HL-7702 cells were purchased from the Institute of Biochemistry and Cell Biology, China Academy of Science, Shanghai. Cordyceps polysaccharide, obtained from Shanghai University of Traditional Chinese Medicine, was dissolved in sterile distilled water. 3-(4,5-dimethylthiazol-2-yl) -2,5-diphenyltetrazolium bromide (MTT) was purchased from Sigma-Aldrich (St. Louis, MO, USA). Dimethlysulfoxide (DMSO) was obtained from Shenggong Biology (Shanghai, China). Mitochondrial membrane potential detection kit, ATP detection kit and Hoechst 33258 were purchased from Beyotime (Jiangsu, China). Dulbecco’s modified Eagle’s medium (DMEM) supplement was obtained from Gibco Invitrogen Co. (Gaithersburg, MD, USA). The fluorescent dye 2′,7′-dichlorodihydro-fluorescein diacetate (H2DCF-DA) was purchased from Invitrogen. Antibodies against Cyt C were from Cell Signaling Technology (Beverly, MA, USA) and Epitomics (Burlingame, CA, USA). Antibodies against Bcl-2 and Bax were purchased from Epitomics. All the other chemicals were of the highest grade of purity available commercially.

HL-7702 cells were routinely cultured in DMEM (Gibco) containing 10% fetal calf serum (FBS; Gibco), 100 U/ml penicillin and 100 mg/ml streptomycin and maintained in a humidified 5% CO₂ atmosphere at 37°C. The medium was changed every two days. Cells were incubated with 400 μM H₂O₂ for 2 h to induce cell apoptosis. In order to study the effects of CPS, cells were pre-incubated with various concentrations of CPS for 2 h, and then H₂O₂ was added to the medium for another 2 h.

The dried powder of cultured cordyceps mycelia was purchased from Traditional Chinese Medicine Limited Liability Company (Jiangxi, China) and was defatted with various concentrations of ethanol. The extract was boiled twelve times in water for 2 h and centrifuged; the supernatant was concentrated and treated with 15 times volume of ethanol for precipitation. The precipitate was suspended in water, dialyzed and lyophilized to yield the crude polysaccharide-enriched fraction. The extraction ratio of crude polysaccharide was 15% and the purity was above 99%. The crude polysaccharide verified by Fehling Reagent, mainly contained reductive sugars such as mannose, galactose glucose, cordycepin, adenosine, arabinose, xylose and fucose, and these monosaccharides composed the polysaccharide.

The stock solution of CPS was prepared by dissolving 10 mg CPS using 10 ml sterile distilled water and was stored at −70°C for future use.

Cell viability was measured by conventional MTT reduction assay. The cultured cells were seeded at an initial density of 5×10⁴ cells/ml in a 96-well plate for 24 h and pre-incubated with 400, 500 and 600 μM CPS for 2 h and exposed to 400 μM H₂O₂ for 2 h. Following incubation, 20 μl MTT stock solution (5 mg/ml) was added into each well at a final concentration of 0.5 mg/ml for another 4 h. The resulting formazan was dissolved in 150 μl DMSO and measured with a microtiter plate reader at a wavelength of 492 nm.

The intracellular ROS was detected by H2DCF-DA, an oxidation-sensitive fluorescent probe. Cells were pre-incubated with CPS for 2 h and exposed to 400 μM H₂O₂ for 2 h. The medium was removed; cells were washed twice with serum-free medium and incubated with H2DCF-DA (10 μM) for 20 min at 37°C in the dark. The fluorescence intensity was monitored on an automatic fluorescence microplate reader with an excitation wavelength of 488 nm and an emission wavelength of 525 nm and observed under a fluorescence microscope. Data were expressed as a percentage of the control.

Intracellular ATP levels were measured by a firefly-luciferase-based ATP detection kit (Beyotime) according to the manufacturer’s instructions. Briefly, cells were seeded into the 24-well plates and washed with phosphate-buffered saline (PBS) three times. The supernatant of samples was collected immediately on ice and measured with an illuminometer. ATP levels were calculated according to an ATP standard curve. Intracellular ATP levels were analyzed as the percentage of the control group.

The intracellular MMP was evaluated by use of the fluorescent, lipophilic and cationic probe, 5,5′,6,6′-tetrachloro-1,1′,3,3′-iodide (JC-1) (Beyotime) according to the manufacturer’s instructions. Briefly, following treatment, the cells were loaded with JC-1 staining solution for 20 min at 37°C and washed three times with JC-1 staining buffer. The fluorescence intensity was measured by a CytoFluor multiwell plate reader with 514 nm for excitation and 529 nm for emission of green (monomer form) fluorescence, and 585 nm for excitation and 590 nm for emission for red (aggregate form) fluorescence. The MMP of cells in each group was evaluated as the fluorescence ratio of red to green and observed by a fluorescence microscope. The data were expressed as the percentage of the control.

Cells were placed on cover slips in 24-well plates and pretreated with CPS for 2 h prior to exposure to 400 μM H₂O₂ for 2 h. After washing with PBS, the slides were fixed in 4% paraformaldehyde for 10 min at room temperature, washed three times with PBS, permeabilized with 0.1% saponin, blocked with 10% normal goat serum and incubated overnight at 4°C with anti-Cyt C. The slides were incubated with FITC-conjugated goat anti-rabbit immunoglobulin (Sigma-Aldrich) for 2 h. Nuclei were stained with Hoechst 33258 (Beyotime). Cover slips were observed under a fluorescence microscope (Leica Microsystems, Wetzlar, Germany) equipped with a Leica DFC420 camera.

Cells were lysed in SDS buffer supplemented with a mixture of protease inhibitors, 1 μg/ml aprotinin and 100 μg/ml phenylmethylsulfonyl fluorides. The cell suspension was incubated on ice for 30 min then centrifuged at 20,000 × g for 15 min at 4°C. The supernatant was collected for further analysis. The protein concentrations of the supernatants were determined using the Bradford method. Cell lysates were denatured for 15 min in 5X sample buffer and separated by SDS-polyacrylamide gel electrophoresis. For western blot analysis, the gel was transferred onto nitrocellulose membranes using a tank transfer system. Blotted membranes were placed in a blocking solution of 5% nonfat milk in Tris-buffered saline Tween-20 (TBS-T). For immunodetection, membranes were incubated for 1 h at room temperature and then incubated overnight at 4°C with the relevant primary antibodies, followed by washing with TBST and incubation with the appropriate horseradish peroxidase-conjugated secondary antibodies. Immunocomplexes were visualized using a commercially available enhanced chemiluminescence kit with exposure of the transfer membrane to X-ray film. The following antibodies were used: anti-Bcl-2, anti-Bax and anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH).

Data were expressed as the means ± SEM from at least three independent experiments. Statistical significance analysis was carried out using Student’s t-test or ANOVA. Mean values were considered to be statistically significant at P<0.05 or P<0.01.

Firstly, in order to investigate the effect of CPS on HL-7702 cells, an MTT assay was used to evaluate the cytotoxicity of cells treated with various concentrations of CPS. As shown in Fig. 1A, CPS at a concentration range from 300 to 800 μM had no significant cytotoxicity on HL-7702 cells and the cytotoxicity was observed when the concentration of CPS increased to 1000 μM. Then, to examine the protective effect of CPS on H₂O₂-induced cell apoptosis, cells were incubated with 400 μM H₂O₂ for 2 h with or without different concentrations of CPS. The findings showed that cell viability decreased to 43.3% after treatment with H₂O₂ for 2 h compared with the control group. However, pretreatment with CPS significantly enhanced cell viability from 43.3 to 58.5, 52.1 and 54.9%, respectively (Fig. 1B), while Vitamin E (VE) serving as a control showed a protection of 56.2%. In addition, the effects were also observed under Hoechst 33258 staining, which revealed contracted nucleus and condensed chromatin fragments. Following treatment with H₂O₂ for 2 h, the number of apoptotic cells increased compared to the control. However, pretreatment with CPS significantly decreased the number of apoptotic cells (Fig. 1C and D). These results suggested that CPS inhibits cell apoptosis induced by H₂O₂, which may be correlated with scavenging free radicals. Therefore, we further measured the eliminating capacity of CPS.

ROS generation is an important indicator of oxidative stress-induced mitochondrial dysfunction. In order to detect the capability of CPS scavenging free radicals, the H2DCF-DA assay was used to detect the generation of intracellular ROS induced by 400 μM H₂O₂ for 2 h. The fluorescence images showed that green fluorescence intensity markedly increased when HL-7702 cells were incubated with 400 μM H₂O₂ for 2 h. However, pretreatment with CPS reduced the green fluorescence intensity (Fig. 2A). Fig. 2B shows the same result: exposure of HL-7702 cells to H₂O₂ led to an increase of the intracellular ROS levels, which was approximately 1.39-fold relative to that of control cells. Pretreatment with CPS and VE inhibited the intracellular ROS level. These results suggested that CPS restrains H₂O₂-induced cell apoptosis by eliminating intracellular ROS, and mitochondrial membranes are key action sites of ROS. Therefore, we further detected the effect of CPS on mitochondrial membrane potential and energy synthesis.

Dissipation of mitochondrial integrity is one of the early events leading to apoptosis (19). To assess whether CPS affects the function of mitochondria, the changes of MMP were analyzed by employing mitochondria fluorescence dye, JC-1, which stains mitochondria in a membrane potential-dependent manner. As shown in Fig. 3A, cells exposed to 400 μM H₂O₂ for 2 h resulted in a significant decrease in aggregate and increase in monomer forms; however, pretreatment with CPS prevented the loss of aggregate and the increase of the monomer forms. In addition, the effects of CPS on H₂O₂-induced MMP disruption were also confirmed by an automatic fluorescence microplate reader. Exposure to H₂O₂ for 2 h gave rise to a decrease in the ratio of aggregate to monomer, and 400, 500 and 600 μM CPS prevented the decrease of the ratio between aggregate and monomer in a concentration-dependent manner. VE also increased the ratio between aggregate and monomer forms (Fig. 3B). These results implied that CPS attenuated H₂O₂-induced MMP dissipation.

In order to determine whether the dysfunction of mitochondrial energy generated occurred in H₂O₂-treated cells, we investigated the changes of intracellular ATP content in the H₂O₂-treated cells with and without various concentrations of CPS (400–600 μM). As shown in Fig. 4A, when HL-7702 cells were treated with 400 μM H₂O₂ for 2 h, ATP concentration markedly decreased to 0.465 μM (the concentration of control group was 4.537 μM). However, pretreatment with CPS (400, 500 and 600 μM) increased the ATP concentrations from 0.456 to 0.799, 1.085 and 1.722 μM, respectively, and VE inhibited the decrease of ATP levels induced by H₂O₂. Our data implied that CPS avoids the mitochondrial energy dysfunction induced by H₂O₂.

Cyt C release from the mitochondria to the cytosol is a critical event in the mitochondrial dysfunction induced by various types of cell stress. We therefore examined whether CPS has a significant role in regulating the release of Cyt C using immunofluorescence staining. As shown in Fig. 5, following treatment with 400 μM H₂O₂ for 2 h, diffuse cytoplasmic staining was detected, implying that Cyt C was released from the mitochondria to the cytosol. Moreover, pretreatment with 600 μM CPS prevented the Cyt C release from the mitochondria to the cytosol caused by H₂O₂. The data demonstrated that Cyt C is a key apoptotic factor in the mitochondrial-dependent apoptotic pathway, and CPS inhibits cell apoptosis by regulating mitochondrial apoptotic factors, such as Cyt C. Consequently, we further observed whether CPS was capable of accommodating other apoptosis-related factors.

Bcl-2 family membranes play critical roles in maintaining mitochondrial integrity and mitochondria-initiated Cyt C release. Previous studies have reported that the ratio of the pro-apoptotic protein Bax to the anti-apoptotic Bcl-2 was correlated with cell apoptosis (20). Our results showed that the protein level of Bax displayed little change and there was a prominent decrease in the protein expression of Bcl-2 after HL-7702 cell treatment with 400 μM H₂O₂, and there was an approximate 1.9-fold increase in the ratio of Bax/Bcl-2 in the H₂O₂ treatment group compared with the control group, as shown by western blot analysis. However, CPS inhibited the H₂O₂-induced increase of Bax/Bcl-2 ratio in a concentration-dependent manner (Fig. 6A–D). The effect of CPS on H₂O₂-induced apoptosis may be, at least in part, mediated by the regulation of Bax and Bcl-2 expression. CPS decreased the ratio of Bax/Bcl-2 and suggested that it may suppress the mitochondrial-dependent apoptosis induced by H₂O₂.