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Biochemical, Molecular, and Genetic Mechanisms

PPAR1 as a Molecular Target of Eicosapentaenoic Acid in Human Colon Cancer (HT-29) Cells^1,2

³ Department of Nutrition and Food Science, Texas A&M University, College Station, TX 77843 and ⁴ Department of Molecular and Biomedical Pharmacology, University of Kentucky, Lexington, KY 40536

^* To whom correspondence should be addressed. E-mail: callred{at}tamu.edu.

	ABSTRACT

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
Diets high in (n-3) PUFA decrease colon cancer development and suppress colon tumor growth, but the molecular mechanism through which these compounds act is largely unknown. We sought to determine whether PPAR1 serves as a molecular link between the physiological actions of eicosapentaenoic acid (EPA) in human colon cancer cells (HT-29). At nutritionally relevant concentrations, EPA stimulated a PPAR response element (PPRE) reporter assay in a dose-responsive manner in HT-29 cells. Cotreatment with GW9662 (GW), a PPAR antagonist, significantly inhibited this effect, whereas overexpressing the receptor enhanced it. EPA also stimulated the PPRE reporter in a PPAR negative cancer cell line (22Rv1) when the cells were cotransfected with a PPAR1 expression plasmid and this effect was again inhibited by GW. Furthermore, in vitro incubation of EPA with PPAR1 enhanced binding of the protein to DNA containing a PPRE. Next, we sought to determine whether EPA or a prostaglandin formed from EPA is the functional ligand of PPAR. Cotreatment in HT-29 and 22Rv1 cells with EPA and acetyl salicylic acid, an inhibitor of cyclooxygenase activity, activated the PPRE reporter at levels similar to EPA alone, suggesting that EPA itself is a ligand of PPAR. Finally, EPA suppressed HT-29 cell growth and this effect was significantly reversed by the addition of GW, suggesting that in part the physiological actions of EPA are the result of PPAR activation. These studies identify PPAR as a molecular mediator of (n-3) PUFA actions in colon cancer cells.

	Introduction

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

Experiments in animals have shown that chemical induction of colon tumors in rats decreased when the animals were fed diets high in fish oil (6–8) or supplemented with (n-3) PUFA (9). In the colonic crypts of rats exposed to azoxymethane, a chemical carcinogen, and fed either a high-corn oil or a high-fish oil diet, the fish oil diet inhibited colon tumor formation associated with increased colon cell apoptosis (10). The 2 most common (n-3) PUFA in fish oil are eicosapentaenoic acid (EPA)⁵ and docosahexaenoic acid (DHA). Rats fed diets supplemented with EPA as the main source of fat had reduced colon tumor formation compared with animals consuming diets with linoleic acid, an (n-6) PUFA (9). Dietary DHA has been shown to reduce formation and growth of polyps in mice (11) and inhibit colon cancer metastasis to the lungs (12). Although there appears to be a link between (n-3) PUFA consumption and reduced colon cancer development and progression, the molecular mechanisms that result in this correlation remain poorly understood. We hypothesize that (n-3) PUFA utilize PPAR1 as a molecular target of their physiological actions in colon cells.

Cloned in 1990 (13), PPAR is a member of the nuclear receptor family that includes the adrenal and sex steroid receptors as well as thyroid, retinoid, and orphan receptors. Three isoforms of PPAR have been identified thus far, termed , β/, and (13–15), and all 3 forms heterodimerize with retinoid X receptor (RXR) (16). Evidence suggests that PPAR plays a role in the development and progression of several different forms of cancers, including colorectal cancer (17–19). PPAR has been shown to be expressed in human colonic mucosa, human colon adenocarcinoma, and colon cancer cell lines (19). While PPAR is expressed in both normal and cancerous colon tissue, the relative level of expression between these tissues is controversial. However, PPAR levels are thought to be higher in colon tumors, making the receptor a favorable target for chemotherapeutic interventions for this disease. Ligands for PPAR include but are not limited to prostaglandins (20), thiazolidinedione (TZD) drugs (21–23), and, more controversially, fatty acids (15,24,25).

Fatty acids have been shown to activate PPAR response element (PPRE)-reporter assays in several different cell lines (15,24,25) and individual fatty acids have been shown to bind all 3 subtypes of PPAR (, , and ) (24). The ability of EPA to stimulate a PPRE reporter construct in human colon cancer cells (HT-29) has not been tested. In these studies, we sought to define the molecular mechanism through which EPA activates the PPRE reporter and to evaluate if EPA is a functional ligand of PPAR in HT-29 cells. Furthermore, we sought to determine whether the physiological effects of EPA treatment on colon cancer (HT-29) cells is the result of direct PPAR activation.

	Materials and Methods

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
    Reagents. EPA was purchased from Nu-Chek Prep. Other reagents used in the fatty acid preparations were purchased from Sigma. Rosiglitazone (Ros) and GW9662 (GW) were purchased from Cayman Chemical. Ros was dissolved in dimethyl sulfoxide (Sigma). Acetylsalicylic acid (ASA; Sigma) and GW were solubilized in ethanol (Aaper Alcohol and Chemical Company).

    Cells and cell culture. Colon adenocarcinoma cells (HT-29) were used in most experiments and were generously provided by Dr. David Kaetzel (University of Kentucky, College of Medicine). The human prostate carcinoma cells (22Rv1) were a gift from Dr. Natasha Kyprianou (University of Kentucky, College of Medicine). HT-29 cells were maintained in DMEM/F-12 containing 10% fetal bovine serum and were grown in medium lacking phenol red at 37°C in a 5% CO₂ atmosphere. Human breast cancer cells and 22Rv1 cells were maintained under similar conditions in DMEM and Roswell Park Memorial Institute-1640 media formulation, respectively.

    EPA preparations. EPA was purchased in pure fatty acid form and then dissolved in hexane. This stock solution was maintained under nitrogen gas at all times and fresh fatty acid preparations were made before every experiment. Appropriate volumes of the stock solution and 6 mol/L NaOH were combined to form fatty acid salt complexes. The fatty acid salts were then dried under nitrogen gas and dissolved in cell culture media containing 10% fetal bovine serum. Once the fatty acid was completely dissolved, the media was pH balanced and filter sterilized through a 0.2-µm syringe filter.

    Plasmids. The PPRE reporter construct, 3XPPRE-TK-pGL3, contains 3 copies of a PPRE sequence (AGGACAAAGGTCA) upstream of the mTK promoter between the XhoI and HindIII restriction enzyme sites of the pGL3 basic vector (Promega). Cytomegalovirus promoter-controlled β-galactosidase (β-GAL) expression vector was a kind gift from Dr. Melinda Wilson (University of Kentucky, College of Medicine). pBluescript cloning vector plasmid was purchased from Stratagene.

    Transfection assays. HT-29 cells were transiently transfected with 3 µg of PPRE reporter and 1 µg of β-GAL plasmid per 24-well plate. For 22Rv1 experiments, we transfected cells with 3 µg PPRE reporter, 1 µg β-GAL, and either 1 µg pBluescript or 1 µg PPAR1 per 24-well plate. Plasmids were transfected into cells using ESCORT transfection reagent (Sigma) over a 4-h period. We subsequently treated cells with the selected PPAR ligands and other reagents for 18 h and then lysed them in 50 µL passive lysis buffer. The quantification of induced firefly (Phontius pyralis) luciferase protein was performed using a Luciferase Assay System kit (cat. no. E1501, Promega) on a Berthold Lumat 9507. β-GAL activity was utilized as a constitutively active reporter. β-GAL activity was measured using a β-GAL Enzyme Assay System (Promega). Mean fold induction was obtained by dividing the Relative Luciferase Unit:β-GAL ratio data from each treatment well by the mean values of the vehicle control appropriate for each treatment. Each set of treatments was performed in replicates of 6 in 3 separate experiments.

    Quantification of PPAR binding to DNA. To determine whether EPA influenced PPAR1 ability to bind to a PPRE, an ELISA-based protocol was used. Prior to using this TransAM PPAR kit (Active Motif), an in vitro reaction was performed. For this reaction, either vehicle (methanol) or 100 µmol/L EPA (final concentration) was incubated with 100 ng each of PPAR1 and RXR recombinant proteins (Protein One) at room temperature for 20 min. The TransAM PPAR kit was then performed according to kit instructions. Briefly, 3-µL aliquots of each reaction were added to the 96-well ELISA plate in triplicate. The wells of the ELISA plates were coated with an immobilized oligonucleotide that contained a PPRE (5'-AACTAGGTCAAAGGTCA-3'). After incubation with the ligand/protein reaction mixture, the wells were exposed to a primary antibody recognizing an accessible epitope on PPAR1 protein and then with a secondary antibody conjugated to horseradish peroxidase. These steps were followed by a colorimetric reaction, which was quantified using spectrophotometry.

    Cell proliferation assay. Twenty four hours prior to the experiment, HT-29 cells were seeded at a density of 50,000 cells per well in 6-well plates. The cells were then exposed to individual treatments for 72 h and the media was changed daily. The cells were then washed with PBS, trysinized, pelleted, and resuspended. A total of 100 µL 0.4% trypan blue was added to the cells immediately before counting manually on a hemocytometer. Studies were conducted in triplicate. Each experimental well (n = 9) was counted in triplicate so that for each treatment, there were 3 wells and 3 counts per treatment per experiment. Data are presented as means ± SEM fold of the vehicle control.

    Statistical analysis. For transfection assays, we tested data by balanced 1-way or 2-way (treatment x day) ANOVA using a custom-designed program operating on StatServer 6.1 (Insightful). After determining the presence of a significant interaction effect, treatment means were then subjected to post hoc testing. We performed Tukey's pairwise comparison test post hoc to identify significant differences between the various treatments within a cell line. For transfection assays with the PPRE reporter plasmid, the PPAR expression vector, and multiple treatments, data were subjected to a balanced 2-way (plasmid x treatment or treatment x day) ANOVA hypothesis testing followed by Tukey's pairwise comparison post hoc testing. In similarly designed experiments with only 2 treatments, data were subjected to an unbalanced 2-way ANOVA and Tukey's pairwise comparison test using SigmaStat (Systat Software) with the help of Dr. Eric Blalock (University of Kentucky). For protein/DNA binding assays, the group means were compared using a 2-tailed heteroscedastic Student's t test. For the cellular proliferation assay, the data were subject to 1-way ANOVA and Tukey's post hoc testing as described above.

	Results

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

 
    EPA enhances PPRE reporter activation in HT-29 cells. Cells were transiently transfected with the PPRE reporter construct and subsequently treated with increasing concentrations of EPA for 18 h. EPA increased PPRE reporter activity in HT-29 cells and the level of reporter activity increased in a dose-responsive pattern (Fig. 1). Reporter activity in the 1 µmol/L and 10 µmol/L EPA-treated cells did not differ from the control but were significantly higher in cells treated with 100 µmol/L and 150 µmol/L EPA (P < 0.05). The 100 µmol/L of EPA is a nutritionally relevant dose, because this concentration is similar to human supplementation with (n-3) PUFA (26) and corresponds to animal studies showing physiological effects of (n-3) PUFA in colonic tissue (27).

View larger version (13K): [in this window] [in a new window]   FIGURE 1  EPA enhances PPRE reporter activation in HT-29 cells. Luciferase:β-GAL ratios are expressed as the fold of that in the vehicle control cells. These data are representative of 3 separate experiments. Values are means ± SEM, n = 12. Bars without a common letter differ, P < 0.05.

View larger version (15K): [in this window] [in a new window]   FIGURE 2  GW suppresses Ros and EPA activation of the PPRE reporter in HT-29 cells. Cells were transiently transfected with a 3XPPRE-TK-pGL3 luciferase reporter vector and treated with 1 µmol/L Ros, 1 µmol/L GW, or the combination of the 2 (Ros+GW) for 18 h (A). In separate experiments, HT-29 cells were again transiently transfected with a 3XPPRE-TK-pGL3 reporter vector and treated with 100 µmol/L EPA, 1 µmol/L GW, or the combination of the 2 (EPA+GW) for 18 h (B). Luciferase activity was normalized to a constitutively active β-GAL reporter. Luciferase:β-GAL ratios are expressed as the fold of that in the vehicle control cells. Values are means ± SEM of 3 experiments, n = 12. Bars without a common letter differ, P < 0.05.

    Overexpression of PPAR1 enhances PPRE reporter activation in HT-29 cells.

View larger version (33K): [in this window] [in a new window]   FIGURE 3  Overexpression of PPAR enhances PPRE reporter activation in HT-29 cells. HT-29 cells were transiently transfected with a 3XPPRE-TK-pGL3 luciferase reporter vector in combination with either a pBluescript plasmid or a PPAR1 expression vector and treated with either vehicle control or EPA. Luciferase:β-GAL ratios are expressed as the fold of that in the vehicle control cells. Values are means ± SEM of 3 experiments, n = 17. Bars without a common letter differ, P < 0.05.

View larger version (13K): [in this window] [in a new window]   FIGURE 4  Influence of EPA and ASA cotreatment on PPRE reporter activation in HT-29 cells. Luciferase:β-GAL ratios are expressed as the fold of that in the vehicle control cells. These data are representative of 3 separate experiments. Values are means ± SEM, n = 12. Bars without a common letter differ, P < 0.05.

    EPA activates PPRE in 22Rv1 cells transiently transfected with PPAR1.

View larger version (33K): [in this window] [in a new window]   FIGURE 5  EPA activates a PPRE reporter in 22Rv1 cells transiently transfected with PPAR1. 18S was used for normalization. 22Rv1 cells were transiently transfected with a 3XPPRE-TK-pGL3 luciferase reporter vector in combination with either a pBluescript plasmid or a PPAR1 expression vector and treated with 100 µmol/L EPA, 50 µmol/L ASA, 1 µmol/L GW, EPA+ASA, or EPA+GW. Luciferase:β-GAL ratios are expressed as the fold of that in the vehicle control cells transfected with pBluescript plasmid. Values are means ± SEM of 3 experiments, n = 12. Within the treatment groups that were transfected with the PPAR1 expression vector, bars without a common letter differ, P < 0.05.

View larger version (16K): [in this window] [in a new window]   FIGURE 6  EPA suppression of cellular proliferation is inhibited by GW. These data are representative of 3 separate experiments. Values are means ± SEM, n = 9 wells. Bars without a common letter differ, P < 0.05.

	Discussion

TOP ABSTRACT Introduction Materials and Methods Results Discussion LITERATURE CITED

It has been proposed that individual fatty acids can bind and activate several different nuclear receptors, including all 3 subtypes of PPAR (15,24,25) and RXR (30). Although we demonstrated that EPA activates a PPRE reporter in a dose-dependent manner in HT-29 cells, a previous study found that DHA, another (n-3) PUFA, and to a lesser extent EPA both suppressed PPRE reporter activation in a different colon cancer cell line (HCT 116) (31). The intriguing possibility exists that EPA may be acting as a selective PPAR1 modulator (SPARM) in these 2 cell lines. Data from our laboratory has demonstrated that a single ligand can function as a SPARM activating a PPRE reporter in 1 colon cancer cell line, but not in another (21). Further investigation is necessary to explore whether EPA is acting as a SPARM in these cells.

Although fatty acids have been shown to activate PPRE reporter assays in a number of different cell types (15,24,25), left unexplored is whether PPAR1 serves as a mediator of these responses. To study this, we have utilized both pharmacological and molecular approaches. In the first approach, we used GW, a PPAR-specific antagonist, to block the agonistic effects of EPA. EPA activation of the PPRE reporter was inhibited by GW. In addition, overexpressing PPAR1 in HT-29 cells increases their sensitivity to EPA. As additional support that EPA is a ligand of PPAR1, we have shown that in an in vitro system, EPA is capable of mediating increased (P < 0.001) DNA binding of the receptor to a PPRE (data not shown). For these studies, recombinant PPAR1 and RXR proteins were incubated with either vehicle or 100 µmol/L EPA. DNA binding to an oligonucleotide containing the same PPRE used in the transfection studies was quantified using an ELISA-based assay specific for PPAR. EPA treatment resulted in almost twice as much (0.37 OD for Veh vs. 0.78 OD for 100 µmol/L EPA-treated) PPAR1 bound to the PPRE. This is critical in demonstrating that EPA, in the absence of any further metabolic alteration, is capable of mediating the structural changes to the receptor that confers response element recognition and binding.

To further evaluate whether EPA can influence PPAR1 ability to activate a PPRE in a cellular context, we employed a human cell line (22Rv1) that is PPAR negative. In these studies, we cotransfected 22Rv1 cells with both the PPRE reporter construct and a PPAR1 expression plasmid and then treated the cells with EPA. In the absence of PPAR1 expression, EPA did not affect PPRE reporter activity, but the reporter was induced in cells cotransfected with PPAR1. In this system, GW again inhibited EPA activation of the reporter. Taken together, these data demonstrate the molecular consequence of EPA exposure in these cells results in the direct transactivation of PPAR1 and the upregulation of the PPRE reporter. Although these data do not rule out the possibility that EPA may be a ligand for other nuclear receptors, including PPAR and , clearly, PPAR1 is transactivated following EPA treatment.

Fatty acids, including EPA, are metabolic precursors of prostaglandins that are formed at the cellular level (3) and individual prostaglandins have been shown to be ligands of PPAR (20,32). Therefore, we sought to determine whether EPA itself, or a prostaglandin formed from EPA, was responsible for the increased activity in the PPRE reporter assay. To do this, we cotreated cells with EPA and ASA. ASA has been shown to effectively inhibit COX, the enzyme that converts fatty acids into prostaglandins (33,34). Many COX-inhibiting compounds also act as a PPAR agonist in PPRE reporter assays (35). In these studies, cells treated with ASA alone, at an optimal dose for inhibiting COX activity, had no effect on PPRE reporter activation. Cells cotreated with EPA and ASA had increased reporter activity similar to that of those treated with EPA alone. This indicates that the conversion of EPA to prostaglandins is not required for PPRE activation. It should be noted, however, that 5-lipoxygenase can act on EPA, resulting in the formation of leukotrienes. Eliminating these compounds as potential PPAR ligands in HT-29 cells has proven problematic, because commercially available inhibitors of lipoxygenase activity have themselves been shown to influence PPRE reporter activation (36). Therefore, whereas some metabolic products of EPA may stimulate the PPRE reporter, these data in combination with the fact that EPA enhanced PPAR1 binding to a PPRE in a cell free system suggest that EPA itself functions as a ligand of the receptor. In addition, the consumption of ASA and other COX inhibitors has been associated with a reduced risk of colon cancer formation and growth (37,38). The mechanism of this protection is thought to be related to a reduction in inflammation due to reduced levels of prostaglandins in those tissues (39,40). Although this is likely still a predominant mode of action, our data suggests the possibility that preventing the metabolism of fatty acids, such as EPA, in these cells allows these compounds to be more readily available as ligands of the PPAR1 receptor.

Finally, we wanted to determine whether the physiological actions of EPA were the result of PPAR1 activation. PPAR ligands influence colon cancer cell growth in vivo and in vitro. Nude mice transplanted with human colon carcinoma (CX-1) cells and subsequently treated with troglitazone, a TZD, for 30 d had reduced tumor growth compared with control animals (19). In HT-29 cells, 15-deoxy-^12,14-prostaglandin J₂, troglitazone, and ciglitazone, another TZD, suppressed DNA synthesis and induced apoptosis (41–43). Our findings demonstrate that EPA inhibits HT-29 cell growth and 40% of this effect is blocked by cotreatment with GW. These data suggest that EPA is suppressing colon tumor cell growth in part by activating PPAR. It is important to note that expression of PPAR is also elevated in HT-29 cells (44) and (n-3) PUFA can activate this form of PPAR (24). Therefore, whether FA activation of PPAR contributes to decreased colon cancer cell proliferation requires further study. However, other studies have demonstrated that (n-3) PUFA function through PPAR. Treatment of breast cancer cells with (n-3) PUFA-enriched LDL resulted in increased synthesis of syndecan-1, a proteoglycan that functions as a tumor suppressor (45) and this effect is also partially reversed by cotreatment with GW. Also, fewer chemically induced colon tumors were observed in rats fed diets containing conjugated linolenic acid and this effect was associated with increased PPAR (46). While it is likely that other mechanisms contribute to the actions of EPA in colon cancer cells, it is clear that at least some of its physiological actions occur as a direct result of PPAR transactivation.

Data presented in these studies demonstrate that a functional PPAR1 receptor is both necessary and sufficient for physiological concentrations of EPA to activate the PPRE reporter construct. Furthermore, our data demonstrates that EPA, rather than a prostaglandin metabolite, is a functional ligand of PPAR1. Most importantly, we found that in part the physiological effects of EPA on cellular proliferation are the result of a PPAR1-mediated pathway. Collectively, these data demonstrate that PPAR1 is a molecular target of EPA in colon cancer cells and are a significant step toward understanding the molecular actions of EPA in this disease.

	ACKNOWLEDGMENTS

 
We thank Dr. David Kaetzel for providing the HT-29 cells and Dr. Natasha Kyprianou for the 22Rv1 cells used in these studies. We also thank Dr. Arnold Stromberg and Dr. Eric Blalock for assistance with statistical analysis and Dr. Bernhard Hennig for assistance in developing the fatty acid delivery methodology.

	FOOTNOTES

 
¹ Supported by grants 5-K12-DA-14040-02, NCRR-P20-RR15592, and R01-CA-95609 from the NIH to M.W.K, and W81XWH-04-1-0532 from the Department of Defense to C.D.A.

² Author disclosures: C. D. Allred, D. R. Talbert, R. C. Southard, X. Wang, and M. W. Kilgore, no conflicts of interest.

⁵ Abbreviations used: ASA, acetyl salicylic acid; β-GAL, β-galactosidase; COX, cyclooxygenase; DHA, docosahexaenoic acid; EPA, eicosapentaenoic acid; GW, GW9662; HT-29, human colon cancer cell; PPRE, PPAR response element; Ros, rosiglitazone; 22Rv1, human prostate carcinoma cell; RXR, retinoid X receptor alpha; SPARM, selective PPAR1 modulator; TZD, thiazolidinedione.

Manuscript received 14 May 2007. Initial review completed 14 June 2007. Revision accepted 5 November 2007.

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