The human T-cell ALL (T-ALL) cell lines CCRF-CEM and Jurkat (American Type Culture Collection) were maintained in RPMI 1640 medium (Gibco; Thermo Fisher Scientific, Inc.) supplemented with 10% fetal bovine serum (HyClone; Cytiva), 1 mM sodium pyruvate (HyClone; Cytiva), 100 U/ml penicillin, and 100 µg/ml streptomycin (Gibco; Thermo Fisher Scientific, Inc.) in a humidified atmosphere containing 5% CO₂ at 37°C.

Stattic (cat. no. HY-13818; MedChemExpress), a selective inhibitor of STAT3, and dimethyl sulfoxide (DMSO), which was used as a vehicle control, were procured from Sigma-Aldrich; Merck KGaA. For cell treatments, Stattic was dissolved in DMSO for stock preparation. In the in vitro experiments, the vehicle control group received a final DMSO concentration of 0.05%, while the 5, 2.5 and 1.25 µM Stattic groups received DMSO concentrations of 0.05, 0.025 and 0.0125%, respectively.

The cytotoxic effects of Stattic on CCRF-CEM and Jurkat cells were assessed using the Cell Counting Kit-8 (CCK-8; cat. no. 96992; Sigma-Aldrich; Merck KGaA) according to the manufacturer's instructions. Briefly, a total of 1×10⁵ cells/well were seeded in 200 µl culture medium in a 96-well cell culture plate, and were then treated at 37°C for 24 h with various concentrations of Stattic (0.625, 1.25, 2.5, 5 and 10 µM) or an equivalent volume of the vehicle control. Subsequently, CCK-8 solution was added, and plates were incubated for another 2–3 h. Absorbance at 450 nm was then measured using a microplate reader (Enspire 2300–0000; PerkinElmer, Inc.).

For western blot analysis, cells were treated prior to collection. In the dose-dependent experiment, cells were treated with DMSO or different concentrations of Stattic (1.25, 2.5 and 5 µM) for 24 h. In the time-course experiment, cells were treated with DMSO or 5 µM Stattic for 8, 16 and 24 h. After treatment, the cells were first washed with PBS prior to collection, and were then lysed in ice-cold Tris buffer (50 mM, pH 7.5) containing the following: 5 mM EDTA, 300 mM NaCl, 0.1% Igepal, 0.5 mM NaF, 0.5 mM Na₃VO₄, 0.5 mM PMSF and antiprotease mixture (Roche Molecular Diagnostics), and centrifuged at 13,000 × g for 10 min at 4°C. Protein concentrations were determined according to the Bradford procedure. Equal amounts of protein (25 µg) were then separated by SDS-PAGE on 10 and 12% gels, and were transferred to PVDF membranes. After blocking with 5% non-fat milk dissolved in PBS at room temperature for 1 h, the membranes were incubated with primary antibodies against p-STAT3 (1:1,000; cat. no. 9145S; Cell Signaling Technology, Inc.), STAT3 (1:1,000; cat. no. 30835S; Cell Signaling Technology, Inc.), pro-caspase-3 (1:1,000; cat. no. 9662S; Cell Signaling Technology, Inc.), cleaved caspase-3 (1:1,000; cat. no. 9662S; Cell Signaling Technology, Inc.), LC3B (1:1,000; cat. no. 83506S; Cell Signaling Technology, Inc.), p62 (1:1,000; cat. no. 5114; Cell Signaling Technology, Inc.), Bcl-2 (1:1,000; cat. no. 26593-1-AP; Proteintech Group, Inc.), PARP-1 (1:1,000; cat. no. sc-74470; Santa Cruz Biotechnology, Inc.), ATG5 (1:1,000; cat. no. sc-133158; Santa Cruz Biotechnology, Inc.), BECN1 (1:500; cat. no. sc-11427; Santa Cruz Biotechnology, Inc.) and β-actin (1:5,000; cat. no. 3700S; Cell Signaling Technology, Inc.), followed by incubation with an antimouse IgG, HRP-linked secondary antibody (1:7,000; cat. no. 7076P2; Cell Signaling Technology, Inc.) or an anti-rabbit IgG, HRP-linked secondary antibody (1:7,000; cat. no. 7074P2; Cell Signaling Technology, Inc.). The ATG5 antibody used in the present study can detect the ATG5-ATG12 conjugate with a molecular weight of ~50 kDa. Bands were visualized using an Enhanced Chemiluminescence Detection Kit (MilliporeSigma) and the Alliance Q9 (Uvitec Ltd.). The protein expression levels were normalized to β-actin. The band intensity was measured using ImageJ software (version 1.50i; National Institutes of Health).

Apoptosis was quantified based on Annexin V-FITC and propidium iodide (PI) double staining. Briefly, 4×10⁵ CCRF-CEM cells and 3×10⁵ Jurkat cells were treated with 5 µM Stattic or DMSO in a 24-well plate for 24 h. After treatment, the cells were collected and stained using the Annexin V-FITC/PI apoptosis kit according to the manufacturer's protocol (Elabscience Bionovation Inc.). Stained cells were analyzed by flow cytometry within 1 h of staining. Fluorescence intensities were measured using a flow cytometer (FACSCanto II; BD Biosciences) with BD FACSDiva software (version 8.0.1; BD Biosciences). Each experiment was performed in duplicate, at least three times independently.

Female NOD/SCID mice (age, 9 weeks; weight, 18–22 g) were procured from the National Laboratory Animal Center (Taipei, Taiwan). The mice were maintained in an individually ventilated caging system, adhering to a 12-h light/dark cycle, at a temperature of 22 ± 2°C and a humidity of 50–70%, with ad libitum access to food and water. All experimental protocols were approved by the Animal Care and Use Committee of the Taichung Veterans General Hospital (IACUC no. La-1132052; Taichung, Taiwan). To create tumors, 1×10⁶ CCRF-CEM cells suspended in a 1:1 mixture of BD Matrigel™ (cat. no. 356231; Corning, Inc.) and RPMI-1640 medium were subcutaneously injected into the flanks of each mouse. On day 1, T-ALL cells were subcutaneously injected into mice under gas anesthesia using isoflurane (induction, 4–5% isoflurane for 3 min; maintenance, 1–3% isoflurane for an additional 1 min). After the procedure, the mice were placed in a warm, dry environment and were continuously monitored for vital signs during recovery. The mice were returned to their original housing only after fully recovering from anesthesia. Post-surgery, the mice were examined at least once daily, monitoring the injection site for wound condition, including any signs of secretion, as well as assessing body weight, eating, urination and defecation. Analgesics were not used, as they could impact the experimental results; instead, physical pain management strategies, such as environmental enrichment items (e.g., toys), were provided. Mice received wooden sticks, paper houses and similar enrichment items. Mice were subsequently randomized into the following five groups (n=6 mice/group): Vehicle control group and four treatment groups, each receiving three different doses of Stattic (7.5, 15 and 30 mg/kg) or DEX (1 mg/kg; Taiwan Biotech Co., Ltd.). Starting from day 6 after tumor cell injection, Stattic and DEX were administered by intraperitoneal injection three times a week; the volume of intraperitoneal injection per mouse was 300 µl. Stattic was dissolved in DMSO for stock preparation. The final concentrations of DMSO within the administered Stattic doses (7.5, 15, and 30 mg/kg) were 0.75, 1.5 and 3%, respectively. For the vehicle control group, the injection consisted of normal saline containing 3% DMSO. Tumor sizes were measured using a digital caliper thrice weekly until the day of sacrifice and tumor volume was calculated using the following formula: Tumor volume=length × width². All mice were euthanized on day 23 for subsequent analyses. Mice were placed into a euthanasia chamber, which was gradually filled with CO₂ at a flow rate of 30% volume/min. After the gas infusion, the mice were observed for 3 min to ensure proper euthanasia; the signs confirming death included the cessation of heartbeat, lack of respiratory activity and pupil dilation.

Data are presented as the mean ± SEM of at least three independent experiments. To compare differences between the treatment and control groups, statistical significance was assessed using one-way ANOVA followed by Tukey's multiple comparisons test. Data were analyzed using GraphPad Prism software (version 6; Dotmatics). P<0.05 was considered to indicate a statistically significant difference.

To assess the cytotoxic effects of Stattic on T-ALL cells, CCRF-CEM (Fig. 1A) and Jurkat cells (Fig. 1B) were treated with increasing concentrations of Stattic (0.625, 1.25, 2.5, 5 and 10 µM) or vehicle control (DMSO) for 24 h. Cell viability was measured using the CCK-8 assay. In CCRF-CEM cells, Stattic treatment resulted in a significant, dose-dependent reduction in cell viability. A statistically significant reduction was observed at 1.25 µM, and cell viability was further decreased at 2.5 µM and higher concentrations. The half maximal inhibitory concentration (IC₅₀) value for CCRF-CEM cells was determined to be 3.188 µM, indicating that these cells were sensitive to Stattic-induced inhibition of viability (Fig. 1A). In Jurkat cells, a similar dose-dependent reduction in viability was observed; however, the inhibitory effect was less pronounced compared with in CCRF-CEM cells. Significant reductions in viability were detected at 5 and 10 µM concentrations. The IC₅₀ value for Jurkat cells was 4.89 µM, suggesting that Jurkat cells are slightly more resistant to Stattic than CCRF-CEM cells (Fig. 1B). These results indicated that Stattic may effectively reduce the viability of CCRF-CEM and Jurkat cells in a dose-dependent manner, with CCRF-CEM cells showing greater sensitivity. The observed differential sensitivity between the two cell lines highlights the potential for Stattic as a targeted therapeutic agent in T-ALL.

To investigate the effect of Stattic on STAT3 signaling, the expression levels of p-STAT3 were we examined in CCRF-CEM (Fig. 2A) and Jurkat cells (Fig. 2B) at 8, 16 and 24 h following treatment with 5 µM Stattic or DMSO. Total STAT3 and β-actin were used as loading controls. In CCRF-CEM cells, Stattic treatment resulted in a reduction in p-STAT3 levels over time. Notably, a significant decrease in p-STAT3 was observed at 24 h, indicating that Stattic effectively suppressed STAT3 phosphorylation with prolonged exposure. However, total STAT3 levels remained stable across all time points, suggesting that Stattic may specifically inhibit STAT3 activation without affecting its overall expression (Fig. 2A). In Jurkat cells, although p-STAT3 levels fluctuated, no statistically significant differences were observed at any of the time points compared with the DMSO-treated controls. Total STAT3 expression remained constant, similar to in CCRF-CEM cells (Fig. 2B). These findings indicated that Stattic may inhibit STAT3 activation more effectively in CCRF-CEM cells than in Jurkat cells, reflecting a differential response between the two T-ALL cell lines. The suppression of STAT3 phosphorylation suggested that Stattic may exert its anti-proliferative effects, at least in part, through the inhibition of STAT3-mediated signaling in CCRF-CEM cells.

Western blot analysis was conducted to investigate the time-dependent effects of Stattic (5 µM) on apoptosis and autophagy markers in two T-ALL cell lines: CCRF-CEM (Fig. 3A) and Jurkat (Fig. 3B). Both cell types were treated for 8, 16 and 24 h with Stattic or with DMSO as a control. In CCRF-CEM cells, an inhibition of pro-caspase-3 expression was observed at 8 and 24 h after Stattic treatment. Furthermore, a significant increase was detected in cleaved caspase-3 expression 16 h after Stattic treatment, indicating the activated apoptotic cascade. Stattic also induced a time-dependent upregulation of LC3B expression at 8, 16 and 24 h, suggesting enhanced autophagic activity. Meanwhile, p62 protein levels showed a decreasing trend at 24 h, although this change was not statistically significant, further supporting autophagy activation. Bcl-2 expression remained relatively stable, and full-length PARP-1 displayed a significant reduction at 8 and 16 h, reflecting apoptotic progression. Markers related to autophagy initiation, such as ATG5-ATG12 conjugate and BECN1, did not exhibit substantial changes during the observation period (Fig. 3A). In Jurkat cells, the apoptotic response to Stattic was less pronounced. While pro-caspase-3 levels remained relatively stable, cleaved caspase-3 exhibited a slight increase at 16 h. LC3B levels also demonstrated a significant increase at 8,16 and 24 h. Similarly, p62 levels showed a decreasing trend at 24 h, indicating autophagy activation, though less prominent compared with in CCRF-CEM cells. Bcl-2 expression was significantly reduced at 8 h but remained unchanged thereafter. Full-length PARP-1 and autophagy-related proteins, including ATG5-ATG12 conjugate and BECN1, showed no significant changes across all time points (Fig. 3B). Together, these results indicated that Stattic could induce both apoptotic and autophagic processes in CCRF-CEM and Jurkat cells, with more robust effects observed in CCRF-CEM cells. The differential responses between the two cell lines highlight the potential variability in the sensitivity of T-ALL subtypes to Stattic treatment.

To further explore the impact of Stattic on STAT3 signaling, CCRF-CEM (Fig. 4A) and Jurkat cells (Fig. 4B) were treated with increasing concentrations of Stattic (1.25, 2.5 and 5 µM) or vehicle control (DMSO) for 24 h. The expression levels of p-STAT3 and total STAT3 were measured by western blotting, with β-actin serving as the loading control. In CCRF-CEM cells, Stattic reduced p-STAT3 levels in a dose-dependent manner. A marked reduction in p-STAT3 was observed at the 5 µM concentration, with statistical significance indicated. However, total STAT3 protein expression remained unchanged across all treatment groups, suggesting that Stattic specifically inhibited STAT3 phosphorylation without affecting total STAT3 levels (Fig. 4A). Similarly, in Jurkat cells, p-STAT3 levels were reduced with increasing Stattic concentrations. A significant decrease was evident in response to the 5 µM concentration, while total STAT3 levels showed no major changes, indicating selective inhibition of phosphorylation by Stattic (Fig. 4B). These results indicated that Stattic may effectively inhibit STAT3 activation in a dose-dependent manner in both CCRF-CEM and Jurkat cells. The suppression of p-STAT3 without affecting total STAT3 levels further supports the role of Stattic as a selective inhibitor of STAT3 signaling, which could underlie its therapeutic potential in T-ALL.

To investigate the effects of Stattic on apoptosis and autophagy, CCRF-CEM (Fig. 5A) and Jurkat cells (Fig. 5B) were treated with increasing concentrations of Stattic (1.25, 2.5 and 5 µM) or vehicle control (DMSO) for 24 h. Protein expression levels of apoptotic markers (pro-caspase-3, cleaved caspase-3) and autophagy markers (LC3B, p62) were assessed by western blotting, with β-actin used as a loading control. In CCRF-CEM cells, cleaved caspase-3 levels showed an increasing trend with higher concentrations of Stattic, indicating enhanced apoptotic activity, although the changes were not statistically significant. LC3B levels were significantly increased at 5 µM concentrations, suggesting induction of autophagy. By contrast, the expression levels of p62, a marker of autophagic flux, remained relatively unchanged across all treatment groups, indicating incomplete autophagic flux. Pro-caspase-3 levels remained stable, further supporting that apoptosis was primarily indicated by the cleaved form (Fig. 5A). In Jurkat cells, a similar trend was observed. Cleaved caspase-3 levels increased slightly with higher Stattic concentrations, but the changes were not statistically significant. LC3B expression was significantly increased in response to 5 µM Stattic, indicating the activation of autophagic processes. However, as in CCRF-CEM cells, p62 levels did not show a significant reduction, suggesting a potential blockade in autophagic flux. Pro-caspase-3 levels also remained constant across the different Stattic concentrations (Fig. 5B). These findings demonstrated that Stattic significantly increased autophagy markers in CCRF-CEM and Jurkat cells, while also showing a trend toward increased apoptosis in both cell lines. The differential expression patterns of LC3B and p62 suggested that Stattic may trigger autophagy but not complete autophagic degradation. The increased levels of cleaved caspase-3 suggest a potential role of Stattic in promoting apoptosis in these T-ALL cell lines, although the changes were not statistically significant.

To further confirm the pro-apoptotic effects of Stattic, flow cytometry was performed to analyze apoptosis in CCRF-CEM (Fig. 6A) and Jurkat cells (Fig. 6B) after treatment with 5 µM Stattic for 24 h. Cells were stained with Annexin V-FITC and PI to differentiate between early and late apoptotic cells, which were detected in quadrants 3 and 2, respectively. The forward scatter (FSC)/side scatter (SSC) plots in Fig. 6A and B show the FSC and SSC characteristics of the cells, which provide information about cell size and granularity, respectively. From the results of the 5 µM Stattic treatment, the FSC/SSC plots in CCRF-CEM (Fig. 6A) and Jurkat (Fig. 6B) cells showed an increase in the proportion of cells with reduced size, indicating cell shrinkage typically associated with apoptosis. These results were obtained after Annexin V/PI staining, highlighting the effects of Stattic on inducing apoptotic changes in both cell lines. In CCRF-CEM cells, Stattic treatment significantly increased both early and late apoptotic populations compared with the control and DMSO groups. The percentage of early apoptotic cells increased significantly upon treatment with 5 µM Stattic. Moreover, late apoptotic cells showed a significant increase in the Stattic-treated group, indicating that Stattic strongly induced the apoptosis of CCRF-CEM cells (Fig. 6A). Similarly, in Jurkat cells, Stattic treatment led to a significant increase in apoptosis. Early apoptotic cells were significantly elevated, and late apoptotic cells increased substantially following treatment with 5 µM Stattic compared with in the control and DMSO-treated groups (Fig. 6B). These results indicated that Stattic effectively promoted both early and late apoptosis in CCRF-CEM and Jurkat cells, with a particularly strong effect on late apoptosis. This supports the role of Stattic as a potent inducer of apoptosis in T-ALL cells, highlighting its therapeutic potential for T-ALL treatment.

Using a xenograft mouse model of T-ALL, CCRF-CEM-xenografted mice were intraperitoneally injected with Stattic three times a week. The results revealed that while the control group showed a progressive growth in tumor volume over a 22-day period, the Stattic groups (15 and 30 mg/kg) and the DEX group (1 mg/kg; positive control), showed a significant reduction in tumor growth; the antitumor effect of Stattic was dose-dependent, with a peak effect observed at 30 mg/kg (Fig. 7A). Excised tumor volumes measured at the end of treatment (day 23) confirmed such a dose-dependent reduction in tumor size (Fig. 7B), and the significant decrease in volume relative to the control group in response to all doses of Stattic (Fig. 7C). In addition, a significant reduction in tumor weight was detected in the 15 and 30 mg/kg Stattic, and 1 mg/kg DEX treatment groups relative to the control (Fig. 7D). These results suggested that Stattic, along with DEX, effectively reduced tumor burden in a dose-dependent manner, with 30 mg/kg Stattic being the most effective dose.