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Hepatocellular carcinoma (HCC) is a common human cancer with high mortality and currently there is no effective chemoprevention or systematic treatment. Recent evidence suggests that COX-2-derived PGE2 and Wnt/β-catenin signaling pathways are implicated in hepatocarcinogenesis. Here we report that ω-3 PUFAs, docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA), inhibit HCC growth through simultaneously inhibition of COX-2 and β-catenin. DHA and EPA treatment resulted in a dose-dependent reduction of cell viability with cleavage of PARP, caspase-3 and caspase-9 in three human HCC cell lines (Hep3B, Huh-7, HepG2). In contrast, arachidonic acid (AA), a ω-6 PUFA, exhibited no significant effect. DHA and EPA treatment caused dephosphorylation and thus activation of GSK-3β, leading to β-catenin degradation in Hep3B cells. The GSK3-β inhibitor, LiCl, partially prevented DHA-induced β-catenin protein degradation and apoptosis. Additionally, DHA induced the formation of β-catenin/Axin/GSK-3β binding complex, which serves as a parallel mechanism for β-catenin degradation. Furthermore, DHA inhibited PGE2 signaling through downregulation of COX-2 and upregulation of the COX-2 antagonist, 15-hydroxyprostaglandin dehydrogenase (15-PGDH). Finally, the growth of HCC in vivo was significantly reduced when mouse HCCs (Hepa1–6) were inoculated into the Fat-1 transgenic mice which express a Caenorhabditis elegans desaturase converting ω-6 to ω-3 PUFAs endogenously. These findings provide important preclinical evidence and molecular insight for utilization of ω-3 PUFAs for the chemoprevention and treatment of human HCC.
Hepatocellular carcinoma (HCC) is the fifth most common human cancer with high mortality and its incidence is rising worldwide. The overall survival of patients with HCC is dismal and currently no efficient secondary prevention or systemic treatments are available. HCC usually develops in the presence of continuous inflammation and hepatocyte regeneration in the setting of chronic hepatitis and cirrhosis (1). Increased cellular turnover and regeneration within the context of a noxious chronically inflamed environment cause accumulation of chromosomal damages, which eventually affect the structure and expression of oncogenes and tumor suppressor genes leading to carcinogenesis. Recent studies have shown that mediators of inflammation, such as prostaglandins (PGs), play an important role in hepatocarcinogenesis (2). For example, increased cyclooxygenase-2 (COX-2) expression has been found in human and animal HCCs and in dysplastic hepatocytes (3–9). Elevated levels of PGs, most notably PGE2, have also been detected in HCC (10). Overexpression of COX-2 or treatment with exogenous PGE2 increases human HCC cell growth and invasiveness (8, 11). The cyclooxygenase (COX) inhibitors, nonsteroidal anti-inflammatory drugs (NSAIDs), inhibit the proliferation and induce apoptosis in cultured HCC cells and in animal models of hepatocarcinogenesis (2), although these inhibitors are known to mediate effects through both COX-dependent and -independent mechanisms.
In addition to upregulation of COX-2, Wnt/β-catenin activation has also been implicated in various stages of hepatic tumorigenesis, including the dysplastic foci, hepatic adenoma, hepatoblastoma and HCC (12–16). Activation of the Wnt/β-catenin pathway occurs in approximately 30 to 40% of HCCs (17). Multiple mechanisms of β-catenin activation or stabilization have been reported in hepatic tumorigenesis, including mutations in the β-catenin gene (Ctnnb1), or components of its degradation machinery such as Axin and GSKβ inactivation (12–16). In mice, hepatic deletion of APC, another degradation component of β-catenin, leads to HCC (18). Recently, upregulation of a member of Wnt receptors, Frizzled-7, has been shown as another possible mechanism of β-catenin activation in HCC (19). In addition, Wnt/β-catenin also plays an important role in regulation of hepatocyte proliferation, survival, liver regeneration and in the maintenance and self-renewal of pluripotent stem cells and progenitor cells (12), hence, they may play a role in the maintenance of the cancer stem cell compartment. Indeed, β-catenin activation has been identified in oval cells (liver stem cells), which might be precursors of a subset of HCC (13). Thus, there appears to be multiple mechanisms of β-catenin activation leading to liver neoplasia. Although PGE2 has recently been shown to activate β-catenin in colon cancer cells (20, 21), it remains unknown whether the COX-2/PG and Wnt/β-catenin signaling pathways converge during hepatocarcinogenesis.
In contrast to the documented carcinogenic effect of the prostaglandins (PGE2 in particular) derived from arachidonic acid (an ω-6 PUFA), there is abundant experimental evidence that the ω-3 PUFAs rich in fish oil, such as docosahexaenoic acid (DHA) and eicosapentaenic acid (EPA), prevent carcinogenesis (22, 23). However, the molecular mechanisms for the anticancer actions of ω-3 PUFAs remain incompletely understood. This study was designed to investigate the effect and mechanism of ω-3 PUFAs in HCC cells. Our results show that DHA and EPA inhibited the growth of three human HCC cells (Hep3B, Huh-7, HepG2), in vitro. The growth of HCC in vivo was also significantly reduced when mouse HCC cells (Hepa1–6) were inoculated into the syngeneic Fat-1 transgenic mice which carry a Caenorhabditis elegans desaturase converting ω-6 to ω-3 PUFAs. Moreover, our data reveal that COX-2-derived PGE2 activates β-catenin signaling pathways in human HCC cells and that ω-3 PUFAs inhibit HCC growth by simultaneously blocking β-catenin and COX-2 signaling pathways. These findings provide important preclinical evidence and molecular framework for utilization of ω-3 PUFAs in the chemoprevention and treatment of HCC.
α-MEM, DMEM, RPMI 1640, fetal bovine serum (FBS), glutamine, antibiotics, and Lipofectamine plus reagent were purchased from Life Technologies, Inc. (Rockville, MD). PGE2 was purchased from Calbiochem (San Diego, CA). The cell proliferation assay reagent WST-1 was purchased from Roche Molecular Biochemicals (Indianapolis, IN). The antibody for human COX-2, 15-hydroxyprostaglandin dehydrogenase (PGDH) were purchased from Cayman Chemical Company (Ann Arbor, MI). The antibodies against human Axin, β-catenin, PARP, caspase-3, caspase-9, and c-Met were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The antihuman β-actin monoclonal antibody was purchased from Sigma (St. Louis, MO). The horseradish peroxidase-linked streptavidin and chemiluminescence detection reagents were from Amersham Pharmacia Biotech, Inc. (Piscataway, NJ). The rabbit antibodies for phospho-Akt (Thr308), Akt, phospho-GSK-3β (Ser9), GSK-3β were purchased from Cell Signaling Technology (Beverly, MA). Mouse monoclonal anti GSK-3β was purchased from Transduction Laboratories and cytochrome c was purchased from BD Bioscience (Franklin Lakes, NJ). The Bio-Rad protein assay system was obtained from Bio-Rad Laboratories (Hercules, CA). The Tris-glycine gels were obtained from Invitrogen Life Technologies, Inc. (Carlsbad, CA). Hr. T. Hla at the University of Connecticut Health Center provided the COX-2 expression plasmid (containing full length of human COX-2 cDNA in sense orientation cloned in mammalian expression vector PCDNA3).
The human hepatocellular carcinoma cell lines (Hep3B, HepG2 and Huh7) were obtained from American Type Culture Collection (Manassas, VA) and cultured according to our previous described methods (8, 11, 24). Briefly, the cells were cultured in EMEM supplemented with 10%FBS, 2 mM L-glutamine, and penicillin/streptomycin. The cells were incubated at 37°C in a humidified CO2 incubator. The experiments were performed when cells reached ~80% confluence and conducted in serum-free medium (with serum deprivation for 24 hr before each experiment).
Cell growth was determined using the cell proliferation reagent WST-1, a tetrazolium salt that is cleaved by mitochondrial dehydrogenases in viable cells. Briefly, 100 µl of cell suspension (containing 0.5−2 × 104 cells) were plated in each well of 96-well plates. After 24 hr culture to allow reattachment, the cells then were treated with specific reagents such as DHA, EPA or Wnt3a-conditioned medium (Wnt3a-CM) for indicated time points. At the end of each treatment, the cell proliferation reagent WST-1 (10 µl) was added to each well, and the cells were incubated at 37°C for 0.5~5 hr. Absorbance at 450 nm was measured using an automatic ELISA plate reader.
Equal amount of cellular protein from the treated cells was incubated with 10 µl of rabbit antihuman Axin polyclonal antibody at 4°C for overnight, followed by addition of 20 µl Protein A/G PLUS agarose (Santa Cruz Biotechnology). The mixture was incubated for 2 hr and then washed three times with the cell lysis buffer [50 mM HEPES (pH 7.55), 1 mM EDTA, 1 mM DTT, and protease inhibitor cocktail tablets from Roche Diagnostics (Basel, Switzerland)]. The final pellets were dissolved in 20 µl 2X protein loading buffer, and the samples were subjected to SDS-PAGE and Western blot analysis using 1:1000 dilution mouse antihuman GSK-3β or β-catenin monoclonal antibodies and enhanced chemiluminescence Western blot detection system (Amersham Pharmacia Biotech, Inc.).
Hep3B cells were exposed to the mixture of Lipofectamine plus reagents and COX-2 expression plasmid (full-lengh human COX-2 cDNA cloned in pcDNA3 vector), pcDNA3 control vector or TCF/LEF-Luc reporter plasmid for 4 hr. Following removal of the transfection mixtures, the cells were cultured in fresh serum-free medium with or without specific treatment as indicated in the text. The expression of COX-2 was verified by immunoblotting.
The cultured cells were seeded at a concentration achieving 80% confluence in 12-well plates for eighteen hours before transfection. The cells were transiently transfected with 0.2 µg/per well translucent TCF/LEF-Luc reporter vector, which was designed to measure the β-catenin transcriptional activity of TCF/LEF responsive genes. After transfection, the cells were treated with specific reagents including DHA or Wnt3a conditioned medium in serum-free medium for 24 hr. The cell lysates were then obtained with 1X reporter lysis buffer (Promega). The luciferase activity was assayed in a Berthold AutoLumat LB953 Luminometer (Nashua, NH) by using the luciferase assay system from Promega. The relative luciferase activity was calculated after normalization of cellular proteins. All values are expressed as percentage of activity induction relative to control activity.
At the end of each indicated treatment, the cells were scraped off the plates and centrifuged, washed twice with cold phosphate-buffered saline (PBS) containing 0.5 mM PMSF and 10 µg/ml leupeptin and resuspended in 5-fold volume of hypotonic buffer consisting of 50 mM HEPES pH 7.55, 1 mM EDTA, 1 mM DTT and protease inhibitor cocktail tablets (Roche Diagnostics GmbH). After sonication, the whole cell lysate was collected by centrifugation at the speed of 15,000g at 4°C for 10 minutes to remove cell debris and stored in aliquots at −80°C until use. The protein concentration in the cell extracts were determined by the BioRad protein assay (Bio-Rad, CA). 30 µg of cellular protein was subjected to SDS-PAGE on 4–20% Tris-glycine gels for PARP, β-catenin, GSK-3β, phosphor-GSK-3β, Akt, phosphor-Akt, c-Met, Axin cytochrome c, caspase-3, caspase-9, COX-2, 15-PGDH or β-actin. The separated proteins were electrophoretically transferred onto the nitrocellulose membrane (BioRad, CA). Nonspecific binding was blocked with PBS-T (0.5% Tween 20 in PBS) containing 5% non-fat milk for 1 hr at room temperature. The membranes were then incubated overnight at 4°C with individual primary antibodies in PBS-T containing 1% non-fat milk at the dilutions specified by the manufactures. Following three washes with PBS-T, the membranes were then incubated with the horseradish peroxidase-conjugated secondary antibodies at 1:10,000 dilution in PBS-T containing 5% non-fat milk for 1 hour at room temperature. The membranes were then washed 3 times with PBS-T and the protein bands were visualized with the ECL Western blotting detection system.
The Fat-1 transgenic mice (from Dr. J.X. Kang of Harvard University) (25). The animals were kept at 22°C under a 12-h light/dark cycle and fed standard mouse chow (Prolab® IsoPro® 5P75 RMH 3000) with water ad libitum. The fatty acid compositions in the Fat-1 transgenic mice and wild type control mice housed at our animal facility are shown in Supplementary Table 1. The mice were kept under specific pathogen-free conditions in standard cages, and used 8–10 weeks old male for this experiment. Each mouse was injected subcutaneously (s.c.) into the area overlying the right flank with 1.5 × 106 mouse hepatocellular carcinoma cells (Hepa1–6) suspended in 100 µl of serum-free medium. After inoculation, the animals were closely monitored for the development of subcutaneous tumor. The tumor size was measured with a caliper every 2 days. Upon sacrifice, the tumor volume was calculated according to the following formula: Tumor volume = L × W2 × 0.5. The animal experiments were carried out according to the protocol approved by the University of Pittsburgh Institutional Animal Care and Use Committee (#0201740B).
Twenty paired human hepatocellular carcinomas and their matched nonneoplastic/nondysplastic liver tissues were analyzed by immunohistochemistry for the expression of COX-2 and β-catenin. Increased cytoplasmic staining for COX-2 and nuclear staining for β-catenin was observed in HCC cells when compared with the non-tumor liver tissue (Supplementary Figure 1). The average staining intensity for COX-2 in HCC is 2.10 ± 0.78, which is significantly higher than that in nontumor liver tissue (0.20 ± 0.09) (p<0.01, Student t test). Whereas COX-2 is expressed exclusively in the cytoplasm of both HCC cells and to a less degree in hepatocytes, the expression pattern of β-catenin between hepatocellular carcinoma cells and non-neoplastic hepatocytes are distinctly different. In nontumorous hepatocytes, β-catenin is expressed exclusively in the plasma membrane with no significant cytoplasmic staining and absence of nuclear staining in all 20 patients. In contrast, in hepatocellular carcinoma cells, nuclear staining for β-catenin was observed in 5/20 patients (25%), with focal cytoplasmic staining and decreased membrane staining, indicating β-catenin nuclear translocation and activation. Thus, COX-2 and β-catenin signaling pathways are active in a significant percentage of human HCCs.
Human HCC cell lines were examined for their response to DHA, EPA and AA treatment. As shown in Figure 1A, treatment of Hep3B cells with two ω-3 PUFAs (30 µM), DHA and EPA, induced a time-dependent reduction of cell viability; in contrast AA, a ω-6 PUFA, had no significant effect. Treatment with 30 µM EPA for 12, 24, 48 and 72 hours induced approximately 45%, 60%, 70% and 75% reduction of viable cells, respectively. DHA appears to have more effect, with approximately 75% reduction of viable cells at 12 hours and more than 90% reduction at 24, 48 and 72 hours. The cells treated with DHA and EPA show morphological features of cell death, characterized by cell shrunken, round and detachment. In contrast, AA treatment did not significantly alter the cell morphology. The effect of DHA and EPA is dose-dependent in all three human HCC cell lines (Hep3B, Huh7 and HepG2 (Figure 1B). The observations that DHA induced the cleavage of PARP, caspase-3 and caspase-9, with concomitant release of cytochrome c from mitochondria to cytosol confirm the induction of apoptosis (Figure 1C). Taken together, these results document induction of apoptosis by ω-3 PUFAs in HCC cells. We have also tested the effect of DHA and EPA in primary cultures of liver parenchymal cells and these compounds were found to have no cytotoxic effect in primary cells (unpublished observations).
Further experiments were performed to assess the mechanisms by which ω-3 PUFAs induce HCC apoptosis. Since β-catenin activity is importantly involved in hepatocarcinogenesis, the potential effect of ω-3 PUFAs on β-catenin protein level and activity was examined. As shown in Figure 2A and 2B, treatment with DHA or EPA reduced the level of β-catenin protein; this effect was time-dependent (observed 1–6 hours after treatment). As c-Met is a β-catenin controlled downstream gene, the potential effect of DHA and EPA on c-Met protein expression was also examined. Indeed, DHA and EPA treatment also reduced the expression of c-Met. In contrast, treatment with AA did not alter β-catenin or c-Met level (Figure 2C).
Since β-catenin regulates gene expression via binding as a transcription factor in complex with the TCF/LEF transcription factor family to the promoter region of target genes, we further examined the effect of DHA on TCF/LEF reporter activity. The TCF/LEF transcription activity was assayed after transient transfection of a luciferase reporter construct under the control of TCF/LEF response element. As shown in Figure 2D, DHA treatment significantly inhibited the TCF/LEF reporter activity (approximately 5 fold, p<0.01). This result further confirms suppression of β-catenin activity by DHA.
The level of β-catenin in cells is tightly controlled by its degradation complex composed of Axin, APC, GSK-3β and β-catenin, in which GSK-3β phosphorylates β-catenin and thus triggers its ubiquitination and subsequent proteosomal degradation. The activity of GSK-3β is regulated by its phosphorylation status, with GSK-3β phosphorylation at Ser-9 being functionally inactive. To determine whether ω-3 PUFAs might induce β-catenin degradation through inhibition of GSK-3β phosphorylation, we examined the phospho-Ser-9-GSK-3β and total GSK-3β protein levels in Hep3B cells treated with PUFAs. As shown in Figure 3A, DHA treatment reduced GSK-3β phosphorylation, whereas it had no effect on the protein level of total GSK-3β. Similarly, EPA treatment also decreased the level of phosphor-GSK-3β, whereas AA had no effect. Since the phosphorylation of GSK-3β is controlled by Akt, we also examined the potential effect of DHA on Akt phosphorylation. Our data showed that DHA had no effect on Akt phosphorylation (Figure 3A). Thus, DHA most likely inhibited GSK-3β phosphorylation through mechanism independent of Akt. Taken together, the above findings provide evidence for GSK-3β dephosphorylation (activation) in ω-3 PUFA-induced degradation of β-catenin in HCC cells.
To further determine the role of GSK-3β in DHA-induced β-catenin degradation, Hep3B cells were pretreated for 1 hr with LiCl prior to DHA treatment to determine the level of β-catenin protein, TCF/LEF reporter activity and cell growth. As shown in Figure 3B, inhibition of GSK-3β by LiCl prevented DHA-induced reduction of β-catenin protein and TCF/LEF reporter activity. Accordingly, LiCl pretreatment also prevented DHA-induced PARP and restored DHA-induced cell death (Figure 3C). These findings further support the role of GSK-3β activation (dephosphorylation) in DHA-induced β-catenin degradation in HCC cells.
The degradation of β-catenin strictly depends upon β-catenin phosphorylation, which occurs in a multiprotein complex containing Axin and GSK-3β and β-catenin. It is believed that in this complex assembled by Axin, GSK-3β phosphorylates the β-catenin primarily when it is bound to Axin. To determine whether DHA alters the assembly of the Axin/GSK-3β/β-catenin complex, immunoprecipitation and western blot experiments were performed to detect the Axin/GSK-3β/β-catenin binding complex. As shown in Figure 4A–C, treatment of Hep3B cells with DHA induced the association of Axin with GSK-3β as well as β-catenin. This effect was observed within 1 hour and persisted at 5 hours. In contrast, AA did not affect the association between Axin and GSK-3β (Figure 4D). These findings indicate that DHA induces the association of Axin with GSK-3β and β-catenin, thereby facilitating the formation of β-catenin destruction complex. Taken together, our data suggest that ω-3 PUFAs induce β-catenin degradation through dephosphorylation of GSK-3β and formation of β-catenin destruction complex in HCC cells.
Since Wnt3a is known to activate β-catenin signaling in cells, further experiments were carried out to determine whether Wnt3a might protect HCC cells from DHA-induced apoptosis. Indeed, treatment of Hep3B cells with Wnt3a conditioned medium partially prevented DHA-induced cell death (Supplementary Figure 2A). The effect of Wnt3a on β-catenin activation was confirmed by the observation that Wnt3a conditioned medium prevented DHA-induced reduction of TCF/LEF transcription activity (Supplementary Figure 2B). These results further demonstrate that DHA inhibits HCC growth at least in part through down-regulation of Wnt/β-catenin signaling pathway.
We next examined whether DHA might also affect the expression of COX-2 in HCC cells. As shown in Supplementary Figure 3, DHA significantly inhibited the COX-2 promoter activity and COX-2 protein expression in HCC cells. These findings suggest that DHA inhibits the expression of COX-2 through suppression of gene transcription.
15-PGDH catalyzes the rate-limiting step of prostaglandin catabolism and thus represents a physiological antagonist of COX-2 (26, 27). Recent emerging evidence suggests that elevated PGE2 in cancers may be the result of enhanced COX-2-mediated PGE2 synthesis as well as reduced 15-PGDH-mediated degradation of PGE2. Therefore, we sought to further determine whether DHA might affect 15-PGDH expression in HCC cells. As shown in Supplementary Figure 4, DHA treatment enhanced the expression of 15-PGDH in a dose-dependent manner in HCC cells (Hep3B, HepG2 and Huh7). These data are consistent with the observation that DHA and EPA inhibit PGE2 production in Hep3B cells (Supplementary Figure 5).
Since DHA reduces PGE2 level through concomitant inhibition of COX-2 and induction of 15-PGDH, we postulate that DHA might also inhibit β-catenin through inhibition of PGE2. To evaluate this hypothesis, further experiments were performed to examine the direct effect of PGE2 on β-catenin activation and to determine whether DHA might prevent PGE2 effect. As shown in Figure 5, PGE2 treatment resulted in dissociation of Axin from GSK-3β and enhanced TCF/LEF reporter activity in Hep3B cells; these effects were significantly blocked by cotreatment with DHA. These findings suggest that suppression of PGE2 by DHA represents another mechanism for β-catenin degradation.
After the in vitro effect of ω-3 PUFAs on HCC cell growth was documented, further experiments were carried out to evaluate the effect of ω-3 PUFAs on HCC growth in vivo. We implanted murine HCC cells (Hepa1-6) into the syngeneic Fat-1 transgenic and control mice (with C57BL/6 genetic background) and examined the growth of the inoculated tumor cells in these animals. The Fat-1 transgenic mice carry a Caenorhabditis elegans desaturase gene that adds a double bond into a saturated fatty-acid hydrocarbon chain and converts ω-6 to ω-3 polyunsaturated fatty acids, resulting in a significant increase in ω-3 PUFAs and reduction in ω-6 PUFAs in all the organs and tissues (25). The Hepa1–6 cell line was chosen because it was derived from hepatocellular carcinoma of C57BL/6 strain and can be grown to form tumors in mice with C57BL/6 genetic background. As shown in Figure 6, there is a marked difference in the tumor size and tumor volume between Fat-1 transgenic (n=10) and wild type mice (n=12). Over an observation period of 14 days, all wild type mice developed a palpable tumor by day 4, whereas only 5 of 10 Fat-1 transgenic mice developed a minor tumor palpable by day 4 day and the mass of all palpable tumor almost disappeared at 12 day. Mice with homozygous mutation for the prostaglandin receptor EP1 (in C57BL/6 background) was used as an additional control, which showed a similar degree of tumor growth as the wild type mice. These findings demonstrate that ω-3 PUFAs inhibit HCC growth, in vivo.
Given the marked reduction of Hepa1–6 cell growth in the Fat-1 transgenic mice, we conducted subsequent experiments to evaluate ω3-PUFA actions in Hepa1–6 cell growth, in vitro. Both DHA and EPA significantly reduced the viability of cultured Hepa1–6 cells (Supplementary Figure 6). Treatment of Hepa1–6 with DHA led to reduction of β-catenin protein as well as TCF/LEF reporter activity (Supplementary Figure 7). In parallel, DHA also inhibited the expression of COX-2 and induced the expression of 15-PGDH in Hepa1–6 cells (Supplementary Figure 8A). Consistent with the latter observations, DHA treatment also inhibited the production of PGE2 in Hepa1–6 cells (Supplementary Figure 8B). Therefore, the effects of ω3-PUFAs in Hepa1–6 cells are similar to those in human HCC cells. These findings demonstrate that ω3-PUFAs inhibit β-catenin and COX-2 signaling in both murine and human HCC cells.
Both the COX-2/PGE2 and Wnt/β-catenin signaling pathways are active in human hepatocellular carcinomas. There is constitutively high expression and activation of COX-2 in human liver cancers and pre-cancerous inflammatory liver diseases; COX-2 activation enhance the production of PGs from AA that subsequently promote hepatic inflammation and neoplasia (2). In parallel, Wnt/β-catenin pathway is also activated in various stages of hepatic tumorigenesis (13–16, 28–31). Therefore, we postulate that therapies aimed at simultaneous disruption of the COX-2/PGE2 and Wnt/β-catenin pathways may produce effective chemopreventive and anti-tumorigenic effects. This study provides important experimental evidence and mechanisms for inhibition of both Wnt/β-catenin and COX-2/PGE2 signaling pathways by ω-3 PUFAs in HCC (Supplementary Figure 9).
Compelling epidemiological and experimental studies have indicated a relationship between PUFAs and the risk of cancer. For example, a high dietary intake of omega-6 polyunsaturated fatty acids, such as linoleic acid (18:2ω-6), is associated with a high risk for colon cancer, whereas high intake of omega-3 PUFAs from fish oils, such as DHA (22:6ω-3) and EPA (20:5ω-3), decreases it (22, 23). Experimental data have shown that the ω-6 fatty acids stimulate carcinogenesis, tumor growth and metastasis, whereas the ω-3 fatty acids exert suppressive effects. In this study, we utilized both in vitro and in vivo models to evaluate the effect of ω-3 PUFAs on HCC growth. Treatment with DHA and EPA induced a dose- and time-dependent growth inhibition and apoptosis in three human HCC cell lines. The induction of apoptosis is confirmed by cleavages of PARP, caspase-3 and caspase-9 and release of cytochrome c. To evaluate the anti-tumor effect of ω-3 PUFAs on HCC growth in vivo, we implanted murine hepatocellular carcinoma cells (Hepa1–6) into the syngeneic Fat-1 transgenic and control mice. The Fat-1 transgenic mice ubiquitously express a Caenorhabditis elegans desaturase, leading to significant increase in ω-3 PUFAs and reduction in ω-6 PUFAs in all the organs and tissues (25, 32). This model was selected because it provides a balanced ratio of ω-6 to ω-3 fatty acids in mouse tissues and eliminates the potential dietary variation associated with long-term feeding of PUFAs. A significant reduction of HCC tumor size and tumor volume was observed in the Fat-1 transgenic mice. These findings provide important in vivo evidence for inhibition of HCC by ω-3 PUFAs. The effect of dietary DHA and EPA on hepatocellular cancer growth remains to be further evaluated.
A prominent mechanism for the chemopreventive action of ω-3 PUFAs is their suppressive effect on the production of AA-derived prostanoids, particularly PGE2 (23, 33). This is important since PGE2 is implicated in multistages of tumorigenesis, including modulation of inflammation, cancer cell proliferation, differentiation, apoptosis, angiogenesis, metastasis and host immune response to cancer cells (2). Our data in this study show that ω-3 PUFAs inhibit COX-2 expression in HCC cells, which is consistent with recently reported downregulation of COX-2 by ω-3 PUFAs in colon cancer cells (34). Moreover, our findings provide novel evidence for induction of 15-PGDH, a rate-limiting key enzyme in prostaglandin catabolism, by ω-3 PUFAs in human cancer cells. The latter observation is noteworthy, since 15-PGDH is a prostaglandin-degrading enzyme that physiologically antagonizes COX-2 and suppresses tumor growth.
In addition to modulation of COX-2 and 15-PGDH by ω-3 PUFAs, our results reveal that degradation of β-catenin is a novel parallel mechanism for ω-3 PUFA-mediated anti-tumor effect. β-catenin is a key molecule in the canonical Wnt pathway that regulates multiple biological functions, including embryogenesis and tumorigenesis (35–38). In the absence of Wnt ligands, cytoplasmic β-catenin associates in a complex with GSK3β, Axin and APC, where it is phosphorylated and targeted for proteosomal degradation. Activation of Wnt signaling causes dissociation of the β-catenin degradation complex, leading to β-catenin accumulation in the cytoplasm and translocation into the cell nucleus. In the nucleus, β-catenin binds the transcription factor T-cell factor (TCF)/lymphoid enhancer factor (LEF) that induce transcription of important downstream target genes implicated in cell proliferation, differentiation, and apoptosis (35–38). Recent evidence has shown that PGE2 induces the cytoplasmic and nuclear accumulation of β-catenin in human colon cancer cells. Castellone and colleagues (20) reported that PGE2 activates its G protein-coupled receptor, EP2, resulting in direct association of the G protein alpha subunit with the regulator of G protein signaling (RGS) domain of axin; this causes release of GSK-3β from its complex with axin, thus leading to β-catenin accumulation. A separate study by Shao et al showed the involvement of cAMP/protein kinase A pathway in PGE2-induced β-catenin accumulation in colon cancer cells (21). The current study provides evidence that PGE2 induces dissociation of GSK-3β from Axin thus preventing β-catenin reduction in HCC cells.
Our data suggest that ω-3 PUFAs induce β-catenin degradation through three interrelated mechanisms. First, we show that DHA and EPA induce a rapid dephosphorylation of GSK-3β in HCC cells, suggesting that GSK-3β activation is involved in ω-3 PUFA-induced β-catenin degradation. This assertion is further supported by the observations that the GSK-3β inhibitor, LiCl, prevents DHA-induced reduction of β-catenin protein and transcription activity and restored DHA-induced cell death. Second, DHA treatment induces the association of Axin with GSK-3β forming β-catenin destruction complex. Third, ω-3 PUFAs suppress PGE2 signaling through concomitant inhibition of COX-2 and induction of 15-PGDH, thus preventing PGE2-induced β-catenin accumulation. The involvement of β-catenin degradation in ω-3 PUFA-induced inhibition of tumor growth is further supported by the observation that Wnt3a conditioned medium partially protects HCC cells from DHA-induced apoptosis.
In summary, this study provides encouraging preclinical evidence and important mechanism for utilization of ω-3 PUFAs in the chemoprevention and treatment of HCC, although the data should be interpreted with caution since the concentration of EPA and DHA used in the cultured cells is relatively high and most likely will not be achieved in vivo. Our findings have significant clinical implications, given that HCC is a common and highly malignant human cancer. It is conceivable that ω-3 PUFAs, either applied alone or in conjunction with current modalities, may represent an effective, nontoxic, and safe chemopreventive and therapeutic agent for patients with HCC or at high risks for development of this devastating tumor.
The authors would like to thank Dr. J. X. Kang at the Harvard Medical School for providing the Fat-1 transgenic mice and thank Dr. Beverly Koller at the University of North Carolina and Dr. John McNeish at Pfizer Inc. for providing the EP1 knockout mice.
The work in the authors’ laboratories is supported in part by National Institutes of Health R01 grants CA134568, CA102325, CA106280 and DK077776 (to T.W.) and DK64207 (to Y.D.) and by KIEST and KOSEF through ISNRC (R13-2007-020-01000-0) (to KL).
No conflicts of interest exist.