In this study, pharmaceutical-grade PTE with a purity of at least 98% was obtained from Shanghai Yuanye Bio-Technology Co., Ltd (Shanghai, China). PTE was dissolved in dimethyl sulfoxide (DMSO) and stored at -20 °C until further use. H-151, a STING inhibitor, was purchased from Sigma-Aldrich (St. Louis, MO, USA) and also dissolved in DMSO and stored at -80 °C. Primary antibodies against various proteins, including glyceraldehyde-3-phosphate dehydrogenase (GAPDH), poly ADP ribose polymerase (PARP), Phospho-STING (Ser366), Phospho-TBK1/NAK (Ser172) (D52C2), TBK1 (E8I3G), Phospho-IRF-3 (Ser386), and STING (#11904), were obtained from Cell Signaling Technology (CST; Danvers, MA, USA). Fluorescein-conjugated secondary antibodies (anti-rabbit and anti-mouse) were purchased from Odyssey (Lincoln, NE, USA). Annexin V/PI staining dye was acquired from BD Biosciences (San Jose, CA, USA). All other reagents and cell culture mediums were obtained from Thermo Fisher Scientific (Waltham, MA, USA), unless stated otherwise.

The human NSCLC cell lines A549 and H358 cell lines were obtained from the American Type Culture Collection (ATCC, Virginia, USA). Both cell lines were cultured in RPMI-1640 medium manufactured by Gibco (Gibco, Grand Island, NY, USA). The medium was supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, and 100 μg/mL streptomycin (Sigma-Aldrich, St. Louis, MO, USA). All cell lines were maintained at 37 °C in a humidified atmosphere containing 5% CO₂.

Cell viability was assessed using the cell counting kit-8 (CCK8; Dojindo, Kumamoto, Japan) method, as described previously (25). A549 and H358 cells were seeded in 96-well plates at 3000 cells/well and treated with different concentrations of PTE (1.25, 2.5, 5, 10, and 20 µM) for 24, 48, and 72 hours. After incubation, 10 µL of CCK-8 reagent was added to each well and incubated for 4 hours. Absorbance was measured at 450 nm using a microplate reader (BioTek, Winooski, VT, USA). All experiments were performed with at least three independent replicates (n=3).

Apoptosis was evaluated using an Annexin V-FITC/propidium iodide (PI) Apoptosis Detection Kit (BD Biosciences, San Jose, CA, USA) (26). Cells were treated with PTE, N-acetylcysteine (NAC), PTE, H-151 or corresponding vehicles for 24 hours, harvested, and stained according to the manufacturer’s protocol. Flow cytometry was performed using a BD FACSAria™ III cell sorter (BD Biosciences), and data were analyzed with FlowJo software.

Cells were harvested, washed with PBS, and fixed in 70% ice-cold ethanol at -20 °C for at least 4 hours. Fixed cells were pelleted, treated with RNase A (100 µg/mL) at 37 °C for 30 minutes to digest RNA, and then stained with propidium iodide (PI, 50 µg/mL) in the dark at room temperature for 15 minutes before analysis (27, 28). Cell cycle distribution was analyzed using a BD FACSAria III flow cytometer. was analyzed using a BD FACSAria III flow cytometer and data were analyzed with Flowjo software.

Cell invasion was assessed using 8 µm pore-size Transwell chambers (Corning, Corning, NY, USA). Briefly, 1 x 10⁴ cells in 200 µL serum-free medium were seeded into the upper chamber, with or without PTE (0-30 µM) or cisplatin (6.67 µM). The lower chamber contained 600 µL complete medium. After 24 hours, non-invading cells were removed with a cotton swab. Invaded cells on the lower surface were fixed with paraformaldehyde for 20 minutes, stained with 0.1% crystal violet (Solarbio, Beijing, China) for 15 minutes at room temperature, and then washed. Membranes were imaged under an inverted microscope (IX73; Olympus, Tokyo, Japan) at 100x magnification and five random fields per membrane were counted.

The protein extraction process followed an established protocol (29). Briefly, total protein was extracted from cells using RIPA lysis buffer (200 µL per well of a 6-well plate) supplemented with protease and phosphatase inhibitors. Protein concentration was determined using a BCA assay kit (Beyotime, China). Equal amounts of protein (30 µg) were separated by 8-12% SDS-PAGE and transferred to nitrocellulose membranes (Millipore, Billerica, MA, USA). After blocking with 5% non-fat milk in TBST for 1 hour at room temperature, membranes were incubated with primary antibodies (diluted 1:1000 in 5% BSA-TBST) overnight at 4 °C. The following primary antibodies were used: GAPDH, PARP, Phospho-STING (Ser366), Phospho-TBK1/NAK (Ser172), TBK1, Phospho-IRF-3 (Ser386), STING, and γ-H2AX (all from Cell Signaling Technology). After washing, membranes were incubated with IRDye-labeled secondary antibodies (LI-COR Biosciences, Lincoln, NE, USA; 1:15000 dilution) for 1 hour at room temperature. Protein bands were visualized using an Odyssey Infrared Imaging System (LI-COR).

Intracellular ROS levels were detected using the fluorescent probe DCFH-DA (Beyotime, Shanghai, China). After treatment with PTE for 24 hours, cells were incubated with 10 µM DCFH-DA for 30 minutes at 37 °C. Fluorescence intensity was measured by flow cytometry (30).

Cells were seeded into 6-well plates at 200 cells per well and treated with PTE (0-40 µM) or cisplatin (6.67 µM) for 24 hours. After 14 days, colonies were fixed and stained with 0.1% crystal violet. Visible colonies were counted manually (31).

Total RNA was extracted using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) (24). cDNA was synthesized using a PrimeScript RT reagent kit (Takara, Dalian, China). qPCR was performed with SYBR Green Premix (Takara) on a QuantStudio 5 Real-Time PCR System (Applied Biosystems, Foster City, CA, USA). The primers used for the qPCR were consistent with those reported in prior literature to ensure the reliability and comparability of the results (32).

Cells were seeded in 6-well plates and grown to 60-70% confluence. Transfected was performed using Lipofectamine 3000 reagent (Invitrogen) according to the manufacturer’s instructions. For each well, 2 ug of control shRNA or STING-specific shRNA using lentiviral vectors (Santa Cruz Biotechnology, Dallas, TX, USA) was mixed with the transfection reagent and added to the cells. The medium was replaced with fresh complete medium 6 hours post-transfection. Knockdown efficiency was verified by Western blot and qPCR after 72 hours.

All animal procedures were approved by the Animal Ethics Committee of Zhuhai Hospital of Integrated Traditional Chinese & Western Medicine (Approval No. EC-C-008-A04-V1.0) and were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (33). This study is reported in accordance with the ARRIVE guidelines. Eight-week-old male BALB/c nude mice were subcutaneously inoculated with 5×10⁵ H358 cells suspended in serum-free media into the right flank (31). When tumor volumes reached 50 to 100 mm³, mice were randomly divided into experimental groups, with five mice in each group. For 21 days, mice received daily intraperitoneal injections of either a carrier solution consisting of 30% polyethylene glycol 400 (PEG400) and 5% Tween 80 in PBS or PTE at a concentration of 10 mg/kg, also dissolved in the same carrier solution. Throughout the treatment period, the overall health of the animals was monitored, including weight fluctuations and any signs of toxicity, such as >20% body weight loss, lethargy, hunched posture, ruffled fur, or reduced mobility, which were not observed in our study.

Tumor growth was measured every three days using calipers, and tumor volume was calculated using the formula: volume = length (mm) × width (mm)² × π/6 (34). At the end of the treatment period, mice were euthanized via exposure to carbon dioxide (CO₂) in a sealed chamber at a flow rate of 20% chamber volume per minute (until respiratory arrest was confirmed). The tumors were then surgically excised from the mice for further analysis. These procedures ensured effective monitoring and evaluation of tumor growth and the impact of PTE treatment on the xenografts, all in accordance with animal welfare guidelines.

The effect of PTE on the anti-tumor immune response was evaluated in an 8-week-old male C57BL/6 mouse model transplanted with LLC1 cells, which establishes a fully immunized lung cancer scenario (35). Mice received subcutaneous injections of LLC1 cells, and upon tumors reaching sizes of 50 to 100 mm³, they were randomly assigned to either the solvent (control) group or the PTE treatment group for a duration of 21 days. Mice were euthanized when the tumor diameter reached 15 mm (predefined humane endpoint) via exposure to carbon dioxide (CO₂) in a sealed chamber at a flow rate of 20% chamber volume per minute (until respiratory arrest was confirmed). After euthanasia, both tumor weight and volume were measured.

Single-cell suspensions were prepared from the tumors. The following anti-mouse antibodies were used: PerCP anti-mouse CD45 Antibody (clone 30-F11), APC anti-mouse CD3 antibody (clone 17A2), PE-cy7 anti-mouse CD8a antibody (clone 53-6.7), FITC anti-mouse CD4 Antibody (clone RM4-5), PE anti-mouse FOXP3 antibody (clone MF-14), Gr-1 (clone RB6-8C5), APC/Cyanine7 anti-mouse/human CD11b Antibody (clone M1/70), PE/Cyanine7 anti-mouse F4/80 Antibody (clone BM8) from BioLegend or eBioscience. For cytokine detection, cells were stimulated with Cell Stimulation Cocktail (plus protein transport inhibitors, eBioscience) for 4 hours, followed by staining for PE/Dazzle™ 594 anti-mouse TNF-α antibody (clone MP6-XT22), APC/Cy7 anti-mouse IFN-γ antibody (clone XMG1.2), and Granzyme B (clone NGZB). The levels of TNF-α, GrzmB, and IFN-γ in tumor-infiltrating CD8⁺ T cells were quantified by measuring the Mean Fluorescence Intensity (MFI) and the percentage of positive cells using flow cytometry (BD FACSAria™ III) and analyzed with FlowJo software. The study specifically focused on the impact of PTE on tumor-infiltrating CD8⁺ T cells, assessing the levels of TNF-α⁺, granzyme B⁺, and IFN-γ⁺ within these cells. Tumor and spleen tissues were homogenized, single cells were isolated, and flow cytometry analysis was performed following treatment with anti-CD16/32 antibodies and staining with specific antibodies per the manufacturer’s instructions. The resulting cell populations were analyzed via flow cytometry.

Additionally, we evaluated the composition of immune cells in the TME post-PTE treatment, including Tregs, DCs, macrophages, and infiltrating CD8⁺ T cells. The levels of TNF-α, granzyme B (GrzmB), and IFN-γ in tumor-infiltrating CD8⁺ T cells were quantified to further understand the immune responses induced by PTE.

The data presented in the study are expressed as the mean ± standard deviation (SD) from three independent experiments. Statistical differences between groups were evaluated using one-way analysis of variance (ANOVA), followed by Bonferroni’s test for pairwise comparisons. Results were deemed statistically significant at the following thresholds: *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001, indicating the level of significance for the observed differences among the experimental groups.

The chemical structure of pterostilbene (PTE) is shown in Figure 1A . We evaluated its effects on cell viability using CCK-8 assays in A549 and H358 NSCLC cells treated with various concentrations of PTE (0-40 µM) for 24, 48, and 72 hours. PTE significantly inhibited cell viability in both cell lines in a concentration- and time-dependent manner ( Figures 1B, C ).

The calculated half-maximal inhibitory concentration (IC₅₀) values indicated differing sensitivities to PTE between the two cell lines. Specifically, the IC₅₀ values were 31.79 µM for A549 and 68.3 µM for H358 at the 24-hour mark, 7.46 µM and 29.3 µM at 48 hours, and 4.66 µM and 6.67 µM at 72 hours, respectively. Based on these findings and to guide future experiments, we selected concentrations of 10 µM, 20 µM, 30 µM, and 40 µM for a 24-hours treatment period as the experimental conditions for further investigations.

The colony formation assay is a crucial method for assessing the long-term growth potential of cancer cells (36). Colony formation assays revealed that PTE treatment for 14 days significantly reduced the number of colonies in A549 and H358 cells in a concentration-dependent manner, with efficacy comparable to cisplatin (6.67 µM; Figures 2A, B ).

Transwell invasion assays further demonstrated that PTE suppressed cell invasion in a dose-dependent fashion ( Figure 2C ). These findings indicated that PTE restrained both proliferative capacity and invasive behavior in NSCLC cells.

Using Annexin V/PI staining and flow cytometry, we observed that PTE treatment induced apoptosis in A549 and H358 cells in a concentration-dependent manner ( Figures 3A–D ). Additionally, cell cycle analysis showed that PTE treatment resulted in G2/M phase arrest ( Figures 3E–H ). These data suggested that PTE promoted apoptotic cell death and disrupted cell cycle progression in NSCLC cells.

Reactive oxygen species (ROS) serve as key mediators of apoptosis (37). Prior research has indicated that PTE induces apoptosis in glioma cells via ROS production (38). In this study, we evaluated ROS levels using the fluorescent probe DCFDA. Flow cytometry analysis revealed a significant increase in ROS levels in both A549 and H358 NSCLC cells following PTE treatment ( Figures 4A, B ). Furthermore, immunofluorescence data supported these findings, showing elevated ROS levels as a result of PTE treatment ( Figure 4C ).

To investigate the relationship between ROS and PTE-induced growth inhibition, we conducted experiments with the ROS scavenger NAC. The CCK8 assay demonstrated that pretreatment with NAC effectively reversed the inhibitory effects of PTE on cell proliferation in both A549 and H358 cells ( Figure 4D ). Additionally, NAC pretreatment significantly reduced PTE-induced apoptosis ( Figures 4E, F ). Collectively, these findings indicated that ROS played a pivotal role in PTE-induced cell death and the inhibition of cell proliferation in NSCLC cells, highlighting the potential of targeting ROS pathways in cancer therapy.

The STING pathway plays a critical role in modulating the TME by regulating both antitumor immunity and cellular proliferation (25, 39). To clarify the relationship between ROS and STING activation, we conducted additional experiments in which cells were pretreated with NAC followed by PTE treatment. Western blot analysis showed that NAC pretreatment markedly suppressed PTE-induced phosphorylation of STING, TBK1, and IRF3 ( Figure 4G ). Additionally, NAC abolished PTE-induced upregulation of CXCL10 and CCL5 mRNA, as measured by qPCR ( Figure 4H ). These results confirmed that ROS generation preceded and was required for STING pathway activation and subsequent chemokine production.

To determine whether STING signaling contributes to the antitumor effects of PTE, we examined pathway activation in A549 and H358 cells after PTE treatment. As shown in Figure 5A , PTE treatment markedly increased phosphorylation of STING, TBK1, and IRF3, indicating activation of the pathway.

STING activation is commonly triggered by cytoplasmic double-stranded DNA (dsDNA), often resulting from genomic stress or DNA damage (39, 40). We therefore evaluated DNA damage responses by measuring γ-H2AX expression. Elevated levels of γ-H2AX were detected in both cell lines following PTE treatment, confirming induction of DNA damage ( Figure 5A ). To further investigate the functional role of STING in PTE-induced effects, we used the selective STING inhibitor H-151 (41). H-151 pretreatment significantly suppressed PTE-induced phosphorylation of TBK1 and IRF3 ( Figure 5A ), and partially restored cell proliferation ( Figure 5B ) and viability ( Figures 5C, D ), supporting the functional involvement of STING in PTE-mediated anticancer activity.

To complement the pharmacological inhibition approach and rule out off-target effects of H151, we performed shRNA-mediated knockdown of STING in A549 and H358 cells. Western blot analysis confirmed successful reduction of STING protein expression in both cell lines ( Figure 5E ), and qPCR verified the downregulation of STING mRNA ( Figure 5F ). Consistent with the H-151 results, STING knockdown significantly reversed PTE-induced inhibition of cell viability ( Figure 5G ) and reduced PTE-induced apoptosis ( Figures 5H, I ). These convergent findings from both pharmacological and genetic approaches strongly confirmed that STING activation was essential for the antitumor effects of PTE. Collectively, these results suggest that PTE effectively activated the STING pathway in NSCLC cells.

To evaluate the anticancer efficacy of PTE in vivo, we established a xenograft model by subcutaneously inoculating H358 cells into male BALB/c nude mice (42, 43). The mice were randomly assigned to two groups: one treatment group that received PTE (10 mg/kg) and a control group administered with the vehicle solution (5% DMSO, 10% Tween 80, 30% PEG400, and 55% normal saline). Both solutions were delivered via daily intraperitoneal injection. Throughout the experimental period, no significant differences in body weight were observed between the PTE -treated and control groups ( Figure 6A ), suggesting that PTE did not induce systemic toxicity at the administered dose.

Notably, PTE treatment significantly suppressed tumor growth compared to the vehicle control. A statistically significant reduction in tumor volume became apparent from day 14 post-treatment onward ( Figure 6B ). At the endpoint, tumors resected from the PTE -treated group were both visibly smaller ( Figure 6C ) and had significantly lower weights than those from the control group ( Figure 6D ). These consistent results across volume and weight measurements robustly demonstrate the inhibitory effect of PTE on NSCLC tumor progression in vivo. In summary, these findings indicated that PTE effectively restrained tumor growth in a xenograft model without eliciting overt adverse effects, underscoring its potential as a promising therapeutic candidate for NSCLC treatment.

Activation of the STING pathway in tumor cells can initiate potent innate and adaptive immune response against tumors (44). To investigate whether PTE exerts its anticancer effects through immunomodulation, we utilized an immunocompetent murine model of NSCLC. Treatment with PTE did not cause significant weight loss in NSCLC mice, as shown by the body weight change curve ( Figure 7A ). From day 15 of treatment onward, we observed a notable trend of tumor suppression in the PTE treatment group ( Figure 7B ), accompanied by significant reductions in tumor volume ( Figures 7B, C ) and overall tumor weight ( Figure 7D ).

To elucidate the immune mechanisms involved, we first measured systemic levels of key antitumor cytokines. At the endpoint of the experiment (day 24), ELISA analysis of mouse plasma revealed that PTE treatment significantly elevated concentrations of IFN-γ and TNF-α ( Figure 7E ), suggesting activation of a Th1-type immune response. To gain deeper insights into the enhanced therapeutic response, we employed flow cytometry to evaluate the anti-tumor immune activity within the TME, and Supplementary Figures S1 , 2 showed the flow cytometric analysis gating strategy. Our analysis revealed an increase in both the activity and abundance of CD8⁺ T cells in the PTE-treated group compared to the control group receiving carrier solution ( Figure 7F ). Notably, CD8⁺ T cells in the tumors of PTE-treated mice exhibited a more robust activation profile, producing significantly higher levels of TNF-α, GrzmB, and IFN-γ compared to the control group ( Figure 7G ).

Moreover, we observed a reduction in the populations of MDSCs, Tregs, and TAMs in the tumors of PTE-treated mice. This finding suggests that PTE not only enhances the activation of anti-tumor CD8⁺ T cells but also alleviates the suppressive immune environment in the TME. Overall, these results indicate that PTE treatment enhances the immune response and significantly improves the functionality of CD8⁺ T cells, thus promoting tumor control and survival in mouse models of NSCLC. Furthermore, PTE’s ability to reduce the expression of Tregs, MDSCs, and tumor-infiltrating macrophages underscores its potential as an effective therapeutic strategy for NSCLC.

Having established the mechanistic basis for PTE’s antitumor efficacy, we next considered its translational potential. Although the primary aim of this study was to elucidate the mechanistic basis of PTE’s antitumor effects, its translational potential justifies a discussion of pharmacokinetic properties. Previous reports indicate that PTE possesses superior pharmacokinetic profiles compared to resveratrol, attributable to its methoxyl groups, which enhance oral bioavailability and metabolic stability (73). Animal studies show that PTE reaches peak plasma concentrations within 15–30 minutes after oral administration and remains detectable for up to 8 hours, distributed in tissues including the lung, liver, and kidney (53). The dose used in our in vivo studies (10 mg/kg, intraperitoneal) yields plasma concentrations consistent with effective concentrations in our in vitro models, supporting the biological relevance of our findings. PTE has demonstrated a favorable safety profile in preclinical studies, with no significant toxicity observed at efficacious doses (74). In our hands, no significant weight loss or behavioral alterations were observed, suggesting limited systemic toxicity. We acknowledge that a detailed PK/PD study was beyond the scope of this initial mechanistic work. Future studies should focus on characterizing the pharmacokinetics, biodistribution, and maximum tolerated dose of PTE in immunocompetent NSCLC models to better bridge the gap between our findings and clinical application. Nano-formulation strategies, such as self-assembled micelles (75), could further improve tumor-specific delivery and minimize off-target effects.

Our proposed dosage of 10 mg/kg was based on prior studies demonstrating efficacy without overt toxicity. To contextualize PTE within the current NSCLC treatment landscape, we discuss its potential to overcome limitations of immune checkpoint inhibitors (ICIs). ICIs, such as anti-PD-1/PD-L1 agents, show limited efficacy in “immune-cold” tumors due to poor T-cell infiltration and immunosuppressive microenvironments (38). Our data suggest that PTE remodels the tumor microenvironment (TME) by activating STING-mediated signaling, leading to increased chemokine production (e.g., CXCL10, CCL5) and enhanced CD8⁺ T-cell recruitment and function. This mechanism aligns with recent strategies aiming to convert “cold” tumors into “hot” ones (48). We propose that PTE could synergize with ICIs by priming the TME—first, inducing immunogenic cell death and innate immune activation via ROS/STING, followed by ICI-mediated reversal of T-cell exhaustion. This sequence could potentially expand the therapeutic window and address ICI resistance. Future work will directly evaluate the efficacy of combining PTE with anti-PD-1 antibodies in immunocompetent NSCLC models. However, to meaningfully translate these preclinical insights, several key challenges and limitations of the current study must be addressed.

Despite the promising findings, this study has several limitations that warrant acknowledgment. First, the in vivo models employed-including H358 xenografts in immunodeficient mice and the LLC1 model in immunocompetent mice-may not fully recapitulate the heterogeneity and immunosuppressive complexity of human NSCLC. The LLC1 model, in particular, is known to be relatively immunogenic (76), which might lead to an overestimation of PTE’s immunomodulatory efficacy. Future investigations should utilize genetically engineered mouse models (GEMMs) that better mimic key molecular subtypes of human NSCLC (e.g., KRAS-driven or KRAS/p53-mutant models (77)) to validate our findings in more clinically relevant settings. Second, while we established a crucial link between ROS and STING activation, the precise mechanistic steps, such as whether ROS primarily causes mitochondrial DNA release or sensitizes the STING pathway to existing cytosolic DNA, remain to be elucidated. Third, the causal relationship between STING activation, chemokine production, and antitumor immunity would be strengthened by functional validation experiments, such as in vivo CD8⁺ T cell depletion or neutralization of key chemokines like CXCL10 and CCL5.

To bridge the gap between our mechanistic insights and clinical application, we outline the following future research directions: (1) Detailed PK/PD and Toxicity Studies: Conduct comprehensive pharmacokinetic, biodistribution, and maximum tolerated dose studies of PTE in immunocompetent NSCLC models. It is crucial to evaluate advanced formulations (e.g., nano-micelles (75)) to enhance tumor targeting and minimize off-target effects. Ultimately, toxicity assessments in non-human primates would be invaluable for de-risking clinical translation. (2) Combination Therapy Trials: Systematically evaluate the synergistic potential of PTE with standard-of-care immune checkpoint inhibitors (e.g., anti-PD-1/PD-L1 antibodies) in a range of immunocompetent models. This should aim to verify our hypothesis that PTE can convert “immune-cold” tumors into “immune-hot” microenvironments (78), thereby overcoming a primary mechanism of ICI resistance. (3) Biomarker Development: Identify predictive biomarkers (e.g., tumor STING pathway activation status, baseline CD8⁺ T cell infiltration, or chemokine profiles) to help identify patient subgroups most likely to benefit from PTE-based therapies.