit. Equal amounts of proteins (40 µg) were resolved by electrophoresis on 12% SDS-PAGE gels and then transferred to PVDF membrane (Millipore, Billerica, MA, USA). The membrane was blocked with 3% BSA at room temperature for 1 h, incubated with primary antibody at 4°C overnight followed by HRP-conjugated goat secondary antibodies at room temperature for 1 h. Protein bands were visualized using enhanced chemiluminescence detection reagents (Bio-Rad, Hercules, CA, USA) (22). The resulting images were scanned using a scanner (Canon PIXMA MP150, Japan).

Wild-type zebrafish (AB strain) embryos were selected to evaluate the anti-angiogenic ability of ETSs in vivo, a quantitative endogenous alkaline phosphatase (EAP) assay was performed as described previously (18). Briefly, healthy, limpid, and regular embryos (24 h post fertilization, 24 hpf) were arrayed in a 96-well plate (1 embryo/well) and treated with ETSs (1–10 µM) until 48 hpf. Embryos were then stained according to the manufacturer's instructions of the phosphatase substrate kit (Pierce, Waltham, MA, USA). The optical density of soluble end product was measured at 405 nm using a UV-VIS spectrophotometer. Vessel growth is presented as the percentage in optical density compared to that of the vehicle group: % vessel formation = (OD treated at 48 hpf - OD vehicle at 24 hpf)/(OD vehicle at 48 hpf - OD vehicle at 24 hpf) × 100%.

All experiments were performed at least three times. Data are presented as the mean ± standard deviation (SD). Statistical analysis was determined using one-way ANOVA.

The chemical structures of ETSs are shown in Fig. 1A. Before assessing the anti-angiogenic properties of ETSs, the MTT assay was used to evaluate their toxic effects on HUVECs in the normal growth medium. As shown in Fig. 1B, various concentrations (0–100 µM) of ETSs were non-toxic to HUVECs. In this regard, the concentrations of ETSs (≤100 µM) used in the subsequent in vitro experiments were considered non-toxic.

VEGF plays an important role during angiogenesis for its mitogenic effect on vascular endothelial cells (23). Therefore, we measured the inhibitory effect of ETSs on VEGF-induced HUVECs. The viability of HUVECs was upregulated significantly from 77.32 to 100% (p<0.005) when VEGF was added (Fig. 1C), while with the addition of ETSs, the viability was suppressed dose-dependently (Fig. 1C). At 25 µM of Comp. 2, cell viability decreased to 97.21% (p<0.05). A stronger inhibitory effect was shown by Comp. 1 with viability of 91.07% (p<0.01). These data suggest that VEGF-induced proliferation of the HUVECs is sensitive to the presence of ETSs (Fig. 1B and C). Next, the anti-angiogenic activities of ETSs were examined in the wild-type zebrafish embryos. As shown in Fig. 1D, the two selected ETSs dose-dependently inhibited the vessel formation in the zebrafish embryos in the range of 1.3–10 µM. At the concentration of 5 µM, both ETSs have exhibited very significant anti-angiogenic activities (p<0.005).

The effects of ETSs on VEGF-mediated HUVECs migration were investigated using a wound-healing assay (Fig. 2A). HUVECs migrated to a clear area when stimulated with VEGF, and the stimulatory effect of VEGF was inhibited by ETSs in a dose-dependent manner (Fig. 2A). Even at 25 µM, the two compounds exhibited very significant inhibitory effects (p<0.005) (Fig. 2A). These data reveal that ETSs inhibit VEGF-stimulated migration of the endothelial cells.

The activation of endothelial cell migration by VEGF involves a massive remodeling of the actin cytoskeleton that reorganizes into transcytoplasmic stress fibers (24). As shown in Fig. 2B, the immunofluorescent intensity of F-actin (green) stained with FITC-conjugated phalloidin was highly increased in the VEGF-stimulated HUVECs, which indicated an increase of the stress fibers formation. However, such stress in the endothelial cells was markedly attenuated in the presence of ETSs (Fig. 2B). These results show that ETSs might suppress the migration of the VEGF-induced HUVECs by inhibiting the formation of the stress fiber.

During angiogenesis, a critical step is the formation of the new capillary tube derived from the migrated endothelial cells (4). Therefore, we investigated whether ETSs could inhibit VEGF-induced endothelial cell tube formation. As shown in Fig. 2C, a rough, but complete tube network was formed from the VEGF-stimulated HUVECs. However, with the treatment of ETSs (100 µM), the tube formation pattern disassembled in a manner similar to that of the vehicle group (p>0.05, data not shown). Quantification of the tube length revealed that 25 µM ETSs could abolish the VEGF-stimulated tube formation significantly (p<0.05) (Fig. 2C), suggesting a potent inhibitory effect of ETSs on the tubulogenesis of the VEGF-induced endothelial cells.

The effects of ETSs on the angiogenic-related gene expression levels in HUVECs were evaluated by the real-time PCR analysis. HUVECs were exposed to ETSs (50 µM) for 24 h, the mRNA levels of VEGFR2 and Tie1/2 were then determined. As shown in Fig. 3A, both ETSs significantly inhibited the mRNA levels of VEGFR2 (p<0.005), the key receptor of VEGF. Only Comp. 1 downregulated the activity of the angiopoietin orphan receptor Tie1 (p<0.01). However, both compounds significantly reduced the mRNA level of angiopoietin receptor Tie2 (p<0.01 or 0.005). These data suggest that ETSs apparently acted as inhibitors of angiogenesis by regulating VEGF/VEGFR2 and Ang2/Tie2 pathways. Thus, the following results from in vivo experiment focused on the two pathways.

To determine the effects of ETSs on the pro-angiogenic-related gene expression levels in vivo, zebrafish embryos were treated with ETSs (10 µM) for 24 h and subjected to real-time PCR analysis. As shown in Fig. 3B, the mRNA levels of both VEGFA and VEGFR2 were downregulated significantly in the presence of ETSs. As for the Ang2/Tie axis, ETSs downregulated the mRNA levels of Ang2 (p<0.01 or p<0.005) and Tie2 receptor (p<0.01), but no significance was found in Tie1 (Fig. 3B). The PCR data from the zebrafish embryos further confirmed their anti-angiogenic activities.

Our previous results proved that ETSs effectively suppressed VEGF-stimulated angiogenesis both in vitro and in vivo. To understand the molecular mechanism of ETS-induced inhibition of angiogenesis, we investigated whether ETSs suppressed the phosphorylation of VEGFR2, the critical receptor tyrosine kinase on the surface of endothelial cells (25). The addition of exogenous VEGF to HUVECs strongly increased the phosphorylation of VEGFR2 at Ser1175 site (Fig. 4). However, pretreatment with ETSs suppressed the VEGF-stimulated VEGFR2 phosphorylation (Fig. 4).

VEGF binding to VEGFR2 activates various downstream signaling molecules with distinct and overlapping functions, which is responsible for endothelial cell proliferation, migration and tube formation (5,26). Therefore, we explored whether ETSs inhibited VEGF/VEGFR2-mediated signaling pathways. Starved HUVECs were pretreated with ETSs for 4 h and subsequently stimulated by VEGF for 20 min. As shown in Fig. 4, an extensive downregulation of the activation of Src/AKT/eNOS signaling pathway was observed at 50 µM ETSs (Fig. 4). ETSs also suppressed the activation of the VEGFR2 downstream signaling molecules, namely, FAK, PLCγ, ERK1/2 and p38. Taken together, these data suggested that ETSs suppressed angiogenesis by inhibiting the VEGF/VEGFR2-mediated Src/AKT/eNOS, FAK, PLCγ/ERK1/2, and p38 signaling pathways.

With the stimulation of VEGF in endothelial cells, the mRNA level of Ang2, another crucial pro-angiogenic factor, increased (11). As shown in Fig. 5, in the presence of VEGF, the Ang2 mRNA level increased significantly compared to the vehicle. Pretreatment by ETSs (50 µM) almost completely reversed the increase induced by VEGF (p<0.001). VEGF induced Ang2 secretion and synthesis, this Ang2 could bind with Tie2 and promote the angiogenic activity of VEGF. The low Ang2 level of ETS-treated group may indicate that Ang2/Tie2 might have taken part in the ETS-induced anti-angiogenesis. To clarify the direct effects of ETSs on Ang2/Tie2 axis, we completed the following experiments.

Ang2 acts as an agonist of Tie2 receptor and leads to the phosphorylation of Tie2. This would result in vascular destabilization and remodeling (22). Hence, we utilized exogenous Ang2 to induce Tie2 phosphorylation, while the activation of Tie2 in the presence of ETSs was evaluated by both immunofluorescent staining and the western blotting. Starved HUVECs were pretreated with ETSs (100 µM) for 4 h and stimulated with Ang2 for 30 min. Compared to the vehicle group (DMSO-treated), Ang2 increased the phosphorylation of Tie which occurred on the cell membrane (Fig. 6A). However, for the group treated with ETSs (100 µM), they showed weaker fluorescence intensity than that with only the Ang2 stimulation, indicative of the inhibition of the phosphorylation of Tie receptor in the presence of ETSs. In the western blotting results (Fig. 6B), the data showed the inhibitory effects of ETSs on Ang2-induced Tie2 phosphorylation.

Ang2 promotes the proangiogenic action of VEGF leading to blood vessel growth. Besides, Ang2 also enhances tumor growth and cancerous metastasis (27). In order to evaluate whether ETSs could inhibit Ang2 stimulated cancer cell migration, we conducted wound-healing analysis on breast cancer MCF-7 cells. Data revealed that stimulation with Ang2 (400 ng/ml) highly enhanced cell mobility, leading to smaller wound area (Fig. 7A). However, ETSs inhibited Ang2-induced MCF7 cells migration dose-dependently from 25 to 100 µM. At 25 µM of ETSs, Comp. 1 exhibited stronger inhibitory effect with larger wound area (p<0.005) than that of Comp. 2 (Fig. 7A).

E-cadherin is a cell-cell adhesion molecule with pivotal roles in tissue formation, epithelial cells behavior, and is linked to the invasion of cancer cells (28). With the stimulation of Ang2 on MCF-7 cells, we observed the loss of E-cadherin (Fig. 7B). Western blot assay of the E-cadherin protein level also confirmed the loss of E-cadherin induced by Ang2 (Fig. 7C). However, ETS treatment suppressed the E-cadherin loss, both immunofluorescent staining and western blot analysis showed the inhibitory effect of ETSs. Besides, the p-AKT level was upregulated after Ang2 stimulation in MCF-7 cells, while the ETS exposed cells showed a sharp reduction of the p-AKT level (Fig. 7C). These data suggested that ETSs inhibited Ang2-induced MCF-7 cell migration by suppressing E-cadherin loss and AKT phosphorylation.