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Originally published In Press as doi:10.1194/jlr.M400348-JLR200 on December 6, 2004 Originally published In Press as doi:10.1194/jlr.M400348-JLR200 on December 6, 2004 Originally published In Press as doi:10.1194/jlr.M400348-JLR200 on October 16, 2004

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Journal of Lipid Research, Vol. 46, 143-153, January 2005
Copyright © 2005 by American Society for Biochemistry and Molecular Biology

Resistin expression in 3T3-L1 adipocytes is reduced by arachidonic acid

Fred Haugen, Naeem Zahid, Knut T. Dalen, Kristin Hollung, Hilde I. Nebb and Christian A. Drevon1

Department of Nutrition, Institute of Basic Medical Sciences, University of Oslo, Oslo, Norway

Published, JLR Papers in Press, October 16, 2004. DOI 10.1194/jlr.M400348-JLR200

1 To whom correspondence should be addressed. e-mail: c.a.drevon{at}basalmed.uio.no


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The resistin gene is expressed in adipocytes and encodes a protein proposed to link obesity and type 2 diabetes. Increased plasma FFA is associated with insulin resistance. We examined the effect of separate FFAs on the expression of resistin mRNA in cultured murine 3T3-L1 adipocytes. The FFAs tested did not increase resistin expression, whereas both arachidonic acid (AA) and eicosapentaenoic acid (EPA) reduced resistin mRNA levels. AA was by far the most potent FFA, reducing resistin mRNA levels to ~20% of control at 60–250 µM concentration. Selective inhibitors of cyclooxygenase-1 and of mitogen-activated protein kinase kinase counteracted AA-induced reduction in resistin mRNA levels. Transient overexpression of sterol-regulatory element binding protein-1a (SREBP-1a) activated the resistin promoter, but there was no reduction in the abundance of ~65 kDa mature SREBP-1 after AA exposure. Actinomycin D as well as cycloheximide abolished the AA-induced reduction of resistin mRNA levels, indicating dependence on de novo transcription and translation.

Our data suggest that reductions in resistin mRNA levels involve a destabilization of the resistin mRNA molecule. An inhibitory effect of AA and EPA on resistin expression may explain the beneficial effect of ingesting PUFAs on insulin sensitivity.

Abbreviations: AA, arachidonic acid; BADGE, bisphenol A diglycidyl ether; C/EBP, CCAAT/enhancer binding protein; COX, cyclooxygenase; DHA, docosahexaenoic acid; EP, E-prostanoid; EPA, eicosapentaenoic acid; FP, F-prostanoid; LA, linoleic acid; LDH, lactate dehydrogenase; LN, {alpha}-linolenic acid; LUC, luciferase; LXR, liver X receptor; MAPK, mitogen-activated protein kinase; MEK, mitogen-activated protein kinase kinase; OA, oleic acid; PA, palmitic acid; PG, prostaglandin; PI3K, phosphatidylinositol 3-kinase; PPAR, peroxisome proliferator-activated receptor; RELM, resistin-like molecule; RXR, retinoic acid X receptor; SA, stearic acid; SREBP, sterol-regulatory element binding protein; TTA, tetradecylthioacetic acid

Supplementary key words fatty acid • insulin resistance • peroxisome proliferator-activated receptor {gamma} • liver X receptor {alpha} • sterol-regulatory element binding protein-1 • polyunsaturated fatty acids • prostaglandin • E-prostanoid 1 • E-prostanoid 4 • F-prostanoid


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
An essential function of the adipose tissue is to secrete adipokines, a group of hormones with effects on whole body metabolism. Adipokines are potentially important for the development of obesity-related diseases such as diabetes, cardiovascular diseases, and cancer (1). Insulin resistance is a hallmark of diabetes, and its development may involve several adipokines, including leptin (2), tumor necrosis factor-{alpha} (3), interleukin-6 (4), adiponectin (5), and resistin (6).

The resistin gene belongs to the resistin-like molecule (RELM) gene family (6–8). In mice, resistin is preferentially expressed in adipose tissue (6, 7), whereas RELM{alpha} is found mostly in adipose tissue and lung (7, 8), RELMß is in the intestine (8), and RELM{gamma} is expressed in hematopoietic tissues (9). In humans, resistin and RELMß homologous genes have been identified, with highest expression in macrophages (10, 11) and the intestine (8), respectively. RELMs encode peptides (105–117 amino acids) with 10 cysteine residues in the C terminus having identical spacing (7, 12). Resistin peptides are linked in pairs with disulfide bonds via the conserved cysteine residues and secreted as homodimers (13).

Administration of recombinant resistin reduced insulin-mediated glucose uptake in 3T3-L1 adipocytes (6) as well as L6 rat skeletal muscle cells (14). Resistin may also inhibit glucose oxidation and insulin-stimulated glycogenesis in isolated muscle from rats (15). Normally, insulin inhibits hepatic glucose production when plasma glucose levels increase in mice, but resistin attenuates this effect of insulin, promoting increased plasma glucose concentration (16). Mice lacking resistin exhibit low blood glucose levels after fasting as a result of reduced hepatic glucose production (17). Transgenic expression of the mouse resistin gene in adipose tissue impaired oxidative and nonoxidative glucose disposal in skeletal muscle and promoted glucose intolerance (15).

The resistin mRNA level was reduced by insulin-sensitizing thiazolidinedione ligands of the transcription factor peroxisome proliferator-activated receptor {gamma} (PPAR{gamma}) in 3T3-L1 adipocytes (6, 18, 19), although the regulation in vivo has been disputed (6, 20, 21). Both suppressors (insulin) and activators (ß-adrenoceptor agonists) of lipolysis have been reported to reduce resistin mRNA levels in 3T3-L1 adipocytes (18, 22, 23). Furthermore, intracellular levels of cAMP (forskolin) involving protein kinase A may modulate the level of resistin mRNA (18, 23).

Increased plasma FFA is associated with insulin resistance (24). However, dietary intake of PUFAs may alter the development of human diseases such as cardiovascular diseases, diabetes, and cancer by affecting enzyme activities, cellular signal transduction, and composition of membrane lipids (25). In addition, PUFAs may influence gene expression by direct interaction with the transcription machinery (26). Fatty acids may modulate the activity of the adipogenic transcription factors PPAR{gamma}, liver X receptor {alpha} (LXR{alpha}), and sterol-regulatory element binding protein (both SREBP-1a and SREBP-1c) (27–30).

Our aim in the present work was to study the effects of selected dietary FFAs on resistin expression in murine 3T3-L1 adipocytes to determine if resistin could represent a link between increased FFA plasma levels and insulin resistance. FFAs in general did not affect resistin mRNA levels. Only stearic acid (SA) promoted a small increase in resistin expression, but we observed a marked suppression of resistin mRNA levels by arachidonic acid (AA) and to a smaller extent by eicosapentaenoic acid (EPA). AA promoted its effects dependent on cyclooxygenase-1 (COX-1) and mitogen-activated protein kinase kinase (MEK) activity. In transfection experiments, SREBP-1a transactivated the resistin promoter, but we found no evidence that AA regulated SREBP-1 activity in 3T3-L1 adipocytes. Our data also suggested an active degradation of resistin mRNA dependent on the de novo synthesis of mRNA and protein.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell culture and supplements
3T3-L1 and COS-1 cells from the American Type Culture Collection (Rockville, MD) were cultured in DMEM supplemented with 10% FBS (Integro, Dieren, Holland), 2 mM L-glutamine, penicillin (50 U/ml), and streptomycin (50 µg/ml) at 37°C in 5% CO2 (31). 3T3-L1 fibroblasts were seeded on 12-well plates, and differentiation was initiated 2 days after confluence (18). After 6 days of differentiation, adipocytes were given fresh growth medium daily for up to 4 days and every other day thereafter. The cells were maintained for up to 8 days or 15 days after induction of differentiation, at which time specific treatments were initiated. Cells incubated under serum-free conditions were pretreated in DMEM containing 2 mM L-glutamine, streptomycin (50 µg/ml), penicillin (50 U/ml), and 1% FFA-free BSA (Sigma) for 24 h. The sodium salts of the fatty acids palmitic acid (PA; 16:0), SA (18:0), oleic acid (OA; 18:1, n-9), linoleic acid (LA; 18:2, n-6), {alpha}-linolenic acid (LN; 18:3, n-3), AA (20:4, n-6), EPA (20:5, n-3), or docosahexaenoic acid (DHA; 22:6, n-3) (Nu-Chek-Prep, Elysian, MN) were dissolved at 50°C to make 6 mM stock solutions in culture medium containing 2.4 mM FFA-free BSA (Sigma, St. Louis, MO) and stored for up to 1 month at –20°C under argon (32). Fatty acid-containing media were dilutions of stock solutions, and the molar ratio of BSA to FFA was 1:2.5 for all cell treatments. Cytotoxicity was controlled by determining lactate dehydrogenase (LDH) activity in the medium with the Cytoxicity Detection Kit (Boehringer Mannheim, Indianapolis, IN) after incubation with the different fatty acids. We used COX inhibitors dissolved in vehicle liquids: indomethacin (ethanol), flurbiprofen (DMSO), and meloxicam (DMSO), as well as the isoform-specific COX inhibitors SC-506 (DMSO) for COX-1 and NS-398 (DMSO) for COX-2 (Calbiochem, La Jolla, CA). MEK and phosphatidylinositol 3-kinase (PI3K) signaling was inhibited with DMSO-dissolved PD98059 and LY294002 (Calbiochem), respectively. Transcription and translation were inhibited with actinomycin D (DMSO) and cycloheximide (DMSO), respectively (Sigma). Other chemicals and the vehicles we used were prostaglandins (PGs; Calbiochem; ethanol), insulin (Sigma; water), 25-OH-cholesterol (Sigma; ethanol), darglitazone (AstraZeneca R&D, Molndal, Sweden; DMSO), T0901317 (Alexis, Lausen, Switzerland; DMSO), bisphenol A diglycidyl ether (BADGE; Fluka, Buchs, Switzerland; DMSO), and tetradecylthioacetic acid (TTA; Rolf K. Berge, University of Bergen, Norway; water). The cells were harvested in TRIzol Reagent (Invitrogen Life Technologies, Carlsbad, CA), and samples were stored at –70°C until RNA isolation.

RNA preparation and Northern blot/RT-PCR analysis
Total RNA was prepared using the TRIzol method, and RNA quality and quantity were determined spectrophotometrically. For Northern blots, RNA (10 µg) was electrophoresed on a 1.0% agarose gel containing 6.7% formaldehyde, transferred onto Hybond-N nylon membranes (Amersham, Little Chalfont, UK) by capillary action, and UV cross-linked (33). Primer sequences for amplifying cDNA from the mouse resistin gene (approved name, Retn) (AGCGGATGAAGAACCTTTCA and GGAGGAGACTGTCCAGCAAT) were designed based on the available mRNA sequence (gi:12228705) using Primer3 software (34). Using 3T3-L1 mRNA as a template, the 185 bp region of resistin cDNA was amplified on a LightCycler with the RNA Amplification Kit (Roche Diagnostics GmbH, Mannheim, Germany). The RT-PCR product was cloned and sequenced before hybridization of membranes (33). Membranes were probed with [{alpha}-32P]dCTP (Amersham)-radiolabeled cDNAs synthesized using a multiple DNA labeling system (Amersham) and purified on ProbeQuant G50 Micro columns (Amersham) (33). The tissue specificity of resistin mRNA detection using the probe described in this paper was the same as that reported previously (18). Blots were stripped and reprobed for other mRNAs: SREBP-1, PPAR{gamma} (33), LXR{alpha} (31), L27 (RPL27; American Type Culture Collection catalog no. 107385), and 36B4 (acidic ribosomal phosphoprotein P0; used here as an internal control). Hybridization signals were detected by the PhosphorImager and quantified with ImageQuant software (Molecular Dynamics, Sunnyvale, CA). For RT-PCR analysis of PG receptor mRNA, exon-spanning primer pairs for E-prostanoid 1 (EP1; ptger1; ACGCACACGATGTGGAAAT and CGCTGCAGGGAGTTAGAGTT; 127 bp), EP2 (ptger2; ATCACCTTCGCCATATGCTC and AGTATGGCAAAGACCCAAGG; 143 bp), EP3 (ptger3; GGATCATGTGTGTGCTGTCC and CCCATCTGTGTCTTGCATTG; 103 bp), EP4 (ptger4; ACCTCAGTTCCGGAAAGTTG and GCTCACGGTTCGATCTAGGA; 132 bp), and F-prostanoid (FP; ptgfr; AAGGCAGATCTCACCACCTG and TTCA-CAGGTCACTGGGGAAT; 137 bp) were designed with Primer3 (34) and mRNA input sequences (gi:7363446, gi:31560647, gi:6755217, gi:6679530, and gi:31560648, respectively). The one-step RT-PCR amplification of PG receptors was performed for each primer pair with Thermoscript Plus reverse transcriptase and Platinum Taq DNA polymerase (Invitrogen) using 1 µg of total RNA from 3T3-L1 adipocytes 15 days after induction of differentiation (24 h BSA-treated, serum-free conditions) or water only (negative control) as a template (annealing temperature, 60°C), and after 40 cycles the entire 25 µl reaction volumes were loaded onto 3% agarose gels and evaluated visually.

Whole cell protein extracts and Western blot analysis
After treatment, 3T3-L1 adipocytes were washed twice with PBS and added 100 µl/well of lysis buffer (10 mM Tris, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.5% Nonidet P-40, and 0.2 mM PMSF) with Complete protease inhibitor cocktail (Roche Diagnostics, Basel, Switzerland). Plates were shaken at 4°C for 30 min, and the resulting lysate was collected in a microtube and centrifuged at 13,000 rpm in a table centrifuge. The protein concentrations in the cleared lysates were assayed using the bicinchoninic acid (BCA) Protein Assay Reagent (Pierce, Rockford, IL). A total of 100 µg of protein from each lysate was boiled for 1 min in SDS buffer (62.5 mM Tris, pH 6.8, 2% SDS, 10% glycerol, and 50 mM DTT) before separation by SDS-PAGE on a Criterion precast 7.5% Tris-glycine gel (Bio-Rad, Hercules, CA) together with a prestained Precision Plus Dual Color protein standard (Bio-Rad) and finally blotted onto a 0.2 µm nitrocellulose membrane (Schleicher and Schuell, Dassel, Germany). The blots were incubated at room temperature in different TBS (pH 7.5) solutions: first, 30 min of blocking with 3% dry milk, then without washing, 2 h incubation with 1:100 dilution of SREBP-1 rabbit polyclonal antibody (K-10; Santa Cruz Biotechnology, Santa Cruz, CA) with 0.005% Tween 20 (Sigma), and then washing three times in 0.05% Tween 20, and finally, 1:4,000 dilution of goat anti-rabbit IgG(H+L)-HRP, mouse/human absorbed antibodies (Southern Biotechnology, Birmingham, AL) for 2 h followed by two washes in 0.05% Tween 20 and TBS alone. SREBP-1 was visualized on film (Eastman Kodak, Rochester, NY) using the ECL+ chemo-luminescence kit (Amersham).

Cloning of the mouse resistin promoter
The available resistin mRNA (gi:12228705) was used as a bait in a Basic Local Alignment Search Tool search against nonredundant and high-throughput genomic sequence databases to identify the mouse resistin promoter (annealed to gi: 9964856). Primers (TACTCGAGTTTTTGGTGTTTGGAGACAGGGTTT and TACTCGAGCACATACACAGACATGGACAGCACA) were designed with Primer3 software (34) to amplify a long fragment of the resistin promoter (5,358 bp; –4638 to +720). The PCR amplification was performed with Pfu Turbo DNA polymerase (Stratagene, La Jolla, CA) as described (31) from mouse genomic DNA (Clontech; BALB/c, #6650-1). The amplified promoter fragment was subsequently cloned into the pPCR-Script vector (Stratagene). A truncated version of this fragment, obtained by restriction cutting with NheI and XhoI generating a 3.3 kb fragment (–2610 to +720 of the resistin gene), was subcloned into the Nhe I/XhoI sites in the pGL3-basic firefly luciferase (LUC) reporter vector (Promega, Madison, WI) to generate the resistin reporter vector. A more truncated version of the resistin reporter (–393 to +720 of the resistin gene) was obtained by restriction digestion with KpnI followed by religation of the reporter vector.

Transfection and LUC assay
COS-1 cells were transiently transfected on six-well plates with firefly LUC reporter (5 µg) and cotransfected with pSV-ß-galactosidase (pCMV-ßgal; 3 µg; for use as an internal standard) and CCAAT/enhancer binding protein-{alpha} (C/EBP{alpha}) expression vector (1 µg) (35) with calcium phosphate precipitation (36). The mouse C/EBP{alpha} expression vector (pCDNA-3.1-C/EBP{alpha}) was described previously (37) and was a gift from O. A. MacDougald (University of Michigan, Ann Arbor, MI). Total DNA concentration was adjusted to 12 µg with empty expression vector and pGL3-basic vector. Undifferentiated 3T3-L1 fibroblasts were plated on 24-well plates and were semiconfluent on the day of transfection. Wells were added to a transfection solution containing 1 µl of LipofectAMINE reagent, 4 µl of LipofectAMINE Plus reagent (Invitrogen Life Technologies), and ~500 ng of DNA (same amount for all wells in separate experiments), including firefly LUC reporter (200 ng) and pTK Renilla LUC expression vector (20 ng; Promega) and different transcription factor expression vectors (10 ng of each if not stated otherwise). The retinoic acid X receptor (RXR{alpha}; pCMX-RXR) and LXR{alpha} (pCMX-LXR) expression vectors have been described (38) and were provided by D. J. Mangelsdorf (University of Texas Southwestern Medical Center, Dallas, TX). The PPAR{gamma} (pSG5 hPPAR{gamma}2) expression vector was a gift from J. Auwerx (CNRS/INSERM/Université Louis Pasteur, Illkirch, France) (31, 39). The SREBP-1a (pCMV-CSA10) expression vector was provided by T. F. Osborne (University of California, Los Angeles, CA) (40). The C/EBP{alpha} expression vector is described above. Total DNA concentration was adjusted using empty reporter and expression vectors. After 3 h of transfection, medium containing supplements of AA or PGs was added and cells were incubated for 24 h. Cells were harvested in 200 µl of lysis buffer, and LUC activities were measured using the dual LUC assay kit (Promega) in a Luminometer (TD-20/20 Luminometer; Turner Designs, Sunnyvale, CA).

Presentation of data and statistical analysis
Relative resistin mRNA levels were calculated as the ratio of resistin and 36B4 signal intensities. The effects of different incubations were expressed relative to the control incubation, which was assigned a value of 100%, and are presented in bars representing means ± SEM of three to five experiments. The statistical significance of differences between incubations was assessed with a two-tailed Student's t-test (* P < 0.05, ** P < 0.01).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
To test whether fatty acids affect the transcriptional level of resistin expression, we differentiated 3T3-L1 adipocytes until day 8 or day 15 and incubated with vehicle (BSA) or with 250 µM OA, LN, AA, or EPA. Initially, we observed that resistin mRNA levels were consistently reduced by incubation with AA in 3T3-L1 adipocytes (Fig. 1A). Judging by the size of the intracellular lipid droplets, cells differentiated for 15 days had more lipids than after 8 days of differentiation (data not shown). The PUFAs we tested (LN, AA, and EPA) appeared to reduce resistin expression at day 8, whereas AA reduced resistin mRNA at day 15 (Fig. 1A). No effect was observed with OA (Fig. 1A). Leakage of LDH into the media after incubation with the different FFAs was 5–6% (day 8) and 2–4% (day 15) of total LDH released after complete cell lysis and did not indicate toxic side effects of the different FFAs. In the subsequent experiments with FFAs, we omitted serum and used the more lipid-loaded and fully differentiated (day 15) 3T3-L1 adipocytes. The level of resistin mRNA was significantly reduced at concentrations of >=30 µM in adipocytes incubated with AA, and expression levels at ~20% of the BSA control were observed at concentrations of 60–250 µM (Fig. 1B). Incubation with 250 µM EPA reduced the level of resistin mRNA to 78 ± 11%, whereas 30 µM SA promoted a significant increase in resistin expression to 122 ± 8% of the control (Fig. 1B). There was no significant effect on resistin expression of PA, OA, LA, LN, and DHA in the range 15–250 µM (Fig. 1B). To determine the time course of the AA effect on resistin expression, we harvested adipocytes at different time points after incubation with 250 µM OA or AA. There was a reduction of resistin mRNA to ~30% of the control after 12 h of incubation with AA and to ~20% after 24 and 48 h, whereas no effect was observed with OA (Fig. 1C).



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  		Fig. 1. Reduction of resistin mRNA by arachidonic acid (AA). Resistin mRNA was monitored in 3T3-L1 adipocytes by Northern blotting using 36B4 as an internal control. A: After 8 days (d 8) or 15 days (d 15) of differentiation, the adipocytes were incubated for 48 h with fatty acid-free BSA vehicle (–) or 250 µM BSA-bound oleic acid (OA), {alpha}-linolenic acid (LN), AA, or eicosapentaenoic acid (EPA) in medium with (+ Serum) and without (– Serum) 10% FBS. Further FFA incubations of adipocytes were performed with 15 day differentiated adipocytes incubated without FBS. B: Resistin mRNA levels in adipocytes incubated with BSA-bound palmitic acid (PA), stearic acid (SA), OA, linoleic acid (LA), LN, AA, EPA, or docosahexaenoic acid (DHA) at 0, 15, 30, 60, 125, and 250 µM for 48 h. Relative resistin mRNA levels were calculated as the ratio of resistin and 36B4 signal intensities related as percentage to the respective negative control stimulated with BSA only. Values shown are mean percentages of three to four experiments. * P < 0.05, ** P < 0.01. C: Relative resistin mRNA levels in adipocytes incubated for 0, 3, 6, 12, 24, or 48 h with 250 µM OA or AA. The plots represent means of two determinations.

 
PGs of the 2-series are synthesized from AA, and we tested if PG synthesis was involved in regulating resistin expression. Unspecific COX inhibitors (indomethacin, flurbiprofen, and meloxicam) neutralized the AA-induced reduction in resistin mRNA (Fig. 2A). Experiments with specific inhibitors of COX-1 (SC-506) and COX-2 (NS-398) revealed that SC-506, but not NS-398, could inhibit the AA effect on resistin mRNA (Fig. 2B). Incubations with AA alone reduced resistin mRNA levels to 6 ± 1% of control, whereas this was reversed to 71 ± 17% of control by SC-506 (Fig. 2B), implying a mechanism involving COX-1. After incubation with AA and NS-398, resistin mRNA was 30 ± 16% of the control, but the AA effect with this inhibitor was not significantly different from the AA effect without the inhibitor added (Fig. 2B). As a control, OA was included in the experiments and had no effect on resistin expression (Fig. 2B). We tested if the effect of AA on resistin expression was mimicked by direct exposure of adipocytes to different concentrations of PGs. At concentrations of 1 and 10 µM, PGE2 downregulated resistin mRNA levels to 53 ± 8% and 19 ± 6% of control, respectively (Fig. 2C), and no effect was seen using 0.001, 0.01, or 0.1 µM. In addition, 0.1, 1, and 10 µM PGF2{alpha} reduced resistin mRNA levels to 37 ± 9%, 24 ± 4%, and 19 ± 3% of control, respectively (Fig. 2C). The COX-1 selective inhibitor SC-506 did not affect the inhibitory effect of PGE2 and PGF2{alpha} on resistin expression (Fig. 2D). The suggested PPAR{gamma} agonist 15deoxy-{delta}12,14-PGJ2 (10 µM) did not have any significant effect on resistin expression (data not shown). Physiological effects of PGs are mediated via interactions with G-protein-coupled receptors at the cell surface (41). By the use of known prostanoid receptor agonists, we examined the involvement of EP and FP receptors in the regulation of resistin mRNA. We incubated the differentiated 3T3-L1 cells with agonists of the different prostanoid receptors in the range 10–14 to 10–4 M and measured resistin mRNA levels by Northern blotting (n = 3–4). The EP1/EP3 agonist sulprostone and the EP2/EP4 agonist misoprostol had an EC50 > 1 µM, whereas the EP2 agonist butaprost did not decrease resistin mRNA levels dose-dependently (data not shown). The FP agonist fluprostenol EC50 was in the range of 0.1–10 nM (data not shown). By RT-PCR analysis (see Materials and Methods), we could detect mRNA encoding EP1, EP4, and FP in 3T3-L1 adipocytes, but not EP2 and EP3 (Fig. 2E). These results indicate that reduced resistin mRNA expression by AA depends on its conversion to PGs and possibly on the subsequent autocrine activation of EP1, EP4, and FP.



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  		Fig. 2. AA-induced reduction in resistin mRNA depends on cyclooxygenase-1 (COX-1) activity and is mimicked by prostaglandins (PGs). Northern blots show resistin and reference 36B4 mRNA in adipocytes incubated with BSA vehicle (–) or 250 µM OA or AA for 24 h. A: Adipocytes were preincubated at 1 h and throughout the fatty acid stimulation with vehicle (Control) or with inhibitors of COX-1 and COX-2: 10 µM indomethacin (Indo), 100 µM flurbiprofen (Flur), and 10 µM meloxicam (Melo). Results shown are representative of two experiments. B: Similarly, cells were treated with vehicle (Control) or isoform-specific inhibitors: 0.3 µM SC-506 (SC; COX-1 inhibitor) and 30 µM NS-398 (NS; COX-2 inhibitor). Columns represent means ± SEM of the relative resistin levels from three to five experiments with the BSA incubations set to 100% (* P < 0.05). C: Relative resistin mRNA levels in adipocytes incubated for 24 h with ethanol vehicle alone or different concentrations of PGE2 and PGF2{alpha} (10–4, 10–3, 10–2, 10–1, 1, or 10 µM). The plots represent means ± SEM of the relative resistin levels from five experiments with the ethanol incubations set to 100% (* P < 0.05, ** P < 0.01). D: Relative levels of resistin mRNA determined by Northern blotting in serum-starved adipocytes incubated in media with 0.1% fatty acid-free BSA supplements. Cells were first treated for 1 h with DMSO vehicle (–) or 0.3 µM SC-506 (SC; COX-1 inhibitor). Incubations with inhibitor continued for another 24 h after the addition of 10 µM PGE2, 10 µM PGF2{alpha} or ethanol vehicle only (set to 100%). Values shown are means ± SEM of three experiments. E: RT-PCR analysis of the E-prostanoid receptors EP1, EP2, EP3, and EP4 and F-prostanoid (FP) mRNA expression in 3T3-L1 adipocytes 15 days after induction of differentiation. One-step RT-PCR was performed using water only (–) or 1 µg of total RNA (+) as a template.

 
We examined if the AA-induced reduction of resistin mRNA expression depends on activation of the mitogen-activated protein kinase (MAPK) and PI3K pathways, both of which are important for signal transduction (e.g., leading to transcription factor activation). The inhibitory effect of AA on resistin expression was reversed by an inhibitor of MEK (PD98059) in the MAPK pathway; resistin mRNA levels were reduced to 84 ± 7% of control by AA together with PD98059, as opposed to 6 ± 1% with AA alone (Fig. 3A). Incubation with a PI3K inhibitor (LY294002) partly inhibited the AA effect (33 ± 4% of control; Fig. 3A). At concentrations >50 µM, LY294002 alone had an inhibitory effect on resistin expression (data not shown), and side effects of the inhibitor in combination with fatty acids cannot be excluded. Control incubations with OA in combination with the inhibitors did not affect resistin expression (Fig. 3A). Furthermore, the inhibitory effects of PGE2 and PGF2{alpha} on resistin expression were partially antagonized by the MEK inhibitor PD98059 (Fig. 3B). This finding indicates that activation via the MAPK pathway is involved in AA-induced reduction of resistin mRNA expression, whereas signaling through the PI3K pathway seems less important.



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  		Fig. 3. AA- and PG-induced reduction in resistin mRNA involves mitogen-activated protein kinase kinase (MEK) activity. A: Northern blots of resistin and reference 36B4 mRNA in adipocytes incubated with BSA vehicle (–) or 250 µM OA or AA for 24 h in the presence of DMSO vehicle (Control), 50 µM PD98059 (PD; MEK inhibitor), or 10 µM LY294002 (LY; phosphatidylinositol 3-kinase inhibitor). Columns represent means ± SEM of relative resistin mRNA levels from three to four experiments with the BSA controls set to 100% (* P < 0.05, ** P < 0.01). B: Relative resistin mRNA levels were determined by Northern blot analysis in adipocytes incubated in media with 0.1% FFA-free BSA. Cells were treated for 1 h in advance with DMSO vehicle (–) or 50 µM PD98059 (PD; MEK inhibitor) and for another 24 h after the addition of 10 µM PGE2, 10 µM PGF2{alpha} or ethanol vehicle only (set to 100%). Values shown are means ± SEM of three experiments (** P < 0.01).

 
As potential mediators of AA-regulated resistin expression, we investigated if the insulin signaling pathway and activity of the nuclear receptors PPAR{gamma} and LXR{alpha} affect resistin expression. In the absence of serum, adipocytes differentiated for 15 days responded to insulin with a dose-dependent reduction in resistin mRNA (Fig. 4A, upper panels). PD98059 reduced the effect of insulin on resistin expression (Fig. 4A, lower panels). Contrary to our earlier experiments performed in the presence of serum (18), serum-starved cells did not respond to darglitazone with decreased resistin expression even at high doses (Fig. 4A, upper panels). In contrast, the LXR agonist T0901317 dose-dependently reduced the expression of resistin mRNA in the absence of serum (Fig. 4A, upper panels). Adding the MEK inhibitor PD98059 did not affect the LXR-induced reduction of resistin mRNA expression (Fig. 4A, lower panels), suggesting that the LXR activator executes its effect independently of the MAPK pathway. In separate experiments, we observed that the LXR agonist T0901317 reduced the resistin mRNA levels to 54 ± 4% of the DMSO-treated control (Fig. 4B). This shows for the first time that LXR{alpha} may regulate resistin expression.



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  		Fig. 4. The liver X receptor {alpha} (LXR{alpha}) agonist T0901317 downregulated resistin mRNA levels. A: Northern blots of resistin and reference 36B4 mRNA from serum-starved adipocytes incubated for 24 h with insulin (0, 0.01, 0.1, 1, 10, and 100 nM), the peroxisome proliferator-activated receptor {gamma} (PPAR{gamma}) agonist darglitazone, or the LXR{alpha} agonist T0901317 (both 0, 0.001 0.01, 0.1, 1, and 10 µM) in the absence (–) or presence (PD) of 50 µM PD98059 dissolved in vehicle. B: Adipocytes were incubated for 24 h with vehicle (–) or 1 µM T0901317. Relative resistin mRNA levels as percentages of the DMSO controls of three experiments are presented as mean percentages ± SEM (* P < 0.05).

 
To further clarify the role of PPAR{gamma} in resistin expression, we tested the combination of AA and darglitazone. The presence of the PPAR{gamma} agonist darglitazone interfered with the inhibitory effect of AA on resistin expression (Fig. 5A). Without darglitazone, AA reduced resistin mRNA to 19 ± 8% of the fatty acid-free control, but when darglitazone was included in the media this effect was significantly changed to 67 ± 10% (Fig. 5B). The PPAR{gamma} antagonist BADGE (42) did not affect the AA-induced reduction in resistin mRNA (data not shown).



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  		Fig. 5. The PPAR{gamma} agonist darglitazone interfered with the inhibitory effect of AA on resistin mRNA levels. A: Northern blots from adipocytes were incubated for 48 h with BSA vehicle (–) or 250 µM OA, LN, AA, or EPA in medium with (+ Serum) and without (– Serum) 10% FBS in the presence of 0.1 µM darglitazone. B: Relative resistin mRNA levels in serum-starved adipocytes after 24–48 h of incubation with BSA or 250 µM AA together with vehicle (–) or 0.1 µM darglitazone. Columns represent percentages of the respective BSA controls given as means ± SEM of four experiments (** P < 0.01).

 
To test if AA-inhibited resistin expression is dependent on the direct interactions of transcription factors with the resistin promoter, we constructed a firefly LUC reporter containing the –2610 to +720 bp and the –393 to +720 bp fragments of the mouse resistin promoter (Fig. 6A). C/EBP{alpha} has been shown to be a positive regulator of the resistin promoter (43, 44), and we tested if the promoter region included in our constructs was able to convey activation by a C/EBP{alpha} expression vector. Transiently overexpressing C/EBP{alpha} in COS-1 cells induced a 60-fold increase in relative LUC activity conferred by the –2610 to +720 bp resistin promoter (Fig. 6B). With this reporter in COS-1 cells, PPAR{gamma}2 overexpression alone did not affect LUC activity, whereas together with darglitazone it induced a 2-fold increase (Fig. 6C). In the same assay, cotransfection with the LXR{alpha} expression plasmid alone or with T0901317 increased LUC activity 2-fold and 3-fold, respectively (Fig. 6C). We tested the capacity of several adipocyte-enriched transcription factors to activate the resistin promoter in undifferentiated 3T3-L1 fibroblasts. Although to a much lesser extent than in COS-1, C/EBP{alpha} overexpression also activated the –2610 to +720 bp resistin reporter construct in 3T3-L1 fibroblasts, resulting in a 3-fold increase in relative LUC activity (Fig. 6D). When we overexpressed mature SREBP-1a in 3T3-L1 fibroblasts, we observed an >12-fold increase in relative LUC activity with the –2610 to +720 bp resistin promoter construct (Fig. 6D). Transfection with RXR{alpha} induced a small (3-fold) increase in relative LUC activity, and the effect of PPAR{gamma}2 or LXR{alpha} in combination with RXR{alpha} was only slightly inhibitory (Fig. 6D). Incubation of fibroblasts with AA or PGE2 did not significantly affect the ability of C/EBP{alpha} or SREBP-1a to induce the resistin reporter (Fig. 6D). However, relative LUC activity induced by SREBP-1a was reduced by ~50% after incubation of the transfected fibroblasts with PGF2{alpha} (Fig. 6D). In undifferentiated 3T3-L1 fibroblasts, SREBP-1a activated the –2610 to +720 and the –393 to +720 resistin constructs in a concentration-dependent manner, inducing up to a 12-fold increase in relative LUC activity for both constructs (Fig. 6E).



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  		Fig. 6. Resistin promoter activity is enhanced with overexpression of CCAAT/enhancer binding protein-{alpha} (C/EBP{alpha}) and sterol-regulatory element binding protein-1a (SREBP-1a). A: The –2610 to +720 bp and the –393 to +720 bp fragments of the mouse resistin promoter were cloned into firefly luciferase (LUC) reporter plasmid. B: The reporter vector alone (pGL3) or the –2610/+720 resistin-firefly LUC construct was cotransfected with the C/EBP{alpha} expression vector or the empty vector (–) together with the ß-galactosidase expression vector (as an internal control) into COS-1 cells. Relative LUC activity was calculated as the ratio of firefly LUC activity to ß-galactosidase activity, and fold induction is relative to the –2610/+720 resistin-firefly LUC construct in the absence of C/EBP{alpha} expression vector. Values shown are means ± SEM of three experiments. C: The –2610/+720 resistin-firefly LUC construct was cotransfected into COS-1 cells with the PPAR{gamma} or LXR{alpha} expression vector. After transfection, the activity of PPAR{gamma} and LXR{alpha} was amplified by incubating cells with vehicle (–) or 1 µM of the appropriate ligand (Lig.), darglitazone or T0901317. Relative LUC activity was calculated as the ratio of firefly LUC activity to ß-galactosidase activity, and fold induction is relative to the –2610/+720 resistin-firefly LUC construct cotransfected with empty expression vector. Bars represent means of two experiments performed in triplicate. D: The –2610/+720 resistin-firefly LUC construct was cotransfected with empty expression vector (Vector) or combinations of transcription factor expression vectors [retinoic acid X receptor (RXR{alpha}), PPAR{gamma}2, LXR{alpha}, C/EBP{alpha}, and SREBP-1a] into 3T3-L1 preadipocytes. Renilla LUC expression vector was included in the transfections as an internal control. Cells were incubated with vehicle (–), 100 µM AA, 10 µM PGE2 (PGE), or 10 µM PGF2{alpha} (PGF). Relative LUC activity was calculated as the ratio of firefly LUC activity to Renilla LUC activity, and fold induction is relative to BSA incubation in the absence of transcription factor expression vectors. Values shown are means ± SEM from a representative experiment of two experiments performed in triplicate. E: The –2610/+720 and –393/+720 resistin-firefly LUC constructs were cotransfected with different amounts (ng/well) of SREBP-1a expression vector in 3T3-L1 preadipocytes as above. Values shown are means ± SEM from a representative experiment of two experiments performed in triplicate.

 
We also examined other potential transcriptional and posttranscriptional processes in the AA-induced reductions in resistin mRNA. Incubation of adipocytes with AA did not change mRNA levels of LXR{alpha}, SREBP-1, and PPAR{gamma} (Fig. 7A), suggesting that transcriptional regulation of these transcription factor genes is of minor importance for resistin expression. SREBP-1 activity is also regulated at the posttranslational level, and cleavage of SREBP-1 is required to release its mature and active form (45). However, in adipocytes, the level of the mature SREBP-1 protein was unchanged after incubation with AA, EPA, or 25-hydroxycholesterol (Fig. 7B). This does not show regulated SREBP-1 maturation in 3T3-L1 adipocytes, and it does not support SREBP-1 maturation as an important factor in AA-regulated resistin expression. The effect of AA on resistin mRNA levels in adipocytes was abolished when the cells were incubated with cycloheximide or actinomycin D for 24 h (Fig. 7C). Surprisingly, there was no reduction in the amount of resistin mRNA in the controls after incubation with actinomycin D for 24 h (Fig. 7C), whereas extending the incubation to 48 h resulted in a marked reduction of resistin mRNA in the controls (Fig. 7D). The suppressive effect on resistin mRNA levels of both AA and T0901317 was more rapid than the effect of actinomycin D alone (Fig. 7D).



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  		Fig. 7. Reduced levels of resistin mRNA are not associated with reduced expression of adipogenic transcription factors but require de novo protein and mRNA synthesis. A: Northern blots showing mRNA encoding resistin, SREBP-1, LXR{alpha}, and PPAR{gamma} after 48 h of incubation of adipocytes with vehicle (–) or PA, OA, LA, LN, AA, EPA, DHA, or tetradecylthioacetic acid (TTA). Bands of internal controls (L27 and 36B4) were included to demonstrate similar loading of wells. B: Immunoblot analysis showing the precursor (P) and mature (M) forms of SREBP-1 extracted from adipocytes incubated for 24 h with vehicle (–) or 250 µM AA, 250 µM EPA, or 5 µg/ml 25-hydroxycholesterol (HOC). The presented blot is representative of three experiments. C: Northern blot analysis of resistin mRNA in adipocytes incubated for 24 h with BSA vehicle (–), 250 µM OA or AA in the absence (Control) or presence of 10 µM cycloheximide (inhibitor of protein synthesis) or 1 µg/ml actinomycin D (inhibitor of RNA synthesis). Data are representative of two experiments. D: Northern blot showing adipocytes incubated with vehicle (–), 250 µM AA, or 1 µM T0901317 (T) in the absence (Control) or presence of 1 µg/ml actinomycin D for 24 or 48 h.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we have searched for a link between the availability of fatty acids and resistin gene expression, with possible implications for the development of insulin resistance. We incubated 3T3-L1 adipocytes with different FFAs, and only SA induced a marginal increase in resistin mRNA. We observed that AA at >=30 µM promoted a marked reduction in resistin mRNA in 3T3-L1 adipocytes independent of serum constituents and the degree of intracellular lipid loading. The strong effect indicates that AA might be a physiological regulator of resistin expression in adipocytes. A much smaller reduction in resistin mRNA was also induced by EPA at 250 µM. In contrast to our data, an earlier study of cultured primary rat adipocytes indicated the inhibition of resistin mRNA expression by OA (46). Our present study reveals a strong inhibition of resistin mRNA expression specifically by AA and does not indicate a regulation by FFAs in general.

Our experiments with isoform-specific COX inhibitors indicate that COX-1 and not COX-2 activity is required for the inhibitory effect of AA on resistin expression. This is in accordance with findings demonstrating that AA inhibits fatty acid synthase and S14 gene expression in 3T3-L1 adipocytes via COX (47). Also, COX-1 is the most abundant isoform in differentiated 3T3-L1 cells, with COX-2 being expressed at a much lower level (48). PGE2 and PGF2{alpha} are secreted from 3T3-L1 adipocytes (49, 50), and we observed that high doses (10 µM) of both of these PGs reduced resistin expression to the same extent as AA, in agreement with the fact that AA is the precursor of PGs. Earlier studies have reported detectable expression of the FP receptor gene (ptgfr) by RT-PCR in 3T3-L1 adipocytes (51). Quite recently, mRNA expression of the four EP receptor genes ptger1, ptger2, ptger3, and ptger4, expressing the EP1, EP2, EP3, and EP4 receptors, respectively, was investigated in 3T3-L1 adipocytes (52). In agreement with those findings, we were able to detect mRNAs encoding FP, EP1, and EP4 but no expression of EP2 and EP3 by RT-PCR analysis of 3T3-L1 adipocytes. Earlier studies show that the Kd values of PGE2 and PGF2{alpha} to their respective murine prostanoid receptors are in the 1–20 nM range (53). The dose-response curve of PG-regulated resistin expression we present here shows an effect of PGF2{alpha} in the physiological range (>10 nM), whereas the effect of PGE2 depends on its superphysiological concentrations (>100 nM). Furthermore, the FP receptor agonist fluprostenol effectively downregulated resistin mRNA at nanomolar concentrations (EC50 ~ 0.1–10 nM). High doses of the EP1/EP3 agonist sulprostone and the EP2/EP4 agonist misoprostol (EC50 > 1 µM) were required, whereas the EP2 agonist butaprost had no effect on resistin mRNA levels. The high doses of PGE2, sulprostone, and misoprostol required to obtain an effect suggest that EP receptors are of small importance for the regulation of resistin mRNA levels, and the effect may be indirect. However, the potent effects of PGF2{alpha} and flurprostenol suggest that FP receptor is a major mediator of PG-regulated resistin gene expression. The alleged biological PPAR{gamma} ligand 15deoxy-{delta}12,14-PGJ2 (54) did not have any effect and therefore seems not to play a role in regulating resistin expression. Furthermore, pharmacological COX-1 inhibitors did not block the effect of PGs on resistin expression, supporting the notion that enzymatic transformation of AA into PGs is required to downregulate resistin mRNA.

Of the prostanoid receptors present in 3T3-L1 adipocytes, EP1 acts via the phospholipase C/inositol trisphosphate pathway, EP4 receptors activate the cAMP/protein kinase A pathway, and FP receptors signal through the heterotrimeric G-protein Gq (55). We also found that a specific MEK inhibitor (PD98059) nearly abolished the AA-induced reduction of resistin expression. This inhibitor also partially blocked the downregulation of resistin mRNA when PG was included in the incubation media. This indicates that MEK activity is required downstream of COX-1 activity for the full AA-mediated downregulation of resistin mRNA. This suggests a mechanism involving the secretion of PGs and subsequent activation of G-protein-coupled receptors at the cell surface and further signaling via the MAPK pathway. Moreover, an inhibitor of PI3K (LY294002) did not block the AA-induced reduction of resistin mRNA. This is in contrast to observations by Song et al. (44) demonstrating that overexpression of PI3K suppressed resistin expression.

The transcription factors PPAR{gamma}, LXR{alpha}, and SREBP-1 are induced during adipogenesis, and their transcriptional activities may be regulated by PUFAs. Previously, we have shown that the PPAR{gamma} agonist darglitazone reduced resistin mRNA in the presence of serum (18). In the present study, we observed that the downregulatory effect of darglitazone was dependent on serum in the culture medium. Under these conditions, we also observed that darglitazone neutralized the inhibiting effect of AA, suggesting an interaction between the two. We first suspected that AA could execute its effect on resistin via an association with the PPAR{gamma} ligand binding pocket and that darglitazone under these conditions functioned as a PPAR{gamma} antagonist. Incubation with BADGE (42), a ligand for PPAR{gamma} that can antagonize thiazolidinedione stimulation of PPAR{gamma} transcriptional activity, did not affect resistin mRNA levels. However, in contrast to darglitazone, BADGE did not counteract the reduction of resistin mRNA induced by AA. Our data do not support the idea that the interference between darglitazone and AA is the result of competition for the PPAR{gamma} ligand binding pocket. PPAR{gamma} ligands exert PPAR{gamma}-independent effects in part through activation of the MAPK pathway (56). On the other hand, MAPK-mediated phosphorylation may reduce PPAR{gamma} transcriptional activity (57), and it is possible that the AA-induced reduction in resistin mRNA is modulated by darglitazone via MAPK signaling affecting PPAR{gamma} transcriptional activity.

In the present study, we also found that the LXR{alpha} agonist T0901317 is a negative regulator of resistin mRNA levels in 3T3-L1 adipocytes, which is in accordance with LXR{alpha} agonists promoting increased expression of mRNA for the glucose transporter GLUT4 and increasing insulin sensitivity (31). This made us speculate that the downregulation of resistin expression by AA was via the LXR{alpha} nuclear receptor. With transient cell transfections, we attempted to narrow the list of candidate transcription factors involved in the AA-induced inhibition of resistin expression. A firefly LUC reporter gene controlled by the resistin proximal promoter was used to test the promoter's responsiveness to various adipocyte-enriched transcription factors. We found that transfection of a C/EBP{alpha} expression vector in COS-1 cells caused more than 60-fold induction of the resistin promoter activity. It has been shown that C/EBP{alpha} can transactivate the resistin promoter (43, 44). In undifferentiated 3T3-L1 fibroblasts, the effect of C/EBP{alpha} was less prominent, possibly because of mechanisms repressing differentiation at the preadipocyte stage. We further tested if the reduction in resistin mRNA levels by PPAR{gamma} and LXR{alpha} ligands could be paralleled in transfection studies. Both PPAR{gamma} and LXR{alpha} overexpression reduced RXR{alpha}-induced resistin promoter activity, but these effects were so small that their biological significance is uncertain. In addition, we did not see any effect of AA treatment on mRNA levels of PPAR{gamma} and LXR{alpha}. In accordance with similar experiments with the human resistin promoter (58), we found that SREBP-1a is a dose-responsive positive regulator of resistin promoter activity in 3T3-L1 fibroblasts. This is particularly interesting because the SREBP-1 transcription factors are involved in PUFA-inhibited gene expression in the liver (59). However, the mRNA levels of SREBP-1 (transcripts of both the -a and -c isoforms) were unaffected by incubation with AA, suggesting that AA does not reduce resistin mRNA levels by inhibiting SREBP-1 transcription. Because SREBP-1 activity requires proteolysis of the membrane-bound precursor and its subsequent translocation to the nucleus (45), we investigated if AA affected the amount of mature SREBP-1 (both the -a and -c isoforms), but our experiments did not support the involvement of a regulated SREBP-1 maturation.

Applying inhibitors of transcription (actinomycin D) and translation (cycloheximide) indicated that both de novo transcription and translation are necessary for the AA-induced reduction of resistin mRNA. Furthermore, inhibiting transcription with actinomycin D per se had no apparent effect on resistin mRNA levels after 24 h, and a reduction was only seen after 48 h. This indicates that the resistin transcript is stable, with a half-life of >24 h. It should also be noted that the reduction in resistin mRNA observed after incubation with AA occurred faster than by blocking transcription with actinomycin D. This suggests that resistin mRNA is actively degraded through a process induced by AA and that AA is not likely to exert its effect by regulating resistin transcription via the gene's promoter. Our results also indicate that resistin mRNA is reduced by the LXR agonist (T0901317) in a similar manner as AA. A mechanism involving the degradation of mRNA is likely to reduce resistin mRNA levels in 3T3-L1 adipocytes not only after AA and LXR agonist incubation but also after exposure to insulin (60).

In conclusion, we observe that AA reduces resistin mRNA levels in 3T3-L1 adipocytes involving COX-1 and MEK activity, and possibly a destabilization of the resistin transcript.


   ACKNOWLEDGMENTS
 
F.H., N.Z., and K.H. were research fellows supported by the Norwegian Cancer Society. K.T.D. was supported by The Medical Faculty, University of Oslo. The authors thank Anne Randi Alvestad, Hege Henriksen, Marianne Sanderud, Assim Dutta-Roy, Janne E. Reseland, Arild C. Rustan, and Trine Ranheim for important technical, administrative, and scientific support. The mouse C/EBP{alpha} expression vector (pCDNA-3.1-C/EBP{alpha}) was a generous gift from O.A. MacDougald (Ann Arbor, MI), the RXR{alpha} (pCMX-RXR) and LXR{alpha} (pCMX-LXR) expression vectors were provided by D.J. Mangelsdorf (Dallas, TX), and the PPAR{gamma} (pSG5hPPAR{gamma}2) expression vector was a gift from J. Auwerx (Illkirch, France). The authors are also grateful for the financial support from the Throne Holst Foundation for Nutrition Research, the Freia Research Foundation, and the Norwegian Odd Fellow Medical-Scientific Research Foundation.

Manuscript received September 14, 2004 and in revised form September 29, 2004.


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