CT (Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) was dissolved (20 mM CT in 100% dimethyl sulfoxide) to obtain a stock solution and stored at −20°C. The stock solution was diluted with cell culture media prior to further use.

Informed consent was obtained from RA patients prior to inclusion in the present study, and the procedure was approved by the Ethical Committee of the Fifth Affiliated Hospital of Harbin Medical University (Daqing, China). Synovial tissue samples were obtained from RA patients (including 3 females and 3 males aged 53-66 years) undergoing synovectomy or joint replacement in the Fifth Affiliated Hospital of Harbin Medical University. The synovial tissues were minced into small pieces and digested with 2 mg/ml collagenase type I at 37°C for 4 h. Subsequent to the enzymatic dispersion of synovial tissues, synovial cells were pelleted by centrifugation (1,200 × g for 3 min at room temperature) and cultured in Dulbecco’s modified Eagle’s medium (DMEM; Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) at 37°C in a humidified atmosphere containing 5% CO₂. After 24 h of incubation, the adherent cells were collected and cultured in DMEM, and are referred to as the primary RA-FLS cells.

The HFLS-RA (cat. no. CBR130803), MH7A (cat. no. CBR130742) and normal synovial HFLS (cat. no. CBR130948) cells were purchased from Saiqi Biological Engineering Co., Ltd. (Shanghai, China). Cells were cultured in DMEM supplemented with 10% fetal bovine serum (Gibco; Thermo Fisher Scientific, Inc.), 100 µg/ml penicillin and 100 U/ml streptomycin in a humidified 5% CO₂ incubator at 37°C.

Primary RA-FLS, HFLS-RA, MH7A and HFLS cells were seeded on 96-well plates (1×10⁴ cells/well) and treated with different concentrations of CT (1, 3, 10, 30 and 100 µmol/l) for 24 h. Prior to proliferation detection, 10 µl CCK-8 reagent (Beyotime Institute of Biotechnology, Shanghai, China) was added into each well and incubated at 37°C for 1.5 h. Finally, the absorbance was measured by a microplate reader (A₄₉₀) and the cell survival rates were calculated.

A total of 5×10⁵ HFLS-RA and MH7A cells/well were seeded in 6-well plates and cultured overnight. Next, the cells were treated with 5 µmol/l CT for different time durations (0, 3, 6, 12 and 24 h). According to the manufacturer’s manual that was provided with the Apoptosis Detection kit (Beyotime Institute of Biotechnology), cells were stained with 10 µl Annexin V-FITC and 5 µl propidium iodide (PI) for 15 min, and the fluorescence intensity of each sample was determined by fluorescence microscopy (Leica Microsystem, Wetzlar, Germany).

A total of 5×10⁵ HFLS-RA were seeded in 6-well plates and cultured overnight. HFLS-RA cells were pretreated with N-acetyl cysteine (NAC, 5 mmol/l, Sigma-Aldrich; Merck KGaA) for 30 min and incubated with CT (5 µmol/l) for 24 h. Then, cells were stained with 10 µl Annexin V-FITC and 5 µl PI for 15 min, and the number of apoptotic cells were detected by flow cytometry (Beckman Coulter, Inc., Brea, CA, USA).

In order to measure the ROS levels, HFLS-RA cells were treated with CT (5 µmol/l) for 0, 3, 6, 12 and 24 h, following which the cells were incubated with 2′,7′-dichlorofluorescein-diacetate (DCFH-DA; Merck, Shanghai, China) for 30 min at 37°C. Next, cells were washed 2-3 times with PBS and harvested by centrifugation at 1,200 × g for 5 min at room temperature. The intracellular ROS levels were subsequently analyzed by flow cytometry.

Cells were lysed in ice-cold radioimmunoprecipitation buffer (50 mmol/l Tris (pH 7.4), 150 mmol/l NaCl, 1% Triton X-100, 1% sodium deoxycholate, 0.1% SDS, 20 mg/ml AEBSF, 0.5 mg/ml pepstatin, 0.5 mg/ml leupeptin and 2 mg/ml aprotinin; (Beyotime Institute of Biotechnology) at 4°C for 30 min by shaking every 5 min and then separated by centrifugation at 4°C and 12,000 × g for 30 min. The protein concentrations were determined using Bradford reagent (Bio-Rad Laboratories, Inc., Hercules, CA, USA). Next, 5X buffer was added to the total supernatants (30 µg), and proteins were resolved by 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Subsequently, the proteins were transferred to nitrocellulose (NC) membranes, which were blocked in 5% skim milk for 2 h at room temperature. The NC membranes were then incubated overnight at 4°C with the following primary antibodies (all purchased from Santa Cruz Biotechnology, Inc., Dallas, TX, USA): Mouse monoclonal antibodies against α-tubulin (1:2,500; cat. no. sc-8035), B-cell lymphoma 2 (Bcl-2; 1:1,500; cat. no. sc-7382), Bcl-2-associated death promoter (Bad; 1:1,500; cat. no. sc-8044), cleaved caspase-3 (1:1,500; cat. no. sc-373730), cleaved poly (ADP-ribose) polymerase (PARP; 1:1,500; cat. no. sc-8007), phosphorylated ERK [p-ERK (Tyr 204); 1:1,500; cat. no. sc-8059], p-JNK (Thr 183 and Tyr 185, 1:1,500; cat. no. sc-6254), JNK (1:1,500; cat. no. sc-7345), p-p38 (1:1,500; cat. no. sc-7973), p-STAT3 (Tyr 705, 1:1,500; cat. no. sc-8059) and STAT3 (1:1,500; cat. no. sc-8019); and rabbit polyclonal antibodies against ERK2 (1:1,500; cat. no. sc-154), p38α/β (1:1,500; cat. no. sc-7972), Akt1/2/3 (1:1,500; cat. no. sc-8312) and p-Akt1/2/3 (Ser 473, 1:2,500; cat. no. sc-7985-R). Following washing with Tris-buffered saline/Tween 20 (TBST), the NC membranes were incubated with horseradish peroxidase (HRP)-labeled goat anti-rabbit IgG (1:5,000; cat. no. ZB-2301) and goat anti-rat IgG (1:5,000; cat. no. ZB-2305) were used as secondary antibodies for 2 h at room temperature. After the membranes were washed again with TBST, an enhanced chemiluminescence reagent (Thermo Fisher Scientific, Inc.) was used to detect the relative intensities of proteins using Amersham imager 600 (AI600; GE Healthcare, Chicago, IL, USA). The band intensities were analyzed by ImageJ software (National Institutes of Health, Bethesda, MD, USA), and the relative intensity of protein expression levels was calculated by SPSS version 18.0 statistical software (SPSS, Inc., Chicago, IL, USA).

Data are presented as the mean ± standard deviation, and all the experiments were replicated three times. Continuous data were analyzed by one-way analysis of variance followed by Tukey’s post hoc tests using SPSS version 18.0 statistical software. P<0.05 was considered to indicate a difference that was statistically significant.

To determine the effects of CT on RA-FLS survival, primary RA-FLS, HFLS-RA, MH7A and normal HFLS cells were treated with different concentrations of CT (1, 3, 10, 30 and 100 µmol/l) for 24 h, and the cell viabilities were evaluated by the CCK-8 assay. As shown in Fig. 1A-C, CT significantly decreased the viabilities of primary RA-FLS, HFLS-RA and MH7A cells in a dose-dependent manner. The half maximal inhibitory concentration (IC₅₀) values of primary RA-FLS, HFLS-RA and MH7A cells were 6.69±0.97, 2.56±0.42 and 7.10±0.94 µmol/l, respectively. Furthermore, CT exhibited low cytotoxic effects on HFLS cells, which had an IC₅₀ value >100 µmol/l; however, significance was observed compared with the control (Fig. 1D).

To determine whether CT induced apoptosis in RA-FLSs, HFLS-RA and MH7A cells were treated with 5 µmol/l CT for different time points (0, 3, 6, 12 and 24 h), and apoptosis was examined by Annexin V-FITC/PI double staining and flow cytometry. As shown in Fig. 2A-D, the fluorescence intensity significantly increased with the increase in the duration of CT treatment. In addition, apoptosis gradually increased in HFLS-RA and MH7A cells in a time-dependent manner (Fig. 2E-H). These results suggest that CT may be a potential inducer of apop-tosis in RA-FLSs.

CT-induced apoptosis was more notable in HFLS-RA cells compared with in MH7A cells; thus, HFLS-RA cells were used as the representative RA-FLS cells in subsequent experiments. As shown in Fig. 3A and B, CT treatment markedly downregulated the expression of Bcl-2, while it upregulated the protein expression levels of Bad, cleaved caspase-3 and cleaved PARP. These results revealed that CT induced apoptosis through the mitochondrial pathway.

To determine whether CT induced apoptosis by regulating the Akt, MAPK and STAT3 signaling pathways, HFLS-RA cells were treated with 5 µmol/l CT for different time points (0, 3, 6, 12 and 24 h), and protein expression levels were detected by western blotting. As shown in Fig. 4A and B, CT exposure increased the protein expression levels of p-p38 and p-JNK, and downregulated the expression levels of p-Akt, p-ERK and p-STAT3, in a time-dependent manner. These results revealed that CT induced apoptosis in HFLS-RA cells via the MAPK, Akt and STAT3 signaling pathways.

To determine whether CT induced intracellular ROS generation, the HFLS-RA cells were treated with 5 µmol/l CT for different durations (0, 3, 6, 12 and 24 h), after which the levels of ROS were detected by flow cytom-etry. As shown in Fig. 5A and B, CT treatment significantly increased ROS generation in a time-dependent manner.

To further investigate whether CT-induced apoptosis was mediated by intracellular ROS, the cells were divided into four groups, as follows: Control, CT, NAC and CT+NAC. As shown in Fig. 5C and D, CT-induced apoptosis was significantly suppressed after scavenging of intracellular ROS by treatment with NAC. Changes in the expression levels of apoptosis-associated proteins were detected by western blotting, which demonstrated that the expression levels of Bcl-2, Bad, cleaved caspase-3 and cleaved PARP were recovered following intracellular ROS scavenging (Fig. 5E and F). These results indicated that the increased levels of ROS were involved in CT-induced apoptosis.

To investigate whether CT induced apoptosis through regulating ROS-mediated Akt, MAPK and STAT3 signaling pathways, HFLS-RA cells were cultured overnight, treated with CT or NAC for 24 h, and divided into four groups as follows: Control, CT, NAC and CT+NAC. Next, the protein expression levels were detected by western blotting. As shown in Fig. 6A and B, as compared with the CT treatment group, upregulation of p-Akt, p-ERK and p-STAT3 levels, and downregulation of p-p38 and p-JNK were detected in the CT+NAC treatment group. These data indicated that CT induced apoptosis through ROS-mediated MAPK, Akt and STAT3 signaling pathways.