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Cytochrome P450 Eicosanoids are Activators of Peroxisome Proliferator-Activated Receptor

Department of Biopharmaceutical Sciences (V.Y.N., Y. H., E.T.L., D.L.K.) and the Liver Center (D.L.K.), University of California, San Francisco, San Francisco, California; and Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas (M.R., J.R.F.)

(Received November 10, 2006; accepted April 9, 2007)

	Abstract

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Cytochrome P450 (P450) eicosanoids regulate vascular tone, renal tubular transport, cellular proliferation, and inflammation. Both the CYP4A -hydroxylases, which catalyze 20-hydroxyeicosatetraenoic acid (20-HETE) formation, and soluble epoxide hydrolase (sEH), which catalyzes epoxyeicosatrienoic acid (EET) degradation to the dihydroxyeicosatrienoic acids (DHETs), are induced upon activation of peroxisome proliferator-activated receptor (PPAR) by fatty acids and fibrates. In contrast, the CYP2C epoxygenases, which are responsible for EET formation, are repressed after fibrate treatment. We show here that P450 eicosanoids can bind to and activate PPAR and result in the modulation of PPAR target gene expression. In transactivation assays, 14,15-DHET, 11,2-EET, and 20-HETE were potent activators of PPAR. Gel shift assays showed that EETs, DHETs, and 20-HETE induced PPAR-specific binding to its cognate response element. Expression of apolipoprotein A-I was decreased 70% by 20-HETE, whereas apolipoprotein A-II expression was increased up to 3-fold by 11,12-EET, 14,15-DHET, and 20-HETE. In addition, P450 eicosanoids induced CYP4A1, sEH, and CYP2C11 expression, suggesting that they can regulate their own levels. Given that P450 eicosanoids have multiple cardiovascular effects, pharmacological modulation of their formation and/or degradation may yield therapeutic benefits.

Regulation of P450 eicosanoid levels is determined by many factors, including the induction or repression of the P450 enzymes responsible for their formation. Large numbers of studies have focused on the induction of rat CYP4A protein and mRNA levels by antihyperlipidemic agents such as clofibrate (Kimura et al., 1989). Rat renal CYP2C23 has also been shown to be inducible by fibrates (Muller et al., 2004), whereas rat hepatic CYP2C11 and CYP2C12, but not CYP2C13, are repressed (Corton et al., 1998). Hepatic and renal sEH expression in rodents is also induced after treatment with clofibrate (Pinot et al., 1995).

Fibrate drugs are part of a diverse group of compounds known as peroxisome proliferators, which also include herbicides and phthalate ester plasticizers (Bishop-Bailey, 2000). Peroxisome proliferators act via peroxisome proliferator-activated receptor (PPAR) (Issemann and Green, 1990). This receptor is a member of the PPAR nuclear receptor family that also consists of the PPAR and PPAR isoforms. PPAR is mainly expressed in the heart, liver, and kidney, whereas the expression of PPAR is predominantly in the adipose tissue (Bishop-Bailey, 2000). The biological role of PPAR as a lipid sensor has been well established. In primary rat hepatocytes, PPAR activation results in the up-regulation of apolipoprotein A-I (apoA-1) and apoA-II and increased expression of ß-oxidation enzymes, fatty acid transport proteins, fatty acid binding proteins, and other genes involved in the control of triglyceride and fatty acid metabolism (Fruchart et al., 2001). In human hepatocytes, however, apoA-I expression is decreased after PPAR activation (Vu-Dac et al., 1998). The species-specific differences in response to PPAR agonists have been attributed to the expression level of PPAR (Palmer et al., 1998), which is significantly lower in humans than in rodents, as well as differences in the regulatory regions of the responsive genes (Vu-Dac et al., 1998; Cheema and Agellon, 2000).

More recently, both PPAR and PPAR have also been found in vascular smooth muscle and endothelial cells (Bishop-Bailey, 2000). In both smooth muscle and endothelial cells derived from human and bovine sources, PPAR inhibits the inflammatory response by repressing NF-B signaling. Expression of genes involved in inflammation, such as interleukin-6, cyclooxygenase 2, and vascular adhesion molecule, is inhibited after PPAR activation (Bishop-Bailey, 2000).

Mechanistically, PPARs are ligand-activated transcription factors that, upon ligand binding, will heterodimerize with the retinoic X receptor and bind to its response element, the peroxisome proliferator response element (PPRE). Ligands for PPAR include fatty acids, eicosanoids, and fibrate drugs (Forman et al., 1997). One of the more potent endogenous activators has been identified as the eicosanoid 8(S)-HETE (Forman et al., 1997). 8(S)-HETE is generated from the lipoxygenase pathway of arachidonic acid metabolism and activates and binds to PPAR at nanomolar concentrations. Other related eicosanoids such as prostacyclin, some prostaglandins, and leukotriene B₄ also activate PPAR (Devchand et al., 1996). A more recent report also showed that -hydroxylated EETs are high-affinity ligands of PPAR (Cowart et al., 2002).

Given that related eicosanoids bind to and activate PPAR, and that P450s and sEH are regulated by peroxisome proliferators, we hypothesize that P450 eicosanoids are also agonists of PPAR. To test whether P450 eicosanoids can functionally activate PPAR, transactivation assays were carried out and demonstrated that 11,12-EET, 14,15-DHET, and 20-HETE are potent activators of PPAR and PPAR. By using gel shift assays, P450 eicosanoids were shown to induce the binding of PPAR to a PPRE. Furthermore, we show that 11,12-EET and 14,15-DHET behave like peroxisome proliferators in that they were able to alter apoA-I and apoA-II mRNA expression in primary rat hepatocytes. Because apoA-I and apoA-II are involved in the transport of high-density lipoprotein, these findings suggest that P450 eicosanoids may play a role in the regulation of triglyceride levels via PPAR. In addition, we show that P450 and sEH mRNA levels were increased in primary rat hepatocytes after treatment with these eicosanoids. These results suggest that P450 eicosanoids may regulate their own levels through a complex autoregulatory mechanism.

	Materials and Methods

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Materials. Wy-14,643 and ciglitazone were obtained from BIOMOL Research Laboratories (Plymouth Meeting, PA) and eicosanoids were purchased from Cayman Chemical (Ann Arbor, MI). The EET agonist 11,12-epoxyeicosa-8(Z)-enoic acid (11,12-EEZE) was synthesized as described previously (Falck et al., 2003). 3,3',5-Triiodo-L-thyronine (T₃) was purchased from Sigma Chemical Co. (St. Louis, MO). CYP4A1 and CYP2C11 primary antibodies were purchased from BD Gentest (Woburn, MA). The sEH antibody was a kind gift from Dr. Bruce Hammock (University of California, Davis, CA), and the CYP2C23 antibody was a kind gift from Dr. Jorge Capdevila (Vanderbilt University, Nashville, TN). Gal4 expression and reporter plasmids were provided by Dr. Thomas Scanlan (University of California, San Francisco, CA) and pCMV-mPPAR and pRS-hRXR plasmids were a generous gift from Dr. Ronald Evans (Salk Institute, La Jolla, CA).

Cell Culture. HepG2 cells were obtained from The American Type Culture Collection (Manassas, VA) and maintained in modified Eagle's medium with Earle's balanced salt solution and contained 10% fetal bovine serum, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids, and penicillin-streptomycin. Sprague-Dawley primary hepatocytes were isolated by the University of California, San Francisco Liver Center Core Facility, were cultured on collagen type I plates (BD Biosciences, Bedford, MA) and overlaid with 0.25 mg/ml Matrigel (BD Biosciences) in Hepatocyte Culture Medium (Cambrex Bio Science Walkersville., Inc., Walkersville, MD). Primary hepatocytes were treated on the third day after isolation. All cells were cultured in 37°C with 5% CO₂.

Transactivation Assays. HepG2 cells were plated in 24-well plates at 8 x 10⁴ cells per well and transfected the next day using Lipofectamine PLUS reagents (Invitrogen, Carlsbad, CA) with 0.1 to 1 ng of Gal4-hPPAR or Gal4-hPPAR, 100 ng of UAS₄-LUC, and 35 ng of pCMV-ßgal. After 24 h, cells were dosed with 50 µM Wy-14,643, 10 µM ciglitazone, or 10 µM P450 eicosanoids in serum-free medium containing up to 0.1% DMSO for 6 h. Cells were lysed using Reporter Lysis Buffer (Promega, Madison, WI) and subjected to one freeze-thaw cycle. A single-tube format luminometer (MGM Instruments, Hamden, CT) was used for obtaining luciferase values using the Luciferase Assay Reagent as substrate (Promega). To determine ß-galactosidase activity, cell lysates were incubated with the substrate o-nitrophenyl-ß-D-galactopyranoside and absorbance was measured at 420 nm. Luciferase activity is expressed relative to ß-galactosidase activity, and -fold activation is calculated relative to DMSO control. Each experiment was carried out in quadruplicate and repeated three times.

Lipid Extraction and Liquid Chromatography-Tandem Mass Spectrometry. A confluent culture of HepG2 cells in a T75 flask was incubated with 10 µM P450 eicosanoids for 6 h. Extraction of lipids from culture medium and cells has been described previously (Fang et al., 2001). In brief, lipids from cell culture medium were extracted twice with water-saturated ice-cold ethyl acetate in a ratio of 1:4 (v/v). For the extraction of intracellular lipids, cells were first trypsinized and washed with phosphate-buffered saline, then homogenized by hand with a Dounce homogenizer. Ice-cold chloroform/methanol (2:1 v/v) was then added and the organic phase was removed. In some cases, hydrolysis of intracellular lipids was carried out by adding methanol containing 5% NaOH and 10% H₂O to the organic phase and incubating the reaction for 1 h at 50°C. Lipids were then extracted with ethyl acetate as described above. The extracted layer was evaporated under nitrogen gas and stored at –80°C. On the day of analysis, lipids were reconstituted in acetonitrile (100 µl). Quantitation of eicosanoids has been described previously (Yu et al., 2004). In brief, analysis of lipids by high-performance liquid chromatography was performed using a 4.6 x 150 mm, 5-µm Luna C18(2) column (Phenomenex, Torrance, CA) and lipids were eluted from the reverse-phase high-performance liquid chromatography column. The effluent was then injected into a Quattro Ultima tandem-quadrupole mass spectrometer (Waters, Milford, MA) and subjected to negative mode electrospray ionization. Multireaction monitoring was used to detect the eicosanoids simultaneously, and data were analyzed using the MassLynx 3.5 software (Waters).

Gel Shift Assays. pCMX-mPPAR (1 µg) and pRS-hRXR (1 µg) were translated using the TNT reticulocyte in vitro translation system (Promega). The binding reaction contained 1 µl of translated PPAR, 0.3 µl of RXR in 10 mM Tris, 150 mM KCl, 6% glycerol, 0.05% Igepal (Sigma), 1 mM dithiothreitol, 2 µg of poly(dI-dC), and P450 eicosanoids with or without 1 µl of PPAR or RXR antibody (Santa Cruz Biotechnology, Santa Cruz, CA). The reactions were incubated for 10 min on ice before 250,000 cpm of -³²P-labeled PPRE was added. The sequence for the consensus PPRE oligonucleotide is 5'-CAAAACTAGGTCAAAGGTCA-3', and the sequence for the mutant oligonucleotide is 5'-CAAAAGTAGCACAAAGCACA-3'. The oligonucleotides were end-labeled with -³²P using T4 polynucleotide kinase. After incubation for 30 min at room temperature, the reaction was separated on a 5% pre-run polyacrylamide gel at 4°C. The gel was dried and radioactive bands were visualized using a PhosphorImager and ImageQuant software (GE Healthcare Biosciences, Chalfont St. Giles, Buckinghamshire, UK).

Real-Time Quantitative PCR. Primary hepatocytes were treated with peroxisome proliferators or eicosanoids for 24 to 48 h. RNA was isolated using TRIzol reagent (Invitrogen) according to the manufacturer's instructions. Reverse transcription was carried out using M-MLV reverse transcriptase (Promega). Primers and probe sets were designed using Primer Express (Applied Biosystems, Foster City, CA) and are listed in Table 1. Probes were labeled with reporter dye, 6-carboxyfluorescein (FAM), at the 5'-end and the dye quencher, Black Hole Quencher, at the 3'-end. The apoA-I, apoA-II, and carnitine palmitoyl transferase 1A (CPT1A) primer and probe sets were Assays-on-Demand purchased from Applied Biosystems. Reactions were run on an ABI Prism 7700, and cycling conditions were 95°C for 10 min, followed by 45 cycles of 95°C for 15 s and 60°C for 1 min. The relative expression of specific transcripts was calculated by the following formula: Relative expression = 2^–Ct, where Ct = (Ct_target –Ct_cyclophilin)_treated – (Ct_target – Ct_cyclophilin)_control.

Statistics. Statistical significance of differences between values was evaluated by an unpaired Student's t test. Significance was set at a p value of <0.05.

	Results

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P450 Eicosanoids Transactivate PPARs. Transactivation assays were carried out to determine whether EETs or DHETs were able to functionally activate PPAR and PPAR (Fig. 1). HepG2 cells were transfected with a Gal4 plasmid containing the ligand-binding domain of either human PPAR or PPAR (Gal4-hPPAR or Gal4-hPPAR) and a luciferase reporter plasmid containing four repeats of the Gal4 response element (UAS₄-LUC). In cells not transfected with hPPAR or hPPAR, only minimal basal activation was observed. When hPPAR-transfected cells were treated with the PPAR-specific activator Wy-14,643, hPPAR was activated greater than 30-fold. Similarly, hPPAR was transactivated by ciglitazone, a PPAR ligand. Significant activation was observed when hPPAR-transfected cells were dosed with 10 µM 11,12-EET, 11,12-EEZE, or 14,15-EET (Fig. 1A). The highest activation was observed with 11,12-EEZE, a monounsaturated 11,12-EET analog. 8,9-EET did not significantly activate either hPPAR or hPPAR.

View larger version (20K): [in this window] [in a new window]   FIG. 1. P450 eicosanoids transactivate hPPAR and hPPAR in HepG2 cells. Transactivation assays were performed in HepG2 cells that were transfected with Gal4-hPPAR or Gal4-hPPAR and the reporter UAS4-LUC. Wy-14,643 (Wy, 50 µM), ciglitazone (Cig, 10 µM), or EETs (10 µM) (A), DHETs (10 µM) (B), or 20-HETE (10 µM) (C) were added to cells in serum-free medium 24 h post-transfection and cells were lysed 6 h later. D, HepG2 cells were transfected with Gal4-TRß and UAS4-LUC and treated with 10 µM 11,12-EET and 10 µM 14,15-DHET as described above. Luciferase activity was normalized to ß-galactosidase values and -fold activation was calculated as increase over the DMSO control (transfected only with UAS4-LUC). The values shown are the mean ± S.D. of a representative experiment performed in quadruplicate. Similar results were obtained in additional experiments. *, significant difference compared to DMSO (p < 0.05).

14,15-DHET was further tested for its ability to transactivate hPPAR and hPPAR in a dose-dependent and saturable manner (Fig. 2). Cells were transfected as described above and treated with 0.1 µM to 100 µM 14,15-DHET. A steep dose dependence was observed between 1 µM and 10 µM 14,15-DHET. Maximal activation was observed at 50 µM for both receptors with EC₅₀ values of 1.5 µM and 2.5 µM for hPPAR and hPPAR, respectively.

View larger version (15K): [in this window] [in a new window]   FIG. 2. Dose-dependent transactivation of hPPAR and hPPAR by 14,15-DHET. Transactivation assays were performed as described in Fig. 1. Transactivation of hPPAR (A) and hPPAR (B) by 0.1 to 100 µM 14,15-DHET is shown in the absence and presence of hPPAR or hPPAR. The values shown are mean ± S.D. of a representative experiment performed in quadruplicate. EC50 values of 1.5 µM for hPPAR and 2.5 µM for hPPAR were estimated using GraphPad Prism (GraphPad Software Inc., San Diego, CA).

Characterization of the Distribution of EETs and DHETs in HepG2 Cells. To characterize the availability of eicosanoids in our cellular system, the distribution of exogenously administered P450 eicosanoids was examined using liquid chromatography-tandem mass spectrometry (Table 2). In cells treated with 10 µM 11,12-EET, approximately 48% was metabolized into 11,12-DHET. Of the 52% 11,12-EET remaining, 32% was detected in the extracellular medium, and 20% was incorporated into phospholipid pools; unbound intracellular 11,12-EET was not detected. The 11,12-DHET formed from 11,12-EET was largely detected in the extracellular medium (47%), with 0.7% found in phospholipid pools and 0.2% as free intracellular lipids. Treatment with 10 µM 14,15-DHET resulted in the predominant distribution of 14,15-DHET extracellularly (99%), with a small percentage bound within lipid pools.

EETs and DHETs Induce PPAR/RXR Binding to a PPRE. Gel shift assays were carried out to determine whether P450 eicosanoids could induce a conformational change in PPAR, resulting in subsequent binding of the PPAR/RXR heterodimer to a PPRE. EETs, DHETs, and 20-HETE induced heterodimer binding, which was not observed with a mutant PPRE or unprogrammed reticulocytes (Fig. 3A). Supershifts with mPPAR and RXR specific antibodies demonstrated that heterodimers consisted of mPPAR and RXR (Fig. 3, B–E). To further validate binding specificity, increasing amounts of cold PPRE were incubated with the complex, which resulted in a dose-dependent decrease in mPPAR/RXR bound to radiolabeled PPRE (Fig. 3F). Likewise, 20-HETE induced mPPAR/RXR-specific heterodimerization and subsequent binding to the PPRE (Fig. 3G).

View larger version (73K): [in this window] [in a new window]   FIG. 3. P450 eicosanoids induce mPPAR/RXR-PPRE binding. Electrophoretic mobility shift assays were performed by incubating in vitro translated mPPAR and RXR with DMSO control, 5 µM Wy-14,643, or 1 µM P450 eicosanoids and 32P-labeled PPRE. The complex was resolved through a 5% nondenaturing polyacrylamide gel and analyzed using a PhosphorImager. Mutant PPRE (Mut) and unprogrammed reticulocyte (Unpgr) were used as controls (A and B). Specificity of the EET (B and D), DHET (C and E), and 20-HETE (G) effects were determined by supershifts with a PPAR (B, C, and G) and RXR (D, E, and G) antibody. N.S. denotes nonspecific binding. Competition assays were performed with increasing amounts of cold PPRE (F and G).

View larger version (27K): [in this window] [in a new window]   FIG. 4. P450 eicosanoids induce PPAR-responsive genes in rat hepatocytes. Primary hepatocytes from Sprague-Dawley rats were treated for 24 h with DMSO, 50 µM Wy, 10 µM 11,12-EET, 14,15-DHET, or 20-HETE. Total RNA was isolated with TRIzol reagent and RNA expression was determined by real-time quantitative PCR. Expression of apoA-I, apoA-II, and CPT1A (A) and CYP4A1, sEH, CYP2C11, and CYP2C23 (B) are normalized to cyclophilin levels and expressed as increase over DMSO control. The values shown are mean ± S.D. of triplicate determinations. *, significant difference compared to DMSO (p < 0.05).

	Discussion

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P450 eicosanoids are identified as novel PPAR and PPAR activators. Of the eicosanoids tested, 11,12-EET, 14,15-DHET, and 20-HETE are the most potent activators of both isoforms. It is recognized that the exogenous concentrations added to culture medium are likely to be higher than intracellular eicosanoid levels. A significant attenuation of PPAR activation by P450 eicosanoids dosed in serum-containing medium is consistent with binding to cellular proteins and reduced free intracellular concentrations (data not shown). Of the exogenously administered 11,12-EET, no free fatty acids were detected intracellularly, and approximately 2 µM was bound in phospholipid pools. This concentration represents one-fifth of the initially administered 11,12-EET dose and suggests that the concentration of free 11,12-EET within the cell is even lower than 2 µM. In vivo, endogenous EET levels in rodent plasma (Z. Yu and D. L. Kroetz, unpublished results) and human liver (Karara et al., 1991) are in the nanomolar range. The ED₅₀ values obtained from transactivation assays in this report were in the low micromolar range; consideration of the ratio of the EET dose to the level of lipid-bound EETs measured intracellularly (5:1) would be consistent with intracellular eicosanoid concentrations in the nanomolar range, making it likely that 11,12-EET can activate PPAR and PPAR in vivo.

The fact that 11,12-EET-derived DHET was detected predominantly in the extracellular medium suggests that, subsequent to metabolism by intracellular sEH, 11,12-DHET was secreted into the medium. Although almost all of the exogenously administered 14,15-DHET was found in the culture medium, this finding does not indicate that 14,15-DHET was not available intracellularly. It is likely that, akin to 11,12-DHET, 14,15-DHET is secreted after entry into the cell. The above phenomenon makes it difficult to determine the free intracellular concentrations of DHETs that are available to activate PPARs in the cellular system used in these studies, but it is likely to be submicromolar.

Regioisomeric P450 eicosanoids may be preferentially metabolized in the cell, thus influencing PPAR activation. Differences in PPAR activation by regioisomeric EET/DHET pairs suggest that the DHET alone cannot account for all of the activity associated with EET treatment. To address the relative contributions of EETs and DHETs, it would be of interest to measure EET transactivation in a cell system devoid of sEH activity. Unfortunately, inhibition of sEH by urea-based and chalcone oxide sEH inhibitors was not possible in these studies since these inhibitors also activated PPARs (V. Y. Ng and D. L. Kroetz, unpublished data; Ng et al., 2006). Further studies would be required to adequately address the relative contribution of EETs and DHETs in the observed activation of PPAR.

11,12-EET and 14,15-DHET transactivated human PPAR and PPAR to a similar extent. Many reports have shown the dual activation of PPARs by polyunsaturated fatty acids. Linoleic, arachidonic, and eicosapentaenoic acids activate murine PPAR and PPAR (Forman et al., 1997), and palmitic, oleic, linolenic, and arachidonic acids are agonists for murine PPAR and PPAR (Kliewer et al., 1997). In recent years, increased efforts to synthesize compounds that act as PPAR/PPAR dual agonists such as ragaglitazar (Brand et al., 2003) and MK-0767 (Doebber et al., 2004) have shown that these compounds have beneficial effects on insulin resistance and display anti-hyperglycemic and hypolipidemic activities. Because P450 eicosanoids are endogenous compounds that are released into the intracellular environment, it is of interest to explore whether P450 eicosanoids play a role in improving insulin sensitivity and maintaining lipid homeostasis.

It has been shown that 10 to 20 µM -hydroxylated EETs transactivated Gal4-mPPAR about 2- to 3-fold over control (Cowart et al., 2002). In the present study, 10 µM 11,12-EET and 14,15-EET activated Gal4-hPPAR more than 10-fold, suggesting that the non--hydroxylated EETs may be more potent activators of PPAR. Itis important, however, to take into account the differences in cell type and the species of PPAR that were used in the assays. 20,14,15-HEET displaced cis-parinaric acid from the ligand-binding domain of PPAR with high affinity (K_i = 3 nM). The corresponding K_i values for EETs and Wy-14,643 are about 10- and 26-fold higher, respectively (Cowart et al., 2002). On the basis of ligand displacement assays, EETs and Wy-14,643 are expected to have lower affinity for PPAR than have the hydroxylated EETs. However, Wy-14,643 was functionally as potent at transactivating full-length PPAR as 20,14,15-HEET, and in assays using Gal4-mPPAR chimeras, Wy-14,643 was 4 times more potent than 20,14,15-HEET. The lack of correlation between cis-parinaric acid displacement constants and transactivation potential makes it difficult to assign relative potencies for PPAR activation to the P450 eicosanoids. The possibility exists that in vivo, EETs are converted to their hydroxylated products in the presence of the CYP4A enzymes. However CYP4A expression was not detected in HepG2 cells, where the present transactivation studies were performed, consistent with a direct effect of EETs on PPAR.

8(S)-HETE, 15-deoxy-^12,14-prostaglandin J₂, and hydroxylated EETs are naturally occurring eicosanoid ligands for PPAR and PPAR (Forman et al., 1997; Kliewer et al., 1997; Cowart et al., 2002). Given the similarity in structure between these established ligands and P450 eicosanoids, and the promiscuous ligand-binding pocket of PPARs, it is not surprising that P450 eicosanoids can also activate PPARs. A major determinant of substrate specificity between PPAR and PPAR is the Tyr314 residue. It will be interesting to model whether the dual agonist P450 eicosanoids can efficiently hydrogen bond with Tyr314 and its equivalent in PPAR. Preliminary docking efforts to examine this interaction were hindered by the large number of configurations that P450 eicosanoids can assume. Structural information will provide additional insight as to why certain P450 eicosanoids are better agonists than others despite their high degree of structural similarity.

Most of the interest in PPAR activation has focused on its role in regulating lipid homeostasis. The degree to which P450 eicosanoids decreased apoA-I mRNA levels in primary rat hepatocytes was similar to or greater than that of the well characterized PPAR agonist Wy-14,643. The regulation of the apoA-I gene by PPAR is species- and ligand-specific (Vu-Dac et al., 1998). In previous rat hepatocyte studies, fenofibrate transcriptionally decreased apoA-I mRNA, whereas benzafibrate, gemfibrozil, and Wy-14,643 had no effect (Staels et al., 1992). The difference in Wy-14,643 effects between the studies could be due to differences in rat strain and the mode of activator exposure. Because P450 eicosanoids cannot be administered in vivo, hepatocytes were dosed after isolation. In contrast, for the fibrate studies, hepatocytes were isolated after in vivo administration of the activators (Staels et al., 1992). Remarkably, 20-HETE was significantly more potent and resulted in a 70% decrease in apoA-I mRNA. It may be possible that the 20-HETE effect is not due solely to PPAR activation. The RXR homodimer (Nagasaki et al., 1994), Rev-erb (Vu-Dac et al., 1998), hepatic nuclear factor-4 (Chan et al., 1993), and saturated fatty acids (Srivastava, 1994) have been implicated in apoA-I regulation. It is important to note, however, that in contrast to rodents, PPAR activation in human hepatocytes leads to an increase in apoA-I expression (Berthou et al., 1996), which is regulated by more complex mechanisms. The effect of P450 eicosanoids on apoA-I expression in humans requires further study.

apoA-II expression in rat hepatocytes was significantly induced after treatment with Wy-14,643 and P450 eicosanoids. In earlier studies, fenofibrate decreased apoA-II mRNA in the rat liver (Staels et al., 1992) and increased hepatic production of apoA-II in humans (Vu-Dac et al., 1995). The conflicting effects of PPAR agonists on apoA-II expression may be related to strain-specific regulation of this gene, as has been previously reported for rat CYP4A (Sewer et al., 1996). In Fischer 344 rats, the lauric acid -hydroxylase activity was induced 1.6-fold, whereas -1 activity decreased 38% in response to lipopolysaccharide treatment. In Sprague-Dawley rats, however, both - and -1 activities decreased. Corresponding CYP4A protein levels were also not consistent between Fischer and Sprague-Dawley rats. In comparing the response of P450s to lipopolysaccharide stimulation between Fischer and Sprague-Dawley rats, the authors (Sewer et al., 1996) found that in general, the results obtained from Fischer rats were more consistent and reproducible than that from Sprague-Dawley rats. It is possible that regulatory regions of the CYP4A gene may be different between these strains, leading to differences in response to the same stimulus. As is the case with apoA-I, we cannot rule out the possibility that P450 eicosanoids mediate their effects via other mechanisms.

The ability of P450 eicosanoids to regulate their own levels via PPAR was examined by looking at their effects on PPAR-responsive genes in the arachidonic acid metabolism cascade (summarized in Fig. 5). mRNA expression of CYP4A and sEH is highly responsive to treatment with fibrates and Wy-14,643 (Kimura et al., 1989; Tollet et al., 1994; Pinot et al., 1995). In this study, moderate differences in CYP4A1 and sEH RNA expression were observed with P450 eicosanoid treatment. One possibility for the minimal effect of P450 eicosanoids on CYP4A1 and sEH in hepatocytes, despite potent transactivation of PPAR in vitro, is an opposing effect of PPAR and PPAR activation on the expression of these genes. Although the role and expression of PPAR has been mainly described in the adipose tissues and macrophages (Bishop-Bailey, 2000), PPAR does exhibit weak expression in murine (Vidal-Puig et al., 1996) and human liver (Semple et al., 2006). Under pathological conditions such as hepatic steatosis and obesity, the hepatic expression of PPAR can also be up-regulated (Vidal-Puig et al., 1996; Gavrilova et al., 2003). It has been shown that the uncoupling protein 1 promoter can be responsive to both PPAR and PPAR (Barbera et al., 2001). Troglitazone, an agonist of both PPAR and PPAR, has opposing effects on PPAR and PPAR expression in mononuclear cells (Aljada et al., 2001). The transrepressive effect of PPAR in macrophages has also been described (Pascual et al., 2005; Ghisletti et al., 2007). Since Wy-14,643 is highly specific for PPAR, the expected induction of CPT1A, CYP4A1, and sEH was observed (Fig. 5), whereas when hepatocytes were treated with P450 eicosanoids, the potentially opposing dual effects elicited by both PPAR and PPAR may have accounted for the moderate increase in the expression of these genes. It will be of interest in future studies to determine whether the dichotomous effects of Wy-14,643 and P450 eicosanoids on PPAR-responsive genes in rat hepatocytes are due to the dual activation of PPAR and PPAR. Experiments in which the expression of each PPAR isoform is silenced could be used to isolate the effect of these two related nuclear receptors.

View larger version (17K): [in this window] [in a new window]   FIG. 5. Relationship between PPAR, P450s, sEH, and eicosanoids. Schematic of the results from this and other studies showing the metabolism of arachidonic acid by P450s and sEH. In this study, PPAR is activated by 20-HETE, EETs, and DHETs (depicted with bold arrows). PPAR may in turn regulate the expression of the enzymes responsible for the formation of these eicosanoids, resulting in a feedback mechanism (indicated by curved arrows). In the present study, expression of CYP4A and sEH genes was moderately increased, whereas CYP2C11 expression was markedly elevated after PPAR activation by P450 eicosanoids (enzymes examined are underlined).

In contrast to CYP4A and sEH, CYP2C11 expression was markedly induced after treatment with P450 eicosanoids. Of note is that the P450 eicosanoids are more potent in inducing CYP2C11 than is Wy-14,643. It has previously been reported that dietary administration of Wy-14,643, gemfibrizol, and di-n-butyl phthalate suppresses hepatic CYP2C11 expression in Fischer 344 and Sprague-Dawley rats (Corton et al., 1998). A similar decrease in CYP2C11 protein levels was observed after treatment of Fisher 344 rat hepatocytes with Wy-14,643 (Corton et al., 1998). The contrasting effects of Wy-14,643 on CYP2C11 expression in the present study and in this earlier study could be due to differences in the administered dose. It is difficult to compare our results with the earlier in vivo studies, in which Wy-14,643 was administered on a ppm basis for up to 13 weeks. However, in the previous rat hepatocyte studies, higher concentrations of Wy-14,643 (100–400 µM) were used than in our study (50 µM) and CYP2C11 down-regulation was not apparent until a concentration of 200 µM. It is possible that at lower concentrations of Wy-14,643, activation of PPAR is observed, whereas at higher concentrations, repression is apparent. A similar concentration-dependent effect has been reported for the PPAR activator, dehydroepiandrosterone (Ripp et al., 2003). Another possibility is that differences in the hepatocyte culture conditions between the two studies influenced the effect of Wy-14,643 on CYP2C11 expression. In the present study, hepatocyte conditions were optimized with respect to CYP4A induction by Wy-14,643, and these conditions may support activation and not repression of CYP2C11 by PPAR activators. As with the response seen with apoA-I and apoA-II, the route of agonist exposure may also play an important role in the effects the agonist can elicit on gene expression.

One of the best characterized roles of PPAR is the induction of adipocyte differentiation (Forman et al., 1995). Activation of PPAR can lead to an increase in proteins involved in lipid storage and metabolism during adipogenesis, such as adipocyte fatty acid binding protein and phosphoenolpyruvate carboxykinase (Tontonoz et al., 1994, 1995). Although we have shown that EETs, DHETs, and 20-HETE activate PPAR, the expression of CYP4A and CYP2C enzymes responsible for their formation have not been described in adipocytes. Therefore, the biological effects of P450 eicosanoids in adipocytes are not clear.

PPAR is well established as a mediator of hepatic lipid homeostasis. Although many studies have demonstrated the responsiveness of hepatic P450s to fibrate treatment, the biological consequences of increased P450s in the liver have not been elucidated. Through activation of PPAR, P450 metabolites of arachidonic acid alter apoA-I and apoA-II expression in hepatocytes and may play a role in regulating triglyceride transport. It will be interesting to examine the overall effect of P450 eicosanoids on fatty acid transport and metabolism by analyzing a panel of genes involved in these processes. P450 levels are often altered in experimental models of hypertension and diabetes. Because P450 eicosanoids may also be moderately autoregulatory by inducing PPAR-responsive P450s and sEH, it may be of therapeutic benefit to modify their levels by targeting PPAR.

	Acknowledgments

 
We acknowledge the help of Colleen Hefner at the University of California, San Francisco (UCSF) Liver Center for the preparation of primary hepatocytes and Dr. David Ginzinger of the UCSF Cancer Center for help with the design of primers and probes used in quantitative reverse transcriptase-PCR experiments.

	Footnotes

 
This work was supported by grants from the National Institutes of Health (Grants HL53994 to D.L.K. and GM31278 to J.R.F.). The UCSF Liver Center Core Facility (Cell and Tissue Biology) was supported by National Institutes of Health Grant P30 DK26743. V.Y.N was supported in part by predoctoral fellowships from the Pharmaceutical Research and Manufacturers of America Foundation (PhRMA) and the American Foundation for Pharmaceutical Education (AFPE).

Article, publication date, and citation information can be found at http://dmd.aspetjournals.org.

doi:10.1124/dmd.106.013839.

ABBREVIATIONS: P450, cytochrome P450; HETE, hydroxyeicosatetraenoic acid; EET, epoxyeicosatrienoic acid; sEH, soluble epoxide hydrolase; DHET, dihydroxyeicosatrienoic acid; PPAR, peroxisome proliferator-activated receptor; apoA, apolipoprotein A; PPRE, peroxisome proliferator response element; Wy-14,643, pirinixic acid, 4-chloro-6-(2,3-xylidino)-2-pyrimidinyl)thioacetic acid; 11,12-EEZE, 11,12-epoxyeicosa-8(Z)-enoic acid; T₃, 3,3',5-triiodo-L-thyronine; DMSO, dimethyl sulfoxide; RXR, retinoid X receptor ; CPT, carnitine palmitoyl transferase; h, human; m, murine; PCR, polymerase chain reaction; MK-0767, ±-5-[(2,4-dioxothiazolidin-5-yl)methyl]-2-methoxy-N-[[(4-trifluoromethyl)phenyl]methyl]-benzamide.

Address correspondence to: Dr. Deanna L. Kroetz, Department of Biopharmaceutical Sciences, University of California San Francisco, 1550 4^th Street, Box 2911, San Francisco, CA 94143-2911. E-mail: deanna.kroetz{at}ucsf.edu

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