A total of 121 human liver tissue samples were collected from patients who underwent surgical resections at the Hepatic Surgery Centre, Tongji Hospital of Huazhong University of Science and Technology (Wuhan, China). Detailed clinicopathological parameters are listed in Table I. The procedure of human specimen collection was approved by the Ethics Committee of Tongji Hospital, Huazhong University of Science and Technology, and the study was conducted according to the Declaration of Helsinki.

The human hepatoma cell line HepG2 and HBV genome-transfected HepG2 (HepG2.2.15) cells were obtained from the China Center for Type Culture Collection (Wuhan, China). Cells were cultured in high glucose Dulbecco's modified Eagle's medium (Gibco, Carlsbad, CA, USA) containing 10% fetal bovine serum (Gibco). Dimethyl sulfoxide (DMSO) and γ-secretase inhibitor N-[N-(3,5-difluorophenacetyl)-l-alanyl]-S-phenylglycine t-butyl ester (DAPT) were purchased from Sigma-Aldrich (St. Louis, MO, USA). DAPT was dissolved in DMSO.

Immunohistochemistry (IHC) was performed as described previously (16). Primary antibodies against HBx and Notch1 were purchased from Merck-Millipore (Billerica, MA, USA) and Santa Cruz Biotechnology (Santa Cruz, CA, USA), respectively.

Western blotting was performed as described previously (17). The primary antibodies and their sources were as follows: anti-Notch1, anti-Hes1, anti-NICD, anti-pERK, anti-ERK, and anti-pAKT (Cell Signaling Technology, Beverly, MA, USA); anti-Jagged1, anti-β-actin, anti-AKT, and anti-DUSP1 (Santa Cruz Biotechnology); anti-HBx (Merck-Millipore); anti-PTEN (Proteintech Group, Chicago, IL, USA). Horseradish peroxidase-conjugated secondary antibodies were purchased from Jackson Immuno Research Laboratories (West Grove, PA, USA).

Double immunofluorescence immunostaining was performed as described previously (18,19). Double-labeling immunofluorescence was used to detect HBx and Notch1 simultaneously. All sections were analyzed by confocal laser-scanning microscopy using a Nikon Digital Eclipse C1 system (Nikon Corp., Japan).

PCR was performed as described previously (20). The primer sequences for HBx were as follows: sense, 5′-GGCTGCTAGGCTGTGCTGCC-3′; antisense, 5′-GTTCCTGTGGGCGTTCACGG-3′. Images of electrophoresed PCR products were acquired using the Alpha Innotech Fluorochem Imaging system. Real-time PCR primers are listed in Table II. Cycle threshold values were reported relative to β-actin mRNA. Expression values were obtained in triplicate and normalized to β-actin expression. Results were calculated as fold induction relative to controls.

Small interfering RNA (siRNA) duplexes targeting human HBx, Hes1, DUSP1, and PTEN sequences and a scrambled siRNA were designed as described previously (5,21-23). All siRNAs were synthesized by Ribobio (Guangzhou, China). Transfection of the siRNA duplexes was performed by jetPRIME (Polyplus-transfection SA, Illkirch, France) according to the manufacturer's instructions.

The cell viability assay was performed as described previously (24). HepG2.2.15 cells (2×10³ per well) were seeded and cultured in 96-well plates for the indicated times. Cell Counting Kit-8 (CCK-8, Dojindo, Japan) was added to the cells, and the optical density value at 450 nm was measured after 2 h.

Luciferase assays were performed as described previously (25). Cell lysates was subjected to luciferase assays using the Dual-luciferase Reporter assay system (Promega) according to the manufacturer's instructions. DUSP1-luc reporter, PTEN-luc reporter, and Hes1 expression vectors were transfected as reported previously (26-28). Relative luciferase activities were determined by a Glomar 20/20 Illuminometer (Promega) and normalized against Renilla luciferase as an internal control.

Quantitative ChIP analysis was performed as described previously (29). Briefly, cells were crosslinked, sonicated, and immunoprecipitated with anti-Hes1 antibodies or IgG (negative control). A mixture of two anti-Hes1 antibodies (H140, Santa Cruz Biotechnology; 4H1, Novus Biologicals) was used for immunoprecipitation. Then, DNA was eluted and crosslinking was reversed, followed by purification and amplification for PCR analysis. The primers used in qPCR are listed in Table III.

Data were analyzed using GraphPad Prism 5.0 (La Jolla, CA, USA) or SPSS 13.0 (Chicago, IL, USA). All experiments were independently performed at least three times. The results are presented as the mean ± standard error of the mean. Comparisons between different groups were evaluated by one-way analysis of variance. P<0.05 was considered as statistically significant.

Immunohistochemical analysis showed that Notch1 was highly expressed in HCC tissues. Notch1 also exhibited expression in adjacent non-tumor and cirrhosis tissues. However, its expression was significantly lower than that in HCC tissues (P<0.01). Moreover, the expression of Notch1 in normal liver tissues was significantly lower than that in the above-mentioned tissues (P<0.001) (Fig. 1A and B). Western blotting showed similar results (Fig. 1C). In most HCC tissues, the expression of Notch1 was highly elevated compared with that in the other tissues. Notch1 was weakly expressed in all normal liver tissues.

Specifically, among 121 HCC tissues, 19 of them had weak expression and 102 of them had high expression. In the 121 corresponding peritumoral tissues, 59 of them had weak expression and 62 of them had high expression (P<0.001) (Table IV).

Table I shows the correlation between the factors and clinicopathological parameters in the 121 HCC patients. As shown in the table, sex, age, and the AFP level were unrelated to Notch1 expression. The level of Notch1 was significantly more elevated in HBV-positive patients (P<0.001), cirrhosis patients, (P<0.05), and patients with large HCCs (P<0.05).

Previous reports have shown the relationship of Notch1 with HBx. Here, we found co-localization of the two proteins. IHC results showed that Notch1 and HBx were co-expressed in HCC tissues (Fig. 2A). Fig. 2B shows the correlation of Notch1 and HBx expression levels. Spearman's rho was 0.584, indicating that Notch1 levels were positively correlated with HBx expression in the 108 HBV-positive HCC tissues. Fig. 2C shows HBx (green) and Notch1 (red) staining in HepG2.2.15 cells. The yellow staining in dual labeling experiments indicated overlapping areas of HBx and Notch1 staining, suggesting co-expression of Notch1 with HBx in HepG2.2.15 cells.

Because a relationship of HBx with Notch1 was found in HCC tissues and cells, we determined whether HBx could regulate the expression of Notch1 in HepG2.2.15 cells in vitro. To test this hypothesis, we compared the expression of HBx and some members of the Notch1 pathway between HepG2 and HepG2.2.15 cells, and between HepG2.2.15-SiCtrl and HepG2.2.15-SiHBx cells. Western blotting and PCR confirmed that HepG2 cells did not express HBx, whereas the HBx gene and protein were expressed in HepG2.2.15 cells (Fig. 3A and B). Next, we tested the expression of some members of the Notch1 pathway, such as Jagged1, Notch1, and Hes1. Western blotting showed that the protein levels of the three proteins were much more elevated in HepG2.2.15 cells than in HepG2 cells (Fig. 3B). Next, we knocked down HBx expression in HepG2.2.15 cells by HBx-specific siRNA. Western blotting and PCR confirmed that HBx expression was significantly decreased by the siRNA (Fig. 3C and D). Western blotting showed that the decrease of HBx expression led to a decrease of Notch1 and Hes1 expression in HepG2.2.15 cells (Fig. 3D), verifying that HBx regulated the expression of Notch1 in vitro.

Our results indicated that Notch1 was highly expressed in HCC tissues, especially in large HCC tissues, and the Notch1 pathway was regulated by HBx, we therefore determined whether inhibition of the Notch1 pathway attenuated cell growth. To inhibit the Notch1 pathway, we used the γ-secretase inhibitor DAPT. HepG2.2.15 cells were treated with increasing concentrations of DAPT for 4 days, and then cell viability was evaluated by CCK-8 assays. As shown in Fig. 3E, inhibition of HBx attenuated the growth of HepG2.2.15 cells. Increasing concentrations and treatment times of DAPT resulted in progressive inhibition of HepG2.2.15 cell viability (Fig. 3F). At day 1, DAPT treatment at various concentrations did not reduce cell viability. However, DAPT treatment for more than 2 days triggered a significant time- and dose-dependent decrease in the viability of HepG2.2.15 cells. We ultimately selected a DAPT treatment concentration of 20 µM for further experiments.

HepG2.2.15 cells treated with 20 µM DAPT for 1 and 6 h were assessed for inhibition levels of the Notch1 pathway by western blot analyses of Jagged1, Notch1, NICD, and Hes1 expression. DAPT treatment significantly decreased the amount of NICD and Hes1 in a time-dependent manner, but did not have any effect on Jagged1 or Notch1 (Fig. 3G). Our data showed that DAPT treatment greatly inhibited the growth of HepG2.2.15 cells. Therefore, we tested expression of ERK and AKT pathways that are closely related to cell proliferation. Western blotting indicated that DAPT greatly reduced the amount of pERK and pAKT in a time-dependent manner, but did not have any effect on total ERK or AKT (Fig. 3H).

Previous reports have revealed the regulatory circuit linking the pathway with DUSP1 expression and ERK activity (21). A link has also been found between the Notch1 pathway and PTEN and AKT activities (28). Our data showed that DUSP1 and PTEN were upregulated after DAPT treatment in HepG2.2.15 cells by western blotting (Fig. 3I and J).

We tested the effect of Hes1 inhibition on DUSP1 and PTEN levels. Treatment of HepG2.2.15 cells with siRNA targeting Hes1 mRNA (SiHes1) effectively reduced the Hes1 mRNA level (Fig. 4A). Growth of HepG2.2.15 cells was inhibited after SiHes1 treatment (Fig. 4B). Next, we found that the mRNA levels of DUSP1 and PTEN were greatly increased after SiHes1 treatment (Fig. 4C and D). Western blot analysis showed the same results. The Hes1 protein level was signifi-cantly reduced and the protein levels of DUSP1 and PTEN were elevated significantly (Fig. 4E).

Based on the above data, we investigated whether Hes1 suppressed DUSP1 and PTEN. We performed DUSP1 and PTEN promoter assays using a luciferase reporter. The basal activities of DUSP1 and PTEN gene promoters were reduced by Hes1. Treatment of HepG2.2.15 cells with DAPT induced DUSP1 and PTEN gene promoters, which was reversed by cotransfection of Hes1 (Fig. 4F and G). We next performed ChIP assays to determine whether Hes1 directly bound to the promoters of DUSP1 and PTEN genes using two mixed antibodies against Hes1. We designed several primers in the promoter regions of DUSP1 and PTEN genes (Fig. 5). ChIP assays showed that Hes1 bound to the regulatory sequences in the promoters of DUSP1 and PTEN genes in HepG2.2.15 cells (Fig. 6A and B). Quantitative ChIP assays revealed significant enrichment of two regions of the DUSP1 and PTEN gene promoters compared with control IgG immunoprecipitation, which was reversed by treatment with DAPT (Fig. 6C and D). These results further reinforce our previous findings that DAPT upregulated DUSP1 and PTEN expression by downregulating Hes1.

We next tested the effect of DUSP1 inhibition on ERK phosphorylation. Treatment of HepG2.2.15 cells with siRNA targeting DUSP1 mRNA (SiDUSP1) effectively reduced DUSP1 mRNA and protein levels, resulting in an elevated pERK level (Fig. 7A and B). We then evaluated the effect of PTEN inhibition on AKT phosphorylation. Treatment with SiPTEN effectively decreased PTEN mRNA and protein levels, which induced an increase of the pAKT level (Fig. 7C and D). The above findings demonstrated that DUSP1 dephosphorylated pERK and PTEN dephosphorylated pAKT.