Patients’ blood and PSCC tissue samples were collected at Aldenora Bello Cancer Hospital, São Luis-Brazil, from 2017 to 2020. This study was approved by the Research Ethics Committee on Humans from the Federal University of Maranhão and by the National Research Ethics Commission (CONEP‐ Brazil, CAAE: 46371515.5.0000.5087).

This study employed experimental research techniques utilizing both in vitro and ex vivo methodologies. Peripheral blood was collected from patients (n=30) and healthy donor volunteers (n=19). The tumor tissue from patients was enzymatically digested to characterize the immune infiltrating cells and to establish ex vivo cultures. HPV genotyping was performed using nested-PCR, following the protocol provided by BioGenetics Molecular Technologies (Uberlândia, Minas Gerais, Brazil; patent number BR102017004615.0), to identify viral subtypes classified as high risk (16, 18, 31, 33, 35, 39, 45, 51, 52, 56, 58, 59, 66, 68, 73, and 82), intermediate risk (MM7, MM8, 26, 30, 34, 53, 54, 55, 61, 62, 64, 67, 69, 70, 71, 72, 74, and 81), and low risk (6, 11, 42, 43, 44, and 57). In addition, we performed DNA sequencing according to our previous protocol (3). The tumor cell culture supernatants were used as tumor microenvironment-like to perform the apoptosis lymphocyte assay. From the peripheral blood mononuclear cells (PBMCs), the monocytes were isolated for in vitro differentiation in mature Dendritic Cells (Mo-DCs) and the lymphocytes were used in proliferation and apoptosis assay. Mo-DCs activation was evaluated through the expression of costimulatory molecules and compared with patients’ groups and healthy donors ( Supplementary Figure 1 ).

Tumor tissues fragments were treated with Collagenase IV (0.056 mg/mL of RPMI 1640 medium) according to the manufacturer (Sigma-Aldrich, Saint Louis,USA) at 37°C for 2 hours (18) After enzymatic digestion, the cells were stained with 0.04% Trypan Blue dye and counted using a Neubauer chamber. Phenotypic analysis by flow cytometer was performed on 2x10⁵ tumor cells, as described below. Additionally, 1x10⁴ tumor cells were cultured in 12-well plates for 5 days. After 5 days, the supernatant was removed from the plate and centrifuged to remove non-adherent cells that remain in suspension in the medium. For the apoptosis assay, the tumor supernatant was added to the lymphocyte culture at a concentration of 10%. This assay was conducted over three days in 96-wells plates containing 100 µL of supplemented medium.

Lymphocyte activation depends on the antigen presentation mechanism by Antigen-Presenting Cells (APCs), mainly represented by DCs. Thus, monocytes obtained from the peripheral blood of patients with HPV-positive and HPV-negative PSCCa were differentiated in vitro into dendritic cells. Buffy coat of peripheral blood (5-10mL) samples from patients and healthy blood donors were used for the isolation of peripheral blood mononuclear cells (PBMCs) and plasm. Dendritic Cells from both groups were produced by standard procedures (18). Briefly, PBMCs were isolated from peripheral blood buffy coats using Ficoll density gradient. After washing with phosphate buffered saline, 1x10⁶ cells/mL were plated and kept in culture with cell culture medium (RPMI 1640, Gibco, New York, USA) supplemented with 10% of fetal bovine serum (FBS), 1% of Antibiotic-antimycotic (Gibco, New York, USA) overnight, to separate adherent cells (monocytes) from non-adherent cells (lymphocytes). The lymphocytes were separated for future proliferation and apoptosis assays. The monocytes were seeded in 12-well plates and cultured in RPMI 1640 medium with 10% FBS added 50 ng/mL⁻¹ recombinant human GM-CSF (PeproTech, Cranbury, USA), and 50 ng/mL⁻¹ IL-4 (PeproTech, Cranbury, USA) to generate immature DCs. On the fifth day of culture, 20 ng/mL⁻¹ TNF-α (PeproTech, Cranbury, USA) was added to the culture to generate mature monocyte-derived dendritic cells (Mo-DCs). After 7 days of culture, the cells were harvested, washed once with PBS, and used for the experiments.

A total of 1x10⁵ cells were incubated, at room temperature, for 30 min, with PBS and antibody: CD14 (BD Biosciences Cat# 555397, RRID: AB_395798), CD86 (BioLegend Cat# 305422, RRID: AB_2074981), HLA-DR (BD Biosciences Cat# 339194, RRID: AB_647443), CD3 (BioLegend Cat# 300312, RRID: AB_314048), CD8 (BioLegend Cat# 300906, RRID: AB_314110), CD4 (BD Biosciences Cat# 555347, RRID: AB_395752), CD19 (BioLegend Cat# 302210, RRID: AB_314240), and CD56 (BD Biosciences Cat# 555516, RRID: AB_395906) ( Supplementary Table 1 ). Following incubation, cells were washed with PBS, and analyzed using an InCyte Guava flow cytometer (Luminex Corporation). Data analysis was performed using FlowJo software (v.10; RRID: SCR_008520).

To assess the ability of Mo-DCs to induce lymphocyte activation, the allogeneic lymphocytes obtained from healthy donors were labeled with 2.5 μM carboxyfluorescein succinimidyl ester (CFSE) (Sigma-Aldrich,Saint Louis, USA) and co-cultured with patients’ or healthy donors’ Mo-DCs for 3 days. Fluorescent signals were analyzed using a flow cytometer, following the manufacturer’s instructions. A minimum of 10,000 events were acquired using a FACSCalibur flow cytometer and analyzed with cytometer software (FlowJo Software, v.10; RRID: SCR_008520).

To simulate the effect of the tumor microenvironment on lymphocyte viability, the ex vivo culture supernatant from the tumor tissue was used as treatment. The healthy donor and patient’s lymphocytes were incubated for three days in a 96-well plate containing 100 µL of RPMI medium supplemented with 10% tumor supernatant. Fluorescence-conjugated Annexin V and propidium iodide were utilized for apoptosis analysis (BD Pharmingen, Franklin Lakes, USA) following the manufacturer’s instructions. After the incubation, the lymphocytes isolated from patients and healthy donors were washed and incubated in Annexin V binding buffer at a 1:20 dilution for 15 minutes at room temperature. Subsequently, the cells were then washed, resuspended in a fresh Annexin V binding buffer, and promptly analyzed by flow cytometry, according to the manufacturer’s guidelines.

Statistical analyses were performed using GraphPad Prism software (v10). Data are presented as the mean ± standard error of the mean (SEM). The Shapiro-Wilk test was used to assess the normality of data distribution. Depending on the distribution (normal or non-normal), appropriate parametric or non-parametric tests were applied. For comparisons between two groups, the parametric t-test or the non-parametric Mann-Whitney U test (both two-tailed and unpaired) were used. For comparisons involving three or more groups, either the parametric one-way ANOVA followed by Tukey’s multiple comparison post hoc test or the non-parametric Kruskal-Wallis test followed by Dunn’s multiple comparison post hoc test was employed. A p-value < 0.05 was considered statistically significant, with significance levels indicated as *p < 0.05, **p < 0.005, and ***p < 0.0005. The sample size for each test and the specific test used in each analysis are detailed in the figure legends.

Most of the patients enrolled in the study have low levels of education. The main occupation was agriculture, and the majority were married. The average age of the patients was 60.18 (26-94 year) years old, and reported smoking and drinking habits (50% and 53.3%, respectively). Phimosis was present in 40%, but this information was not available for 40% of the patients. The most frequent site of the tumor was the glans (46.6%) or the glans associated with other areas (39.9%).

All tumors were squamous cell carcinoma with 56.5% HPV-related (Warty carcinoma: 33.3%; Basaloid: 6.6%; Warty-basaloid: 16.6), 26.6% non-HPV related, and 16.6% had no information. Histological grades I (16.6%) and II (16.6%) were the most frequent, however, this information was not available for 40% of tumors. The size of the tumor for most patients (70%) was 0.7 to 5.4 cm, and the tumors were defined as pT1 (13.3%), pT2 (20%), pT3 (23.3%), and pT4 (3.3%). However, 36.6% of this information was not available in the medical records. All tumor samples were tested for HPV DNA detection using nested-PCR, with 66.6% testing positive. Among the HPV-positive samples used in immunological assays, HPV16 was the most common genotype, present in 66.7% of cases, including one case of co-infection with HPV6. Additionally, 33.3% of the samples were positive for HPV18, defined as high-risk oncogenic HPV (hrHPV). Table 1 provides an overview of the sociodemographic, clinical, and pathological characteristics of the patients.

The expression of specific markers CD3+ for T lymphocytes, CD19+ for B lymphocytes, CD56+ for NK cells, and CD14+ for monocytes/macrophages was used to assess immune cell infiltration within the tumor. The results revealed that the frequency of T lymphocytes was significantly lower in hrHPV-positive (37.96 ± 6.78) compared to HPV-negative PSCC (63.89 ± 5.12, p=0.0104). An inverse trend was observed in the frequency of NK cells, with a higher frequency in hrHPV-positive (3.44 ± 2.26) compared to HPV-negative cases (1.27 ± 0.54, p=0.2698). No significant differences (p≥0.05) were observed in the average frequencies of B lymphocytes and monocytes/macrophages between the two groups ( Figure 1 ).

Comparing the frequency of Mo-DCs positive for CD86 between patients and healthy donors, it was observed that there was a significant reduction in cells obtained from hrHPV-positive patients (13.10 ± 3.03; healthy donors: 69.81 ± 9.10, p=0.0025). However, despite a trend towards reduction, the frequency was not significant in HPV-negative patients (49.36 ± 13.92, p= 0.3180). As for the frequency of Mo-DCs expressing co-stimulatory molecules, the results showed that hrHPV-positive patients have a lower proportion of positive cells compared to cells obtained from HPV-negative patients, but without significance ( Figure 2A ). The intensity of CD86 molecule expression on the membrane of Mo-DCs was lower in PSCC patients, both hrHPV-positive (101.8 ± 22.30; p=0.0293) and HPV-negative (145.2 ± 44.68; p=0.0490), compared to healthy donors (415.0 ± 103.4) ( Figure 2B ). The ability of Mo-DCs to induce lymphocyte proliferation was lower in cells from hrHPV-positive patients compared to healthy donors (Proliferation Index respectively: 1.33 ± 0.07; 2.08 ± 0.22, p=0.0275). No difference was observed in the proliferation index between HPV-negative patients and healthy donors. There was also no significant difference in the ability to induce lymphoproliferation between patient groups; however, the data showed a trend towards reduction in cells obtained from hrHPV-positive patients compared to HPV-negative patients (Proliferation Index respectively: 1.33 ± 0.07; 1.80 ± 0.21, p=0.1668) ( Figure 2C ). Interestingly, these findings correlate with the CD86 expression and frequency of positive cells.

Assessing the impact of the tumor microenvironment on the induction of apoptosis in lymphocytes, the results showed that lymphocytes from healthy donors (38,80 ± 0,6938) exhibited a lower initial apoptosis rate compared to lymphocytes from hrHPV-positive (42,07 ± 0,1687, p=0,0067) and HPV-negative (41,90 ± 0,6351, p=0,0229) patients when cultured in medium. However, when lymphocytes were cultured in the presence of tumor supernatant, lymphocytes from healthy donors showed a significant increase in apoptotic cells (42.53 ± 0.40, p= 0.0018), compared with healthy donors. Interestingly, lymphocytes from hrHPV-positive patients had a reduction in the frequency of apoptotic cells when cultured with tumor supernatant (38.37 ± 0.78), when compared to culture with medium only (42.07 ± 0.16, p=0.0019) ( Figure 3A ). The data remained consistent across groups in different treatments when evaluating late apoptosis ( Figure 3B ). On the other hand, the frequency of viable cells was higher in lymphocytes from healthy donors (47.10 ± 1.00) cultured in the medium compared to hrHPV-positive (43.70 ± 0.98, p=0.0392) and HPV-negative patients (42.20 ± 0.11, p=0.0496). However, there was a reduction in viability of lymphocytes from healthy donors (42.53 ± 0.40) when treated with tumor supernatant (47.10 ± 1.00, p=0.0392) ( Figure 3C ). The data on viability and initial apoptosis complement each other, both in lymphocytes from healthy donors and patients.