The fruiting body of Omphalia lapidescens was provided by Fang Hui Chun Tang (Hangzhou, China). A total of 200 mg of the fruiting body of Omphalia lapidescens was washed three times with distilled water at 4°C and placed into 0.875 ml of cold PVP extraction buffer (15% 1.0 M Tris-HCl, pH 8.0; 2% PVP; 25% glycerol) on ice for 4 h. The samples were then centrifuged (12,000 × g, 20 min) at 4°C, and the supernatant was collected and retained for purification.

A SephadexG-50 column (GE Healthcare Life Sciences, Little Chalfont, UK), pre-equilibrated with 50 mM Tris-HCl buffer (pH 8.5), was employed to purify the sample (1 ml sample for each experiment). The absorbance was measured at 280 nm, and the flow rate was 0.2 ml/min, producing three peaks. In accordance with a previously described protocol (15), the second A_{280 nm} peak fraction was ultrafiltrated using a Millipore ultrafiltration tube (EDM Millipore, Billerica, MA, USA). Finally, the protein was sterilized by filtration using a 0.22 µm filter, and then stored at −20°C for later use.

The human gastric cancer cell line SGC-7901 was provided from Zhejiang Provincial Center for Disease Control and Prevention (Zhejiang, China). SGC-7901 cells were cultured in RPMI-1640 medium (cat no. GNM31800; Hangzhou Genom Biomedical Technology Co., Ltd., Hangzhou, China; http://www.genom.com.cn/) supplemented with 5% (v/v) fetal bovine serum (cat no. 22011-8612; Zhejiang Tianhang Biotechnology Co., Ltd., Zhejiang, China), 100 U/ml penicillin and 100 µg/ml streptomycin at 37°C, 5% CO₂.

Cultured SGC-7901 cells were detached with 0.25% trypsin during the logarithmic growth phase. The suspension of SGC-7901 cells containing 1×10⁶/ml cells was seeded into a 96-well plate and incubated at 37°C with 5% CO₂ for 24 h in preparation for an MTS assay. Subsequently, pPeOp (30, 60 and 90 µg/ml), 100 µg/ml5-FU (cat no. 51-21-8; Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) or PVP (90 µg/ml) diluted with RPMI-1640, which had been sterilized through a 0.22 µm filter, were added to the cells and incubated for 24 h at 37°C. A CellTiter 96^®AQueous One Solution Assay kit (cat no. G3580; Promega Corporation, Madison, WI, USA) was used for the MTS assay, according to the manufacture protocol. According to the manufacturer's protocol, 20 µl MTS solution (Promega Corporation) was added to each well, briefly agitated for 30 sec and incubated at 37°C for 60 min. Absorbance at 490 nm was measured using an automatic microwell plate reader (Thermo Labsystems, Helsinki, Finland). The light microscope used to visualize cells was the Nikon eclipse ti (Nikon Corporation, Tokyo, Japan) at a magnification of ×100.

Cellular migration was investigated using a wound-healing assay. SGC-7901 cells (1×10⁶/ml) were seeded into 6-well plates and cultured under standard aforementioned conditions for ~24 h. When cells were ~80% confluent, a wound was scratched into the cell layer with a 200 µl pipette tip. Cells were then treated with 30, 60 or 90 µg/ml pPeOp, 90 µg/ml PVP or 100 µg/ml 5-FU; for 24 h at 37°C. Cells were then washed with pre-warmed PBS (37°C) to remove cell debris and allowed to migrate for 24 h prior to image capture.

SGC-7901 cells were located in 6-well plates and incubated under standard aforementioned conditions for 24 h in preparation for a cell cycle assay. Then, cells were treated with pPeOp (30, 60 or 90 µg/ml), PVP (100 µg/ml) or 5-FU (100 µg/ml) under standard aforementioned conditions for 24 h. Cultured cells were harvested with EDTA-free trypsin (0.25%) and washed twice with ice-cold PBS. Subsequently, cells (1×10⁶ /ml) were incubated with 0.5 ml propidium iodide (PI)/RNase staining buffer (cat no. 550825; BD Biosciences, San Jose, CA, USA) for 15 min at room temperature (25°C) in the dark prior to being fixed with 1% paraformaldehyde (Beijing Solarbio Science & Technology Co., Ltd., Beijing, China) at 25°C for 30 min, and stored in 70% ethanol at −20°C for 15 min. Cell cycle progression analyses were performed using a Beckman FC500 (Beckman Coulter, Inc., Brea, CA, USA) flow cytometer with red fluorescent light at a wavelength of 488 nm. Win Cycle software (version 32; Beckman Coulter, Inc.) were used for analyses.

FITC Annexin V Apoptosis Detection kit I (cat no. 556547; BD Biosciences) was used to assay cell apoptosis. Cultured cells, subsequent to treatment with pPeOp (30, 60 or 90 µg/ml), PVP (100 µg/ml) or 5-FU (100 µg/ml) under standard aforementioned conditions for 24 h, were collected with EDTA-free trypsin (0.25%) and washed twice with ice-cold PBS. Subsequently, cells (1×10⁶/ml) were resuspended in 100 µl 1Xbinding buffer solution (BD Biosciences), to which 5 µl of fluorescein isothiocyanate/Annexin V (BD Biosciences) and 5 µl PI (BD Biosciences) were added. Cells were then gently vortexed and incubated for 15 min at room temperature (25°C) in the dark. Incubated cells were washed again with 1X binding buffer and resuspended in 200 µl of 1X binding buffer. PI (5 µl) and 400 µl of 1X binding buffer were added to each sample, and samples were analyzed using a Beckman FC500 flow cytometer (Beckman Coulter, Inc.). The software used to analyze apoptosis was CXP analysis (version 32; Beckman Coulter, Inc.).

MMP2, MMP9, cyclin D1, cyclin A2, cyclin B1, CDK1, CDK2, CDK4, Bcl-2, p53, caspase-3, JAK1, JAK2 and STAT3) were analyzed by RT-qPCR. In brief, cells (1×10⁶ /ml), subsequent to treatment with pPeOp (30, 60 or 90 µg/ml), PVP (100 µg/ml) or 5-FU (100 µg/ml) under standard aforementioned conditions for 24 h, were trypsinized and total RNA was isolated using TRIzol reagent (Invitrogen; Thermo Fisher Scientific, Inc., Waltham, MA, USA). cDNA was synthesized using a Maxima First Strand cDNA Synthesis kit (Thermo Fisher Scientific, Inc.), according to the manufacturer's protocol, which was used as a template for qPCR. Taq^®PCR Master Mix (cat no. A6001; Promega Corporation) for qPCR, and performed using a Step One Plus Real-Time PCR System (Applied Biosystems; Thermo Fisher Scientific, Inc.), according to the manufacturer's protocol. Data were normalized to the expression level of a GAPDH reference gene. Fold-changes in the amplified cDNA were calculated by the (2^−ΔΔCq) method (16). Primer sequences used are presented in Table I. All primers were purchased from Sangon Biotech Co., Ltd. (Shanghai, China). RT-qPCR was performed using an Eppendorf Realplex-4 Real Time PCR Machine (Eppendorf, Hamburg, Germany). The PCR amplification conditions were as follows: One cycle at 95°C pre-denaturing for 2 min, followed by 40 cycles at 95°C denaturing for 15 sec, and annealing temperature of 62.9°C (GAPDH, JAK1, JAK2, STAT3, caspase-3, p53, Bcl-2), 57.8°C (MMP2, MMP9, cyclin A2, cyclin B1, cyclin D1) or 61°C (CDK1, CDK2, CDK4) for 1 min. At the end of the PCR cycle, a dissociation curve was performed with StepOne™ Software (version 2.2.2; Applied Biosystems; Thermo Fisher Scientific, Inc.) to confirm amplification of a single product. The results are represented as the fold-change in gene expression relative to GAPDH.

MMP2, MMP9, cyclin D1, cyclin A2, cyclin B1, CDK1, CDK2, CDK4, Bcl-2, p53, caspase-3, JAK1, JAK2 and STAT3 were analyzed by western blot analysis. Cells (1×10⁶), subsequent to treatment with pPeOp (30, 60 or 90 µg/ml), PVP (100 µg/ml) or 5-FU (100 µg/ml) under standard aforementioned conditions for 24 h, were collected following treatment with PBS (1 ml, 30 min); centrifuged for 10 min at 200 × g and the supernatant was discarded. The pellet was then incubated with 200 µl lysis buffer (cat no. C0201; Beyotime Institute of Biotechnology, Shanghai, China) for 30 min on ice, and agitated every 10 sec. Lysates were separated using centrifugation at 4°C for 15 min at 10,000 × g, the supernatant was collected, and protein determined by BCA Protein Assay kit (cat no. KGP902; Nanjing KeyGen Biotech Co., Ltd., Nanjing, China). Protein (40 µg per lane) was loaded onto a 12% standard polyacrylamide gel and resolved by SDS-PAGE. Resolved proteins were subsequently transferred onto an Immobilon^®-P Transfer Membrane (cat no. IPVH00010; Merck KGaA), saturated with 5% milk in Tris-buffered saline and 0.5% Tween-20 (TBST) at 25°C for 2 h. Antibodies against JAK1 (cat no. ab138005; 1:1,000 dilution), JAK2 (cat no. ab32101; 1:1,000 dilutions) and STAT3 (cat no. ab76315; 1:200,000 dilutions) were purchased from Abcam (Cambridge, UK). Antibodies against MMP2 (cat no. 13132; 1:1,000 dilutions), MMP9 (cat no. 3852; 1:1,000 dilutions), cyclin A2 (cat no. 4656; 1:2,000 dilutions), cyclin B1 (cat no. 4138; 1:1,000 dilutions), cyclin D1 (cat no. 2978; 1:1,000 dilutions), CDK1 (cat no. ab32384; 1:1,000 dilutions), CDK2 (cat no. 2546; 1:1,000 dilutions), CDK4 (cat no. 12790; 1:1,000 dilutions), Bcl-2 (cat no. 2870; 1:1,000 dilutions), p53 (cat no. 2527; 1:1,000 dilutions) and caspase-3 (cat no. 14220; 1:1,000 dilutions) were purchased from Cell Signaling Technology, Inc. (Danvers, MA, USA). Anti-GAPDH (cat no. YM3029; 1:5,000 dilutions), used as a control, was purchased from Immuno Way Biotechnology Company (TX, USA). Horseradish peroxidase (HRP)-conjugated goat anti-rabbit immunoglobulin G (cat no. A0208; 1:1,000 dilutions) or goat anti-mouse immunoglobulin G (cat no. A0216;1: 1,000 dilutions) were used as secondary antibodies. All secondary antibodies were purchased from the Beyotime Institute of Biotechnology. The membranes were incubated overnight with primary antibodies (described above) at 4°C. Membranes were washed three times with TBST. Membranes that were incubated with GAPDH antibodies were then incubated with goat anti-mouse IgG secondary antibody (described above), while membranes which were incubated with MMP2, MMP9, cyclin D1, cyclin A2, cyclin B1, CDK1, CDK2, CDK4, Bcl-2, p53, caspase-3, JAK1, JAK2 and STAT3 antibodies were then incubated with goat anti-rabbit IgG HRP secondary antibodies (described above) for 2 h at 4°C, and washed three times with TBST. Membranes were visualized on film in a dark room using ECL substrate solution (cat no. P0018A; Beyotime Institute of Biotechnology) and results quantified using ImageJ software (version 1.8.0; National Institutes of Health, Bethesda, MA, USA).

All experiments were conducted in triplicate. Data are presented as the mean ± standard deviation. SPSS software (version 20.0; IBM Corp., Armonk, NY, USA) was used for statistical analyses. The MTS cell viability data were subjected to one-way analysis of variance (ANOVA), followed by Fisher's least significant difference (LSD) post-hoc test. Flow cytometry-based analyses were also analyzed using one-way ANOVA, followed by Fisher's LSD. Western blot results were quantitated using Image J software and analyzed by one-way ANOVA, followed by Fisher's LSD. P<0.05 was considered to indicate a statistically significant difference.

Initially, the effect of pPeOp on the proliferation of SGC-7901 gastric cancer cells was investigated. Cells treated with PVP (90 µg/ml, 24 h) maintained a normal morphology comparable to that of untreated cells (Fig. 1). However, when SGC-7901 cells were treated for 24 h with graded concentrations of pPeOp (30, 60 and 90 µg/ml), cells decreased in size and presented a spherical appearance (Fig. 1A). In addition to the morphological changes, the viability was also affected. The survival rates of cells after 24 h of treatment were as follows: PVP (100 µg/ml), 95.26±5.21%; 5-FU (100 µg/ml), 60.13±3.13%; pPeOp (30 µg/ml), 70.97±6.18%; pPeOp (60 µg/ml), 58.44±5.74%; pPeOp (90 µg/ml) 35.22±2.33%. The half maximal inhibitory concentration was 64.326 µg/ml. An increase in the concentration of pPeOp resulted in a proportional and significant increase in the rate of inhibition (P<0.05; Fig. 1B). These results suggested that pPeOp inhibits the viability of SGC-7901 cells.

A wound healing assay was used to assess the migration ability of SGC-7901 cells following treatment with pPeOp. As presented in Fig. 2A, compared with the normal and negative normal treatments, the migration ability of SGC-7901 cells was markedly decreased following pPeOp treatment.

Invasion and metastasis of carcinomas are closely associated with extracellular matrix (ECM) degradation (17,18). MMPs are well characterized and demonstrate proteolytic activity in the ECM, and have recently emerged as key molecules involved in mediating tumor invasion and metastasis (19). Therefore, MMP2 and MMP9 were selected for further studies. MMP2 and MMP9 levels were evaluated by RT-qPCR and western blot analysis. The results demonstrated that MMP2 and MMP9 were significantly downregulated at the mRNA and protein levels compared with the normal control, in proportion to the pPeOp dosage (P<0.05; Fig. 2B and C). However, compared with the normal group, cells treated with PVP did not show any significant difference in gene or protein expression profiles (P>0.05; Fig. 2B and C). Thus, the results suggest that pPeOp inhibits the ability of SGC-7901 cells to migrate.

The influence pPeOp has on the progression of SGC-7901 cells through the cell cycle was investigated using flow cytometry of cells post-treatment with either normal/5-FU/PVP or graded concentrations of pPeOp (Fig. 3A). The percentage of cells in G2/M phases decreased, whereas the proportion of cells in G0/G1, S phases increased in the pPeOp-treated group compared with the control (P<0.05, Fig. 3A). This suggests that pPeOp blocks DNA replication, thereby arresting cells in the S phase.

To verify the data obtained by flow cytometry, how pPeOp treatment influenced the expression of several cell cycle regulators, including cyclin A2, cyclin B1, cyclin D1, CDK1, CDK2, and CDK4 was investigated using RT-qPCR and western blot analysis. The results indicated that protein and mRNA expression levels of all six genes were significantly decreased post-treatment with pPeOp compared with the normal control (P<0.05; Fig. 3B and C). Furthermore, no significant differences were observed between untreated cells and cells treated with PVP (P>0.05; Fig. 3B and C). Collectively, these results suggest that pPeOp arrests cell cycle progression of SGC-7901 cells.

Flow cytometry was used to investigate how treatment with normal/5-FU/PVP or varying dosages of pPeOp affects the proportions of SGC-7901 cells undergoing apoptosis (Fig. 4A). The results demonstrated that pPeOp treatment induced SGC-7901 cancer cells to undergo apoptosis in a dose-dependent manner. Cells treated with 5-FU (100 µg/ml for 24 h) demonstrated a high rate of apoptosis (21%; Fig. 4A). In addition, cells treated with pPeOp (90 µg/ml for 24 h) experienced a markedly increased rate of apoptosis (35.1%; Fig. 4A), surpassing that of 5-FU treatment.

Subsequently, the molecular events underlying this modulation in apoptosis were investigated by determining the expression of apoptosis-associated factors, namely caspase-3, p53 and Bcl-2, at the mRNA and protein levels. The results indicated that pPeOp significantly upregulated the expression of caspase-3 and p53, and markedly downregulated the expression of Bcl-2 (P<0.05; Fig. 4B and C). There was no significant difference in the expression of these regulators between normal and PVP-treated cells (P>0.05; Fig. 4B and C). These results collectively demonstrate that pPeOp is effective for inducing SGC-7901 cells to apoptosis, superseding even a conventional chemotherapeutic agent.

The aforementioned experiments highlight the effects that pPeOp exhibits on a variety of important biological programs, including cell proliferation, migration, apoptosis and cell cycle progression. However, the mechanism underlying these effects remains poorly understood. According to the results of a microarray assay, numerous differentially expressed genes were detected (data not shown). Following Kyoto Encyclopedia of Genes and Genomes analysis, the JAK-STAT signaling pathway was the most notable regulated signaling pathway influenced by pPeOp proteins (data not shown). Therefore, it was hypothesized that pPeOp may exert its anti-tumor effects through its inhibition of the JAK-STAT signaling pathway in SGC-7901 human gastric cells.

In order to investigate this hypothesis, SGC-7901 cells were treated with different concentrations of pPeOp for 24 h and the expression of JAK1, JAK2 and STAT3 mRNA and proteins were explored using RT-qPCR and western blotting. The results demonstrated that the expression of JAK1, JAK2 and STAT3 mRNAs and proteins were significantly decreased post-treatment with 60 and 90 µg/ml pPeOp, compared with the control (P<0.05; Fig. 5A and B). These results provide evidence for the anti-tumor effects of pPeOp being attributed to the inhibition of JAK-STAT signaling.