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/RXR Pathway in Chickens and HD11 Cell Lines1
Department of Animal Science, University of California, Davis, CA 95616
2 To whom correspondence should be addressed. Email: kcklasing{at}ucdavis.edu.
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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, ß, and 
, and retinoic acid X receptor (RXR) and mRNA levels. In Expt. 1, macrophages were collected from broiler chicks fed 3 or 6% dietary fat (g/100 g) with 0, 25, and 50 mg lutein/kg feed for 23 d. In Expt. 2, using a 3 x 3 factorial, eicosapentaenoic acid (EPA) at 0, 15 and 50µmol/L and lutein at 0, 10 and 100µmol/L were applied to HD11 cell culture for 24 h. In both experiments, cells were stimulated with lipopolysaccharide before RNA isolation. Lutein interacted with fat in Expt. 1 and with EPA in Expt. 2 to affect mRNA levels of iNOS, PPAR, and RXR in chicken macrophages and HD11 cells, respectively (P < 0.05). At 3% dietary fat or up to 15 µmol/L EPA in the medium, increasing lutein increased the iNOS mRNA. However, at 6% dietary fat or 50 µmol/L EPA, lutein did not cause a rise in iNOS mRNA. Increasing lutein in the medium from 0 to 100 µmol/L decreased iNOS mRNA. Increasing lutein with high fat (6%) or EPA (15 µmol/L EPA) increased PPAR and RXR mRNA levels. Lutein increased PPAR mRNA levels in both macrophages (P < 0.01) and HD11 (P = 0.01) cells and RXR (P < 0.01) mRNA levels in macrophages. GW9662, a PPAR antagonist, prevented (P < 0.01) the lutein-induced iNOS mRNA downregulation in HD11 cells. LG101208, a RXR antagonist, prevented (P < 0.01) iNOS upregulation induced by 10 µmol/L lutein and iNOS mRNA downregulation induced by 100 µmol/L lutein. We conclude that lutein and EPA interact through the PPAR and RXR pathways to modulate iNOS mRNA.
KEY WORDS: • lutein • fat • eicosapentaenoic acid • nuclear receptors
Xanthophylls, a group of carotenoids, have diminished or no provitamin A activity. Lutein, a xanthophyll lacking vitamin A activity, is enriched in the lens of the eye and in the retinal macular region (1). Derivatives of another xanthophyll, canthaxanthin, activate nuclear hormone receptors (NR)3 (2). Xanthophylls modulate immunity and cancer cell proliferation in humans (1). Among leukocytes, macrophages are particularly sensitive to modulation by xanthophylls (3).
Macrophages induced by lipopolysaccharide (LPS) produce nitric oxide using inducible nitric oxide synthase (iNOS). Inappropriate upregulation of iNOS is associated with cancers and inflammatory disorders (4). We showed previously that dietary lutein and fat interact to modify nitrite production in LPS-stimulated macrophages (5). High levels of dietary lutein increase nitrite production; however, high levels of fat reverse the stimulatory effect of lutein.
Nuclear factor (NF) 
B, a transcription factor with highly regulated activity, increases iNOS gene expression. PPAR/retinoic acid X receptor (RXR) complexes act through sequestration of essential coactivators, receptor mutual antagonism, or by cross-coupling to inhibit NFB (6).
PUFA and eicosanoids activate PPAR. Higher levels of dietary (7) or plasma fatty acids (8) upregulate different PPAR isomers through PPAR coactivator 1 protein (8). Modulation of gene activity by PPAR requires the presence of ligated RXR (9). Ligand-bound PPAR/RXR heterodimerize and bind coactivators, and this complex further binds to response elements (RE) in DNA to increase downstream transcription (10). A carotenoid derivative, 9 cis retinoic acid, PUFA (11), and many other synthetic compounds bind to the RXR ligand binding domain. In addition, the possibility of xanthophylls activating NR (2) and antioxidant RE during transcription (12) was explored. The presumed oxidation product of xanthophylls, acyclo-retinoic acid, transactivates a reporter gene containing the retinoic acid RE (RARE) but with a lower potency than retinoic acid (13). RXR activating agents upregulate RXR isomer mRNA and protein (14). We hypothesized that lutein and fat interact to affect macrophage iNOS production through PPAR-RXR heterodimers, with PUFA or their metabolic products acting as a ligand for PPAR, and lutein or its metabolic products activating RXR. Because PPAR-RXR heterodimers act to suppress NFB and therefore iNOS (10), depressed iNOS may serve as an indirect marker for increased PPAR-RXR heterodimers.
Lutein (15) and PUFA (9) are commonly supplemented in human nutrition as well as in poultry diets (5). Avian species possess the same enzymes as humans for metabolizing dietary lutein and are excellent nonprimate animal models for metabolic studies of xanthophylls (16). The avian model is useful because hens can be depleted of lutein so that the chicks receive all of their lutein via the diet (17). Therefore, the following study examined the interaction of different dietary lutein and fat levels on macrophage iNOS, PPAR, ß, and , and RXR and gene expression in chickens hatched from carotenoid-depleted eggs. Protein levels and mRNA expressions are highly correlated for PPAR isomers (18); because commercial antibodies for chicken PPAR and RXR isoform proteins are not available, the mRNA levels of the iNOS, PPAR, ß, and , and RXR and genes were measured as indicators. Additional in vitro studies using HD11 cells (a chicken macrophage cell line) examined the underlying mechanism by which lutein and fat interact to influence iNOS. Furthermore, GW9662, a PPAR-specific antagonist, and LG101208, a RXR-specific antagonist, were used to probe further the role of these nuclear receptors in mediating the interaction between lutein and fat.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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0.1 ng lutein, zeaxanthin, or canthaxanthin/kg diet) (17) and chicks were hatched from carotenoid-depleted eggs. Birds and experimental design. Broiler C-chicks (n = 108; 1 d old; Cobb x Cobb) of mixed sex were distributed randomly to 1 of the 6 treatments using a 2 x 3 factorial design with 2 levels of fat (3 and 6%, by weight) and three levels of lutein (0, 25, and 50 mg/kg feed). Each treatment contained 3 replicate pens of 6 chicks/pen. The fat was a 1.00:0.86 mixture of sunflower oil and refined menhaden fish oil (Omega Proteins). This ratio was chosen to meet the minimum NRC linoleic acid requirement (19) of chicks fed the diet with 3% fat. Isocaloric and isonitrogenous rice-soybean meal–based diets were formulated for the groups fed 3 and 6% fat. The diets were made isocaloric by varying the cornstarch and cellulose contents (Table 1). The basal diet contained 28 ng lutein/kg feed. Lutein (Oro Glo Dry, Kemin Industries) was added at 0, 25, and 50 mg/kg feed. The chicks were raised in Petersime brooder battery cages, and experimental diets and water were consumed ad libitum from the day of hatch. These experiments were reviewed by the UC Davis Animal Use and Care Committee to ensure adherence to animal care guidelines.
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Macrophage culture and stimulation. Monocytes (1 x 107) from 2 birds/pen (pooled within pen), in 2 mL of RPMI-1640, were supplemented with 1% penicillin:streptomycin (media) and 10% lutein-free chicken serum (LFCS) and incubated in 6-well culture plates at 41°C in a humidified atmosphere with 5% CO2. LFCS was prepared from chickens fed a lutein-free diet (Table 1). Adherent cells at 4 h of incubation were stimulated with LPS (1 mg/L; Sigma Chemicals, # L7261) in 2 mL of medium containing 5% LFCS for 9 h before extracting their total RNA; 9 h was chosen because at this time, iNOS mRNA level increases in LPS-stimulated chicken macrophages (21).
The chicken HD11 macrophage cell line was used for in vitro experiments. Cells were depleted of their lutein content by resuspending them in media supplemented with 5% LFCS and incubating at 41°C in a humidified atmosphere (5% CO2, 95% air). Preliminary trials were conducted with arachidonic acid, eicosapentaenoic acid (EPA), docosahexaenoic acid (DHA), and their combinations to determine which combination(s) interacted with lutein to modify macrophage nitrite production. Among the fatty acids studied, EPA showed a significant interaction with lutein in modifying macrophage nitrite production (data not shown); hence, further experiments in cell cultures were done with EPA. Stock solutions (10 mmol/L) of lutein (Sigma, X6250) and EPA (Nu-Chek-Prep) as the free fatty acid, with 4 µmol/L BHT, were prepared in dimethyl sulfoxide (DMSO) and ethanol, respectively.
Interaction between lutein and EPA in HD11 cell culture.
A factorial design with 3 levels of lutein (0, 10, and 100 µmol/L) and EPA (0, 15, and 50 µmol/L) was applied to HD11 cell cultures with 3 replications/treatment. Cells (1 x 107) were grown in 1 mL of medium with 5% LFCS and appropriate levels of lutein and EPA. Regardless of treatment, all of the wells had 1% DMSO and 0.5% ethanol. After 24 h, the medium was replaced with 1 mL of medium containing 5% LFCS and LPS (0.5 mg/L). A time course was employed to determine the optimum time for evaluating expressions of iNOS, PPAR, ß, and , and RXR and mRNA levels as influenced by LPS at 0, 30 min, 1, 2, 4, 6, 8, and 12 h after LPS stimulation. Except for PPARß and iNOS, all of the genes were downregulated at 4 h of LPS stimulation and remained depressed until 8 h. At 8 h, iNOS expression increased. PPARß expression was not stimulated by LPS. Because all genes that responded had altered expression levels at 8 h, the cell cultures were stimulated with LPS for 8 h before extracting total RNA.
Nuclear receptor antagonists.
HD11 cells (1 x 107) were incubated with the treatment medium (3 x 3 factorial; with lutein at 0, 10, and 100 µmol/L; EPA at 0, 15, and 50 µmol/L) for 24 h. At 24 h, the medium was replaced with 1 mL of medium containing 5% LFCS, LPS (0.5 mg/L), and either 0 or 10 µmol/L GW9662 (Sigma M6191), a PPAR antagonist in DMSO. After 6 h, the medium was replaced with medium containing 5% LFCS and LPS (0.5 mg/L). mRNA was collected at 9 h after the initial LPS addition. In another experiment, LG101208 (0 or 10 nmol/L; Ligand Pharmaceuticals), a RXR antagonist in DMSO, was added in place of GW9662 for a period of only 30 min.
RNA isolation and RT-PCR.
Cells (1 x 107) were detached from wells with a cell scraper, total RNA was isolated, and the RT reaction was run as described previously (22). Quantitative real-time PCR analysis of iNOS, PPAR, ß, and , and RXR and mRNA was performed with a ABI PRISM 7700 sequence detection system (Applied Biosystems). Primers were developed for iNOS, PPAR, ß, and , and RXR and based on the published sequences with the following respective Genbank accession numbers, Q90703, NP_001001464, NP_990059, NM_001001460, XP_415426 and P28701, respectively. The forward and reverse primers for iNOS were AGTGGTATGCTCTGCCTGCT and CCAGTCCCATTCTTCTTCC; for PPAR: CAATGCACTGGAACTGGATG and CGTCAGGATGGTTGGTTTG; for PPARß: CATGGAGCCCAAGTTTGAGT and CGGAGGATGTTGTCTTGGAT; for PPAR: GGGCGATCTTGACAGGAA and GCCTCCACAGAGCGAAAC; for RXR: GATGCGAGACATGCAGATG and GTCGGGGTATTTGTGCTTG, and for RXR: CAAACACATCTGTGCCATCTG and GATGAGGCAGTCCTTGTTGTC, respectively.
Preliminary experiments confirmed that all primer pairs produced only one product at their predicted size; when sequenced (Davis Sequencing) they had 99% homology to their respective gene transcripts (22). The 25-µL final PCR reaction volume contained 0.5 µL RT product and 2.5 µL 10X SYBR Green buffer, 2 µL dNTP, 0.125 µL TAQGOLD, and 0.25 µL Amperase (Applied Biosystems 4804886) together with 300 nmol/L of forward and reverse primers and 3 mmol/L MgCl2. In optimizing RXR, the forward primer was added at 900 nmol/L and in optimizing RXR, MgCl2 was added at 4.5 mmol/L. The PCR cycle was set at 50°C for 2 min followed by a denaturation step at 95°C for 10 min followed by 40 cycles of denaturing, annealing, and extension at 95°C for 15 s and 60°C for 1 min. The melting profile of each sample was analyzed as reported earlier (22) after every PCR run to confirm PCR product specificity.
Quantification of mRNA by real-time PCR was done as reported previously (22). All data were normalized to the mRNA level of the reference group (group fed 0 mg lutein and 3% fat in the in vivo experiment; group with 0 µmol/L lutein and 0 µmol/L EPA in the cell culture experiments) and reported as the fold-change from the reference. Fold-change from the reference was calculated as ES(40-Ct Sample)/ER(40-Ct Reference), where ES and ER are the sample and reference PCR amplification efficiencies, respectively, as determined in the log-linear phase using the LinRegPCR program (23).
In silico analysis.
Using the NUBISCAN matrix (24), NR RE in direct repeat (DR) 1, DR2, DR3, and DR4 in the upstream 2000 bp of the PPAR chicken gene (4916652 to 4937426 bp chromosome WASHUC1:12) and PPAR chicken gene (67845090 to 67879306 bp chromosome WASHUC1:1) were identified using the suggested general NR matrix or specific PPAR and PPAR matrix and score cut-off point of 0.70. Direct repeats consist of hexameric nucleotide sequences separated by 
1 nucleotide.
Statistical analysis. Treatments were applied to the birds and cells in a completely randomized design. A 2-way ANOVA (JMP software) was used to examine the interactive and main effects of dietary or media treatments on the dependent variables. A 3-way ANOVA (JMP software) was used to examine the interactive and main effects of dietary or media or antagonist treatments on the dependent variables. Interactions were removed from the model when the observed P-value for an interaction was >0.20. When interaction or main effects were significant (P < 0.05), differences between means were determined using Tukey's least-square means comparison.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Macrophage iNOS mRNA. Lutein and fat interacted (P = 0.01) to affect iNOS mRNA levels in chicken macrophages (Fig. 2). Macrophages from chicks fed 0 mg lutein with either 3 or 6% fat had the lowest iNOS level. Chicks fed either 25 or 50 mg dietary lutein with 3% fat had more iNOS mRNA than those fed 0 mg lutein, but this did not occur when the fat level was 6%.
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and RXR mRNA.
Lutein and fat interacted to affect PPAR (P = 0.05) and RXR (P = 0.03) mRNA levels in chicken macrophages (Fig. 3). Macrophages from chicks fed 50 mg lutein and 6% fat had higher (P = 0.05) PPAR mRNA levels than those from chicks administered any other treatment. Macrophages from chicks fed 6% fat and either 25 or 50 mg lutein had higher RXR mRNA levels than those from chicks fed 0 or 25 mg lutein and 3% fat. In the group fed 3% fat, macrophages from birds fed 50 mg lutein had higher (P = 0.03) RXR mRNA than the group fed 0 mg lutein.
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, PPARß, and RXR mRNA.
Lutein and fat did not interact to affect and fat level did not affect PPAR or RXR mRNA levels in chicken macrophages. Lutein increased both PPAR (P < 0.01) and RXR (P < 0.01) mRNA levels (Table 2). PPAR and RXR mRNA levels were significantly greater in chickens fed 50 mg lutein than in those fed 0 mg lutein. PPARß mRNA levels in chicken macrophages were not affected by lutein, fat, or their interaction.
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HD11 cell iNOS mRNA. Lutein and EPA interacted (P < 0.01) to affect HD11 cell iNOS mRNA levels (Fig. 4). HD11 cells treated with 0 µmol/L lutein had lower (P < 0.01) iNOS mRNA than cells treated with 10 µmol/L lutein when the EPA concentration was 0 or 15 µmol/L, but 50 µmol/L EPA suppressed iNOS mRNA, resulting in iNOS levels similar to those from cells treated with 0 µmol/L lutein. Cells treated with 100 µmol/L lutein had lower iNOS mRNA levels than cells treated with 0 or 10 µmol/L lutein. EPA did not affect iNOS expression when lutein was present at 0 or 100 µmol/L.
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and RXR mRNA.
Lutein and EPA interacted to increase PPAR (P < 0.01) and RXR (P = 0.01) mRNA levels in HD11 cells (Fig. 5). HD11 cells treated with 0 µmol/L lutein had lower PPAR and RXR mRNA than those treated with 50 µmol/L lutein. Increasing the lutein to 100 µmol/L in the absence of EPA did not affect the expression of PPAR or RXR, but in the presence of 15 or 50 µmol/L EPA, lutein increased PPAR and RXR expression. EPA in the absence of lutein did not change PPAR or RXR expression, but at 100 µmol/L lutein, 50 µmol/L EPA increased PPAR and RXR mRNA levels compared with the group treated with 0 µmol/L EPA.
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, PPARß, and RXR mRNA.
PPAR mRNA levels in HD11 cells were not affected by EPA, or an EPA x lutein interaction. Lutein increased (P = 0.01) PPAR mRNA levels in HD11 cells (Table 2). HD11 cells incubated with 100 µmol/L lutein had a higher PPAR mRNA level than those incubated with 0 µmol/L lutein (P = 0.01). PPARß and RXR mRNA levels in HD11 cells were not affected by lutein (P = 0.31, P = 0.87), EPA (P = 0.22, P = 0.73), or their interaction (P = 0.67, P = 0.93), respectively.
iNOS expression in the presence of a PPAR antagonist.
Cells treated with 10 µmol/L GW9662 (PPAR antagonist) expressed 14-fold more iNOS than the group treated with 0 µmol/L GW9662 (P < 0.01; data not shown). Because of this large difference in iNOS mRNA, the data for the 10 µmol/L GW9662 were normalized within the antagonist treatment to the 0 µmol/L lutein with 0 µmol/L EPA group. The antagonist, EPA, and lutein interacted (P < 0.01) to affect iNOS expression in HD11 cells (Fig. 6). The addition of 10 µmol/L PPAR antagonist reversed the iNOS-depressing effect of 100 µmol/L lutein, and this effect was most pronounced at 15 and 50 µmol/L EPA. The iNOS mRNA–depressing effect of 50 µmol/L EPA in the 10 µmol/L lutein group was not affected by the PPAR antagonist.
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and gene promoter region.
In the PPAR promoter region, one specific PPAR DR1 sequence (AGCTGAAAGGCCA, Score-0.701, P = 0.025, 
350 bp upstream) was identified. In the PPAR promoter region, one DR1 sequence (GGCTCAcAGCACA, Score-0.73, P = 0.05, 400 bp upstream), two DR2 sequences (AGAACAgaAGGGCA, Score-0.81, P = 0.01, 400 bp upstream; AGAACAatAGTGCA, Score-0.79, P = 0.02, 1500 bp upstream), and one DR4 sequence (AGTACAtgtcAGATCA, Score-0.865, P = 0.01, 1600 bp upstream), which are general for NR, were identified. |
DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Fatty acids and their metabolic products are ligands for PPAR. PPAR are active only as heterodimers with RXR (9). We hypothesize that lutein or its metabolites could activate RXR through its antioxidant properties because vitamin E (25) and the redox state of a cell (26) change RXR activity (25). If lutein activates RXR, the effect of lutein-stimulating iNOS (Figs. 2, 4), observed only when the fat or EPA was at lower levels, could be explained from the net balance of the NFB-iNOS pathway as described below. Fatty acids suppress the NFB- iNOS pathway through PPAR (9). At a high lutein concentration, the RXR could be active. This would facilitate PPAR activity and depress iNOS mRNA. At low lutein concentrations, the RXR would not be active and would result in suppression of PPAR activity, thereby stimulating iNOS mRNA.
High lutein and high EPA increased PPAR mRNA levels (Figs. 3A, 5A). The presence of the DR1 sequence specific for PPAR in the PPAR promoter indicates that PPAR are sensitive to PPAR. In addition, we found DR2 and DR4 sequences in the PPAR promoter region. This indicates that PPAR is also highly sensitive to RXR. Lutein increased RXR (discussed below) in macrophages and RXR in HD11 cells and macrophages. This change, together with increased ligands (in the form of oxidized fatty acids from increased fatty acids) for PPAR, could have increased PPAR mRNA synergistically, as reported earlier (7). Here, we report that for fatty acids to increase PPAR, a RXR agonist is needed.
Retinoic acid and several unsaturated fatty acids specifically bind and activate RXR (27). This indicates that RXR are fatty acid sensors in vivo and can influence RXR-mediated gene transcription. Oxyretinoic acid, a metabolic breakdown product of xanthophylls, was proposed to activate NR (2). In our study, cells with high lutein and EPA had higher RXR expression than those with no lutein and EPA (Figs. 3B, 5B). Thus, increased fatty acids and lutein interact to increase RXR mRNA levels.
PPAR is a receptor for fatty acids and regulates their cellular uptake and ß-oxidation (28). To our knowledge, the ability of lutein to increase PPAR has not been reported. The presence of a PPAR DR1 sequence in the PPAR promoter region suggests that PPAR would be upregulated in response to the upregulation of PPAR. PPAR was upregulated with high lutein (Table 2); hence, we propose that the lutein upregulation of PPAR is through upregulation of PPAR.
Feeding chickens higher levels of lutein increased RXR mRNA levels (Table 2), but HD11 cells treated with high lutein did not have altered RXR levels. It is not clear why the lutein effect was absent in the HD11 cells but was present in vivo. Rexinoids, by regulating the activity of multiple nuclear receptor heterodimers, can generate integrated control of complex metabolic pathways in ways that are not replicated by individual nuclear receptor ligands; hence, RXR may be governed by more complex pathways in vivo than in vitro.
Both in vivo and in vitro, there were significant interactive effects of lutein and fat or EPA on iNOS, PPAR, and RXR mRNA levels and a significant independent effect of lutein on PPAR. Increases in PPAR and RXR with high fat or EPA and lutein were accompanied by decreased iNOS mRNA levels. Upregulation of PPAR/RXR leads to transcriptional regulation of PPAR/RXR-responsive genes (28). Hence, we hypothesized that lutein and EPA/fat affect iNOS via PPAR/RXR heterodimers, with higher levels of PPAR/RXR suppressing iNOS. A synergistic effect of PPAR and RXR ligands on reporter gene activation was reported previously (29). To test this hypothesis, we used a PPAR-specific antagonist, GW9662, and a RXR-specific antagonist, LG101208, to determine whether the interactive effect of lutein and EPA could be reversed. We found that blocking either NR ameliorated the interaction between lutein and EPA.
Earlier reports confirmed the iNOS-stimulating effect of GW9662, wherein this PPAR antagonist stimulates iNOS through NFB (30). Similarly, LG101208 stimulates TNF production in human myelomonocytic cell lines (31). Natural and synthetic PPAR and RXR ligands inhibit the LPS induction of inflammatory genes. Two different mechanisms were proposed: one inhibits NFB-dependent transcription through a PPAR-dependent mechanism (10), and one is independent of PPAR (32). We observed that the action of lutein on iNOS was reversed by an RXR antagonist (Fig. 7), suggesting that lutein activates RXR. To examine further the role of lutein on RXR, we determined the effect of an RXR antagonist on 9 cis retinoic acid (RXR agonist)–mediated iNOS suppression in LPS-stimulated HD11 cells and found that 9 cis retinoic acid at 0.5 µmol/L depressed iNOS expression. This iNOS depression was reversed by RXR antagonist (data not reported).
In our experiment, the addition of lutein at 10 µmol/L increased iNOS expression, whereas it was suppressed at 100 µmol/L. The iNOS-stimulating effect of 10 µmol/L lutein was reversed by increasing EPA or RXR antagonist. The iNOS-depressing effect of 100 µmol/L lutein was reversed by increasing the EPA or RXR antagonist or the PPAR antagonist. The EPA effect of suppressing iNOS at 10 µmol/L lutein was not reversed by the PPAR antagonist but was reversed by the RXR antagonist. We propose that lutein and EPA interact through PPAR/RXR heterodimers to modify LPS-stimulated iNOS expression. Increasing lutein and EPA in the presence of each other increased PPAR and RXR mRNA. At lower concentrations of lutein and EPA, PPAR and RXR levels would be expected to be upregulated, but the RXR could be unliganded. This unliganded RXR in the PPAR/RXR heterodimer would act to stimulate the NFB pathway (33), thus increasing the iNOS. This appears to be dependent on the RXR pathway because an RXR antagonist prevented 10 µmol/L lutein from stimulating iNOS production. Increasing EPA decreased the stimulating effect of 10 µmol/L lutein on iNOS mRNA. Fatty acids act as a ligand for RXR (11), suggesting that EPA, by liganding RXR, decreases iNOS. This was further confirmed when the EPA-depressing effect on iNOS at low lutein levels was reversed by an RXR antagonist, but not by a PPAR antagonist. Early reports that unliganded RXR responses on downstream targets were independent of PPAR (10,32), would also explain our results. But because the lutein level is increased further, the RXR in the PPAR/RXR heterodimers could be expected to be activated and suppress iNOS (29). This was confirmed when either RXR or PPAR antagonists reversed the iNOS-depressing effect of 100 µmol/L lutein as well as the effects of EPA at 100 µmol/L lutein. Because a PPAR antagonist reversed the iNOS-depressing effect of high lutein and EPA, it appears that high concentrations of lutein and EPA act through RXR/PPAR to suppress iNOS. Because the iNOS-stimulating effect of low lutein was not influenced by the PPAR antagonist but was stimulated by the RXR antagonist, this part is dependent on RXR but independent of PPAR.
We conclude that lutein and fat or EPA act through the PPAR and RXR pathway to change iNOS expression, and that this effect is dependent on the dose of lutein and fat or EPA. Lutein showed RXR ligand–like activity. Because of the strong interaction between dietary lutein and fat, both of these nutrients should be considered when examining the immunomodulatory properties of diets. Functional assays with macrophages recovered from wild-type or tissue-specific PPAR null mice would provide a more compelling molecular and mechanistic basis for the regulation of the pathway iNOS via the PPAR and RXR pathways.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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3 Abbreviations used: DHA, docosahexaenoic acid; DMSO, dimethyl sulfoxide; DR, direct repeats; EPA, eicosapentaenoic acid; iNOS, inducible nitric oxide synthase; LFCS, lutein-free chicken serum; LPS, lipopolysaccharide; NF, nuclear factor; NR, nuclear hormone receptor; RARE, retinoic acid response element; RE, response element; RXR, retinoic acid X receptor.
Manuscript received 30 September 2005. Initial review completed 15 November 2005. Revision accepted 6 March 2006.
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LITERATURE CITED |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
1. Ribaya-Mercado JD, Blumberg JB. Lutein and zeaxanthin and their potential roles in disease prevention. J Am Coll Nutr. 2004;23:567S–87.
2. Nikawa T, Schulz WA, van den Brink CE, Hanusch M, van der Saag P, Stahl W, Sies H. Efficacy of all-trans-beta-carotene, canthaxanthin, and all-trans-, 9-cis-, and 4-oxoretinoic acids in inducing differentiation of an F9 embryonal carcinoma RAR beta-lacZ reporter cell line. Arch Biochem Biophys. 1995;316:665–72.[Medline]
3. Zhao W, Han Y, Zhao B, Hirota S, Hou J, Xin W. Effect of carotenoids on the respiratory burst of rat peritoneal macrophages. Biochim Biophys Acta. 1998;1381:77–88.[Medline]
4. Surh YJ, Chun KS, Cha HH, Han SS, Keum YS, Park KK, Lee SS. Molecular mechanisms underlying chemopreventive activities of anti-inflammatory phytochemicals: down-regulation of COX-2 and iNOS through suppression of NF-[kappa]B activation. Mutat Res. 2001;480–481:243–68.
5. Selvaraj RK, Koutsos EA, Calvert CC, Klasing KC. Dietary lutein and fat interact to modify macrophage properties in chicks hatched from carotenoid depleted or repleted eggs. J Anim Physiol Anim Nutr. 2005;90:70–80.
6. Kelly D, Campbell JI, King TP, Grant G, Jansson EA, Coutts AGP, Pettersson S, Conway S. Commensal anaerobic gut bacteria attenuate inflammation by regulating nuclear-cytoplasmic shuttling of PPAR-gamma and RelA. Nat Immunol. 2004;5:104–12.[Medline]
7. Panadero M, Herrera E, Bocos C. Different sensitivity of PPARalpha gene expression to nutritional changes in liver of suckling and adult rats. Life Sci. 2005;76:1061–72.[Medline]
8. Watt MJ, Southgate RJ, Holmes AG, Febbraio MA. Suppression of plasma free fatty acids upregulates peroxisome proliferator-activated receptor (PPAR) alpha and delta and PPAR coactivator 1alpha in human skeletal muscle, but not lipid regulatory genes. J Mol Endocrinol. 2004;33:533–44.
9. Hwang D. Fatty acids and immune responses—a new perspective in searching for clues to mechanism. Annu Rev Nutr. 2000;20:431–56.[Medline]
10. Valledor AF, Ricote M. Nuclear receptor signaling in macrophages. Biochem Pharmacol. 2004;67:201–12.[Medline]
11. Lengqvist J, Mata De Urquiza A, Bergman AC, Willson TM, Sjovall J, Perlmann T, Griffiths WJ. Polyunsaturated fatty acids including docosahexaenoic and arachidonic acid bind to the retinoid X receptor alpha ligand-binding domain. Mol Cell Proteomics. 2004;3:692–703.
12. Ben-Dor A, Steiner M, Gheber L, Danilenko M, Dubi N, Linnewiel K, Zick A, Sharoni Y, Levy J. Carotenoids activate the antioxidant response element transcription system. Mol Cancer Ther. 2005;4:177–86.
13. Sharoni Y, Danilenko M, Dubi N, Ben-Dor A, Levy J. Carotenoids and transcription. Arch Biochem Biophys. 2004;430:89–96.[Medline]
14. Chen Q, Catharine Ross A. Retinoic acid regulates cell cycle progression and cell differentiation in human monocytic THP-1 cells. Exp Cell Res. 2004;297:68–81.[Medline]
15. Castenmiller JJ, West CE. Bioavailability and bioconversion of carotenoids. Annu Rev Nutr. 1998;18:19–38.[Medline]
16. Khachik F, de Moura FF, Zhao DY, Aebischer CP, Bernstein PS. Transformations of selected carotenoids in plasma, liver, and ocular tissues of humans and in nonprimate animal models. Invest Ophthalmol Vis Sci. 2002;43:3383–92.
17. Koutsos EA, Clifford AJ, Calvert CC, Klasing KC. Maternal carotenoid status modifies the incorporation of dietary carotenoids into immune tissues of growing chickens (Gallus gallus domesticus). J Nutr. 2003;133:1132–8.
18. Shaban Z, El-Shazly S, Abdelhady S, Fattouh I, Muzandu K, Ishizuka M, Kimura K, Kazusaka A, Fujita S. Down regulation of hepatic PPARalpha function by AhR ligand. J Vet Med Sci. 2004;66:1377–86.[Medline]
19. National Research Council. Nutrient requirements of poultry. 9th edition. Washington, DC: National Academy Press; 1994
20. Repnik U, Knezevic M, Jeras M. Simple and cost-effective isolation of monocytes from buffy coats. J Immunol Methods. 2003;278:283–92.[Medline]
21. Hussain I, Qureshi MA. The expression and regulation of inducible nitric oxide synthase gene differ in macrophages from chickens of different genetic background. Vet Immunol Immunopathol. 1998;61:317–29.[Medline]
22. Humphrey BD, Stephensen CB, Calvert CC, Klasing KC. Glucose and cationic amino acid transporter expression in growing chickens (Gallus gallus domesticus). Comp Biochem Physiol A Mol Integr Physiol. 2004;138:515–25.[Medline]
23. Ramakers C, Ruijter JM, Deprez RH, Moorman AF. Assumption-free analysis of quantitative real-time polymerase chain reaction (PCR) data. Neurosci Lett. 2003;339:62–6.[Medline]
24. Podvinec M, Kaufmann MR, Handschin C, Meyer UA. NUBIScan, an in silico approach for prediction of nuclear receptor response elements. Mol Endocrinol. 2002;16:1269–79.
25. Azzi A, Gysin R, Kempna P, Munteanu A, Negis Y, Villacorta L, Visarius T, Zingg JM. Vitamin E mediates cell signaling and regulation of gene expression. Ann N Y Acad Sci. 2004;1031:86–95.[Medline]
26. Demary K, Wong L, Liou JS, Faller DV, Spanjaard RA. Redox control of retinoic acid receptor activity: a novel mechanism for retinoic acid resistance in melanoma cells. Endocrinology. 2001;142:2600–5.
27. Steineger HH, Arntsen BM, Spydevold O, Sorensen HN. Gene transcription of the retinoid X receptor alpha (RXRalpha) is regulated by fatty acids and hormones in rat hepatic cells. J Lipid Res. 1998;39:744–54.
28. Kota BP, Huang TH, Roufogalis BD. An overview on biological mechanisms of PPARs. Pharmacol Res. 2005;51:85–94.[Medline]
29. Benson S, Padmanabhan S, Kurtz TW, Pershadsingh HA. Ligands for the peroxisome proliferator-activated receptor-gamma and the retinoid X receptor-alpha exert synergistic antiproliferative effects on human coronary artery smooth muscle cells. Mol Cell Biol Res Commun. 2000;3:159–64.[Medline]
30. Lim SY, Jang JH, Surh YJ. Induction of cyclooxygenase-2 and peroxisome proliferator-activated receptor-gamma during nitric oxide-induced apoptotic PC12 cell death. Ann N Y Acad Sci. 2003;1010:648–58.[Medline]
31. Mou L, Lankford-Turner P, Leander MV, Bissonnette RP, Donahoe RM, Royal W. RXR-induced TNF-alpha suppression is reversed by morphine in activated U937 cells. J Neuroimmunol. 2004;147:99–105.[Medline]
32. Giri S, Rattan R, Singh AK, Singh I. The 15-deoxy-delta12,14-prostaglandin J2 inhibits the inflammatory response in primary rat astrocytes via down-regulating multiple steps in phosphatidylinositol 3-kinase-Akt-NF-kappaB-p300 pathway independent of peroxisome proliferator-activated receptor gamma. J Immunol. 2004;173:5196–208.
33. Monden T, Yamada M, Nihei Y, Kishi M, Tomaru T, Ishii S, Hashida T, Shibusawa N, Hashimoto K, et al. Unliganded RXR acts as an inhibitory factor on troglitazone-induced activation. Life Sci. 2004;76:731–41.[Medline]
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