The independent tissue microarrays were purchased (LP803 and LP804; US Biomax, Inc., Rockville, MD, USA). The tissue microarrays consisted of 110 laryngocarcinoma tissues, five normal laryngeal tissues and five samples of normal adjacent laryngeal tissue (NAT), which included 106 men and 14 women. Detailed tissue microarray information is presented in Tables I and II.

PD-L1 antibody (cat. no. 13684; Cell Signaling Technology, Inc., Danvers, MA, USA) was used for immunohistochemical staining, and the method of immunohistochemical staining was performed as previously described (17). To further analyze the immunohistochemical staining results, all TMAs were scored for frequency (0–4) and intensity (0–3) under an Olympus BX51 microscope (Olympus Corp., Tokyo, Japan). The scores of frequency were assigned when 0–25, 26–50, 51–75 and 76–100% of the tumor cells stained positive. The scores of intensity (0–3) respectively represented: 0, negative; 1, weak; 2, moderate; and 4, strong. The composite expression scores (CES) utilized the following formula: CES = intensity × frequency.

Two HNSCC cell lines (Cal-27 and FaDu) were purchased from Type Culture Collection of the Chinese Academy of Sciences (Shanghai, China). All cells were cultured in Dulbecco's modified Eagle's medium (DMEM; Thermo Fisher Scientific, Inc., Waltham, MA, USA) with 10% fetal bovine serum (FBS; Gibco; Thermo Fisher Scientific, Inc.), 20 mg/ml ampicillin and 20 mg/ml kanamycin at 37°C in an incubator with 5% CO₂.

To generate the stable PD-L1-overexpressing (PD-L1^over) and PD-L1 knockdown (PD-L1^RNAi; lentivirus transduction particles containing PD-L1 shRNA) cell lines, as well as negative control (NC) groups (PD-L1^{over NC} and PD-L1^{RNAi NC}), lentivirus transduction particles containing GFP label (cat. nos. GOCL3581014115 and GICL2481014115; Shanghai GeneChem Co., Ltd., Shanghai, China) were transfected (multiplicity of infection=20) into Cal-27 and FaDu cells (2×10⁵ cells/well), and the stable transfected Cal-27 and FaDu cell lines were selected by culturing for 1 week in complete medium (DMEM with 10% FBS, ampicillin and kanamycin), which also contained puromycin (2 µg/ml).

Cell proliferation was determined using Cell Counting Kit-8 (CCK-8; Dojindo Molecular Technologies, Inc., Kumamoto, Japan) and EdU incorporation (cat. no. C10310-1; Guangzhou RiboBio Co., Ltd., Shanghai, China) assays, according to the manufacturer's instructions. For the CCK-8 assay, cells were seeded in 96-well plates at 1×10³ cells/well in DMEM with 10% FBS, and treated with rapamycin (10 nM) for the total culture period of 72 h. The absorbance was measured at 450 nm by a microplate reader (Perkin Elmer) at 0, 24, 48 and 72 h. For the EdU proliferation assay, tumor cells (5×10⁶ cells/well) were plated into 6-well plates in DMEM with 10% FBS, and directly labeled using the Cell-Light™ EdU Apollo^® 567 in vitro Imaging kit, according to the manufacturer's instructions.

For the colony formation assay, cells were seeded into 6-well plates (200 cells/well) and incubated in complete medium for 12 days at 37°C. The 6-well plates were washed with PBS and stained with 0.1% crystal violet at room temperature for 15 min. Colonies which consisted of >50 cells were counted under an Olympus IX51 microscope (Olympus Corp.).

Total RNA was isolated with TRIzol™ reagent (Thermo Fisher Scientific, Inc.). RNA (1 µg) was reverse transcribed using the Super RT Reverse Transcriptase reagent kit (Beijing CoWin Biotech Co., Ltd., Beijing, China) according to the manufacturer's instructions. qPCR was conducted in a 25 µl reaction system, using the 7500 Fast Real-Time PCR System (Applied Biosystems; Thermo Fisher Scientific, Inc.) and amplified with transcript-specific primers and SYBR^®-Green Master Mix (Thermo Fisher Scientific, Inc.), according to the manufacturer's instructions. Relative gene expression was calculated using the 2^−ΔΔCq method (18), with GAPDH as the internal control. PD-L1 (cat. no. HQP008443) and GAPDH (cat. no. HQP006940) primers were purchased from GeneCopoeia, Inc. (Rockville, MD, USA). The primer sequences were as follows: PD-L1 forward, 5′-TAGAATTCATGAGGATATTTGCTGTCTT-3′ and reverse, 5′-TAGGATCCTTACGTCTCCTCCAAATGTG-3′; GAPDH forward, 5′-TGACTTCAACAGCGACACCCA-3′ and reverse, 5′-CACCCTGTTGCTGTAGCCAAA-3.

Female BALB/c nude mice (n=20; 4 weeks old; 16–18 g) were purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd. (Beijing, China) and underwent adaptive feeding 1 week before the experiment. Mice were housed at constant temperature (20–25°C) and humidity (40–70%) in a 12 h light/dark cycle, with free access to sterile water and standard chow. The nude mice were randomly divided into four groups (PD-L1^{over NC}, PD-L1^over, PD-L1^{RNAi NC} and PD-L1^RNAi; n=5 each). Cal-27 cells were selected to establish subcutaneous xenotransplanted tumor model since Cal-27 cells are more superior than FaDu cells in establishing a subcutaneous xenotransplanted tumor model. Cells (2×10⁶) were suspended in PBS (200 µl cell suspension) and injected into the right side of the mice's backs. Xenograft tumor diameters were measured every week, and tumor volumes were calculated using the following equation: Volume = 1/2 × length × width². The maximum tumor size was 20 mm. Nude mice were sacrificed and tumors surgically removed 12 weeks after inoculation.

Cal-27 and FaDu cells were harvested and lysed in radioimmunoprecipitation assay lysis buffer (Thermo Fisher Scientific, Inc.) supplemented with protease and phosphatase inhibitors (Roche Applied Science, Penzberg, Germany). Protein concentration was determined by the bicinchoninic acid protein assay. Lysates (20 µg of protein loaded per lane) were resolved by 10% SDS-PAGE, transferred to polyvinylidene difluoride membranes and immunoblotted with specific primary antibodies (all 1:800) overnight at 4°C against PD-L1 (cat. no. 9234T; Cell Signaling Technology, Inc.), protein kinase B (Akt; cat. no. 4691T; Cell Signaling Technology, Inc.), phosphorylated (p)-Akt^S473 (cat. no. 4060T; Cell Signaling Technology, Inc.), 70 kDa ribosomal protein S6 kinase 1 (P70S6K; cat. no. 2708S; Cell Signaling Technology, Inc.), p-P70S6K^T389 (cat. no. 9234T; Cell Signaling Technology, Inc.) and GAPDH (cat. no. 5174S; Cell Signaling Technology, Inc.). Following immunoblotting with IRDye^® goat-anti rabbit IgG flourescence secondary antibodies (dilution 1:20,000; cat. no. 926-32211; LI-COR Biosciences, Lincoln, NE, USA) at room temperature for 1 h, the membranes were scanned by an Odyssey infrared imaging system (LI-COR Biosciences).

All values are expressed as the mean ± standard deviation of three independent experimental repeats. Statistical analyses were performed in SPSS 19.0 (SPSS, Inc., Chicago, IL, USA), using one-way analysis of variance with Tukey's post hoc test. P<0.05 was considered to indicate a statistically significant difference.

In order to examine the expression of PD-L1 in HNSCC, NAT and normal tissues, TMAs of HNSCCs stained for PD-L1 were analyzed, and the CES of every clinical sample was measured. PD-L1 expression was compared and associated with clinical characteristics, including tumor grade and TNM staging.

PD-L1 expression in tumor tissue was significantly higher than in normal tissue and NAT (P<0.01), with no significant difference in PD-L1 expression between normal tissue and NAT (P>0.05; Fig. 1A). T-stage analysis revealed that although the expression of PD-L1 in normal tissue was significantly higher in T1 (P<0.01), T2 (P<0.05) and T4 (P<0.05), there was no statistical significance in PD-L1 expression in T3 (P>0.05; Fig. 1B). It is likely that a larger number of samples are required to fully investigate the relationship between T-stage and PD-L1, as the sample sizes of T1 and T3 were insufficient, and the expected increase in PD-L1 expression with T stage progression was not observed. For the N-stage, representative images were presented in Fig. 1C. The CES of the PD-L1 protein expression revealed that there was no significant differences between the N-stages (P>0.05). As N-stage indicates regional lymph node metastasis, these results suggest that PD-L1 may have no relationship with the regional metastasis of HNSCC. Finally, the expression of PD-L1 among different tumor grades was compared respectively, demonstrating that an increased tumor grade was associated with increased PD-L1 expression (Fig. 1D).

To explore the function of PD-L1, stable PD-L1-overexpressing (PD-L1^over), PD-L1 knockdown (PD-L1^RNAi) and negative control cells (PD-L1^{over NC}, PD-L1^{RNAi NC}) were generated, and the expression of PD-L1 was detected by RT-qPCR and western blotting. The stably transfected Cal-27 and FaDu cells were established and representative micrographs showed the immunofluorescence of GFP in cells (Fig. 2A). It was demonstrated that the lentiviral transduction particles successfully altered PD-L1 protein (Fig. 2B) and gene (Fig. 2C) expression. PD-L1^RNAi HNSCC cell proliferation was significantly reduced, compared with its respective control group, with the PD-L1^over cells exhibiting the highest rate of proliferation (Fig. 3A). Furthermore, the EdU proliferation assay revealed a similar trend, in which the PD-L1-overexpressing cells exhibited the highest red fluorescence (Fig. 3B). Furthermore, the colony forming ability of PD-L1-overexpressing cells was remarkably increased compared with the PD-L1^{over NC} group, and colony numbers in the PD-L1 knockdown group were significantly reduced, compared with the PD-L1^{RNAi NC} group (Fig. 3C).

To investigate the potential mechanism by which PD-L1 promoted cell growth in HNSCC cells, the expression levels of proteins associated with mTOR signaling and cell proliferation were detected by western blotting (Fig. 4A). Compared with their corresponding control groups, p-P70S6K^T389 and p-Akt^S473 expression was significantly increased on the PD-L1^over group, but significantly decreased in the PD-L1^RNAi group (Fig. 4B).

To validate whether the expression of PD-L1 affected tumor growth in vivo, Cal-27 cells were used to establish a xenograft mouse model. At the 12 week end point, the tumor growth curve (Fig. 5A) showed that the tumor volume in the Cal-27-PD-L1^over group was significantly larger than the control group (P<0.01), and the tumor volume in the PD-L1^RNAi group was the smallest (P<0.05). The tumors were removed and measured on week 12 (Fig. 5B), and the average volume was calculated (Fig. 5C). These results demonstrated that PD-L1 accelerated tumor growth, suggesting an important role for PD-L1 in regulating the tumor cell growth in vivo. No significant differences in animal weight were detected (Fig. 5D).

As PD-L1 upregulated the expression of proteins involved in mTOR signaling, the effects of mTOR inhibitor rapamycin were investigated. Cellular proliferation was significantly inhibited in the PD-L1^over group following treatment with mTOR inhibitor. PD-L1^over Cal-27 cells were the most sensitive to rapamycin in the four groups, and the PD-L1^RNAi tumor cells were the most tolerant to rapamycin-mediated proliferation inhibition (Fig. 6A-C). The result obtained in Cal-27 and FaDu cells were consistent. Although there was no significant statistical difference between PD-L1^RNAi and PD-L1^{RNAi NC} groups in FaDu cells, it was still indicated that PD-L1 exerted some regulatory action on tumor cell proliferation, and mTOR inhibitor was effective in preventing the PD-L1-driven proliferative effects. As presented in Fig. 4, PD-L1 knockdown reduced the activation of mTORC1 and mTORC2. In response to rapamycin treatment, p-P70S6K^T389 expression was markedly suppressed, and p-P70S6KT³⁸⁹ could mediate a negative feedback loop on phosphoinositide 3-kinase (PI3K)/Akt which was de-repressed by rapamycin, and elevated the level of p-Akt^S473 (Fig. 7A and B). The proportion of p-P70S6K^T389 appeared the most markedly decreased in PD-L1^over+RP cells, compared with PD-L1^over cells; and p-Akt^S473 of PD-L1^RNAi, which increased, was more than PD-L1^RNAi+RP (Fig. 7B), further indicating that PD-L1 may have promoted cell growth through mTOR signaling upregulation.