Connective tissue growth factor (CTGF) is a matricellular protein related to hepatic fibrosis. This study aims to clarify the roles of CTGF in hepatocellular carcinoma (HCC), which usually develops from fibrotic liver. CTGF was overexpressed in 93 human HCC compared with nontumorous tissues, primarily in tumor cells. Increased CTGF expression was associated with clinicopathologic malignancy of HCC. CTGF was upregulated in hepatoma cells in hepatocyte-specific Kras-mutated mice (Alb-Cre KrasLSL-G12D/+). Hepatocyte-specific knockout of CTGF in these mice (Alb-Cre KrasLSL-G12D/+ CTGFfl/fl) decreased liver tumor number and size. Hepatic stellate cells (HSC) were present in both human and murine liver tumors, and α-SMA expression, a marker of HSC activation, positively correlated with CTGF expression. Forced expression of CTGF did not affect growth of PLC/PRF/5 cells, a hepatoma cell line with little CTGF expression, but facilitated their growth in the presence of LX-2 cells, an HSC line. The growth of HepG2 cells, which express high levels of CTGF, was promoted by coculture with LX-2 cells compared with monoculture. Growth promotion by LX-2 cells was negated by an anti-CTGF antibody in both culture and xenografts. Coculturing LX-2 cells with HepG2 cells drove LX-2-derived production of IL6, which led to STAT-3 activation and proliferation of HepG2 cells. An anti-CTGF antibody reduced IL6 production in LX-2 cells and suppressed STAT-3 activation in HepG2 cells. In conclusion, our data identify tumor cell–derived CTGF as a keystone in the HCC microenvironment, activating nearby HSC that transmit progrowth signals to HCC cells, and this interaction is susceptible to inhibition by an anti-CTGF antibody.

Significance: Protumor cross-talk between cancer cells and hepatic stellate cells presents an opportunity for therapeutic intervention against HCC.

Graphical Abstract:http://cancerres.aacrjournals.org/content/canres/78/17/4902/F1.large.jpg. Cancer Res; 78(17); 4902–14. ©2018 AACR.

Connective tissue growth factor (CTGF), also known as CCN2, is a member of the CCN (CCN1-6) family proteins (1). CTGF is a secreted matricellular protein that interacts with various molecules in the extracellular matrix (ECM). CTGF contains an N-terminal secretory peptide followed by four conserved domains with sequence homologies to insulin-like growth factor-binding proteins, the von Willebrand factor C (VWC) domain, a motif related to thrombospondin, and a C-terminal domain that contains a cysteine-knot motif (1–3). CTGF has various biological functions, including cell adhesion, migration, proliferation, differentiation, and ECM production, and participates in the development of many organs under normal physiologic conditions (2). CTGF is pathologically viewed as a central mediator of tissue remodeling and fibrosis of various organs, including the lung, heart, liver, and kidney (1–4). In addition to fibrotic diseases, CTGF has also been reported to be associated with the progression of various malignant diseases, such as breast, pancreatic, and gastric cancers (5–8).

Hepatocellular carcinoma (HCC) is a major cancer type and is the third most common cause of cancer-related mortality worldwide (9). One study has reported that this is not the case, but in general, it has been reported that CTGF is highly expressed in HCC tissues (10–12). CTGF is expressed in several cell types in the liver, including hepatocytes, cholangiocytes, hepatic stellate cells (HSC), fibroblasts, and sinusoidal endothelial cells (4, 8, 10, 13–15). The types of cells among HCC tissues that express CTGF have not been clarified. CTGF increases the proliferation of cancer-associated fibroblasts (10) and the migration of macrophages (14). In addition, it has been reported that CTGF acts directly on hepatoma cells and increases DNA synthesis (11), cell-cycle progression (13), invasion and migration abilities (13), and resistance to doxorubicin and TRAIL-induced apoptosis (11). However, these reported actions were mostly based on in vitro experiments, and the significance of CTGF in vivo is completely unknown.

Here, we clarified that CTGF was upregulated in HCC compared with surrounding nontumorous tissues and that CTGF is expressed primarily in cancer cells. Using a genetically engineered mouse model, a hepatocyte-specific knockout of CTGF suppressed the activation of HSCs and the progression of liver cancer. Through the specific inhibition of CTGF function with an anti–CTGF-neutralizing antibody, which is currently being tested in clinical trials for other diseases (3, 16, 17), we further demonstrated that the growth-promoting effect of CTGF is mediated by tumor–stroma interactions between cancer cells and HSCs. This report provides the first demonstration that CTGF produced in cancer cells activates HSCs in the tumor microenvironment and accelerates the progression of liver cancer.

Human samples

Liver samples were collected from both tumorous and nontumorous liver tissues of patients with HCC who underwent hepatectomy. Written informed consent was obtained from all patients. The study protocol conformed to the ethical guidelines of the Declaration of Helsinki. Approval for the use of resected samples was obtained from the Institutional Review Board (IRB) Committees at Osaka University Hospital (IRB No. 13556 and 15267).

Mice

C57BL/6/129 background mice carrying a lox P-stop-lox P (LSL) termination sequence with the KrasG12D point mutation allele (KrasLSL-G12D/+) were kindly provided by the National Cancer Institute (Bethesda, MD). Hepatocyte-specific Kras-mutated mice (KrasLSL-G12D/+Alb-Cre; KrasG12D mice) were generated by mating KrasLSL-G12D/+ mice and heterozygous Alb-Cre transgenic mice that expressed the Cre recombinase gene under the promoter of the albumin gene (4). C57BL/6/J background mice that carried two floxed CTGF alleles (CTGFfl/fl) have been previously described (18). The CTGFfl/fl mice were genotyped using the following primers for the ctgf allele: 5′-ACAATGACATCTTTGAGTCC-3′ and 5′-AGTCTAATGAGTTCGTGTCC-3′. We generated hepatocyte-specific CTGF-knockout Kras-mutated mice by mating KrasG12D mice and CTGF-floxed mice, and littermates generated were used to compare the phenotypes among experimental groups. NOD/Shi-scid/IL-2Rγ (null) (NOG) mice have been previously described (19). Only male mice were used in the experiments. The Institute of Experimental Animal Sciences of Osaka University Graduate School of Medicine specifically approved these studies, and the mice were maintained in a specific pathogen-free facility and were treated humanely.

Cell culture

The human hepatoma cell lines HepG2, Huh7, PLC/PRF/5, and HLF were obtained from the JCRB/HSRRB cell bank in 2012, 2015, 2015, and 2011, respectively. The human HSC line LX-2 was kindly provided by professor Eiji Miyoshi (Department of Molecular Biochemistry and Clinical Investigation, Osaka University Graduate School of Medicine, Osaka, Japan) in 2015, and the cell authentication was performed by the JCRB/HSRRB cell bank. Mycoplasma testing was performed for all cell lines using the MycoAlert Mycoplasma Detection Kit according to the manufacturer's recommended protocols. The latest date the cells were tested is December 5, 2017. The cells were cultured at 37°C with 5% CO2 in DMEM (Sigma-Aldrich) supplemented with 10% fetal calf serum and antibiotics unless otherwise indicated. All experiments were performed using cells with the passage number of less than 30. The Transwell insert system (Corning) was used in the coculture experiments.

Cell proliferation was analyzed via the WST-8 assay (Nacalai Tesque). LY294002, a PI3K inhibitor, FR180204, an extracellular signal-regulated kinase (Erk) inhibitor, and U0126, a mitogen-activated protein kinase/Erk kinase (Mek) inhibitor, were purchased from Sigma-Aldrich. Recombinant human EGF protein was purchased from Thermo Fisher Scientific. Recombinant human CTGF protein was obtained from FibroGen. Anti–IL6-neutralizing antibody was purchased from Thermo Fisher Scientific. Kras siRNA, IL6 siRNA, CTGF siRNA, STAT-3 siRNA, and control siRNA (Thermo Fisher Scientific) were transfected into the cells using Lipofectamine RNAiMAX (Thermo Fisher Scientific) according to the reverse transfection protocol. The CTGF cDNA plasmid vector or the control empty plasmid vector (OriGene) was transfected using Lipofectamine 3000 (Thermo Fisher Scientific) according to the manufacturer's recommended protocols. To establish a cell line that stably overexpresses CTGF, transfected cells were selected over 4 weeks with G418 (Nacalai Tesque) treatment, and several single colonies were isolated after selection. The expression levels of CTGF were compared among these colonies, and the one with the highest CTGF expression was used in the subsequent experiments. CTGF shRNA plasmid (Santa Cruz Biotechnology) or control shRNA plasmid (Santa Cruz Biotechnology) was transfected according to the manufacturer's protocol. Puromycin (Thermo Fisher Scientific) treatment was used to select the stably transfected cells.

Anti–CTGF-neutralizing antibody

The fully recombinant human monoclonal IgG1 anti-CTGF antibody FG-3019 and a nonspecific human IgG control antibody (hIgG) were kindly provided by FibroGen.

Experimental protocol for xenograft tumor models

Under general anesthesia, 7- to 9-week-old male NOG mice were subcutaneously inoculated in the bilateral flank portions with 1 × 107 hepatoma cells with or without 1 × 107 LX-2 cells suspended in Matrigel (Corning). To evaluate the effect of CTGF inhibition, 40 mg/kg FG-3019 or hIgG was intraperitoneally administered twice per week from the day of inoculation. The length and width of the xenograft tumors were measured twice per week, and the tumor volume was calculated according to the following formula: tumor volume = 1/2 × (major axis) × (minor axis)2 (20).

Histologic analyses

Liver sections were routinely stained with hematoxylin and eosin. Immunohistochemical analyses were performed using paraffin-embedded liver sections and a CTGF antibody (Santa Cruz Biotechnology), an α-SMA antibody (Abcam), and a PCNA antibody (Cell Signaling Technology). PCNA-positive cells were counted in high-power fields (HPF) at 400-fold magnification. Positive areas of α-SMA staining within the liver sections were quantified as the α-SMA area divided by total tissue area under an HPF at 400-fold magnification using the image-analysis software WinROOF (Mitani Corporation), as previously described (21–23). Tumor area/total tissue area ratio in macroscopically nontumorous liver tissues was quantified at 40-fold magnification using WinROOF (Mitani Corporation). Double immunofluorescence staining was performed using frozen liver sections and a CTGF antibody conjugated to Alexa Fluor 488 (Santa Cruz Biotechnology) and an α-SMA antibody conjugated to Alexa Fluor 594 (Santa Cruz Biotechnology). We used 4′,6-diamidino-2-phenylindole to label the nuclei.

RNA isolation and quantitative real-time reverse-transcription PCR

Total RNA was extracted from cell lines or liver tissues using the RNeasy Mini Kit (QIAGEN) according to the manufacturer's recommended protocols, and cDNA was produced by reverse transcription as previously described (21). The mRNA expression of specific genes was quantified by real-time RT-PCR using TaqMan Gene Expression Assays (Applied Biosystems) as follows: mouse-ctgf (Mm01192933_g1), mouse-afp (Mm00431715_m1), mouse-glypican-3 (Mm00516722_m1), mouse-acta2 (Mm00725412_s1), mouse-β-actin (Mm02619580_g1), human-ctgf (Hs00170014_m1), human-kras (Hs00364284_g1), human-β-actin (Hs01060665_g1), human-il 6 (Hs00985639_m1), and human-acta2 (Hs00426835_g1). All expression levels were normalized to those of β-actin.

Western blot analysis

Liver tissue was lysed in lysis buffer [1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate (SDS), protease inhibitor cocktail (Nacalai Tesque), phosphatase inhibitor cocktail (Nacalai Tesque), and PBS pH 7.4]. Equal amounts of protein were electrophoretically separated using SDS polyacrylamide gels and transferred onto a polyvinylidene fluoride membrane. The following antibodies were used for immunodetection: anti-Ras, anti–phospho-Erk, anti–phospho-Akt, anti–phospho-STAT-3, anti–β-actin (Cell Signaling Technology), anti–α-SMA (Abcam), and anti-CTGF (Santa Cruz Biotechnology). ImageJ software (NIH, Bethesda, MD) was used to quantify the bands, and the expression levels were normalized to those of β-actin.

ELISA

Commercial ELISA kits for CTGF (PeproTech) and IL6 (R&D systems) were used to quantify their concentrations in cell culture supernatants according to the manufacturer's recommended protocols.

Statistical analysis

Differences between unpaired groups with normal or nonnormal distributions were compared using the Student t test or the Mann–Whitney U test, respectively. The Wilcoxon signed-rank test was used to compare two paired samples. Correlations were assessed using the Pearson product–moment correlation coefficient. The χ2 test was used to analyze categorical data. Parametric or nonparametric multiple comparisons were performed using one-way ANOVA, followed by the Tukey–Kramer post hoc test or Kruskal–Wallis test, followed by the Steel–Dwass test, respectively. P values less than 0.05 were considered to indicate statistical significance.

CTGF is highly expressed in human HCC tissues and is related to the malignant characteristics of HCC

The CTGF gene expression levels in tumorous tissues and surrounding nontumorous tissues were compared among 93 human liver samples from patients who underwent surgical hepatectomy for HCC (Supplementary Table S1). CTGF gene expression was significantly upregulated in tumorous tissues compared with nontumorous tissues (Fig. 1A). A Western blot analysis also showed upregulation of CTGF protein in tumorous tissues (Fig. 1B). Positive immunohistochemical staining for CTGF was observed primarily in the cytoplasm of hepatoma cells (Fig. 1C). Immunohistochemistry using serial sections revealed AFP expression in CTGF-positive cells in tumor tissues (Supplementary Fig. S1A). In tumor tissues, α-SMA–positive HSCs were detected by immunohistochemistry, and α-SMA mRNA was upregulated in these tissues compared with nontumorous liver tissues (Fig. 1D). Alpha-SMA expressions in tumorous tissues were positively correlated with that of CTGF (r = 0.21, P < 0.05). In double immunofluorescence staining, α-SMA–positive cells expressed CTGF, but comprised only a small fraction of the total CTGF expression in the tumor (Supplementary Fig. S1B). We further investigated the association between CTGF gene expression levels and tumor characteristics. According to the median value of the ratio of CTGF mRNA expression in tumorous tissues to nontumorous tissues, we divided 93 patients into a CTGF high-expression group (CTGF mRNA expression ratio > 1.3, N = 46) and a low-expression group (CTGF mRNA expression ratio ≤ 1.3, N = 47) and compared the tumor characteristics between the two groups. The CTGF high-expression group showed higher positive rates of des-γ-carboxy prothrombin, tumor multiplicity, portal invasion, and malignant macroscopic tumor classification, which suggests more malignant characteristics of HCC (Table 1). In addition, the CTGF high-expression group exhibited higher α-SMA expression in tumorous tissues (Fig. 1E).

Figure 1.

CTGF is upregulated in human HCC, and CTGF mRNA expression levels are associated with those of α-SMA, a marker of HSC activation. Liver tissues were obtained from patients with HCC who underwent surgical hepatectomy. A, CTGF mRNA expression levels in tumorous tissues (T) and nontumorous tissues (NT; N = 93). B, Western blot analysis for the expression of CTGF in the livers of representative 5 patients with HCC and CTGF protein expression in the tumorous and nontumorous tissues (N = 25). C, Immunohistochemistry for CTGF expression in HCC. A representative image from 10 HCC samples is presented. D, Immunohistochemistry for α-SMA in HCC and α-SMA mRNA expression levels in tumorous tissues and nontumorous liver tissues (N = 93). A representative immunohistochemical image from four HCC samples is shown. E, α-SMA mRNA expression levels in tumorous tissues in the CTGF low-expression group and in the high-expression group. CTGF-High, T/NT CTGF expression ratio > 1.3 (N = 46); CTGF-Low, T/NT CTGF expression ratio ≤ 1.3 (N = 47). The Wilcoxon signed-rank test was used to compare the expression levels between tumorous and nontumorous tissues in each patient (A, B, and D). The horizontal bars denote medians. *, P < 0.05.

Figure 1.

CTGF is upregulated in human HCC, and CTGF mRNA expression levels are associated with those of α-SMA, a marker of HSC activation. Liver tissues were obtained from patients with HCC who underwent surgical hepatectomy. A, CTGF mRNA expression levels in tumorous tissues (T) and nontumorous tissues (NT; N = 93). B, Western blot analysis for the expression of CTGF in the livers of representative 5 patients with HCC and CTGF protein expression in the tumorous and nontumorous tissues (N = 25). C, Immunohistochemistry for CTGF expression in HCC. A representative image from 10 HCC samples is presented. D, Immunohistochemistry for α-SMA in HCC and α-SMA mRNA expression levels in tumorous tissues and nontumorous liver tissues (N = 93). A representative immunohistochemical image from four HCC samples is shown. E, α-SMA mRNA expression levels in tumorous tissues in the CTGF low-expression group and in the high-expression group. CTGF-High, T/NT CTGF expression ratio > 1.3 (N = 46); CTGF-Low, T/NT CTGF expression ratio ≤ 1.3 (N = 47). The Wilcoxon signed-rank test was used to compare the expression levels between tumorous and nontumorous tissues in each patient (A, B, and D). The horizontal bars denote medians. *, P < 0.05.

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Table 1.

Tumor characteristics of the CTGF high-expression and low-expression groups

CharacteristicsCTGF Low-expression groupCTGF High-expression groupP value
AFP (negative/positive) 24/23 15/31 0.074 
DCP (negative/positive) 16/31 7/39 0.024 
Tumor number (1–5/6+) 43/4 34/12 0.025 
Maximum tumor diameter (cm) 3.8 (0.8–20.0) 4.0 (1.0–18.0) 0.48 
Macroscopic tumor classification (SN-IM/SN/SN-EG/CMN/IF) 1/20/15/5/6 0/12/12/11/11 0.021 
Portal invasion (negative/positive) 40/7 31/15 0.044 
Intrahepatic metastasis (negative/positive) 35/12 26/20 0.069 
TNM stage (I/II/III/IVa/IVb) 8/24/7/8/0 4/18/14/9/1 0.082 
CharacteristicsCTGF Low-expression groupCTGF High-expression groupP value
AFP (negative/positive) 24/23 15/31 0.074 
DCP (negative/positive) 16/31 7/39 0.024 
Tumor number (1–5/6+) 43/4 34/12 0.025 
Maximum tumor diameter (cm) 3.8 (0.8–20.0) 4.0 (1.0–18.0) 0.48 
Macroscopic tumor classification (SN-IM/SN/SN-EG/CMN/IF) 1/20/15/5/6 0/12/12/11/11 0.021 
Portal invasion (negative/positive) 40/7 31/15 0.044 
Intrahepatic metastasis (negative/positive) 35/12 26/20 0.069 
TNM stage (I/II/III/IVa/IVb) 8/24/7/8/0 4/18/14/9/1 0.082 

NOTE: The cutoff values for AFP and DCP are 20 ng/mL and 40 mAU/mL, respectively. Maximum tumor diameter is presented as the medians (ranges).

Abbreviations: AFP, α-fetoprotein; CMN, confluent multinodular type; DCP, des-γ-carboxy prothrombin; IF, infiltrative type; SN, simple nodular type; SN-EG, simple nodular type with extranodular growth; SN-IM, small nodular type with indistinct margin.

CTGF is upregulated in liver tumors of KrasG12D mice

We examined the relationship between liver tumors and CTGF using genetically modified mice that develop liver tumors. KrasG12D mice developed liver tumors at high rates after 4 months of age, and the number and diameter of the tumors increased with age (Fig. 2A and B). In the liver tissues of KrasG12D mice, Ras protein accumulation was confirmed, and the Ras signaling pathway was activated, as assessed by increased Erk and Akt phosphorylation (Fig. 2C). The expression of p-Erk was more prominent in tumors than in nontumorous tissues from the KrasG12D mice. A gene expression analysis revealed that CTGF and tumor markers for HCC, such as AFP and glypican-3, were significantly upregulated in tumors compared with nontumorous tissues from control mice and KrasG12D mice (Fig. 2D). A Western blot analysis showed that CTGF was overexpressed in tumors from KrasG12D mice compared with nontumorous tissues from KrasG12D mice and control mice (Fig. 2C). Immunohistochemical staining for CTGF demonstrated expression primarily in hepatoma cells (Fig. 2E). An analysis of immunohistochemical staining in serial sections showed a positive reaction for AFP in CTGF-positive cells in tumor tissues (Supplementary Fig. S1C). Immunohistochemistry also revealed α-SMA–positive HSCs in liver tumors (Fig. 2F). Alpha-SMA was upregulated in tumorous tissues compared with nontumorous liver tissues from control mice and KrasG12D mice (Fig. 2C and F). In tumorous tissues, α-SMA gene expression was positively correlated with that of CTGF (Fig. 2F). Double immunofluorescence staining α-SMA and CTGF suggested that although α-SMA–positive cells were also positive for CTGF, CTGF derived from α-SMA–positive cells accounted for only a small fraction of the total CTGF expressed in the tumor (Supplementary Fig. S1D).

Figure 2.

Hepatocyte-specific Kras-mutated mice (KrasG12D mice) develop liver tumors in which CTGF is upregulated similarly to human HCC. KrasG12D mice and their littermate Alb-Cre transgenic control mice were sacrificed at the indicated age in months. A, Representative images of the livers of 9-month-old mice. B, Macroscopic tumor incidence rate, tumor number, and maximum tumor diameter according to age in months (N = 7–22 for each group). C, Western blot analysis for the expression of Ras, phospho-Erk, phospho-Akt, CTGF, α-SMA, and β-actin in the livers of 9- to 11-month-old mice. D, Gene expression levels of glypican-3, AFP, and CTGF in tumorous (T) liver tissues and nontumorous (NT) liver tissues from KrasG12D mice and control mice from 9 to 11 months of age (N = 6–12 per group). E, Immunohistochemistry for CTGF in the liver tissues of 9-month-old mice. A representative image from five samples is presented. F, Immunohistochemistry for α-SMA in liver tumors, α-SMA mRNA expression levels in tumorous liver tissues and nontumorous liver tissues in KrasG12D mice and control mice (N = 7–10 per group), and the correlation between α-SMA mRNA expression levels and CTGF mRNA expression levels in tumorous tissues from KrasG12D mice (N = 20). Representative immunohistochemical images from five samples are shown. In a box plot, box denotes 25th and 75th percentiles, and top or bottom whiskers indicate the values higher than 75th percentiles or lower than 25th percentiles, respectively. In bar graphs, bars and top whiskers represent mean values and SDs, respectively. *, P < 0.05.

Figure 2.

Hepatocyte-specific Kras-mutated mice (KrasG12D mice) develop liver tumors in which CTGF is upregulated similarly to human HCC. KrasG12D mice and their littermate Alb-Cre transgenic control mice were sacrificed at the indicated age in months. A, Representative images of the livers of 9-month-old mice. B, Macroscopic tumor incidence rate, tumor number, and maximum tumor diameter according to age in months (N = 7–22 for each group). C, Western blot analysis for the expression of Ras, phospho-Erk, phospho-Akt, CTGF, α-SMA, and β-actin in the livers of 9- to 11-month-old mice. D, Gene expression levels of glypican-3, AFP, and CTGF in tumorous (T) liver tissues and nontumorous (NT) liver tissues from KrasG12D mice and control mice from 9 to 11 months of age (N = 6–12 per group). E, Immunohistochemistry for CTGF in the liver tissues of 9-month-old mice. A representative image from five samples is presented. F, Immunohistochemistry for α-SMA in liver tumors, α-SMA mRNA expression levels in tumorous liver tissues and nontumorous liver tissues in KrasG12D mice and control mice (N = 7–10 per group), and the correlation between α-SMA mRNA expression levels and CTGF mRNA expression levels in tumorous tissues from KrasG12D mice (N = 20). Representative immunohistochemical images from five samples are shown. In a box plot, box denotes 25th and 75th percentiles, and top or bottom whiskers indicate the values higher than 75th percentiles or lower than 25th percentiles, respectively. In bar graphs, bars and top whiskers represent mean values and SDs, respectively. *, P < 0.05.

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The Ras/Mek/Erk pathway regulates CTGF expression in hepatoma cells

We examined the relationship between the Ras signaling pathway and CTGF expression through in vitro and in silico analyses. The stimulation of Huh7 cells, which are Kras wild-type hepatoma cells, with EGF activated Erk and Akt downstream of Ras and increased gene expression and secretion of CTGF (Supplementary Fig. S2A). In HepG2 cells, which contain mutant Kras, CTGF was downregulated by siRNA-mediated Kras knockdown (Supplementary Fig. S2B). PI3K inhibition and Mek/Erk inhibition specifically downregulated p-Akt and p-Erk, respectively (Supplementary Fig. S2C). CTGF expression in HepG2 cells was also decreased by Mek and Erk inhibitors but not by a PI3K inhibitor (Supplementary Fig. S2D) 6 hours after their addition, whereas the cell viability was not affected (Supplementary Fig. S2D). Furthermore, to investigate an association between CTGF gene expression levels and the activity of the Ras/Mek/Erk pathway in patients with HCC, we performed a single-sample gene set enrichment analysis (ssGSEA) using a publicly available microarray dataset of patients with HCC. The ssGSEA revealed a positive correlation between CTGF expression and Ras/Mek/Erk pathway activity in the liver (Supplementary Fig. S2E). These results suggest that Ras/Mek/Erk pathway activation is associated with CTGF upregulation in human liver tissue.

Hepatocyte-specific CTGF deficiency inhibits tumor development and progression in KrasG12D mice

To analyze the functions of CTGF in HCC, we generated hepatocyte-specific CTGF-deficient KrasG12D mice and compared the phenotypes of three groups at 8 months: (1) KrasG12D CTGF+/+ mice (CTGF+/+ KrasLSL-G12D/+Alb-Cre), (2) KrasG12D CTGF+/− mice (CTGFfl/+ KrasLSL-G12D/+ Alb-Cre), and (3) KrasG12D CTGF−/− mice (CTGFfl/fl KrasLSL-G12D/+ Alb-Cre). The incidence rate of macroscopic liver tumors was lower in KrasG12D CTGF−/− mice than in KrasG12D CTGF+/+ mice, although this difference was not statistically significant (Fig. 3A and B). Importantly, KrasG12D CTGF−/− mice showed a significant reduction in the number of macroscopic tumors, tumor size, liver/body weight ratio, and histologic tumor/nontumor area ratio compared with KrasG12D CTGF+/+ mice (Fig. 3B and C). The number of PCNA-positive cancer cells in these liver tumors was lower in KrasG12D CTGF−/− mice than in KrasG12D CTGF+/+ mice, which suggests the attenuation of tumor proliferation in KrasG12D CTGF−/− mice (Fig. 3D). Gene and protein expression of CTGF in these liver tumors was decreased in KrasG12D CTGF−/− mice compared with KrasG12D CTGF+/+ mice (Fig. 3E). The number of positive areas of immunohistochemical staining for α-SMA was decreased in liver tumors in KrasG12D CTGF−/− mice compared with KrasG12D CTGF+/+ mice (Fig. 3F). KrasG12D CTGF+/− mice exhibited intermediate phenotypes (Fig. 3). These results indicate that CTGF produced by hepatoma cells is involved in the development and progression of liver tumors, as well as the activation of HSCs in tumorous tissues.

Figure 3.

Hepatocyte-specific CTGF deficiency suppresses the development and progression of liver tumors in KrasG12D mice. KrasG12D CTGF+/+, KrasG12D CTGF+/−, and KrasG12D CTGF−/− littermates were sacrificed at 8 months of age, and the tumor characteristics were compared among these three groups. A, Representative images of the livers. B, Macroscopic tumorigenesis rate, tumor number, and maximum tumor diameter (N = 11–15 per group). C, Representative images of hematoxylin and eosin staining (×40) of the macroscopically nontumorous liver tissues and quantified tumor area/total tissue area ratio (N = 7–8 per group). Arrows, tumors. D, Representative immunohistochemistry images for PCNA (×400) and the number of PCNA-positive cells per HPF in liver tumors (N = 8–10 per group). E, Gene and protein expression levels of CTGF in tumorous liver tissues (N = 8–10 per group). F, Representative immunohistochemistry images for α-SMA (×400) and α-SMA–positive areas per HPF in liver tumors (N = 10–13 per group). In a box plot, box represents 25th and 75th percentiles, and top or bottom whiskers indicate the values higher than 75th percentiles or lower than 25th percentiles, respectively. In bar graphs, bars and top whiskers indicate mean values and SDs, respectively. *, P < 0.05.

Figure 3.

Hepatocyte-specific CTGF deficiency suppresses the development and progression of liver tumors in KrasG12D mice. KrasG12D CTGF+/+, KrasG12D CTGF+/−, and KrasG12D CTGF−/− littermates were sacrificed at 8 months of age, and the tumor characteristics were compared among these three groups. A, Representative images of the livers. B, Macroscopic tumorigenesis rate, tumor number, and maximum tumor diameter (N = 11–15 per group). C, Representative images of hematoxylin and eosin staining (×40) of the macroscopically nontumorous liver tissues and quantified tumor area/total tissue area ratio (N = 7–8 per group). Arrows, tumors. D, Representative immunohistochemistry images for PCNA (×400) and the number of PCNA-positive cells per HPF in liver tumors (N = 8–10 per group). E, Gene and protein expression levels of CTGF in tumorous liver tissues (N = 8–10 per group). F, Representative immunohistochemistry images for α-SMA (×400) and α-SMA–positive areas per HPF in liver tumors (N = 10–13 per group). In a box plot, box represents 25th and 75th percentiles, and top or bottom whiskers indicate the values higher than 75th percentiles or lower than 25th percentiles, respectively. In bar graphs, bars and top whiskers indicate mean values and SDs, respectively. *, P < 0.05.

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Forced CTGF expression does not affect hepatoma cell growth but increases their growth in the presence of HSCs

To examine the influence of CTGF on the proliferation of hepatoma cells, we established CTGF-overexpressing PLC/PRF/5 cells, which originally showed the lowest endogenous levels of CTGF expression and secretion among several hepatoma cell lines (Huh7, HLF, HepG2, PLC/PRF/5, and Hep3B; Supplementary Fig. S3A and S3B). The in vitro growth of CTGF-overexpressing PLC/PRF/5 cells was not different from that of the parental or mock-transfected PLC/PRF/5 cells (Fig. 4A). After PLC/PRF/5 cells were xenografted into NOG mice, the growth was not found to be different between tumors derived from CTGF-overexpressing PLC/PRF/5 cells and those derived from mock-transfected PLC/PRF/5 cells, which is similar to the results obtained from the in vitro experiments (Fig. 4B). We subsequently investigated the influence of CTGF on the proliferation of hepatoma cells in the presence of HSCs. During the coculture of a human HSC cell line with LX-2 cells in a Transwell system, CTGF-overexpressing PLC/PRF/5 cells proliferated faster than mock-transfected PLC/PRF/5 cells (Fig. 4C). In a xenograft model, the tumor volumes of CTGF-overexpressing PLC/PRF/5 cell–derived tumors were significantly larger than those of mock-transfected PLC/PRF/5 cell–derived tumors when the cells were coinjected with LX-2 cells (Fig. 4D). Although α-SMA–positive cells were present along with the fibrous bands in xenograft tumors mixed with LX-2 cells (Fig. 4E), predominant cell population in tumors was composed of PLC/PRF/5 cells. It is therefore suggested that CTGF contributes to hepatoma cell growth in the presence of HSCs both in vitro and in xenograft models.

Figure 4.

Forced expression of CTGF does not affect the growth of PLC/PRF/5 cells alone but increases their growth in the presence of LX-2 cells. The growth of PLC/PRF/5 cells was evaluated in vitro using a WST-8 assay or xenograft tumor models at the indicated time points. For xenograft models, mock-transfected (PLC/PRF/5-mock) or CTGF-overexpressing PLC/PRF/5 cells (PLC/PRF/5-CTGF), with or without LX-2 cells, were subcutaneously injected into the left and right flanks of NOG mice. The mice were sacrificed, and xenograft tumors were enucleated 36 days after inoculation. Sequential tumor volume and representative images of the xenograft tumors are presented. A, Growth of parental, mock-transfected (PLC/PRF/5-mock), or CTGF-overexpressing PLC/PRF/5 cells (PLC/PRF/5-CTGF). B, Xenograft model of mock-transfected or CTGF-overexpressing PLC/PRF/5 cells. C, Growth of mock-transfected PLC/PRF/5 cells or CTGF-overexpressing PLC/PRF/5 cells in coculture with LX-2 cells. Mock-transfected PLC/PRF/5 cells or CTGF-overexpressing PLC/PRF/5 cells were cocultured with the same number of LX-2 cells in a Transwell system. D, Xenograft model of mock-transfected or CTGF-overexpressing PLC/PRF/5 cells mixed with the same number of LX-2 cells. E, Immunohistochemistry for α-SMA in xenograft tumors composed of mock-transfected or CTGF-overexpressing PLC/PRF/5 cells and LX-2 cells. Arrowheads, α-SMA–positive cells. N = 4 (A and C) or 9 (B and D) per group. In line graphs, plots with whiskers indicate mean values ± SDs. *, P < 0.05 vs. the control.

Figure 4.

Forced expression of CTGF does not affect the growth of PLC/PRF/5 cells alone but increases their growth in the presence of LX-2 cells. The growth of PLC/PRF/5 cells was evaluated in vitro using a WST-8 assay or xenograft tumor models at the indicated time points. For xenograft models, mock-transfected (PLC/PRF/5-mock) or CTGF-overexpressing PLC/PRF/5 cells (PLC/PRF/5-CTGF), with or without LX-2 cells, were subcutaneously injected into the left and right flanks of NOG mice. The mice were sacrificed, and xenograft tumors were enucleated 36 days after inoculation. Sequential tumor volume and representative images of the xenograft tumors are presented. A, Growth of parental, mock-transfected (PLC/PRF/5-mock), or CTGF-overexpressing PLC/PRF/5 cells (PLC/PRF/5-CTGF). B, Xenograft model of mock-transfected or CTGF-overexpressing PLC/PRF/5 cells. C, Growth of mock-transfected PLC/PRF/5 cells or CTGF-overexpressing PLC/PRF/5 cells in coculture with LX-2 cells. Mock-transfected PLC/PRF/5 cells or CTGF-overexpressing PLC/PRF/5 cells were cocultured with the same number of LX-2 cells in a Transwell system. D, Xenograft model of mock-transfected or CTGF-overexpressing PLC/PRF/5 cells mixed with the same number of LX-2 cells. E, Immunohistochemistry for α-SMA in xenograft tumors composed of mock-transfected or CTGF-overexpressing PLC/PRF/5 cells and LX-2 cells. Arrowheads, α-SMA–positive cells. N = 4 (A and C) or 9 (B and D) per group. In line graphs, plots with whiskers indicate mean values ± SDs. *, P < 0.05 vs. the control.

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The growth-promoting effect of HSCs on hepatoma cells with high CTGF expression is abolished by inhibition of CTGF

In HepG2 cells, which showed the highest endogenous expression and secretion of CTGF (Supplementary Fig. S3A), the application of an anti–CTGF-neutralizing antibody FG-3019 did not affect cell growth (Fig. 5A). HepG2 cells grew faster when cocultured with LX-2 cells than when cultured alone (Fig. 5B). The proliferation of HepG2 cells was facilitated by coculture with LX-2 cells when the cells were exposed to control hIgG, but the growth-promoting effect of the coculture with LX-2 cells was abolished after exposure to an anti–CTGF-neutralizing antibody (Fig. 5C). In a xenograft model, the volumes of tumors derived from HepG2 cells coinjected with LX-2 cells were larger than those derived from HepG2 cells injected alone in the hIgG group (Fig. 5D). In contrast, in the anti–CTGF-neutralizing antibody group, the volumes of tumors derived from HepG2 cells did not increase even when these cells were coinjected with LX-2 cells (Fig. 5D). Similar to the experiments with an anti–CTGF-neutralizing antibody, hepatoma cell–specific CTGF inhibition by siRNA also abolished enhanced hepatoma cell growth by the coculture with HSCs in vitro (Supplementary Fig. S4A). In xenograft models, shRNA-mediated CTGF knockdown in hepatoma cells abolished the growth-promoting effect of HSCs (Supplementary Fig. S4B and S4C). Enhanced tumor growth induced by the coexistence of hepatoma cells and HSCs is therefore suggested to be CTGF-dependent.

Figure 5.

The growth of HepG2 cells is accelerated by coculture with LX-2 cells, which is abolished by inhibition of CTGF. The growth of HepG2 cells was evaluated in vitro using a WST-8 assay or xenograft tumor models at the indicated time points. For the coculture experiments, HepG2 cells were incubated in Transwell plates and cocultured with the same number of LX-2 cells. A, Growth of HepG2 cells exposed to 100 ng/mL FG-3019 or hIgG in monoculture. B, Growth of HepG2 cells in monoculture or in coculture with LX-2 cells. C, Growth of HepG2 cells in monoculture or in coculture with LX-2 cells after exposure to 100 ng/mL FG-3019 or hIgG. D, Xenograft model of HepG2 cells alone or HepG2 cells with LX-2 cells. HepG2 cells alone and HepG2 cells mixed with the same number of LX-2 cells were injected into the left and right flanks of mice. Inoculated mice were divided into the hIgG and the anti–CTGF-neutralizing antibody administration groups. From the day of inoculation, 40 mg/kg hIgG or FG-3019 was intraperitoneally administered twice per week. The mice were sacrificed, and the xenograft tumors were enucleated 31 days after inoculation. Representative images of enucleated tumors and sequential tumor volumes are presented. N = 4 (A–C) or 5–6 (D) per group. In line graphs, plots with whiskers represent mean values ± SDs. *, P < 0.05 vs. the control.

Figure 5.

The growth of HepG2 cells is accelerated by coculture with LX-2 cells, which is abolished by inhibition of CTGF. The growth of HepG2 cells was evaluated in vitro using a WST-8 assay or xenograft tumor models at the indicated time points. For the coculture experiments, HepG2 cells were incubated in Transwell plates and cocultured with the same number of LX-2 cells. A, Growth of HepG2 cells exposed to 100 ng/mL FG-3019 or hIgG in monoculture. B, Growth of HepG2 cells in monoculture or in coculture with LX-2 cells. C, Growth of HepG2 cells in monoculture or in coculture with LX-2 cells after exposure to 100 ng/mL FG-3019 or hIgG. D, Xenograft model of HepG2 cells alone or HepG2 cells with LX-2 cells. HepG2 cells alone and HepG2 cells mixed with the same number of LX-2 cells were injected into the left and right flanks of mice. Inoculated mice were divided into the hIgG and the anti–CTGF-neutralizing antibody administration groups. From the day of inoculation, 40 mg/kg hIgG or FG-3019 was intraperitoneally administered twice per week. The mice were sacrificed, and the xenograft tumors were enucleated 31 days after inoculation. Representative images of enucleated tumors and sequential tumor volumes are presented. N = 4 (A–C) or 5–6 (D) per group. In line graphs, plots with whiskers represent mean values ± SDs. *, P < 0.05 vs. the control.

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CTGF induces IL6 secretion from HSCs, which promotes the growth of hepatoma cells

To investigate the mechanisms underlying the growth-facilitating effect of HSCs mediated by CTGF, we focused on IL6, which has been reported to be produced by HSCs in the HCC microenvironment, where it facilitates tumor progression (24). IL6 was secreted not from HepG2 cells but from LX-2 cells in monoculture (Fig. 6A). The secretion levels of IL6 from LX-2 cells were increased by the recombinant CTGF supplementation (Fig. 6B; Supplementary Table S2). Moreover, the concentration of IL6 in the supernatant was elevated by coculture with HepG2 cells compared with LX-2 cells alone (Fig. 6A). The growth-facilitating effect of LX-2 cells on HepG2 cells was abolished by the siRNA-mediated knockdown of IL6 in LX-2 cells, and this effect was accompanied by the downregulation of p-STAT-3 in HepG2 cells (Fig. 6C). Anti–IL6-neutralizing antibody treatment also decreased the growth of HepG2 cells cocultured with LX-2 cells and downregulated p-STAT-3 in HepG2 cells (Supplementary Fig. S5A). The growth-facilitating effect of LX-2 cells was canceled by STAT-3-knockdown in HepG2 cells (Supplementary Fig. S5B). When HepG2 cells and LX-2 cells were cultured together, the application of an anti–CTGF-neutralizing antibody also decreased the concentration of IL6 in the supernatant and downregulated p-STAT-3 in HepG2 cells (Fig. 6D), which resulted in the inhibition of enhanced cell growth due to coculture with LX-2 cells (Fig. 5B). Consistent with these results, p-STAT-3 expression in liver tumors in KrasG12D CTGF−/− mice was lower than that in KrasG12D CTGF+/+ mice (Supplementary Fig. S6).

Figure 6.

CTGF induces IL6 production in HSCs to promote hepatoma cell growth. The concentration of IL6 in the culture supernatant was measured by ELISA after a 24-hour incubation. Cell growth was evaluated in vitro via a WST-8 assay. A, Concentration of IL6 in the supernatant of HepG2 cells alone, LX-2 cells alone, and the coculture of HepG2 cells and LX-2 cells. The total volume of supernatant was normalized among the three groups. B, IL6 concentration in the supernatant of LX-2 cells incubated with or without 5 nmol/L recombinant CTGF protein (rhCTGF). C, HepG2 cells were incubated in monoculture or in Transwell coculture with LX-2 cells transfected with IL6 siRNA or control siRNA. Seventy-two hours before the start of the coculture, LX-2 cells were transfected with IL6 siRNA or control siRNA. The cell viability, IL6 concentration in the supernatant, and protein expression in HepG2 cells were evaluated 24 hours after the start of the coculture. D, HepG2 cells were incubated in monoculture or in Transwell coculture with LX-2 cells and exposed to 100 ng/mL FG-3019 or hIgG. The IL6 concentration in the supernatant and protein expression in HepG2 cells were evaluated after a 24-hour incubation. N = 4 per group. In bar graphs, bars and top whiskers denote mean values and SDs, respectively. *, P < 0.05 vs. the control.

Figure 6.

CTGF induces IL6 production in HSCs to promote hepatoma cell growth. The concentration of IL6 in the culture supernatant was measured by ELISA after a 24-hour incubation. Cell growth was evaluated in vitro via a WST-8 assay. A, Concentration of IL6 in the supernatant of HepG2 cells alone, LX-2 cells alone, and the coculture of HepG2 cells and LX-2 cells. The total volume of supernatant was normalized among the three groups. B, IL6 concentration in the supernatant of LX-2 cells incubated with or without 5 nmol/L recombinant CTGF protein (rhCTGF). C, HepG2 cells were incubated in monoculture or in Transwell coculture with LX-2 cells transfected with IL6 siRNA or control siRNA. Seventy-two hours before the start of the coculture, LX-2 cells were transfected with IL6 siRNA or control siRNA. The cell viability, IL6 concentration in the supernatant, and protein expression in HepG2 cells were evaluated 24 hours after the start of the coculture. D, HepG2 cells were incubated in monoculture or in Transwell coculture with LX-2 cells and exposed to 100 ng/mL FG-3019 or hIgG. The IL6 concentration in the supernatant and protein expression in HepG2 cells were evaluated after a 24-hour incubation. N = 4 per group. In bar graphs, bars and top whiskers denote mean values and SDs, respectively. *, P < 0.05 vs. the control.

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In the present study, using a genetically modified murine model, we demonstrated that CTGF is highly expressed in human HCC tissues and is involved in liver tumor formation. In addition, we showed evidence supporting the hypothesis that CTGF promotes tumor growth via its interaction with HSCs within the tumor microenvironment. HCC is a vasculature-rich cancer that arises predominantly from fibrotic liver, and it is known that VEGF, Ang-2, and PDGF produced by hepatoma cells or other cell types causes endothelial cell proliferation (25, 26). It is also known that HGF, FGF, and TGFβ produced by stromal cells induce the proliferation of hepatoma cells (27, 28). The present study provided the first demonstration of a previously unknown role of CTGF in the association of hepatoma cells and HSCs.

Our ssGSEA revealed that CTGF expression was correlated with activation of the Ras/Mek/Erk pathway in human liver tissues (Supplementary Fig. S2E). EGF activated the Ras signaling pathway in Huh7 cells, which led to CTGF upregulation (Supplementary Fig. S2A). HepG2 cells, which harbor mutant Ras, produced high levels of CTGF, whereas inhibition of the Ras signaling pathway decreased CTGF production (Supplementary Fig. S2B and S2D). The activation of Ras might not be sufficient for high expression of CTGF because the CTGF expression levels in nontumorous liver tissues of KrasG12D mice were not significantly different from those of control mice (Fig. 2D). However, because the expression of CTGF was clearly increased in liver tumors that developed in these mice, activation of the Ras signaling pathway would induce CTGF expression in the tumor cells.

In the present study, we applied KrasG12D mice as a model of liver tumors. One of the limitations of this model is the uncommon driver gene. Although various gene mutations were reported to be observed in human HCC (29), the frequency of Kras mutations is not high and has been reported to be approximately 5% to 7% (30). However, the Ras signaling pathway is frequently activated in human HCC. For example, Ito and colleagues demonstrated activation of MAPK/Erk in 58% of surgically resected human HCC (31). Calvisi and colleagues reported increased phosphorylation of Raf, Mek, Erk, and Akt compared with surrounding nontumorous liver tissues in all 35 cases of human HCC examined (32). Indeed, Raf kinase downstream of Ras is an important molecular target of HCC, and sorafenib, a Raf kinase inhibitor, is approved by the FDA as a molecular targeting agent for HCC. The Erk/Akt pathway was constitutively activated in the liver tissues of KrasG12D mice, which is consistent with activation of the Ras signaling pathway in human HCC. In agreement with the observations in human HCC, tumors in these mice produced high levels of CTGF. Importantly, the hepatocyte-specific knockout of CTGF significantly reduced the number and size of liver tumors in these mice. Thus, CTGF produced by hepatocytes and hepatoma cells contributes to tumor development and progression.

Hepatic fibrosis is not observed in KrasG12D mice, despite the fact that most HCCs develop from fibrotic livers. Hepatic fibrosis is characterized by the excessive accumulation of ECM in the liver (33). ECM is mainly produced by HSCs, which account for 5% to 8% of the cells in the liver and are key regulators of liver fibrosis (34–38). Although the causative relationship between liver fibrosis and hepatocarcinogenesis has been incompletely understood, there is abundant evidence that liver fibrosis and HSCs contribute to the initiation of HCC (24, 33, 34). Owing to the absence of background hepatic fibrosis, KrasG12D mice might not be appropriate for focusing on cancer initiation. Nevertheless, we consider this model is still useful for analyzing the roles of CTGF in cancer cell–HSC interactions in HCC microenvironment, because CTGF was upregulated in cancer cells and activated HSCs were present in tumors, similar to human HCC.

CTGF has been reported to promote DNA synthesis and cell-cycle progression in hepatoma cells (11, 13). In our experiments, CTGF did not directly affect hepatoma cell growth but accelerated it in the presence of HSCs in both culture and xenograft models. HSCs also infiltrate tumorous tissues and exist as α-SMA–positive cells in perisinusoidal areas of HCC tissues (36, 37). Activated HSCs secrete several humoral factors and accelerate tumor proliferation, invasion, and angiogenesis (24, 34–38). Coulouarn and colleagues reported that the cross-talk between hepatoma cells and activated HSCs promoted the production of VEGF and MMP9 from HSCs, which leads to vascular angiogenesis and tissue remodeling (39). In the present study, we revealed that CTGF produced by tumor cells participated in HSC activation and HCC progression. Furthermore, CTGF-mediated cross-talk between hepatoma cells and activated HSCs induced IL6 production from HSCs, which was involved in hepatoma cell growth. Paracrine IL6 production by inflammatory cells was reported to be deeply involved in early hepatocarcinogenesis (40). IL6 has been reported to be produced by HSCs in the HCC microenvironment, where it facilitates the progression of HCC (24). CTGF-induced IL6 secretion from HSCs might be one of the causes of accelerated hepatoma cell growth. Meanwhile, our secretome analysis revealed that LX-2 cells secreted several humoral factors other than IL6 after the stimulation with recombinant CTGF protein (Supplementary Table S2). Although our results indicated that IL6 was related to the CTGF-mediated interaction between cancer cell and HSCs, other molecules might also be involved in this interaction.

The application of an anti–CTGF-neutralizing antibody alone did not affect the growth of hepatoma cells (Fig. 5A). In contrast, an anti–CTGF-neutralizing antibody decreased hepatoma cell growth in the presence of HSCs both in vitro and in xenograft models (Fig. 5C and D). An anti–CTGF-neutralizing antibody was thus shown to prevent CTGF-mediated tumor–stroma interactions between hepatoma cells and HSCs and to successfully inhibit tumor growth. Several pharmacologic inhibitors of CTGF that block the biological action of CTGF have been explored (41). An anti–CTGF-neutralizing antibody (FG-3019) is capable of specifically binding to CTGF with reasonable affinity and can exhibit an inhibitory effect (3). An anti–CTGF-neutralizing antibody has been demonstrated to be effective in animal models of several malignant diseases, such as pancreatic cancer, ovarian cancer, leukemia, and melanoma, and a phase II clinical trial is currently being conducted in patients with pancreatic cancer (3, 6, 41–45). The effectiveness of an anti–CTGF-neutralizing antibody on HCC was previously unknown. Due to the tumor-facilitating actions of HSCs, HSCs are currently considered a novel therapeutic target in HCC (24). Our results suggest that an anti–CTGF-neutralizing antibody might be a novel therapeutic agent that can be used to inhibit the actions of HSCs in HCC.

In conclusion, CTGF produced by hepatoma cells exerts tumor-promoting effects through tumor–stroma interactions between tumor cells and HSCs. Approaches using CTGF-targeted biologics might be able to attenuate this effect.

No potential conflicts of interest were disclosed.

Conception and design: H. Hikita, T. Kodama, R. Sakamori, H. Yokoi, T. Tatsumi, T. Takehara

Development of methodology: M. Shigekawa, R. Sakamori, H. Yokoi, T. Tatsumi

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): Y. Makino, T. Kodama, R. Yamada, R. Sakamori, H. Eguchi, M. Mukoyama

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): Y. Makino, H. Hikita, T. Kodama, R. Yamada, R. Sakamori, E. Morii, T. Tatsumi

Writing, review, and/or revision of the manuscript: Y. Makino, H. Hikita, E. Morii, M. Mukoyama, T. Tatsumi, T. Takehara

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): H. Yokoi, M. Mukoyama, S. Hiroshi

Study supervision: H. Hikita, R. Sakamori, T. Tatsumi, T. Takehara

This work was partially supported by a Grant-in-Aid for Scientific Research (JP26670381 and JP26253947 to T. Takehara) and a Grant-in-Aid for Young Scientists (JP17K15943 to Y. Makino) from the Ministry of Education, Culture, Sports, Science, and Technology, Japan, a research grant from Bristol-Myers Squibb (to T. Takehara), and a Grant-in-Aid for Research from the Japan Agency for Medical Research and Development (JP18fk0210021, JP18fk0310108, JP18fk0210018, and JP18fk0210026). The authors thank FibroGen for kindly providing the anti–CTGF-neutralizing antibody FG-3019.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

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Supplementary data