All experimental procedures involving animals and human tissues were approved by the Ethics Committee of Guangzhou Development District Hospital (approval no. F2024-030; Guangzhou, China).

All animal experiments were approved by the Institutional Animal Care and Use Committee of Guangzhou Development District Hospital (Guangzhou, China). Mice were housed in a specific pathogen-free facility, maintained in a controlled environment (22±2°C, 50 ± 10% humidity, 12 h light/dark cycle) and provided ad libitum access to food and water. All efforts were made to minimize animal suffering. For human studies, this research utilized a retrospective design. Peripheral blood samples were retrospectively collected from healthy donors, and matched tumor tissue samples were obtained from patients with CRC, respectively, following the provision of written informed consent in accordance with the Declaration of Helsinki. The study protocol, including the retrospective use of these samples, was approved by the Institutional Review Board of the Guangzhou Development District Hospital (Guangzhou, China).

Peripheral blood samples were collected from healthy donors via venipuncture and drawn into sterile ethylenediaminetetraacetic acid-coated tubes on ice. Clinical tumor specimens were collected from 11 patients with colon cancer who received ICI therapy at Guangzhou Development District Hospital (Guangzhou, China) between July 2022 and December 2024. Patients were enrolled based on the following criteria: i) Histologically confirmed diagnosis of colon adenocarcinoma; ii) scheduled to undergo or having undergone first-line ICI therapy (either as monotherapy or in combination); iii) availability of pre-treatment tumor tissue samples and complete clinical follow-up records; iv) aged ≥18 years. Key exclusion criteria included: i) Prior history of other active malignancies ≤5 years; ii) severe concurrent autoimmune diseases or active infections; iii) receipt of any other form of anticancer therapy (such as chemotherapy or radiotherapy) ≤4 weeks prior to sample collection; v) incomplete clinical or pathological data. Surgical resection or core needle biopsy was used to obtain tumor tissues, which were immediately placed in ice-cold RPMI-1640 medium (Gibco; Thermo Fisher Scientific, Inc.). Based on the Response Evaluation Criteria for Solid Tumors (34), tumor responses were evaluated using contrast-enhanced computed tomography. Patients whose target lesions exhibited either complete response (CR) or partial response (PR) were categorized as responders (n=5), whereas those with stable disease (SD) or progressive disease (PD) were classified as non-responders (n=6). CR was defined as the disappearance of all target lesions and PR as a ≥30% reduction in their total diameter. SD was characterized by insufficient shrinkage to qualify as a partial response and PD was defined as a minimum 20% increase in the target lesions. The clinical cohort consisted of 7 men and 4 women, with a median age of 62 years (range, 48–71 years).

The CRC cell lines MC38 (cat. no. iCell-m032; Mirror Point (Shanghai) Cell Technology Co., Ltd) and HT-29 (cat. no. iCell-h078; Mirror Point (Shanghai) Cell Technology Co., Ltd) were maintained according to the manufacturer's protocols. They were cultured with activated T cells (specifically, in vitro-activated human peripheral blood-derived T cells and isolated murine CD8⁺ T cells from OT-1 mice, as detailed in the ‘T-cell separation’ section) at 37°C in a 5% CO₂ incubator. MC38 cells were genetically modified to overexpress SIINFEKL via a lentiviral vector system (cat. no. MC38-OVA; iCell Biotech). Cells were cultured in DMEM (Gibco; Thermo Fisher Scientific, Inc.), supplemented with 10% fetal bovine serum (FBS; Gibco; Thermo Fisher Scientific, Inc.), 100 U/ml penicillin, 100 µg/ml streptomycin and maintained in a humidified incubator at 37°C with 5% CO₂. For the treatment experiments, cells were either cultured individually or co-cultured with activated T cells at a ratio of 3:1 for 48 h in the presence of Exe-9 (GLP-1R antagonist; cat. no. E7269; MilliporeSigma) at concentrations of 10 and 50 nM, whereas parallel cultures received phosphate-buffered saline (PBS) as the vehicle control.

Human peripheral blood mononuclear cells were isolated from whole blood of healthy donors via density gradient centrifugation using Lymphoprep (cat. no. 10970; STEMCELL Technologies). Specifically, blood was collected in BD CPT tubes (cat. no. 362782; BD Biosciences) containing Ficoll™ density gradient medium (Ρ=1.077 g/cm³) and sodium citrate. Tubes were incubated at room temperature for 20 min to stabilize the temperature, then centrifuged at 1,650 × g for 20 min at 20°C with the brake on. This formed a density barrier separating PBMCs and serum. The PBMC fraction was gently mixed with the serum by inversion and transferred to 15 ml tubes for further processing. and subsequently cultured in CTS™ AIIM V™ SFM medium (cat. no. A3021002; Gibco; Thermo Fisher Scientific, Inc.) supplemented with ImmunoCult™ Human CD3/CD28/CD2 T Cell Activation Reagent (cat. no. 10970; STEMCELL Technologies) and recombinant 1,000 U/ml human interleukin (IL)-2 (cat. no. 11848-HNAY1; Sino Biological, Inc.) for 7 days to obtain activated human T cells. CD8⁺ T cells were isolated from the spleens of OT-1 mice (cat. no. 003831; THE JACKSON LABORATORY), aged 6–8 weeks old with a body weight of 18–25 g using the EasySep™ Mouse CD8⁺ T Cell Isolation Kit (STEMCELL Technologies) in accordance with the manufacturer's instructions. Collected cells were cultured briefly in complete T cell medium comprising RPMI-1640 (Gibco; Thermo Fisher Scientific, Inc.) supplemented with 10% FBS, 20 mM HEPES, 1 mM sodium pyruvate, 0.05 mM 2-mercaptoethanol, 2 mM L-glutamine and 50 U/ml streptomycin/penicillin, with recombinant mouse IL-2 (cat. no. CK24; Novoprotein Scientific, Inc.) and incubated for 5 days.

After treatment, the adherent cancer cells were washed thrice with PBS, fixed with 100% methanol at room temperature for 10 min and stained with a 0.5% crystal violet solution at room temperature for 30 min. After staining, the wells were rinsed with PBS to remove excess dye and subsequently solubilized by adding 10% acetic acid for 10 min. Optical density (OD) was measured at 570 nm using a BioTek Epoch 2 microplate reader (BioTek Instruments; Agilent Technologies, Inc.) to quantitatively determine cell viability.

To quantify interferon (IFN)-γ and tissue necrotic factor (TNF)-α levels, the supernatants of the colon cancer cell culture were collected after a 48 h incubation period at 37°C and centrifuged at 13,000 × g for 5 min at 4°C. The supernatants were analyzed using the Mouse IFN-γ enzyme-linked immunosorbent assay (ELISA) Kit (cat. no. PI508; Beyotime Biotechnology), Mouse TNF α ELISA Kit (cat. no. ab285327; Abcam), Human IFN-γ ELISA Kit (cat. no. PI521; Beyotime Biotechnology) and human TNF α ELISA Kit (cat. no. ab181421; Abcam), in strict accordance with the manufacturer's instructions. Briefly, 100 µl each standard and sample was dispensed into pre-coated wells of a 96-well microplate and incubated for 2 h at room temperature. After washing with 0.05% Tween-20 in PBS, 100 µl horseradish peroxidase (HRP)-conjugated detection antibody (provided within the respective ELISA kits was added.

The antibody in the aforementioned IFN-γ reagent kit was used directly (utilizing the pre-prepared formulation provided with the kit, which required no additional dilution), while the antibody in the TNF-α reagent kits was used at a 1:100 dilution; both antibody application protocols followed the specifications in the respective manufacturers' instructions; cat. no. 80220; Alpha Diagnostic Intl. Inc.) was added and the plate was incubated for an additional 1 h at room temperature. After a series of washes, 100 µl tetramethylbenzidine substrate solution (MilliporeSigma) was added and the enzymatic reaction proceeded in the dark for 20 min. The reaction was terminated by the addition of 50 µl 2 M sulfuric acid and the absorbance was measured at 450 nm. Cytokine concentrations were determined by generating standard curves using recombinant cytokine standards provided in the kits.

Male BALB/c mice (6–8 weeks old; 22–25 g) were purchased from Beijing Vital River Laboratories Animal Technology Co., Ltd. and subcutaneously injected with 1×10⁶ MC38 cells (day 0) into the left flank. When tumor volumes reached ~150 mm³, the mice were randomized into treatment groups (n=10). To assess the antitumor efficacy of Exe-9, mice were randomly allocated to either the Exe-9 group (n=10) or the control group (n=10). To evaluate the synergistic effect of Exe-9 combined with Anti-PD-1, mice were randomly assigned to four groups: Control (n=10), Exe-9 (n=10), Anti-PD-1 (n=10) and Exe-9 + Anti-PD-1 (n=10). Mice in the Exe-9 group received intratumoral injections of Exe-9 at a concentration of 25 nmol/kg in saline on days 10, 12, 14 and 16. Mice in the Anti-PD-1 group were administered 100 µg of anti-PD-1 antibody (cat. no. BE0146; Bio X Cell, Lebanon) via intraperitoneal injection on days 10, 13, 16 and 19. The control mice received equivalent volumes of sterile saline or isotype immunoglobulin G (IgG; cat. no. 1054; Bio X Cell), as appropriate. To further elucidate whether the antitumor effects of Exe-9 were mediated by the modulation of tumor immunity, an additional experiment was conducted in male BALB/c nude mice (Beijing Vital River Laboratories Animal Technology Co., Ltd.) using the same group size (n=10 per group) following the same injection schedule as the Exe-9 group.

Prior to the invasive procedures, the mice were anesthetized via inhalation of isoflurane (3–4% for induction and 1.5–2% for maintenance in oxygen). Mice demonstrating severe distress signs at the end of the experiment were humanely euthanized via CO₂ inhalation, followed by cervical dislocation.

Tumor length and width were recorded at the same intervals as the injections using digital calipers to generate tumor growth curves. According to institutional ethical guidelines, the maximum tumor volume permitted was 1,500 mm³ and the maximum diameter did not exceed 20 mm in any dimension. In the present study, the largest tumor measured was ~1,200 mm³ in volume and 17 mm in diameter. The mice were euthanized on day 20; tumor tissue and spleen were collected for further assays. Euthanasia was performed by gradual CO₂ inhalation at a displacement rate of 30–40% volume per min of the chamber, followed by cervical dislocation to ensure mortality. The tumors were harvested and weighed to assess their anticancer efficacy every 2 days starting from day 10.

Excised mouse or patient tumor tissues were fixed in 10% neutral-buffered formalin for 24 h at room temperature and processed for paraffin embedding. Paraffin blocks were sectioned at 5 µm thickness and mounted onto slides. Sections were deparaffinized in xylene and rehydrated using a graded ethanol series (100, 95 and 70%) before rinsing in distilled water. After heating in a 10 mM citrate buffer (pH 6.0) at 92°C for 15 min, slides were incubated in 3% hydrogen peroxide for 10 min at room temperature. For the detection of the intracellular/membrane protein GLP-1R, sections were permeabilized with 0.1% Triton X-100 in PBS for 10 min at room temperature. Blocking was performed with 5% normal goat serum (cat. no. 5425; Cell Signaling Technology, Inc.) for 30 min at room temperature and the sections were incubated overnight at 4°C with a primary anti-CD8 (cat. no. PA007381.r2b; Syd Labs) or anti-GLP-1R antibody (cat. no. EPR21819; Abcam) diluted 1:100 in blocking solution. After three washes in PBS, a biotinylated goat anti-mouse IgG secondary antibody (1:200; cat. no. ab6789; Abcam), was applied for 1 h at room temperature. Immunoreactivity was visualized using the VECTASTAIN^® Elite ABC HRP Kit (Vector Laboratories, Inc.) with diaminobenzidine, following the manufacturer's protocol. Subsequently, the sections were counterstained with hematoxylin (MilliporeSigma) at a concentration of 1% for 2 min at room temperature, dehydrated using graded ethanol and xylene solutions and cover slipped. The stained slides were examined and imaged under an Olympus BX53 light/fluorescence microscope (Olympus Corporation). Quantitative analysis of IHC staining was performed using the Alpathwell Pathology Image Analysis System (version 3.5; Servicebio Technology Co., Ltd.).

GLP-1R staining was semi-quantified using the H-score method (35). For each case, five tumor fields (×400 magnification) were randomly selected within the tumor area. The percentage of tumor cells demonstrating weak (1+), moderate (2+) or strong (3+) staining intensity was recorded (P1-P3; total, 0–100%) and the H-score was calculated as (1×P1 + 2×P2 + 3×P3), yielding a range of 0–300. Two board-certified pathologists independently scored all slides in a blinded manner. In cases of initial disagreement, the two pathologists first re-examined the corresponding slides together to reach a consensus. If a consensus could not be achieved, a third senior pathologist was consulted to make the final determination.

Spleens harvested from Exp-9 treated mice or the naive control were mechanically dissociated and filtered through a 70 µm cell strainer to obtain a cell suspension. Cells were then plated at a density of 1×10⁵ cells per well in ELISpot plates pre-coated with anti-IFN-γ capture antibody, as provided in the Mouse IFN-γ ELISpot Kit (cat. no. 3321-4APT-2; Mabtech). For restimulation, cells were incubated with either irradiated MC38 cells or the major histocompatibility complex (MHC)-I restricted AH1 peptide derived from MC38 cells (MilliporeSigma) at a final concentration of 1 µg/ml. After being cultured for 20 h at 37°C, the plates were washed with PBS and incubated with a biotinylated anti-IFN-γ detection antibody for 2 h at room temperature. After subsequent washing steps, an HRP conjugate was added and the reaction was developed using a 3-amino-9-ethylcarbazole substrate solution (cat. no. SK-4200; Vector Laboratories, Inc.) at room temperature in the dark for 15 min. The enzymatic reaction was stopped by rinsing the plates with distilled water and the spots were counted using an automated ELISpot reader (ELR08; Cellular Technology Limited).

Total RNA was extracted from freshly frozen tissue samples using TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc.), following the manufacturer's instructions. RNA purity and concentration was assessed using a NanoDrop™ 2000 spectrophotometer (Thermo Fisher Scientific, Inc.). Subsequently, 1 µg total RNA was reverse transcribed into cDNA using the PrimeScript™ RT reagent Kit (Takara Bio, Inc.). The quantitative real-time PCR was performed on an ABI 7500 Real-Time PCR System (Applied Biosystems, Inc.; Thermo Fisher Scientific, Inc.) using SYBR^® Premix Ex Taq II (Takara Bio, Inc.) The PCR protocol comprised three stages: Initial denaturation at 95°C for 30 sec to activate the DNA polymerase and denature templates; 40 cycles of amplification, each consisting of denaturation at 95°C for 5 sec followed by a combined annealing/extension at 60°C for 30 sec (the temperature of this step was optimized based on the primer Tm); and a melting curve analysis (95°C for 15 sec; 60°C for 1 min; then gradual heating to 95°C) to confirm amplicon specificity and exclude nonspecific products. The primers used to detect GLP-1R and GAPDH for normalization are listed in Table SI. PCR conditions were set as: 95°C for 30 sec, followed by 40 cycles of 95°C for 5 sec and 60°C for 30 sec. Relative gene expression was calculated using the 2^−ΔΔCq method (36). All primers were designed for human genes and validated by National Center for Biotechnology Information Basic Local Alignment Search Tool (National Institutes of Health).

Tissue samples from patients with CRC were homogenized and lysed using radioimmunoprecipitation assay buffer supplemented with 1% protease inhibitor cocktail and 1% phosphatase inhibitor cocktail (MilliporeSigma) on ice for 30 min. Lysates were clarified via centrifugation at 12,000 × g for 15 min at 4°C and protein concentrations were determined using a BCA protein assay kit (Thermo Fisher Scientific, Inc.). Equal amounts of proteins (30 µg per lane) were resolved on 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gels and transferred onto polyvinylidene difluoride membranes (MilliporeSigma) using a Bio-Rad Trans-Blot Turbo system. The membranes were blocked with 5% (w/v) skim milk in Tris-buffered saline containing 0.1% Tween-20 (TBST) for 1 h at room temperature. Membranes were incubated overnight at 4°C with primary antibodies targeting GLP-1R (1:500). After washing, the membranes were incubated with HRP-conjugated secondary IgG (1:2,000, Beyotime Biotechnology) for 1 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence detection kit (Thermo Fisher Scientific, Inc.) and imaged using the ChemiDoc XRS+ Imaging System (Bio-Rad Laboratories, Inc.). Densitometric band intensity analysis was performed using the ImageJ 2.0 software (National Institutes of Health), with GAPDH used for normalization. The antibodies used for western blotting analysis are listed in Table SII.

All data were analyzed using the GraphPad Prism software (version 9.0; Dotmatics). Continuous variables were expressed as mean ± SEM and the Shapiro-Wilk test was employed to assess data normality. For normally distributed data, one-way ANOVA followed by Tukey's honest significant difference post hoc test was used for group comparisons, whereas for non-normally distributed data, the Kruskal-Wallis test with Dunn's multiple comparison test was applied. Pearson's correlation analysis was performed to evaluate the correlation between tumor volume and the number of IFN-γ-secreting cells, as well as between GLP-1R expression and the immunotherapy response. P<0.05 was considered to indicate a statistically significant difference.

The colon cancer cell lines HT-29 and MC38-OVA were cultured either individually or co-cultured with activated T cells for 48 h in the presence of the GLP-1R antagonist Exe-9 at 10 or 50 nM or PBS as the vehicle control. The viability of HT-29 (Fig. 1A) and MC38-OVA (Fig. 1B) cells was assessed. In the absence of T cells, Exe-9 treatment did not significantly alter cancer cell viability, as no statistically significant differences were detected between the PBS- and Exe-9-treated groups. By contrast, when co-cultured with activated T cells, a significant dose-dependent reduction in cell viability was observed, indicating enhanced T-cell-mediated cytotoxicity in the presence of Exe-9. In parallel, ELISA measurements of the culture supernatants revealed that levels of IFN-γ (Fig. 1C) and TNF-α (Fig. 1D) increased following Exe-9 treatment compared with the control, with a dose-dependent effect evident particularly in the MC38-OVA cultures. Collectively, these results suggest that Exe-9 enhances the killing efficiency of CD8⁺ T cells against colon cancer cells, potentially by modulating the cytokine milieu that governs T-cell-mediated cytotoxicity.

To evaluate the effects of the GLP-1R antagonist on tumor growth, male mice were subcutaneously injected with MC38 tumor cells. When tumors reached ~150 mm³ on day 10, Exe-9 was administered intratumorally at days 10, 12, 14 and 16, while control mice received equivalent volumes of saline. Tumor tissues were collected on day 20 for analysis (Fig. 2A). Notably, the tumors in Exe-9-treated mice were markedly smaller compared with those in the control group (Fig. 2B). The tumor volume measurements recorded at each treatment point demonstrated a continuous increase in both groups. However, the Exe-9 group exhibited a markedly lower growth rate (Fig. 2C) compared with the control group. At the endpoint, tumor volume (Fig. 2D) and weight (Fig. 2E) in the control group were significantly greater compared with those in the Exe-9-treated group. IHC analysis revealed that tumors from Exe-9-treated mice displayed markedly higher CD8⁺ T-cell infiltration compared with those from the control group (Fig. 2F), suggesting enhanced antitumor immunity. To investigate the role of T cells in this response, the experiment was replicated using nude mice lacking functional T cells. In this model, no significant differences in tumor size were observed between the Exe-9 and saline-treated groups (Fig. 2G); however, the tumor volume continued to increase in both groups throughout the treatment period (Fig. 2H and I). Tumor weights collected on day 20 were also not significantly different between the two groups (Fig. 2J). These findings suggested that the antitumor effects of Exe-9 depend on the presence of functional T cells, potentially through the enhancement of CD8⁺ T-cell-mediated immunity.

To investigate the effect of the GLP-1R antagonist on tumor-specific CD8⁺ T-cell responses, splenocytes were collected from mice and restimulated for 20 h with either MC38 tumor cells or MHC-I-restricted AH1 peptide. The number of IFN-γ-secreting cells was quantified using ELISpot assays. Exe-9-treated mice exhibited significantly higher numbers of IFN-γ-secreting cells compared with the control mice under MC38 (Fig. 3A) and AH1 (Fig. 3B) restimulation conditions. Correlation analysis between tumor volume and IFN-γ-secreting cells revealed distinct patterns. In the Exe-9-treated group, a negative correlation trend was observed between them following MC38 restimulation, although the trend was not statistically significant (r=−0.6302; P=0.0508; Fig. 3C). However, a significant negative correlation was identified when the AH1 peptide was restimulated (r=−0.6439; P=0.0445; Fig. 3D), suggesting that Exe-9 treatment enhanced tumor-specific CD8⁺ T-cell activation. By contrast, splenocytes of mice from the control group consistently exhibited low numbers of IFN-γ-secreting cells, regardless of tumor volume or type of restimulation antigen (Fig. 3C and D). These results revealed that Exe-9 promotes the generation of tumor-specific CD8⁺ T cells capable of robust IFN-γ production. These results further suggest that the enhanced antitumor response induced by Exe-9 is likely mediated by an increase in antigen-specific CD8⁺ T-cell activity.

A combination treatment experiment with a GLP-1R antagonist and an ICI was performed in mice bearing MC38 tumors. Tumors were established to an average volume of ~150 mm³ by day 10 post-injection. Mice were treated intratumorally with Exe-9 on days 10, 12, 14 and 16 or injected intraperitoneally with anti-PD-1 on days 10, 13, 16 and 19. A combination of both treatments or a control with saline and IgG, was administered at the same time points. The tumors were collected on day 20 for further analysis (Fig. 4A). The control group exhibited the largest tumors, followed by the Exe-9, then the Anti-PD-1 group and the Exe-9 + Anti-PD-1 group with the smallest tumors (Fig. 4B). While the tumor volumes in the control, Exe-9 and Anti-PD-1 groups continuously increased throughout the treatment period, the combination of Exe-9 and Anti-PD-1 resulted in effective tumor control, with no significant changes in tumor volume observed (Fig. 4C). By day 20, the tumor volumes were quantified as shown in Fig. 4D and revealed the same trend as shown in the representative images in Fig. 2B. Consistent with these observations, the tumor weights at the endpoint also demonstrated significant differences among groups. Specifically, the tumor weight in the combination treatment group (Exe-9 + Anti-PD-1) was significantly reduced compared with that in the Anti-PD-1 monotherapy group (P<0.001; Fig. 4E). These results illustrate that Exe-9 enhanced the antitumor efficacy of anti-PD-1 in vivo, indicating a potential synergistic interaction between GLP-1R antagonism and immune checkpoint inhibition in suppressing colon cancer growth.

To investigate the clinical relevance of GLP-1R expression in tumor immunotherapy, tumor samples from 11 patients with CRC treated with ICIs were collected and categorized into responder and non-responder groups based on therapeutic outcomes. Quantitative PCR analysis revealed that GLP-1R expression levels were significantly higher in tumor tissues from non-responders compared with those from responders (P<0.05; Fig. 5A). Stratifying patients into high and low GLP-1R expression groups based on the median expression level revealed a positive correlation between high GLP-1R expression and non-responsiveness to therapy (r=0.8718; P=0.0005; Fig. 5B). These findings were corroborated by IHC and western blotting analyses, revealing that GLP-1R protein expression was significantly higher in tumor tissues from non-responders compared with those from responders (Fig. 5C and D). These results indicate that elevated GLP-1R expression is associated with reduced efficacy of anti-PD-1 immunotherapy in patients with CRC.