A total of 75 patients with HCC (51 males and 24 females) who underwent resection of HCC at the Second Affiliated Hospital of Xi'an Jiaotong University (Xi'an, China) between January 2013 and January 2017 were included in the present retrospective study. Data was also collected from the medical files of the patients. Liver cancer and adjacent normal tissues (≥5 cm from the tumor) were collected from the 75 patients and frozen at −180°C for further analysis. Serum samples (n=75) were collected from 4 ml peripheral blood and stored at 4°C in a 5-ml tube. After centrifugation at 3,000 × g for 10 min at 4°C, the obtained serum (1 ml) was stored at −80°C. Serum samples from the control group were collected from 66 healthy subjects without any disease, matched according to sex and age. The patients did not receive any chemotherapy, radiotherapy or immunotherapy prior to surgery. The liver cancer and adjacent normal tissues were collected for immunohistochemical analysis. According to the Ethics Committee of the Second Affiliated Hospital of Xi'an Jiaotong University (Xi'an, China), all participants provided written informed consent. Clinical data (including age, sex, stage, tumor size, intrahepatic metastasis, portal vein invasion and Edmondson grade) were collected from the medical records of each patient (Table I). All the specimens were graded using the Edmonson method (23). According to the differentiation degree of tumor cells, HCC tissues were categorized into grades I–III. Grade I was defined as low-grade HCC, grade II as medium-grade HCC and grade III as high-grade HCC. Furthermore, the TNM stage was defined according to the American Joint Committee on Cancer TNM Staging for Liver Tumors as follows (24): TX, primary tumor cannot be assessed; T0, no evidence of primary tumor; T1, solitary tumor without vascular invasion; T2, solitary tumor with vascular invasion or multiple tumors <5 cm in size; T3a, multiple tumors >5 cm in size; T3b, single tumor or multiple tumors of any size, involving a major branch of the portal vein or hepatic vein; and T4, tumor(s) with direct invasion of adjacent organs, other than the gallbladder, or with perforation of visceral peritoneum.

The patients were followed up from January 2016 to January 2020, initially every 2 months and at least every 3–6 months following surgery. Furthermore, the liver function and serum α-fetoprotein (AFP) levels were examined, and abdominal ultrasonography was performed. If recurrence was suspected, CT or MRI scan was performed immediately. OS and recurrence-free survival (RFS) times were defined as the interval between surgery and death or recurrence, respectively. If recurrence was not diagnosed, the patient was examined at the date of death or at the last follow-up evaluation. All of the aforementioned investigations were performed as previously described (25,26).

Serum samples were collected from the peripheral blood samples, centrifuged at 4,000 × g for 10 min at −4°C and then stored at −80°C for further analysis. According to the manufacturer's instructions, Ang II (cat. no. ADI-900-204; Enzo Life Sciences, Inc.) and S1P levels (cat. no. abx585002; Abbexa Ltd.) were assessed using their respective ELISA kits. Each analysis was performed in triplicate.

As previously described (27), total protein was extracted using RIPA lysis buffer (cat. no. PP1901; BioTeke Corporation), and protein concentration was determined using a BCA protein detection kit (cat. no. P0010; Beyotime Institute of Biotechnology). Protein samples (25 µg/lane) were separated via 10% SDS-PAGE and transferred onto PVDF membranes (Bio-Rad Laboratories, Inc.), then blocked with 5% skimmed milk in TBS buffer containing 0.1% Tween-20 at 25°C for 2 h, and incubated with specific antibodies against AT1R (1:500; cat. no. ab124734), S1PR1 (1:500; cat. no. ab77076) and β-actin (1:1,000; cat. no. ab200658) at −4°C overnight (all Abcam). β-actin was used as the internal control. Subsequently, membranes were incubated with an HRP-conjugated goat anti-rabbit IgG H&L secondary antibody (1:5,000; cat. no. ab7090; Abcam) at 37°C for 1 h. Enhanced chemiluminescence reagents (Pierce; Thermo Fisher Scientific, Inc.) were used to visualize the blots, and the optical densities of the bands were scanned and quantified using Syngene Gene Tools (Syngene Europe; model G box chem hr16; serial no. SYDR4/2327). A total of three independent experiments was performed.

mRNA levels were detected as previously described (27). Total RNA was extracted using the TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc.). cDNA was synthesized from total RNA (1 µg) using a PrimeScript™ RT master mix kit (Takara Bio, Inc.) according to the manufacturer's instructions. qPCR was performed using a SYBR Premix Ex Taq™ II Perfect Real-Time kit (Takara Bio, Inc.) on an ABI qPCR System (Applied Biosystems; Thermo Fisher Scientific, Inc.) according to the manufacturer's instructions. Each sample was run in triplicate. Primers for AT1R, S1PR1 and β-actin were designed using the Beacon designer 4.0 software (version 4.0; Premier Biosoft International). The primer sequences were as follows: AT1R forward, 5′-AGACTGGCCTTCTCTGGA-3′ and reverse, 5′-CACCGAGGAATACGCTTT-3′; S1PR1 forward, 5′-CACGCTTTCTGTGGCTTGGA-3′ and reverse, 5′-CGACGATGGCGCTCCAACA-3′; and β-actin forward, 5′-TGGCACCCAGCACAATGAA-3′ and reverse, 5′-CTAAGTCATAGTCCGCCTAGAAGCA-3′. The RT-PCR products were observed using electrophoresis and 2% agarose gels with ethidium bromide. A melting point dissociation curve was used to indicate that only a single product was shown. For qPCR, relative mRNA levels were analyzed using the 2^−ΔΔCt method (28). Data were standardized to β-actin mRNA levels. A total of three independent experiments were performed.

Liver cancer tissue samples were collected from 75 patients with HCC following hepatic resection at the Second Affiliated Hospital of Xi'an Jiaotong University between January 2013 and January 2017. During the same period, adjacent normal liver tissue samples were also obtained. All specimens (adjacent normal and liver cancer tissues) were investigated using immunohistochemistry. Tissue samples were fixed in 10% formalin at room temperature for 48 h and embedded in paraffin. Next, 5-µm sections were deparaffinized for antigen retrieval in citric acid buffer (pH 6.0; 95°C for 20 min). Sections were washed 3 times with PBS for 3 min each, 3 times with xylene for 5 min each, 2 times with 100% ethanol for 10 min each, 2 times with 95% ethanol for 10 min each and finally 2 times with double-distilled water for 5 min each. Slides were treated with 3% hydrogen peroxide at room temperature for 10 min to block endogenous peroxidase activity. After blocking in 1% BSA (Sigma-Aldrich; Merck KGaA) at room temperature for 20 min, the slides were incubated with rabbit anti-human AT1R (1:50; cat. no. ab124734; Abcam) and S1PR1 (1:100; cat. no. ab77076; Abcam) antibodies overnight at 4°C. Subsequently, the slides were incubated with 2 µg/ml biotinylated anti-rabbit IgG secondary antibody (XianFeng Bioengineering Institute) at room temperature for 40 min. Next, the sections were stained using a Standard Ultra-Sensitive ABC Peroxidase Staining kit (Thermo Fisher Scientific, Inc.), incubating the tissue sections with the ABC Reagent (Reagent A, 2 ml; Reagent B, 2 ml) at room temperature for 30 min. Bright-field images (magnification, ×200 and ×400) were captured using an Axio Scan Z1 light microscope (Zeiss GmbH). Immunostaining results were independently observed and interpreted by two pathologists blinded to the clinical data. Weak and strong staining were distinguished via the color of brown-yellow granules. When the brown-yellow granules were dark in the HCC tissues, this was considered as strong staining (high expression), while when the brown-yellow granules were weak in the HCC tissues, this was considered as weak staining (low expression).

Quantification of HBsAg was performed using an automated chemiluminescent microparticle immunoassay using the ARCHITECT i2000 system (Abbott Pharmaceutical Co., Ltd.). All procedures were performed according to the manufacturer's protocol (29). The sensitivity and the specificity of this assay are 100 and 99.76%, respectively, according to the manufacturer's protocol.

Data are expressed as the mean ± standard deviation. A Fisher's exact test was performed to determine whether AT1R and S1PR1 expression levels in HCC were associated with clinicopathological characteristics. A two-tailed paired or unpaired Student's t-test was used to determine significant differences between groups. The Kaplan-Meier method was used for survival analysis, and differences in survival were investigated using the log-rank test. These statistical analyses were performed using SPSS software (version 19.0; IBM Corp.). Correlation between Ang II and S1P, and between AT1R and S1PR1 in patients with HCC were analyzed using Pearson's correlation coefficient (GraphPad Prism 8; GraphPad Software, Inc.). P<0.05 was considered to indicate a statistically significant difference.

The clinical characteristics and laboratory parameters of patients with HCC are summarized in Table I. The results indicated that AT1R and S1PR1 expression levels were not significantly associated with patient sex, age, levels of HBsAg or serum AFP, or tumor size. However, AT1R and S1PR1 expression levels in HCC were associated with intrahepatic metastasis (P=0.017 and P=0.008, respectively), portal vein invasion (P=0.033 and P=0.015, respectively), Edmondson grade (P=0.017 and P=0.008, respectively) and TNM stage (P=0.004 and P=0.015, respectively).

The Ang II and S1P levels in the serum collected from the 66 healthy donors and 75 patients with HCC were determined, and the results revealed that both the serum levels were significantly higher in patients with HCC compared with those in healthy donors (Fig. 1).

Next, AT1R and S1PR1 protein expression levels were investigated in HCC tissues from 75 patients using immunohistochemistry. Positive staining for AT1R and S1PR1 was displayed as brown-yellow granules, which were primarily located in the cell membrane or cytoplasm. The cells with positive staining exhibited a clear tissue cell structure, where uniform brown-yellow granules were present in the cell membrane or cytoplasm and staining was markedly higher compared with that in the background. In human normal liver tissue, AT1R and S1PR1 were weakly expressed, whereas they were strongly expressed in human HCC tissue (Fig. 2).

To further assess AT1R and S1PR1 protein and mRNA expression levels in HCC, liver cancer and adjacent normal tissue obtained from 3 representative samples (according to IHC staining) among the 75 HCC samples were subjected to western blot and RT-qPCR analyses. AT1R and S1PR1 protein expression levels in HCC tissue were significantly higher compared with those in normal liver tissue (P<0.05; Fig. 3A and B). In addition, RT-qPCR was performed to quantify AT1R and S1PR1 mRNA expression levels in HCC tissues. AT1R and S1PR1 mRNA expression levels in HCC tissue were significantly higher compared with those in normal liver tissue (P<0.05; Fig. 3C and D), which was consistent with western blot analysis results. In summary, western blot, RT-qPCR and immunohistochemical analyses demonstrated that AT1R and S1PR1 expression was increased in HCC tissue.

The serum levels of Ang II and S1P in 66 healthy donors and 75 patients with HCC were determined using ELISA. A positive correlation was identified between Ang II and S1P in patients with HCC (P=0.002), but not in healthy donors (P=0.228; Fig. 4A and B). Furthermore, the correlation between AT1R and S1PR1 protein expression levels in the 75 HCC and normal adjacent tissues was also investigated and there was no correlation between AT1R and S1PR1 levels in the normal liver samples (P=0.081); however, there was a correlation in HCC tissue samples (P=0.001; Fig. 4C and D).

Kaplan-Meier survival curves were used to determine the effects of AT1R and S1PR1 protein expression levels via immunohistochemical analyses on the prognosis of patients with HCC. The results revealed that OS and RFS rates were significantly lower in patients with high AT1R expression levels compared with those in patients with low AT1R levels (P=0.0250 and P=0.0366, respectively; Fig. 5A and B). The results also revealed that OS and RFS rates were significantly lower in patients with high S1PR1 expression levels compared with those in patients with low S1PR1 levels (P=0.0401 and P=0.0359, respectively; Fig. 5C and D). The present results indicated that the 50-month survival rate of patients with HCC with high AT1R and S1PR1 expression was low. These results demonstrated that AT1R and S1PR1 upregulation was significantly associated with patient death and HCC recurrence.