sylvestris (L.) Hoffm. Dried A. sylvestris roots were extracted with 10 volumes of methanol (v/w) and the sublayer organic phase was concentrated in a rotary vacuum evaporator (Eyela, Tokyo, Japan) and lyophilized. The residue was dissolved in dimethyl sulfoxide (DMSO), filtered, and analyzed using a Shimadzu HPLC system (Shimadzu Corporation, Kyoto, Japan) consisting of an LC-20AR pump, an SCL 10A system controller and an SPD-20A UV-VIS detector. Semi-preparative HPLC for purification of the methanol extract of dried A. sylvestris roots: preparative reversed-phase HPLC was performed using a Shimpack PRC-ODS column (250×20 mm I.D., 5 µm; Shimadzu). The mobile phase was a mixture of two liquids distributed by (A) water and (B) acetonitrile at a flow rate of 1.0 ml/min. The elution program commenced at 95% A: 5% B followed by a linear gradient for 60 min to 5% A: 95% B. The sample injection volume was 1 ml, and the detection by UV was set at a wavelength of 210 nm. Chemical identification was performed by comparing the retention times and mass spectra of the targeted peaks with those of the standard sample. Anthricin (standard sample; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA) was dissolved in DMSO at 10 nM, and the anthricin sample prepared from the methanol extract of dried A. sylvestris roots was dissolved in DMSO at 10 mg/ml.

All the antibodies were purchased from Cell Signaling Technology, Inc. (Beverly, MA, USA), except for β-actin (AB Frontier, Seoul, Korea). A live and dead assay kit and 4,6-dianmidino-2-phenylindole (DAPI) were purchased from Molecular Probes (Carlsbad, CA, USA) and Roche Diagnostics (Bromma, Sweden), respectively. 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) and propidium iodide (PI) were purchased form Sigma-Aldrich (St. Louis, MO, USA). RPMI-1640 medium and 100 U penicillin-streptomycin solution were purchased from WelGene (Deagu, Korea). Fetal bovine serum (FBS) was purchased from Corning Inc. (Corning, NY, USA). A PE-Annexin V apoptosis detection kit was purchased from BD Biosciences (Franklin Lakes, NJ, USA).

A549 human NSCLC cells were purchased from the Korean Cell Line Bank (Seoul, Korea). Cells were cultured in RPMI-1640 medium supplemented with 10% FBS, 100 U penicillin-streptomycin at 37°C in a 5% CO₂-humidified incubator.

The cytotoxicity of anthricin was assessed using the MTT assay. In brief, the cells were seeded into a 12-well plate (5×10⁵ cells/ml) and allowed to adhere overnight. The cells were then treated with either vehicle DMSO or anthricin (10, 20, 50, 100 and 200 nM) for 24 and 48 h. Following incubation, 100 µl of MTT solution (5 mg/ml) was added to each well and the plate was incubated for 4 h at 37°C. The resulting formazan crystals were dissolved in DMSO, and the optical density (OD) was assessed at 570 nm using a microplate reader (Bio-Tek Instruments, Winooski, VT, USA). The cell viability rate was calculated using the following equation: means of OD_treated/means of OD_control × 100%.

A549 cells were seeded in a 4-well chamber slide at a density of 1×10⁵ cells/well. After overnight incubation, the cells were treated with different concentrations of anthricin (0, 10, 20 and 50 nM) for 24 h. Following incubation, the treated cells were washed with phosphate-buffered saline (PBS) twice and fixed with 4% formaldehyde in PBS at room temperature for 10 min. Calcein-AM and ethidium bromide homodimer-1 were used to stain live and dead cells, respectively, according to the manufacturer's instructions. For DAPI staining, the treated cells were washed, fixed with 4% formaldehyde in PBS at room temperature for 10 min, and stained with 300 nM DAPI for 20 min at room temperature. The stained cells were washed thrice with PBS and observed under a fluorescence microscope (Eclipse TE2000; Nikon Instruments, Melville, NY, USA). Cells exhibiting condensed and fragmented nuclei were considered to have undergone apoptosis.

A549 cells were treated with anthricin at concentrations of 0, 10, 20 and 50 nM for 24 h, harvested, and washed with ice-cold PBS twice. For the apoptosis analysis, the cells were stained using a PE-Annexin V/7-AAD apoptosis detection kit (BD Biosciences) according to the manufacturer's protocol and subsequently analyzed by flow cytometry (BD FACSCalibur). The quantitative data was expressed as density plots using WDI software. Non-stained cells (Annexin V and 7-AAD negative) were considered viable, cells stained with Annexin V positive and 7-AAD negative were regarded as early apoptotic cells, and cells stained with both Annexin V and 7-AAD were considered late apoptotic or already dead cells. For cell cycle analysis, after cells were treated with anthricin for 24 h, as previously described, they were harvested and washed in ice-cold PBS. The cell pellets were fixed with 4% formaldehyde in PBS for 15 min and stained with 50 µg/ml propidium iodide and 100 µg/ml RNase for 20 min. The data were analyzed using Attune NxT Acoustic Focusing Cytometer (Thermo Fisher Scientific, Inc., Waltham, MA, USA).

A549 cells were treated with anthricin at concentrations of 0, 10, 20 and 50 nM for 24 h. The cells were lysed with protein extraction reagent (iNtRON Biotechnology, Sungnam, Korea) for 30 min on ice. The supernatant was transferred to a new tube after centrifugation at 13,000 × g for 15 min at 4°C (Sorvall Centrifuge, Bad Homburg, Germany). Then, the protein concentrations were quantified using the BCA protein assay (Pierce; Thermo Fisher Scientific, Inc., Waltham, MA, USA) with bovine serum albumin (BSA) as a standard. Equal amounts of protein (20 µg/lane) were separated by 8 or 15% SDS-PAGE gels under reducing conditions, followed by electrophoretic transfer onto polyvinylidene difluoride (PVDF) membranes (Bio-Rad Laboratories, Hercules, CA, USA). The membranes were blocked with 5% BSA for 1 h. The membranes were incubated at 4°C overnight with the following primary antibodies diluted to 1:1,000: Bax (cat. no. 2772), Bcl-2 (cat. no. 2872), cleaved caspase-8 (cat. no. 9496), cleaved caspase-3 (cat. no. 9661), cleaved caspase-9 (cat. no. 7237), PARP (cat. no. 9542), Cdc2 (cat. no. 28439), Cdc25C (cat. no. 4688), p-IGF1R(Y1131) (cat. no. 9750), IGF1R (cat. no. 3021), PI3K (cat. no. 4292), p-PI3K (cat. no. 4228), AKT (cat. no. 9272), p-AKT (cat. no. 9271), mTOR (cat. no. 2972), p-mTOR (cat. no. 2971) (all from Cell Signaling Technology, Beverly, MA, USA) and β-actin (1:5,000; cat. no. YIF-LF-PA0207A; AB Frontier, Seoul, Korea). After washing with Tris-buffered saline and Tween-20 (TBST) thrice, the membranes were incubated for 1 h at room temperature with HRP-labeled secondary antibodies, anti-rabbit IgG (cat. no. 7074) and anti-mouse IgG (cat. no. 7076) diluted to 1:5,000 from Cell Signaling Technology. The proteins were detected using Immobilon Western Chemiluminescent HRP Substrate (ECL; Millipore, Bedford, MA, USA) and visualized using a MicroChemi 4.2 device (DNR Bioimaging Systems, Jerusalem, Israel). The density of each band was quantified using ImageJ software and β-actin was used as the internal control.

Male BALB/c nude mice at four weeks of age were purchased from Damool Science (Daejeon, Korea). Animals were housed in microisolator ventilated cages with environmentally controlled temperature (21±1°C) and humidity (55±5%), and a reversed 12-h light-dark cycle. Water and food were autoclaved and provided ad libitum. All animal experiments were conducted in accordance with the National Institutes of Health (NIH) Care and Use of Laboratory Animals (21) and were approved by the Chosun University Institutional Animal Care and Use Committee (CIACUC2016-S0040). After one week of acclimatization to the laboratory environment, the mice were subcutaneously inoculated with 0.1 ml of PBS containing A549 cells (1×10⁷ cells/mouse) into the right ear. When the xenograft tumors were palpable and the tumor volume reached ~100 mm³, the mice were randomly assigned to the control and treatment groups (n=3). Mice received either 10% DMSO in normal saline (control) or anthricin (10 and 20 mg/kg body weight) by intraperitoneal injection every three days. The tumor volume and body weight of the animals were assessed every five days. The tumor volume (V) was calculated by measuring two perpendicular diameters (a, length; b, width) using an electronic digital caliper and the formula used according to the National Cancer Institute (22) was as follows: V (mm³) = (a × b²)/2. At the end of the treatment period, all mice were sacrificed under CO₂ and the transplanted tumors were excised and weighed.

All data were derived from at least three independent experiments (except the in vivo study). Results were expressed as the means ± standard deviation (SD). One-way ANOVA followed by Dunnett's t-test was employed for multiple comparisons using GraphPad Prism (GraphPad Software, Inc., La Jolla, CA, USA). Statistical significance was set at P<0.05.

sylvestris roots by HPLC. The chromatograms revealed the presence of anthricin in the methanol extract of A. sylvestris roots (Fig. 1). It was identified by comparing the retention time and UV spectra of the standard sample. The retention time of anthricin was 49.20 min. In Fig. 1B and C, the first peak was ignored as the solvent peak.

To assess the effect of anthricin on cell viability, the cells were treated with anthricin (0, 10, 20, 50, 100 and 200 nM) for the indicated time-points and were analyzed by MTT assay. As shown in Fig. 2, anthricin markedly increased the cytotoxicity of A549 cells in a dose- and time-dependent manner. Since anthricin exhibited cytotoxicity at 10 nM in 24 h, further experiments were performed at 10, 20 and 50 nM.

To determine whether cytotoxicity of anthricin was related to apoptosis, we investigated induction of apoptosis of A549 cells by anthricin using a live and dead assay, DAPI staining, FACS analysis, and western blotting. As shown in Fig. 3A, cells stained with ethidium bromide homodimer-1 (dead dye, red color) gradually increased with anthricin treatment dose-dependently. In addition, anthricin-treated cells exhibited a significant increase in the apoptotic cell population compared with the control. Cells stained brightly by nuclear condensation were considered as apoptotic cells. Based on the apoptosis phenomenon of anthricin observed in the live and dead assay and DAPI staining (Fig. 3A and B, respectively), we performed FACS analysis for the quantification of apoptotic cells. A549 cells were treated with anthricin as previously indicated and then double-stained with PE-Annexin V/7-AAD. The percentage of Annexin V-positive apoptotic cells increased to 13.52, 14.7 and 23.94% at 10, 20 and 50 nM anthricin, respectively, compared with the control (0.09%) (Fig. 3C). Western blotting was performed to evaluate whether A549 cell apoptosis induced by anthricin was dependent on caspases. As shown in Fig. 3E, the cleavage of caspase-8, −3 and −9 significantly increased in a dose-dependent manner and thus the cleavage of native PARP (116 kDa) into its small fragment PARP (89 kDa) increased. Furthermore, anthricin-mediated apoptosis was associated with the outer mitochondrial membrane in a dose-dependent manner. Anti-apoptotic Bcl-2 protein levels decreased but anti-survival Bax protein levels increased (Fig. 3D). Collectively, these results revealed that anthricin-induced apoptosis of A549 cells may be mediated by the activation of caspase in the extrinsic death receptor and intrinsic mitochondrial-dependent apoptotic signaling pathways.

Flow cytometric analysis revealed that the percentage of cells in the G2/M phase increased in anthricin-treated cells compared with the control (Fig. 4A and B). This peak markedly increased at 20 and 50 nM anthricin (~81.05 and 68.69%, respectively). In addition, the results for the detected sub-G1 group indicated that anthricin induced apoptosis of A549 cells. Western blot analysis was performed to examine the expression of G2/M-boundary regulatory proteins, including Cdc2 and Cdc25C. As shown in Fig. 4C, the protein expression of Cdc2 and Cdc25C significantly decreased 24 h after anthricin treatment in a dose-dependent manner, indicating that anthricin induces G2/M phase arrest in A549 cells.

IGF1R is upregulated in various cancers, including breast, prostate, and lung cancers, and mediates cell cycle progression and prevention of apoptosis in hematopoietic cells (23). Thus, we examined the effects of anthricin on the IGF1R/PI3K/Akt signaling pathways in A549 cells. After 24 h of exposure to anthricin (0, 10, 20 and 50 nM), the protein expression levels of p-IGF1R (Y1131) were reduced to 65, 22 and 3%, at 10, 20 and 50 nM anthricin, respectively (Fig. 5). In addition, anthricin treatment reduced the protein expression levels of p-PI3K (P85) and p-Akt (Ser473), which IGF1R targets downstream in a dose-dependent manner (Fig. 5). As mTOR is a downstream target of the Akt gene, the expression of p-mTOR (Ser2448) was reduced due to the inhibition of Akt (Fig. 5). These results revealed that anthricin induced apoptosis by inhibiting the IGF1R/PI3K/Akt signaling pathways.

Based on the aforementioned in vitro anticancer effect, we investigated the inhibitory effects of anthricin on A549 xenografts. Mice were injected (i.p.) with 10 and 20 mg/kg of anthricin every three days for 30 days. The tumors removed from these animals are shown in Fig. 6C, and their size and weight have been provided in Fig. 6C and D, respectively. The tumor growth of A549 xenografts were significantly inhibited by anthricin treatment, and the inhibitory rates were 37.34 and 52.17%, at 10 and 20 mg/kg anthricin, respectively, compared with that in the vehicle-treated animals (Fig. 6B). In addition, the tumor mean weights were 883.33±41.55, 606.66±18.04 and 496.67±23.84 mg in the control and after treatment with 10 and 20 mg/kg anthricin, respectively; the inhibition rates were 32 and 44% (Fig. 6D). There was no significant difference in body weight between the groups (Fig. 6A) and no sign of skin rash or diarrhea. All animals appeared to be in a decent state despite treatment. These results revealed that anthricin exhibits potential as a novel antitumor therapeutic agent against lung cancer.