Active PI3Kα (100 ng) was pre-incubated with various doses of HS-159 or LY294002 for 5 min in kinase reaction buffer (25 mM MOPS, pH 7.0; 5 mM MgCl₂ and 1 mM EGTA) and 10 μg L-α-phosphatidylinositol. The L-α-phosphatidylinositol was sonicated in water for 20 min before being added to the reaction to facilitate micelle formation. The reaction was initiated by adding 10 μM ATP and allowed to run for 180 min at room temperature. To terminate the kinase reaction, an equal volume of Kinase-Glo^® buffer (Promega, Madison, WI) was added. After 10 min, luminescence of the plates was measured with a GloMax plate reader.

Human liver cancer (HepG2, Huh-7 and Hep3B) cells were purchased from the Korean Cell Line Bank (KCLB, Seoul, Korea). Huh-7 cells were cultured in Roswell Park Memorial Institute 1640 (RPMI-1640) medium with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin. HepG2 and Hep3B cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin. Human umbilical vein endothelial cells (HUVECs) were grown in a gelatin-coated 75-cm² flask with M199 medium containing 20 ng/ml basic fibroblast growth factor (bFGF), 100 U/ml heparin and 20% FBS. All cell cultures were maintained at 37°C in a CO₂ incubator with a controlled humidified atmosphere composed of 95% air and 5% CO₂.

Cell viability was evaluated using a 3-(4,5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide (MTT) assay. Briefly, cells were seeded at a density of 1×10⁴ cells/well in a 96-well plate and incubated for 24 h. The medium was then removed and the cells were treated with either DMSO as a control or various concentrations (0.1–10 μM) of HS-159 for 48 h at 37°C. Next, 10 μl of MTT solution (2 mg/ml) was added to each well and the plate was incubated for another 4 h at 37°C. The medium was then removed and the formazan crystals that had formed were dissolved in DMSO (200 μl/well) with constant shaking for 5 min. Absorbance of the solution was then measured with a microplate reader at 540 nm. This assay was conducted in triplicate.

Total cellular proteins were extracted with lysis buffer containing 1% Triton X-100, 1% Nonidet P-40 and a 1% protease and phosphatase inhibitor cocktail containing aprotinin (10 mg/ml), leupeptin (10 mg/ml; ICN Biomedicals, Asse-Relegem, Belgium), phenylmethylsulfonyl fluoride (1.72 mM), NaF (100 mM), NaVO₃ (500 mM) and Na₄P₂O₇ (500 mg/ml; Sigma-Aldrich, St. Louis, MO). The proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto nitro-cellulose membranes. The blots were incubated with the appropriate primary antibodies followed by incubation with secondary antibodies conjugated to horseradish peroxidase. Antibody binding was detected with an enhanced chemiluminescence reagent (Amersham Biosciences). Antibodies against p-mTOR (Ser²⁴⁴⁸), mTOR, p-Akt (Ser⁴⁷³), Akt, p-P70S6K (Thr³⁸⁹), P70S6K, PARP, cleaved caspase-3, hypoxia inducible factor-1α (HIF-1α), vascular endothelial growth factor (VEGF) and β-actin were purchased from Cell Signaling Technology Inc. (Danvers, MA).

Huh-7 cells were plated on 18-mm cover glasses in RPMI-1640 medium and incubated for 24 h at 37°C so that ∼70% confluence was reached. The cells were then incubated in the presence or absence of 10 μM HS-159, washed twice with PBS and fixed in an acetic acid: ethanol solution (1:2, v/v) for 10 min at −20°C. The cells were blocked in 1.5% horse serum in PBS for 30 min at room temperature and then incubated overnight at 4°C with primary antibodies against p-Akt, p-mTOR and p-P70S6K (Cell Signaling) in a humidified chamber. After washing twice with PBS, the cells were then incubated with rabbit TRITC secondary antibody (1:100, Dianova, Germany) for 1 h at room temperature. The cells were also stained with 4, 6-diamidino-2-phenylindole (DAPI) to visualize the nuclei. The slides were then washed twice with PBS and covered with fluorescent mounting medium (Dako North America Inc., Carpinteria, CA) before viewing with a confocal laser scanning microscope (Olympus, Tokyo, Japan).

Huh-7 cells were plated in 100-mm culture dishes with RPMI-1640 medium at ∼70% confluence and incubated for 24 h at 37°C. The next day, the cells were treated with either DMSO as a control or various concentrations (0.01–10 μM) of HS-159. Floating and adherent cells were collected and fixed overnight in cold 70% ethanol at 4°C. After washing with PBS, the cells were subsequently stained with 50 μg/ml propidium iodide (PI) and 100 μg/ml RNase A for 1 h at room temperature in the dark. Flow cytometry was then performed with a FACSCalibur flow cytometer (Becton-Dickinson, San Jose, CA) to determine the percentage of cells in specific phases of the cell cycle (sub-G₁, G₀/G₁, S and G₂/M). The data were analyzed using FlowJo software (Tree Star, Inc.). All the experiments were performed in triplicate.

Huh-7 cells were plated on an 18-mm cover glass with RPMI-1640 medium at ∼70% confluence and incubated for 24 h at 37°C. The cells were then incubated in the presence or absence of 10 μM HS-159 for 24 h. The cells were then fixed in cold 2% para-formaldehyde (PFA), washed with PBS and stained with 2 μg/ml of DAPI (Sigma-Aldrich) for 20 min at 37°C. The stained cells were examined under a fluorescence microscope for evidence of nuclear fragmentation. A TUNEL assay was subsequently performed using a commercial kit (Chemicon, Temecula, CA) according to the manufacturer’s instructions.

Mitochondrial membrane potential was assessed with a mitochondria staining kit (MitoPT, Immunohistochemistry Technologies, Bloomington, MN) using 5, 5’, 6, 6’-tetrachloro-1, 1’, 3, 3’-tetraethylbenzimidazol-carbocyanine iodide (JC-1), which indicates potential-dependent accumulation in mitochondria. Huh-7 cells were plated on 18-mm cover glass in RPMI-1640 medium and incubated for 24 h at 3°C. When confluence reached ∼70%, the cells were incubated in the presence or absence of 10 μM HS-159 at 37°C for 6 h. Next, 100 μl of a JC-1 solution (12.5 μg/ml final concentration) was added to each well and the plate was incubated for 30 min at 3°C. After washing twice with PBS, the cells were stained with DAPI to visualize the nuclei. The slides were then washed twice with PBS and covered with DABCO before viewing with a confocal laser scanning microscope (Olympus). The results [green/red (G/R) fluorescence ratio] were expressed as a percentage of the control.

Matrigel (10 mg/ml, 200 μl; BD Biosciences) was polymerized for 30 min at 37°C. HUVECs were suspended at a density of 2.5×10⁵ cells/well in M199 medium supplemented with 5% FBS. Aliquots (0.2 ml) of the cell suspension were added to each well coated with Matrigel and incubated with or without different concentrations of HS-159 for 24 h at 37°C. Morphological changes of the cells and tube formation were observed with a phase-contrast microscope. The cells were captured at magnification ×200.

HUVECs were plated on 60-mm culture dishes at ∼90% confluence. A razor blade was then used to create a wound 2 mm in width in the cell mono-layer and the injury line was marked. After wounding, the detached cells were removed with serum-free medium and the remaining HUVECs were further incubated in M199 medium supplemented with 5% FBS, 1 mM thymidine (Sigma-Aldrich) and HS-159 (0.5 μM). The cells were allowed to migrate for 18 h before being rinsed with serum-free medium and fixed in absolute methanol. Cell migration was quantitated by counting the number of cells that had moved beyond the injury line.

Data are expressed as the mean ± standard deviation (SD). Statistical analysis was performed using an ANOVA and unpaired Student’s t-test. P-values ≤0.05 were considered statistically significant. All calculations were performed using SPSS software for the MS Windows operating system (version 10.0; SPSS, Chicago, IL).

A new PI3Kα inhibitor, HS-159 (N-(5-(3-(3-cyanophenyl)imidazo[1,2-a]pyridin-6-yl)pyridin-3-yl)benzenesulfonamide) was synthesized as described in our previous study (Fig. 1A) (25). Three-dimensional (3D) coordinates in the X-ray crystal structure of PI3Kα in the resting form (PDB code: 2RD0) were selected as the receptor model for docking simulations. After removing the solvent molecules, hydrogen atoms were added to each protein atom. We used the Auto Dock program for studying docking between PI3Kα and HS-159 because the outperformance of its scoring function over those of the others had been shown in several target proteins. The predicted binding mode of HS-159 in the ATP-binding site of PI3Kα is shown in Fig. 1B. The imidazopyridine of HS-159 maintained stable hydrogen-bonding in the hinge region (Val851). The pyridyl group was anchored by hydrogen bonds to Asp933 and interacted with gatekeeper residue Tyr836. Two stable hydrogen bonds were established between the cyano group of C3 and Thr856. These hydrogen bonds were received from the enzyme and seemed to play a crucial role as an anchor for HS-159 binding. HS-159 could be further stabilized in the ATP-binding site by forming hydrophobic interactions with Met772, Pro778, Lys802, Val850, Ser854, Gln859 and Ile932. Judging from the overall structural features derived from our docking simulations, the inhibitory activity of HS-159 is likely to stem from multiple hydrogen bonds and hydrophobic interactions simultaneously established in the ATP-binding site of PI3Kα. We also found that HS-159 more strongly inhibited PI3Kα activity (EC₅₀ 1.892 nM) compared to LY294002, a conventional specific PI3K inhibitor (EC₅₀ 488.8 nM, Fig. 1C).

Since HS-159 inhibited PI3Kα activity, we next determined whether this drug also affected downstream signal transduction factors that promote PI3K-mediated cell survival. When Huh-7 cells were treated with various concentrations of HS-159 for 6 h, the phosphorylation of Akt and its downstream components such as mTOR and P70S6K was effectively suppressed in a dose-dependent manner (Fig. 2A). We further assessed the effect of HS-159 on Akt, mTOR and P70S6K using a fluorescent imaging system. Results from this assay again showed that HS-159 strongly suppressed phosphorylation of these proteins in Huh-7 cells, similar to the western blotting results (Fig. 2B).

Next, we further evaluated the anti-proliferative effects of HS-159 on HCC cells using an MTT assay. Huh-7, HepG2 and Hep3B cells were exposed to various concentrations (0.1–10 μM) of HS-159 for 48 h. HS-159 markedly reduced viability of the three HCC cell lines in a dose-dependent manner (Fig. 3). These results indicated that HS-159 exerts potent anti-proliferative effects on HCC cells.

Cell apoptosis and growth are correlated with cell cycle progression (26). In the present study, HS-159 (0.1-10 μM) inhibited the growth of HCC cells in a dose-dependent manner (Fig. 3). Thus, we performed flow cytometry to monitor changes of the cell cycle profile induced by HS-159. Data from this analysis revealed that treatment with 5 and 10 μM HS-159 increased the number of cells in a sub-G₁ stage, indicative of apoptosis and decreased the fraction of cells in the G₀/G₁ phase in Huh-7 cells (Fig. 4A).

HS-159-induced apoptosis was also evaluated by DAPI and TUNEL staining to characterize nuclear morphology. As shown in Fig. 4B, cells treated with 10 μM HS-159 had morphological features characteristic of apoptotic cells such as bright nuclear condensation and perinuclear apoptotic bodies observed with DAPI staining. HS-159-induced apoptosis was also confirmed by detecting DNA fragmentation using a TUNEL assay. In addition, we performed JC-1 staining to measure the effect of HS-159 on changes in mitochondrial membrane potential that correlate with the intrinsic pathway of apoptosis. As shown in Fig. 4C, the cytoplasm of control cells showed heterogeneous staining with both red and green fluorescence coexisting in the same cells. Consistent with mitochondrial localization, red fluorescence (corresponding to high mitochondrial membrane potential, ψ_m) was mostly found in granular structures distributed throughout the cytoplasm. Treatment with 10 μM HS-159 decreased the red fluorescence while frequent clusters of mitochondria were still observed. HS-159 induced marked changes in ψ_m as evident from the disappearance of red fluorescence or increased green fluorescence in most cells. We next performed western blotting to evaluate the activation of caspase-3 and PARP after treatment with HS-159. As expected, HS-159 increased the levels of cleaved caspase-3 and PARP in Huh-7 cells in a dose-dependent manner (Fig. 4D). Taken together, these results showed that HS-159 induces apoptotic cell death in Huh-7 cells.

A low oxygen level is characteristic of solid tumors and a negative prognostic factor for cancer patient survival. The response of cancer cells to hypoxia not only drives neo-angiogenesis, but also enhances cell survival and malignant phenotype development. HIF-1α is a major regulator of cellular adaptive responses to hypoxia as well as a major transcriptional modulator of angiogenic factors such as VEGF (27). We therefore evaluated the effect of HS-159 on the expression of HIF-1α and VEGF in Huh-7 cells under hypoxia conditions. The cells were treated with various concentrations of HS-159 cultured for 16 h with 100 μM CoCl₂ for mimic hypoxic conditions. As shown in Fig. 5A, the expression of HIF-1α and VEGF was increased after exposure to hypoxia and subsequently suppressed by HS-159 treatment, respectively.

In endothelial cells, PI3K/Akt signaling actively mediates processes associated with angiogenesis including cell migration and capillary tube formation (28). Therefore, we performed capillary tube formation and migration assays to determine whether HS-159 suppresses the angiogenic activity of HUVECs. HS-159 inhibited the formation of vessel-like structures resulting from HUVEC elongation and alignment (Fig. 5B). HS-159 also markedly inhibited cell migration (Fig. 5C). These results indicate that HS-159 is a potential anti-angiogenic reagent that inhibits the tube formation and migration of HUVECs.