Baicalein was purchased from Sigma-Aldrich Co. (St. Louis, MO, USA). Dulbecco’s modified Eagle’s medium (DMEM) was purchased from Invitrogen (Carlsbad, CA, USA). Fetal bovine serum (FBS), penicillin, and streptomycin were ordered from Gibco-BRL (Rockville, MD, USA). The apoptosis detection kit was from Nanjing KeyGen Biotech. Co. Ltd. (Nanjing, China). MTT (3-(4,5-dimethyl-2-thiazole)-2,5-diphenyltetrazolium bromide) was purchased from Sigma Chemicals Co. (St. Louis, MO, USA). Caspase inhibitor z-VAD-fmk and anti-cytochrome c were purchased from Beyotime (Haimen, China), anti-MEK1 and anti-Phospho-MEK1 (Thr386) anti-Phospho (Thr386) MEK1 (p-MEK1) were from Millipore Co. (Billerica, MA, USA). Anti-ERK1/2, anti-Phospho (Thr202/Tyr204) ERK1/2 (p-ERK1/2), anti-Phospho-MEK1/2 (p-MEK1/2), anti-caspase-3, anti-Bad and anti-Phospho-Bad(Ser112) antibodies were purchased from Cell Signaling Technology Inc. (Danvers, MA, USA).

Male BALB/c nude mice (4-week-old) were purchased from the Beijing Experimental Animal Center and maintained in the Laboratory Animal Center of Xi’an Jiaotong University, in accordance with the University Institutional Animal Care and Use Committee. HepG₂ cells (2×10⁶) suspended in 200 μl of DMEM were injected subcutaneously into the right inguinal area of the 6-week-old male nude mice. All animals developed palpable tumors. Mice were divided into two groups (n=6 per group): group I, treatment with vehicle DMSO as the control group; group II, treatment with 20 mg/kg/day Baicalein via oral adminitration. Treatments were started one week after the injection of HepG₂ HCC cells. Resulting tumors were measured using a vernier caliper every two days following the tumor cell injections, and tumor volumes were calculated using the formula: volume = (length x width²)/2 and expressed as mean size ± standard error.

Human HCC cell lines (HepG₂, BEL-7402, SMMC-7721) and human normal liver cell line (HL-7702) were purchased from Shanghai Institute of Cell Biology (Shanghai, China). Cells were cultured in DMEM supplemented with 10% FBS, 100 U/ml penicillin, 100 μg/ml streptomycin, and 2 mmol/l glutamine. All cells were incubated at 37°C with 5% CO₂.

The full-length pcDNA3.1 (Invitrogen, Paisley, UK) MEK1 vector was made by cloning of the full-length PCR product of MEK1 with KOD^® DNA polymerase (Toyobo, Osaka, Japan). All the plasmid sequences were confirmed by DNA sequencing. For transient transfection experiments, cells were plated 24 h before transfection in a 6-well plate at a density of 2×10⁵. Lipofectamine 2000 (Invitrogen) was used to perform transfection with 4.0 μg pcDNA3.1(+)-MEK1 vector or 4.0 μg pcDNA3.1(+) empty vector (as a negative control) according to the manufacturer’s protocol.

Cell viability was determined by a colorimetric 3-(4,5-dimethylthiazol-2-yl) 2,5-diphenyltetrazolium bromide (MTT) assay as previous reported (11). In brief, after treatment of cells with or without the indicated agent and/or serum for 48 h, the cells were washed twice with PBS and incubated with 0.5 mg/ml MTT (Sigma) for 4 h. The reagent was absorbed by living cells and eventually formed an insoluble blue formazan product. After the incubation period, cells were washed with PBS, solubilized with dimethyl sulfoxide (DMSO), and quantified using a microplate reader at the absorbance of 550 nm. The inhibition rate was determined using SPSS software (version 17.0, SPSS Inc, Chicago, IL, USA).

Apoptotic and/or necrotic cells were evaluated by Annexin V binding and propidium iodide (PI) uptake using an Annexin V-FITC/PI kit as previously described (12). Briefly, tumor cells were plated at a density of 1×10⁵ cells per well into 6-well plates for 24 h. The cells were treated with various concentrations of Baicalein (0, 40, 80 and 120 μM) and incubated at 37°C for 24 and 48 h. The cells were washed with cold PBS and resuspended in Annexin V binding buffer. The cells were stained with Annexin V-FITC for 15 min, washed, and then stained with PI. The samples were analyzed by flow cytometer with CellQuest software.

Loss of MMPΔΨm was assessed by flow cytometry, using a fluorescent indicator Rh123, as previously described (13,14). Briefly, cells were treated with Baicalein at different concentrations (0, 20, 40 and 60 μM) for 24 h. Then, Rh123 working solution was added to the culture at a final concentration of 2 μg/ml and then incubated in the dark at 37°C for 30 min. Cells were then washed with PBS, and fluorescence of Rh123 was detected immediately using a FACSCalibur, at an excitation wavelength of 488 nm and an emission wavelength of 525 nm.

Cell lysates were prepared by incubating 2×10⁶ cells/ml in extraction buffer (25 mM Tris-HCl, pH 7.5, 20 mM MgCl₂, and 150 mM NaCl, 1% Triton X-100, 25 μg/ml leupeptin, and 25 μg/ml aprotinin) for 30 min on ice. Lysates were centrifuged at 12,000 x g for 15 min. Cellular extracts (30 μg) were then incubated in a 96-well microtitre plate with 20 ng Ac-DEVD-pNA (caspase-3 activity) or Ac-LEHD-pNA (caspase-9 activity) (Beyotime) for 2 h at 37°C. Caspase activity was measured by cleavage of the Ac-DEVD-pNA or Ac-LEHD-pNA substrate to pNA, the absorbance of which was measured at 405 nm. Relative caspase activity was calculated as a ratio of emission of treated cells to untreated cells.

Western blot analysis was executed as previously described (15). Whole-cell extracts were prepared from Baicalein-treated or control-treated cells cultured in 6-well plates. After incubation, cells were harvested and resuspended in lysis buffer, washed with ice-cold PBS and lysed in extraction buffer (40 mmol/l Tris-HCl, pH 7.5, 150 mmol/l KCl, 1 mmol/l EDTA, 1% Triton X-100, 100 mmol/l NaVO₃, 1 mmol/l PMSF) supplemented with the protease inhibitor cocktail. The protein (50 μg) was separated on 10% SDS-PAGE and transferred onto PVDF membranes. The membranes were blocked with 5% non-fat milk in Tris-buffered saline (TBS) at 37°C, and then incubated with rabbit anti-MEK1 antibody (1:1,000), rabbit anti-p-MEK1 antibody (1:1,000), mouse anti-ERK1/2 antibody (1:1,000), rabbit anti-p-ERK1/2 antibody (1:1,000), rabbit anti-Bad antibody (1:1,000), rabbit anti-p-Bad antibody (1:1,000) or mouse anti-β-actin antibody (1:500) in TBS containing 5% non-fat milk for 12 h at 4°C. Horseradish peroxidase-linked anti-mouse IgG (1:5,000) or horseradish peroxidase-linked anti-rabbit IgG (1:5,000) was used as a secondary antibody (in TBS containing 5% non-fat milk for 3 h at room temperature), and antigen-antibody complexes were detected using an enhanced chemiluminescence kit (Amersham, ECL Plus, Freiburg, Germany). Densitometry values for western blot analysis and antibody array experiments were estimated by the ImageQuant TL software (GE Healthcare, Buckinghamshire, UK) and expressed as arbitrary units (a.u.). Multiple film exposures were used to verify the linearity of the samples analyzed and to avoid saturation of the film.

The expressions and intracellular localizations of MAPK/ERK in HCC and mice xenograft were examined immunohistochemically. Antigen retrieval was performed by microwave oven for 15 min in TEG buffer (10 mM Tris, 0.5 mM ethylene glycol tetraacetic acid, pH 9.0). Incubation with primary antibody for 60 min at room temperature was followed by detection of the primary antibody using the Advance^TM HRP system (Dako). The chromogen 3,3′-diaminobenzidine was applied and all the staining was performed using the Autostainer Plus Link Instrument (Dako). After washing, the slides were counterstained with Meyer’s hematoxylin for 30 sec. The following antibodies were used: p-MEK1/2 (dilution factor 1:100), pERK1/2 (dilution factor 1:100), PCNA (dilution factor 1:100). All antibodies mentioned above were from Cell Signaling Technology.

Xenograft tumors were resected and fixed in formalin for 24 h, and imbedded in paraffin and 5-micron of sections were prepared. TUNEL assay was performed using an apoptag peroxidase in situ apoptosis detection kit (Chemicon International, Temecula, CA, USA). Briefly, the sections were digested using proteinase K and the endogenous peroxidase activity was blocked using 3% hydrogen peroxide in PBS. The sections were then placed in equilibration buffer and incubated with working strength of TdT enzyme in a humidifying chamber at 37°C for 1 h. The reaction was terminated with a stop/wash buffer provided with the kit. The apoptotic nuclei were stained by direct immunoperoxidase detection of digioxigenin-labeled DNA in test sections.

Data are presented as the mean ± standard errors from at least three independent experiments and analyzed using Student’s t-test. p<0.05 was considered statistically significant. All statistical tests and corresponding p-values were two sided.

In order to investigate whether or not Baicalein has any differential cytotoxicity to HCC and normal liver cells as CIE does (5,6). We examined the cytotoxic activity of Baicalein in three HCC lines (HepG₂, BEL-7402 and SMMC-7721) and one normal liver cell line (HL-7702). The anti-tumor effects of Baicalein were examined by an MTT assay after treatment with 20 to 120 μM of Baicalein for 48 or 72 h. As shown in Fig. 2A, the viability of HepG₂, BEL-7402 and SMMC-7721 cells was significantly reduced by Baicalein treatment in a time- and dose-dependent manner, whereas the normal liver cell line (HL-7702) was hardly affected. To further examine the cytotoxicity of Baicalein in HepG₂ HCC cells and normal liver HL-7702 cells, 5-FU was used as a treatment benchmark in comparison with Baicalein. While 5-FU had a 50% inhibiting concentration (IC₅₀) of 1.05 mM in HepG₂ cells, Baicalein had an IC₅₀ value of 68.32 μM (Fig. 2B). As shown in Fig. 2B, the inhibitory effect of Baicalein on HL-7702 normal liver cells at its IC₅₀ concentration was significantly lower than that of 5-FU at its IC₅₀ concentration. Baicalein had an inhibition rate of 5.43%±1.00%, whereas 5-FU inhibited more than 35% on normal liver cells at its IC₅₀ concentration in normal liver cells (Fig. 2B).

To explore the mechanisms by which Baicalein inhibits HCC growth, HepG₂ HCC cells were first examined by phase contrast microscopy for any apoptotic characteristics after incubation with Baicalein at different concentrations (0, 40, 80, 120 μM) for 24 h. As shown in Fig. 2C, the control-treated cells showed a typical polygonal and intact appearance, whereas the Baicalein-treated cells displayed cellular shrinkage (40, 80 μM), rounding (120 μM), poor adherence (120 μM) and round floating shapes (120 μM). To determine the effect of Baicalein on apoptosis in detail, HepG₂ HCC cells were treated with different concentrations of Baicalein (40, 80 or 120 μM), for 24 h and then subjected to an Annexin V analysis on flow cytometry. As shown in Fig. 3A, Baicalein induced marked apoptosis in HepG₂ cells in a concentration-dependent manner. After short treatment for 24 h, the numbers of early apoptotic cells accompanied some late apoptotic cells were significantly increased in the Baicalein-treated HepG₂ cells when compared with control-treated HepG₂ cells (Fig. 3A). After longer treatment for 48 h, the numbers of late apoptotic and necrotic cells were also dramatically increased along with early apoptotic cells in the Baicalein-treated HepG₂ cells (Fig. 3A).

The decrease of mitochondrial transmembrane potential (MMPΔΨm) has been reported as an early event of apoptosis (16) and can be detected by the decline of rhodamine 123 fluorescence. To determine whether or not Baicalein-induced apoptosis involves the MMPΔΨm, we used a fluorescent indicator Rh123 to detect the MMPΔΨm in HepG₂ cells that were treated with 20–60 μM of Baicalein for 24 h. As shown in Fig. 3B, after exposure to different doses of Baicalein, the cells exhibited dose-dependent decline of Rho123 staining. At the doses of 40 and 60 μM, Baicalein-treated cells had significant lower values of rhodamine 123 fluorescence (3,258.11±355.90, 2,705.45±276.17) than the control (4,703.24±698.91, p<0.05), further indicating that Baicalein can induce apoptosis in liver cancer cells (Fig. 3B).

To further determine whether apoptosis induced by Baicalein was a mitochondrial-dependent pathway, we tested whether cytochrome c could be released from the mitochondria into the cytoplasm. As shown in Fig. 3C, levels of cytochrome c release from the mitochondria increased dose-dependently in the Baicalein-treated HepG₂ cells at concentrations ranging from 40 to 120 μM. To further investigate whether apoptosis induced by Baicalein was a caspase-dependent pathway, we tested whether mitochondrial-related caspases were activated by Baicalein treatment. Our research showed that caspase-9 and caspase-3 activities were highly increased dose-dependently on exposure to Baicalein in HepG₂ cells (Fig. 3E). Furthermore, we treated HepG₂ cells with Baicalein in the presence of 10 M pan-caspase inhibitor (z-VAD-fmk) or DMSO (as a control). MTT assay showed that z-VAD-fmk partially attenuated Baicalein-induced inhibition on HepG₂ cells (Fig. 3D), suggesting that apoptosis induction is an important cause for Baicalein-induced growth inhibition in HCC.

Western blot analysis has been utilized to evaluate the effect of Baicalein on phosphorylation levels of MEK and ERK in HepG₂ cells. As shown in Fig. 4A and 4C, Baicalein inhibited MEK1 and ERK1/2 phosphorylation at a concentration-dependent manner in HepG₂ cells. The phosphorylation level of Bad (Ser 112), which is an anti-apoptosis protein activated by the MEK/ERK pathway in tumor cells (17), was also measured 24 h after Baicalein treatment. Baicalein reduced levels of phosphorylated Bad of Ser 112 in a dose-dependent manner (Fig. 4A and 4C).

To determine whether this Baicalein-induced growth inhibition depends on the MEK-ERK pathway, HepG₂ cells were transfected with a plasmid pcDNA3.1(+)-MEK1 expressing human MEK1. Ectopic expression of MEK1 led to an enhanced activity of MEK-ERK pathway indicated by increased phosphorylation of MEK1 and ERK1/2 (18) (Fig. 4B). Importantly, HepG₂ cells with ectopic expression of MEK1 (higher MEK-ERK activity) became relatively resistant to Baicalein-induced growth inhibition (Fig. 4). Overexpression of MEK1 partially attenuated Baicalein-induced inhibition of ERK1/2 phosphorylation (Fig. 4C). Overexpression of MEK1, in part, blocked Baicalein-induced growth inhibition in vitro (Fig. 4D). These data suggest that inhibition of MEK-ERK is one of critical mechanism by which Baicalein inhibits HCC cells.

In the animal study, the control group received diluent vehicle treatment only, whereas the treatment group received Baicalein 20 mg/kg/day. This in vivo dosage was selected by our pilot experiments that showed significant tumor inhibition but without significant side effects. The mice were treated with Baicalein daily for 21 days. As shown in Fig. 5A, Baicalein-treated mice exhibited a statistically significant tumor volume reduction (p<0.01) compared with the control group. The average tumor volume of control and treatment group were 3.25±0.56 cm³ and 1.02±0.40 cm³, respectively. After treatment for 21 days, the mice were sacrificed, xenograft tumors were resected and the tumor weight of xenograft were measured. As shown in Fig. 5B and 5C, tumor sizes and weights in Baicalein-treated mice were dramatically smaller than those in control-treated mice. The control-treated mice had a median tumor weight of 2.12 g, whereas the Baicalein-treated mice had a median tumor weights of 0.41 g (Fig. 5B).

In order to confirm the in vitro observation of Baicalein-induced apoptosis (Fig. 3), Baicalein-induced apoptosis in xenograft tumors was evaluated with the terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay. As shown in Fig. 6A, Baicalein-treated tumors had greater TUNEL-postive cells than control-treated tumors. In agreement with the in vitro observations, p-MEK1/2 and p-ERK1/2 expression were markedly inhibited in Baicalein-treated tumors as illustrated by immunohistochemical analysis (Fig. 6A) and western blot analysis (Fig. 6B and 6C). MEK-ERK signaling associated Bad phosphorylation (Serine 112) was also decreased by Baicalein treatment (Fig. 6B and 6C). Above data confirm the in vitro results and show that Baicalein treatment can significantly suppress HCC tumor growth and MEK-ERK signaling, and can induce apoptosis in vivo.