This article is mentioned in:

Introduction

Decoy receptor 3 (DcR3), a soluble molecule belonging to the tumor necrosis factor receptor superfamily (TNFRSF), was first identified as a decoy receptor of Fas ligand (FasL) and inhibitor of FasL-induced apoptosis (1). DcR3 also neutralizes the biological effects of two other TNFSF members, namely, LIGHT (TNFSF14) and TNF-like molecule 1A (TL1A/TNFSF15) (2–4). DcR3 can be defined as an immunomodulator on the basis of its neutralizing effects on FasL, LIGHT and TL1A (4–7). DcR3 is upregulated in tumor cells and inflammatory diseases (8–11). DcR3 in serum can be used as a biomarker to predict cancer invasion and progression of inflammation (12–15). DcR3 also acts as an effector molecule to modulate cell function through non-decoy activities, including the effects on cell adhesion and differentiation (16–18).

Liver cancer is a common malignancy worldwide. Cancerous liver tissues yield high DcR3 expression, and this expression is correlated with tumor differentiation, serosal invasion and liver metastases (13,19,20). Nevertheless, the precise mechanisms of DcR3 in liver cancer progression and metastasis remain unclear.

E-cadherin, a classical member of the cadherin superfamily, is a calcium-dependent cell-cell adhesion glycoprotein (21). The E-cadherin-catenin complex plays a key role in cellular adhesion (22,23). The loss of E-cadherin function or expression has been implicated in cancer progression and metastasis (24,25). In the present study, DcR3 treatment caused HepG2 cell cytoskeleton remodeling, inhibited E-cadherin expression, and promoted cell migration. Immunohistochemical analysis revealed that E-cadherin and DcR3 exhibited an opposite expression trend in liver carcinoma tissues. The present study also demonstrated the functional mechanism of DcR3 in cancer cell migration and provides a theoretical basis for the use of a DcR3 antagonist to treat liver cancer.

Materials and methods

Clinical samples

Tissue samples from three patients with hepatic carcinoma and biliary tract disease were collected during surgical resection performed at the Shenzhen Second People's Hospital (Shenzhen, China). Liver tumor and non-tumor liver tissues were fixed and immediately used to prepare tissue slices. All of the samples were obtained with patient consent and approval of the Institutional Animal Care and Use Committee, Shenzhen Institutes of Advanced Technology.

Cell culture and transfection

The human hepatocarcinoma cell line HepG2 or the normal liver cell line L02 were obtained from the Shanghai Institute of Cell Biology (Shanghai, China). Both cell lines were maintained in Dulbecco's modified Eagle's medium (DMEM), supplemented with 10% fetal bovine serum (FBS) (Gibco, Carlsbad, CA, USA), and 100 µg/ml each of penicillin-streptomycin (HyClone, Logan, UT, USA) in 5% CO2 at 37°C.

In 6-well plates, 2×105 cells/well were cultured overnight and transfected with 2 µg PLVX–IRES-ZsGreen-DcR3 plasmid with Lipofectamine® 2000 (Invitrogen, Carlsbad, CA, USA). The cells transfected with an empty vector were used as a blank control.

Western blot analysis

HepG2 cells were treated with 3 µg/ml DcR3-Fc or IgG1 as control, or transfected with the indicated plasmids. At 48 h after transfection or treatment, cells were harvested and determined by the antibodies indicated in the figures. Cell pellets were lysed in RIPA (Thermo Fisher Scientific Inc., Waltham, MA, USA), 1 mM phenylmethylsulfonyl fluoride (PMSF) (Beyotime, Shanghai, China) and 1% protease inhibitor cocktail (Thermo Fisher Scientific, Inc.). Lysates were normalized for total protein (25 µg) and loaded on 8–12% sodium dodecyl sulfate-polyacrylamide gel, electrophoresed, and transferred to a polyvinylidene fluoride (PVDF) membrane (Millipore, Kenilworth, NJ, USA), followed by blocking with 5% skimmed milk at room temperature for 1 h. The membrane was incubated with primary antibodies overnight at 4°C, and rinsed with Tris-buffered saline with Tween-20. The blots were then incubated with horseradish peroxidase (HRP)-conjugated secondary antibody (KPL) for 1 h at room temperature. Detection was performed using EMD Millipore Luminata™ Western HRP Chemiluminescence Substrates (WBLUR0500). Nuclear and cytoplasmic extracts were isolated with NE-PER™ Nuclear and Cytoplasmic Extraction Reagents (78833) purchased from Thermo Fisher Scientific. The defined sections of the film were scanned for image capture and quantification using Adobe Photoshop software (CS4; (Adobe Systems, Inc., San Jose, CA, USA) and ImageJ software (National Institutes of Health, Bethesda, MD, USA).

RNA isolation and real-time quantitative PCR

Total RNA was isolated using TRIzol reagent (Invitrogen) according to the manufacturer's instructions. RNA samples were reverse transcribed with oligo (dT) and M-MLV Reverse Transcriptase (Takara, Tokyo, Japan). A mixture of 1 µg RNA, 4 µl 5X RT mix, 1 µl primer mix, and nuclease-free water were made up to a 20-µl volume. The reverse transcription step was as follow: 37°C for 15 min; 85°C for 5 sec, and then stored at −20°C. Real-time quantitative PCR analysis was performed with specific primers for human E-cadherin (forward, 5′-TGGAGGAATTCTTGCTTTGC-3′ and reverse, 5′-CGTACATGTCAGCCAGCTTC-3′) in a CFX96 Touch™ Real-Time PCR Detection System (Bio-Rad, Hercules, CA, USA) with SYBR qPCR mix (Takara). Relative levels of gene expression were determined using GAPDH as the control (forward, 5′-ATCTGGCACCACACCTTCTAC-3′ and reverse, 5′-CAGCCAGGTCCAGACGCAGG-3′). SYBR-Green PCR Master Mix 2 µl, forward and reverse primers 200 nM, cDNA template 100 ng, and ddH2O up to 10 µl volume was mixed together. PCR conditions consisted of the following: 95°C for 3 min for denaturation; 95°C for 5 sec for annealing; and 60°C for 40 sec for extension, for 40 cycles. The threshold cycle for each sample was selected from the linear range and converted to a starting quantity by interpolation from a standard curve generated on the same plate for each set of primers. The E-cadherin mRNA levels were normalized for each well to the GAPDH mRNA levels using the 2−ΔΔCt method.

Immunofluorescent assay

L02 or HepG2 cells were washed with phosphate-buffered saline (PBS) and fixed at room temperature with 4% polyformaldehyde for 10 min, permeated with 0.1% Triton X-100 for 7 min, blocked for 30 min with 1% BSA, and incubated sequentially with the indicated primary and secondary antibodies. 4,6-Diamidino-2-phenylindole (DAPI) (Beyotime) was used to label the nuclei. Phalloidin-Rhodamine (Thermo Fisher Scientific) was used for F-actin staining.

Flow cytometry

L02 or HepG2 cells were collected and washed with PBS, fixed and permeabilized with Fix/Perm solution (BioLegend, San Diego, CA, USA) before intracellular staining. After 15 min, the cells were washed twice with Perm/Wash buffer, and incubated with the DcR3 antibody at 4°C for 1 h. Cells were washed with PBS and incubated with the FITC-goat anti-rabbit antibody at 4°C for 30 min. Cells were washed with PBS twice. The intracellular fluorescence of FITC was detected by FCM after excitation at 488 nm. Fluorescence emissions at 530 nm from 10,000 cells were collected, amplified and scaled to generate a single-parameter histogram.

Immunochemistry

The sample sections were deparaffinized and rehydrated. After boiling in a microwave oven, the antigen was retrieved with a 0.01 M sodium citrate buffer (pH 6.0) at a sub-boiling temperature for 20 min. The following steps were performed with the SP kit (9001; ZSGB-BIO, Beijing, China). Shortly, the sections were incubated with 3% hydrogen peroxide for 10 min to block endogenous peroxidase. After 15 min of pre-incubation in 5% normal goat serum to prevent non-specific staining, the samples were incubated with the antibody to DcR3 (Abcam, Cambridge, UK) at 4°C overnight. Secondary antibody was added and incubated for 30 min. The sections were incubated in horseradish enzyme-labeled chain avidin solution for 30 min at room temperature. Color was developed with a diaminobenzidine (DAB) substrate kit. Counterstaining was performed with hematoxylin.

Wound healing assay

Confluent cell cultures were grown on 6-well plates. Wounds were made with the tip of a micropipette. DcR3-Fc was added to the culture medium at a final concentration of 3 µg/ml. IgG1 was added as a control. Wound closure speed was analyzed as indicated in the legend.

Transwell assay

HepG2 cells were treated with 3 µg/ml DcR3-Fc or IgG for 24 h, and then were trypsinized and resuspended in DMEM without FBS before plating on the upper layer of the Transwell with an 8-µm pore-size membrane at a cell density of 1×104. DMEM containing 5% FBS was added to the lower layer. After 15 h, the cells remaining on the top surface were scratched off. The cells on the lower surface were fixed in methanol, stained with 0.5% crystal violet (Beyotime), and images were captured under a microscope. The intact Transwell was dissolved in 33% acetic acid, and the supernatant was detected for absorption values with a spectrophotometer at 590 nm.

Materials

DcR3-Fc and human IgG1 proteins were purchased from Sino Biological, Inc. (Beijing, China). Anti-DcR3, anti-E-cadherin and anti-IκBα antibodies were purchased from Abcam. Anti-p65 was purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-tubulin was purchased from Abmart (Shanghai, China).

Statistical analysis

All experiments were repeated at least three times or noted otherwise. Data are expressed as mean ± SD. The t-test was performed for inter-group comparisons. Values with p<0.05 were considered to show significant differences.

Results

DcR3 regulates colony scattering of HepG2 cells and decreases E-cadherin expression

HepG2 cells were examined using a colony scattering assay to analyze the function of DcR3 in regulating cell migration. The colony scattering assay, mimicking certain aspects of tumor invasion, reveals the ability of epithelial tumor cells to detach from colonies in culture. The cells were plated at very low density, and the morphological characteristics of the colonies were evaluated 5 days after plating. The colonies were compact in the control or IgG1 groups, and >90% of the cells in a colony contained cell-cell junctions. By contrast, the cells were scattered in the DcR3 treatment group, and <20% of the cells formed junctions (Fig. 1A). In the cells detached from the scattered colonies, numerous protrusions were formed on the membrane edge of these cells (Fig. 1A, lane 3, arrows). Philloidin staining revealed that DcR3 promoted actin remodeling and revealed a scattered phenotype (Fig. 1B). This finding indicated that DcR3 triggered changes in cell morphology and enhanced the ability of cells to detach from the colonies.

Figure 1. DcR3 regulates colony scattering and inhibits E-cadherin expression. Cells were plated at a low density and treated with 3 µg/ml DcR3-Fc or control IgG1. The colony morphology was analyzed 4 days after plating. (A) Representative images of colony formation. Rows 1–3 indicate colony scattering under different magnification. Scale bar: row 1, 150 µm; row 2, 50 µm and row 3, 20 µm. (B) Representative images of actin remodeling as observed by phalloidin staining. Scale bar, 20 µm. (C) HepG2 cells were treated with 3 µg/ml DcR3-Fc. Forty-eight hours later, the E-cadherin mRNA level was determined by qRT-PCR. Data are from 3 indepentdent experiments; *p<0.05. (D) HepG2 cells were treated as described in C, and the protein level of E-cadherin was detected using western blot analysis. Representative images from two independent experiments are shown. (E) HepG2 cells were transfected with the control or DcR3-expressing vectors. At 48 h after transfection, cells were harvested and whole-cell lysates were detected by the indicated antibodies. Representative images from two independent experiments are shown.

DcR3 treatment disrupts colony scattering and causes cytoskeleton remodeling in HepG2 cells. To investigate the role of DcR3 in the regulation of cell-cell adhesion, we detected whether E-cadherin, a key molecule in the regulation of intercellular adhesion, was regulated by DcR3. The mRNA of E-cadherin was significantly downregulated by DcR3 treatment (Fig. 1C). The same effect was observed at the protein level (Fig. 1D). DcR3 expression also inhibited E-cadherin expression (Fig. 1E). Thus, DcR3 is a negative regulator of E-cadherin.

DcR3 and E-cadherin expression levels are inversely correlated in hepatocarcinoma cell lines and tissues

To understand the significance of the role of DcR3 in hepatocarcinoma, we detected DcR3 and E-cadherin expression levels in the normal liver cell line L02 and the hepatoma cell line HepG2 by performing immunofluorescent assays. The DcR3 expression level was higher in the HepG2 cells than that noted in the L02 cells (Fig. 2A). To confirm this observation, we detected the DcR3 expression using flow cytometry. The DcR3 expression level in HepG2 cells was higher than that in the L-02 cells (Fig. 2B). Protein and mRNA quantification showed that E-cadherin was downregulated in the HepG2 cells as compared with that in the L-02 cells (Fig. 2C and D). Importantly, immunohistochemical staining showed that DcR3 was almost undetectable in the non-tumor liver tissues (from patients with biliary tract disease) but was upregulated in the liver cancer tissues (Fig. 2E). Inversely, E-cadherin was located in the cell junction in non-tumor tissue, but was almost undetectable in liver cancer tissue (Fig. 2F). Therefore, hepatocarcinomas exhibited low E-cadherin expression but high DcR3 levels.

Figure 2. DcR3 and E-cadherin expression in hepatocarcinoma cell lines and tissues. DcR3 expression in L02 and HepG2 cells was determined using (A) immunofluorescence assay (scale bar, 50 µm) and (B) flow cytometry. E-cadherin protein or mRNA expression in L02 and HepG2 cells was determined using (C) western blot analysis and (D) qRT-PCR; *p<0.05. Representative images of (E) DcR3 expression in normal and liver carcinoma tissues and (F) E-cadherin. Scale bar, 200 µm.

DcR3 promotes cancer cell migration

Considering that E-cadherin plays a crucial role in cell adhesion and migration, we analyzed the effect of DcR3 on the migratory ability of HepG2 cells. The wound healing assay demonstrated that the addition of the DcR3-containing supernatant caused a strong increase in cell migration at 24 and 48 h (Fig. 3A and B). To confirm this trend, we detected the cell migratory ability using a Transwell assay. We observed that DcR3 greatly enhanced HepG2 cell migration (Fig. 3C-E). These results also indicated that DcR3 plays a positive role in cancer cell migration.

Figure 3. DcR3 treatment promotes cell migration. (A) Representative images of wound healing assay as described in Materials and methods. (B) Quantitative analysis of three independent experiments as presented in A; *p<0.05. (C) Representative images of the Transwell assay as described in the Materials and methods. (D and E) Quantitative analysis of three independent experiments as presented in C; *p<0.05, **p<0.01, ***p<0.001.

DcR3 induces p65 cytoplasm-nuclear translocation

To investigate whether DcR3 inhibits E-cadherin expression via its decoy function, we treated HepG2 cells by simultaneously adding DcR3 and its ligand FasL, LIGHT or TL1A, to the culture medium. The addition of these ligands did not affect the function of DcR3 in the regulation of E-cadherin expression (Fig. 4A). Thus, DcR3 inhibited E-cadherin expression via its non-decoy function.

Figure 4. DcR3 deregulates IκBα expression and induces p65 nuclear translocation. (A) HepG2 cells were treated with 3 µg/ml control IgG1 or 3 µg/ml DcR3-Fc alone or with 3 µg/ml FasL, LIGHT and TL1A. After 48 h, cells were harvested and E-cadherin expression was detected by western blot analysis. (B) HepG2 cells were treated with 3 µg/ml control IgG1 or 3 µg/ml DcR3-Fc. IκBα expression was detected by western blot analysis at 48 h after treatment. (C) HepG2 cells were treated as described in B, p65 expression in the cytoplasm and nucleus was determined by western blot analysis. p65 in the cytoplasm (C) or the nucleus (N) were normalized to lamin A/C or tubulin, respectively. Representative images from 2 independent experiments are shown. (D) Schematic illustration of the role of DcR3 in regulating cell migration.

To investigate the mechanism of the function of DcR3 in E-cadherin regulation, we explored the involved signaling pathway. DcR3 regulates the NFκB signaling pathway in monocytes. E-cadherin is also a target of p65 (26,27). Consequently, we investigated whether DcR3 activates NFκB signaling in HepG2 cells. DcR3 significantly downregulated IkBα expression (Fig. 4B). Furthermore, DcR3 treatment of HepG2 cells markedly increased the nuclear translocation of the NF-κB subunit p65 (Fig. 4C). These data suggest that the NFκB signaling cascade is an essential component in the involvement of E-cadherin expression for the DcR3-mediated migration response.

Discussion

DcR3 can be defined as a novel immunosuppressant on the basis of its neutralizing effects on FasL, LIGHT and TL1A. DcR3 is expressed by tumor cells from various lineages, including adenocarcinomas of the colon, rectum (28,29), lung (30) and gastric cancer (31), hepatocellular carcinoma (13,20) and in chronic liver diseases (19), which frequently lead to cancer formation. Increased DcR3 levels in serum or tissues were found to be correlated with poor prognosis and resistance to treatment in some cancer patients (13). In addition to its neutralizing effect, DcR3 also acts as an effector molecule to modulate cell function via non-decoy activities, including the regulation of DC and macrophage differentiation that leads to Th2 polarization (32,33), M2 macrophage differentiation (34), and cytoskeleton remodeling (16,17).

DcR3 can induce actin reorganization in human monocytes, and this protein triggers multiple signaling molecules, such as PKC and phosphatidylinositol 3-kinase (PI3K) (35). Furthermore, DcR3 induces NFκB-mediated expression of ICAM-1, VCAM-1 and IL-8 by monocytes; consequently, their binding to endothelial cells is enhanced (17). DcR3-Fc was found to act on THP-1 monocytes and differentiated macrophages to increase the expression level of integrin α4. Thus, cell aggregation and proliferation are promoted and apoptosis is reduced (18). DcR3 is upregulated in cancer cells; thus, these observations suggest its important roles in modulating the migration and trafficking of monocytes/macrophages in the tumor microenvironment.

E-cadherin plays an important role in cell adhesion by forming adherent junctions to bind cells within tissues. The loss of E-cadherin expression has been defined as a hallmark of EMT. EMT is a process by which epithelial cells lose their cell polarity and cell-cell adhesion, and they gain migratory and invasive properties to become mesenchymal stem cells. DcR3 was found to induce IκB kinase activation, IκB degradation, and p65 nuclear translocation in human microvascular endothelial cells (17). E-cadherin is a target of p65, which represses E-cadherin expression and enhances the epithelial-to-mesenchymal transition of mammary epithelial cells via ZEB-1 and ZEB-2 (36). In addition, TGF-β is one of the most critical factors involved in EMT regulation. TGF-β promotes EMT through both Smad-dependent and Smad-independent manner (37). The relationship between DcR3 and the TGF-β pathway will be the research of interest in a future study. A model of the function of DcR3 in cell migration regulation is shown in Fig. 4D. DcR3 controls the expression of E-cadherin and the p65 translocation of HepG2 cells. Thus, it elicits double effects in tumor metastasis regulation and immune modulation. The blocking of DcR3 may be applied as an effective therapeutic strategy to prevent tumor metastasis.

In conclusion, the present study demonstrated that DcR3 is overexpressed in hepatic carcinoma tissues and cell lines. DcR3-Fc treatment inhibited E-cadherin expression, enhanced tumor cell migration in vitro, and promoted p65 nuclear translocation. These findings revealed the mechanism underlying the ability of DcR3 to regulate cell migration. Therefore, DcR3 may be a potential target for the gene therapy of hepatic carcinoma.

Acknowledgements

The present study was supported by the Shenzhen Basic Research Program (JCYJ20150630114942293, JCYJ20140416180228582 and JCYJ20160229201353324), the Nature Science Foundation of China for Young Scholar grant (81501356), the Shenzhen Peacock Next-generation Monoclonal Antibody Drug Research and Development Program (1110140040347265), the Fourth Group of Talents in Guangdong Province (2014–1), the Shenzhen Engineering Laboratory (2014–1677), and the Shenzhen Technology Study program (JSGG20160229202150023).

References