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J. Biol. Chem., Vol. 280, Issue 51, 42464-42475, December 23, 2005
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123**415
From the
Departments of Anesthesiology, **Environmental Health Sciences and Center for Free Radical Biology, University of Alabama at Birmingham, Alabama 35294, the ¶Cardiovascular Research Institute, Morehouse School of Medicine, Atlanta, Georgia 30310, and the ||Department of Chemistry, University of Oregon, Eugene, Oregon 97403
Received for publication, April 18, 2005 , and in revised form, September 23, 2005.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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50% greater than that of nitrated linoleic acid, with the combined free and esterified blood levels of these two fatty acid derivatives exceeding 1 µM. OA-NO2 is a potent ligand for peroxisome proliferator activated receptors at physiological concentrations. CV-1 cells co-transfected with the luciferase gene under peroxisome proliferator-activated receptor (PPAR) response element regulation, in concert with PPAR
, PPAR
, or PPAR
expression plasmids, showed dose-dependent activation of all PPARs by OA-NO2. PPAR showed the greatest response, with significant activation at 100 nM, while PPAR and PPAR were activated at 300 nM OA-NO2. OA-NO2 also induced PPAR -dependent adipogenesis and deoxyglucose uptake in 3T3-L1 preadipocytes at a potency exceeding nitrolinoleic acid and rivaling synthetic thiazo-lidinediones. These data reveal that nitrated fatty acids comprise a class of nitric oxide-derived, receptor-dependent, cell signaling mediators that act within physiological concentration ranges. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The ability of .NO and .NO-derived species to oxidize, nitrosate, and nitrate biomolecules serves as the molecular basis for how .NO influences the synthesis and reactions of bioactive lipids (3-5). Interactions between .NO and lipid oxidation pathways are multifaceted and inter-dependent. For example, .NO regulates both the catalytic activity and gene expression of prostaglandin H synthase (6). Conversely, leukotriene products of lipoxygenases induce nitric-oxide synthase-2 expression and increase inflammatory .NO production (7). The free radical reactivity of .NO lends an ability to inhibit the autocatalytic chain propagation reactions of lipid peroxyl radicals during membrane and lipoprotein oxidation (8). Of relevance, reactions between .NO-derived species, unsaturated fatty acids, and lipid oxidation intermediates yield a spectrum of fatty acid oxidation and nitration products (3). Recently, the nitroalkene derivative of linoleic acid (LNO2) was detected in human blood at concentrations sufficient to induce biological responses (500 nM; Refs. 9-12). Compared with other .NO-derived species such as nitrite (
), nitrosothiols (RSNO), and heme-nitrosyl complexes, LNO2 alone represents the single most abundant pool of bioactive oxides of nitrogen in the healthy human vasculature (9, 13-16).
In vitro studies have shown that LNO2 mediates cGMP-dependent vascular relaxation, cGMP-independent inhibition of neutrophil degranulation and superoxide formation, and inhibition of platelet activation (10-12). Recently, LNO2 has been shown to exert cell signaling actions via ligation and activation of peroxisome proliferator-activated receptors (PPARs) (17), a class of nuclear hormone receptors that modulates the expression of metabolic, cellular differentiation, and inflammatory-related genes (18, 19).
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ligand activity that acts within physiological concentrations (17), (b) an ability to decay in aqueous conditions to release .NO (20), and (c) reactivity as an electrophile, motivated a search for other nitrated fatty acids that might serve related signaling actions. Herein, we report that nitroalkene derivatives of all principal unsaturated fatty acids are present in human blood and urine. Of the fatty acid content in red cells, linoleic acid and oleic acid comprise 8 and 18% of total, respectively (21). Due to its prevalence and structural simplicity, oleic acid was evaluated as a potential candidate for nitration. The synthesis, structural characterization, and cell signaling activities of 9- and 10-nitro-9-cis-octadecaenoic acids are described (nitrated oleic acid, OA-NO2; Fig. 1). OA-NO2 regioisomers were measured in human blood and urine at levels exceeding those of LNO2. Furthermore, OA-NO2 activates PPAR with a greater potency than LNO2. These data reveal that nitrated unsaturated fatty acids represent a class of lipid-derived, receptor-dependent signaling mediators. |
MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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and anti-
-actin antibodies were from Santa Cruz Biotechnology (Santa Cruz, CA); anti-aP2 antibody was from Chemicon International Inc. (Temecula, CA). Synthesis of OA-NO2—Oleic acid and [13C18]oleic acid were nitrated as described (9, 12), with modifications. Oleic acid, HgCl2, phenylselenium bromide, and NaNO2 (1:1.3:1:1, mol/mol) were combined in THF/acetonitrile (1:1, v/v) with a final concentration of 0.15 M oleic acid. The reaction mixture was stirred (4 h, 25 °C), followed by centrifugation to sediment the precipitate. The supernatant was recovered, the solvent evaporated in vacuo, the product mixture redissolved in THF (original volume), and the temperature reduced to 0 °C. A 10-fold molar excess of H2O2 was slowly added with stirring to the mixture, which was allowed to react in an ice bath for 20 min followed by a gradual warming to room temp (45 min). The product mixture was extracted with hexane, the organic phase collected, the solvent removed in vacuo, and lipid products solvated in CH3OH. OA-NO2 was isolated by preparative TLC using silica gel HF plates developed twice in a solvent system consisting of hexane/ether/acetic acid (70:30:1, v/v). The region of silica containing OA-NO2 was scraped and extracted (23). Based on this synthetic rationale, two regioisomers are generated: 9- and 10-nitro-9-cis-octadecenoic acids (generically termed OA-NO2). Preparative TLC does not adequately resolve the two isomers. [13C18]OA-NO2 was synthesized using [13C18]oleic acid as a reactant. All nitrated fatty acid stock solutions were diluted in MeOH, aliquoted, and stored under argon gas at -80 °C. Under these conditions, OA-NO2 isomers remain stable for >3 months.
The nitroalkene positional isomers are described as cis throughout this article based on the configuration of the carbon skeleton, which correlates the cis alkene stereochemistry in the nitroalkenes with the corresponding cis alkene stereochemistry in naturally occurring oleic acid. The IUPAC nomenclature of the nitroalkenes has the opposite stereochemical terminology, because it focuses on the relationship of the higher priority nitro group to the carbon substituents on the alkene. For example, the 9-nitro isomer has the carbon chains cis to each other on the nitroalkene, but the official IUPAC nomenclature designates this compound as E (or trans) because the nitro group on C-9 and the carbon chain on C-10 have the E (entgegen) or trans relationship to each other.
Quantitation of Synthetic OA-NO2—The concentrations of synthetic OA-NO2 stock solutions were determined using chemiluminescent nitrogen analysis (Antek Instruments, Houston, TX), a quantitative measure of nitrogen content in synthetic and biological samples (24, 25). Briefly, purified synthetic nitroalkene preparations were subjected to complete pyrolysis (>1000 °C). The nitrogen-containing OA-NO2 reacts with O2 to ultimately yield .NO at a ratio of one mole .NO for every mole of nitrogen present in OA-NO2. The generated .NO reacts with O3 to yield nitrogen dioxide (.NO2,O2, and hv, the latter of which is sensitively detected with a photomultiplier). Concentrations were calculated using caffeine as standard.
Stability of OA-NO2 and LNO2—The relative stabilities of OA-NO2 and LNO2 in MeOH and phosphate buffer (100 mM KiPO4 containing 100 µM DTPA, pH 7.4) were determined by electrospray ionization triple quadrupole mass spectrometry (ESI MS/MS) using the quantitative methodology detailed below. OA-NO2 and LNO2 (3 µM each) were incubated at 37 °C in either MeOH or phosphate buffer, and aliquots were taken over time. The aliquots were extracted as described (23), with 1 µM [13C18]LNO2 added during the monophase stage of the extraction procedure as an internal standard, and analyzed for non-degraded OA-NO2 and LNO2. In aqueous buffer, nitrated lipids degrade more rapidly than in organic solvents (20); thus, their stability in phosphate buffer was measured over 2 h. The stability of nitrated fatty acids solvated in MeOH at 37 °C was measured over the course of 1 month.
OA-NO2 Spectrophotometric Characterization—OA-NO2 stock solution concentrations derived from chemiluminescent nitrogen analysis were utilized to determine dilution concentrations for subsequent spectral analysis. An absorbance spectrum of OA-NO2 from 200-450 nm was generated using 23 µM OA-NO2 in phosphate buffer (100 mM, pH 7.4) containing 100 µM DTPA. The extinction coefficients (
) for OA-NO2 and the isotopic derivative [13C18]OA-NO2 were measured (
270) using a UV-visible spectrophotometer (Shimadzu, Japan). Absorbance values for increasing concentrations of OA-NO2 and [13C18]OA-NO2 were plotted against concentration to calculate .
NMR Spectrometric Analysis of OA-NO2—1H and 13C NMR spectra were acquired using a Varian INOVA 300 and a 500 MHz NMR and recorded in CDCl3. Chemical shifts are in units (ppm) and referenced to residual proton (7.26 ppm) or carbon (77.28 ppm) signals in deuterated chloroform. Coupling constants (J) are reported in Hertz (Hz).
Structural Characterization of OA-NO2 by ESI MS/MS—Qualitative analysis of OA-NO2 by ESI MS/MS was performed using a hybrid triple quadrupole-linear ion trap mass spectrometer (4000 Q trap, Applied Biosystems/MDS Sciex). To characterize synthetic and endogenous OA-NO2, a reverse-phase HPLC methodology was developed using a 150 x 2 mm C18 Phenomenex Luna column (3 µm particle size). Lipids were separated and eluted from the column using a gradient solvent system consisting of A (H2O containing 0.1% NH4OH) and B (CNCH3 containing 0.1% NH4OH) under the following conditions: 20-65% B (10 min); 65-95% B (1 min; hold for 3 min) and 95-20% B (1 min; hold for 3 min). OA-NO2 was detected using a multiple reaction monitoring (MRM) scan mode by reporting molecules that undergo a m/z 326/279 mass transition consistent with the loss of the nitro group ([M - (HNO2)]-). Concurrent with MRM determination, enhanced product ion analysis (EPI) was performed to generate characteristic and identifying fragmentation patterns of eluting species with a precursor mass of m/z 326. Zero grade air was used as source gas, and nitrogen was used in the collision chamber.
Red Blood Cell Isolation and Lipid Extraction—Peripheral blood from fasting, apparently healthy human volunteers was collected by venipuncture into heparinized tubes (UAB Institutional Review Board-approved protocol no. X040311001). Blood was centrifuged (1200 x g; 10 min), the buffy coat was removed, and erythrocytes were isolated. Lipid extracts were prepared from red cells and plasma and directly analyzed by mass spectrometry (23). Care was taken to avoid acidification during extraction to prevent artifactual lipid nitration due to the presence of endogenous nitrite (9). In experiments using urine as the biological specimen (UAB Institutional Review Board-approved protocol no. X040311003), extraction conditions were identical.
Detection and Quantitation of OA-NO2 in Human Blood and Urine—Quantitation of OA-NO2 in biological samples was performed as described (9), with modifications. Matched blood and urine samples were obtained after >8 h fasting; urine was collected from the first void of the day. During the monophase stage of the lipid extraction (23), [13C18]OA-NO2 was added as internal standard to correct for any losses. Nitrated fatty acids were then analyzed by HPLC ESI MS/MS in the negative ion mode. Lipids were eluted from the HPLC column using an isocratic solvent system consisting of CH3CN:H2O:NH4OH (85:15:0.1, v/v) so that the two OA-NO2 regioisomers co-elute. During quantitative analyses, two MRM transitions were monitored: m/z 326/279 (OA-NO2) and m/z 344/297 ([13C18]OA-NO2), transitions consistent with the loss of the nitro group from the respective precursor ions. The areas under each peak were integrated, the ratios of analyte to internal standard areas were determined, and OA-NO2 was quantitated using Analyst 1.4 quantitation software (Applied Biosystems/MDS Sciex). Data are expressed as mean ± S.D. (n = 10; 5 female and 5 male).
To address whether artifactual synthesis of OA-NO2 occurred during sample preparation and extraction, control studies were performed as described (9). Briefly, [13C18]oleic acid was added as a reporter molecule prior to red cell and plasma lipid purification and analysis, which permitted the MS detection of possible 13C-labeled OA-NO2 formation. Also, 200 µM was included in initial lipid extractions to determine whether separations or analysis-induced nitration reactions might be supported by physiological levels that can exceed 200 nM (14, 15). In no case did we detect artifactual nitration of oleic acid due to sample processing and analysis.
Qualitative Analysis of Nitro and Nitrohydroxy Adducts of Fatty Acids—Using HPLC ESI MS/MS in the negative ion mode, blood and urine samples were evaluated for the presence of nitroalkene derivatives other than LNO2. HPLC separations using the qualitative gradient elution methodology were performed similarly to those used to characterize OA-NO2, with some modifications. Alternative MRM transitions were used to detect other potential nitroalkene derivatives. Theoretical MRM transitions were determined for the CID-induced loss of the nitro group from nitrated palmitoleic (16:1-NO2), linolenic (18:3-NO2), arachidonic (20:4-NO2), and eicosapentaenoic (20:5-NO2) acids. MRM transitions for nitrohydroxy adducts were also monitored: 16:1(OH)-NO2; 18:1(OH)-NO2; 18:2(OH)-NO2; 18:3(OH)-NO2, 20:4(OH)-NO2, and 20:5(OH)-NO2.
In Vitro Formation of OA-NO2—Three different conditions were examined for an ability to induce nitration of oleic acid: acidic nitration, treatment with peroxynitrite, and treatment with MPO in the presence of H2O2 and nitrite. Briefly, for acidic nitration, oleic acid (1 mM) and sodium nitrite (100 µM) were prepared in phosphate buffer (50 mM, pH 7.2) in the presence of 2% sodium cholate (26). The pH was adjusted to 3.0, and the reaction mixture was incubated with stirring (40 min; 25 °C). The reaction was stopped by solvent extraction, and OA-NO2 levels were measured by HPLC ESI MS/MS. For peroxynitrite-induced nitration, oleic acid (1 mM) was suspended in phosphate buffer (100 mM, pH 7.2), and ONOO- was infused via syringe pump into a stirred chamber (100 µM/min; 15 min) (26). Decayed ONOO- (pH 7.4, 10 min) was added as a control. Products were extracted and analyzed for OA-NO2. For MPO-induced nitration, oleic acid (1 mM) was incubated in phosphate buffer (100 mM; pH 7.2) in the presence of MPO (50 nM), sodium nitrite (100 µM), and hydrogen peroxide (100 µM) as described (27). The reaction proceeded for 90 min with additional aliquots of hydrogen peroxide added at 30-min intervals. The reaction was stopped by lipid extraction, and OA-NO2 was measured by HPLC ESI MS/MS. Significance of difference between treated and control groups was determined using a one-tailed, paired Student's t test.
PPAR Transient Transfection Assay—CV-1 cells from the ATCC (Manassas, VA) were grown to 85% confluence in DMEM/F-12 supplemented with 10% FBS and 1% penicillin-streptomycin. Twelve hours before transfection, the medium was removed and replaced with anti-biotic-free medium. Cells were transiently co-transfected with a plasmid containing the luciferase gene under the control of three tandem PPAR response elements (PPRE) (PPRE x 3 TK-luciferase and PPAR, PPAR, or PPAR expression plasmids, respectively (provided by Ron Evans, Salk Institute). In all cases, a green fluorescence protein (GFP) expression plasmid was co-transfected as the control for transfection efficiency. Twenty-four hours after transfection, cells were returned to Opti-MEM (Invitrogen) for 24 h and then treated as indicated for another 24 h. Reporter luciferase assay kits from Promega (Madison, WI) were used to measure the luciferase activity according to the manufacturer's instructions with a luminometer (Victor II, PerkinElmer Life Sciences). Luciferase activity was normalized by GFP units. Each condition was performed in triplicate for each experiment (n 
3).
3T3-L1 Differentiation and Oil Red O Staining—3T3-L1 preadipocytes were propagated and maintained in DMEM containing 10% FBS. To induce differentiation, 2-day post-confluent preadipocytes (designated day 0) were cultured in DMEM containing 10% FBS plus 1 and 3 µM OA-NO2 for 14 days. The medium was changed every 2 days. Rosiglitazone (3 µM) and oleic acid (3 µM) were used as positive and negative controls, respectively. Differentiated adipocytes were stained with oil red O as described previously (28).
[3H]2-Deoxy-D-Glucose Uptake Assay in Differentiated 3T3-L1 Adipocytes—[3H]2-Deoxy-D-glucose uptake was analyzed as described previously (29). 3T3-L1 preadipocytes were grown in 24-well tissue culture plates, 2-day post-confluent monolayers were treated with 10 µg/ml insulin, 1 µM dexamethasone, and 0.5 mM 3-isobutyl-1-methylxanthine in DMEM containing 10% FBS for 2 days, then cells were maintained in 10 µg/ml insulin in DMEM containing 10% FBS for 6 days (medium was changed every 3 days). Eight days after induction of adipogenesis, test compounds in DMEM containing 10% FBS were added for an additional 2 days (medium was changed every day). The PPAR-specific antagonist GW9662 was added 1 h before other additions. After two rinses with serum-free DMEM, cells were incubated for 3 h in serum-free DMEM and rinsed at room temperature three times with freshly prepared KRPH buffer (5 mM phosphate buffer, 20 mM HEPES, 1 mM MgSO4, 1 mM CaCl2, 136 mM NaCl, 4.7 mM KCl, pH 7.4). The buffer was replaced with 1 µCi/ml of [3H]2-deoxy-D-glucose in KRPH buffer for 10 min at room temperature. Cells were then rinsed three times with cold PBS, lysed overnight in 0.8 N NaOH (0.4 ml/well), neutralized with 26.6 µ l of 12 N HCl, and 360 µl of lysate was added to Scintisafe PlusTM for radioactivity determination by liquid scintillation counting.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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NMR Analysis of OA-NO2—The structure of synthetic OA-NO2 (a 1:1 mixture of C-9- and C-10-regioisomers) was analyzed by 1H and 13C NMR. NMR splitting patterns are designated as s, singlet; d, doublet; t, triplet; q, quartet; m, multiplet; and br, broad. 1H NMR (CDCl3): 11.1 (br s, 1H), 7.06 (dd, 1H, J = 7.8 Hz), 2.55 (t, 2H, J = 7.6 Hz), 2.35 (m, 2H), 2.20 (q, 2H, J = 7.3 Hz), 1.61 (m, 2H), 1.47 (m, 4H), 1.32-1.25 (m, 16H), 0.87 (t, 3H, J = 7.0 Hz).
The 1H spectrum and proposed assignments of diagnostic peaks are presented in Fig. 3A: 11.1 (COOH), 7.06 (C-9 or C-10, alkene proton, each a triplet from coupling to neighboring methylene CH2, with regioisomers superimposed on each other, appearing on one NMR spectrometer at 300 MHz as a doublet of triplets and on the other at 500 MHz as a quartet, in actuality a superimposed pair of triplets), 2.55 (C-8 or C-11, allylic methylene neighboring nitro group; nitroalkene more electron-withdrawing than carbonyl), 2.35 (C-2 methylene neighboring carbonyl), 2.20 (C-8 or C-11, allylic methylene opposite nitro group); 0.87 (C-18 terminal methyl).
The signal for the alkene CH is sufficient to assign the stereochemistry of the alkene. E-Nitroalkenes have characteristic chemical shifts of approximately 7.0 ppm, while Z-nitroalkenes have characteristic chemical shifts of approximately 5.8 ppm (30-32). The only alkene CH observed in the 1:1 mixture of 9- and 10-nitro isomeric OA-NO2 are centered on 7.06 ppm as superimposed signals from each isomer, 9-nitro and 10-nitro. No other alkene peaks are present. Thus, both the 9-nitro and the 10-nitro isomers have the E-configuration (referred to as cis-isomers herein, as detailed above). The remaining regions of the spectrum also overlap and are similar for each isomer.
Spectral Characterization of Synthetic OA-NO2—The absorbance spectrum of OA-NO2 was acquired in phosphate buffer in the presence of the iron chelator DTPA (Fig. 4A). The maximum at 270 nm was ascribed to photon absorption by pi electrons in the nitro functional group. Extinction coefficients for OA-NO2 and [13C18]OA-NO2 were determined by plotting absorbance (270) versus concentration, giving m = AU·cm-1·mM-1 and a calculated = 8.22 and 8.23 cm-1·mM-1, for OA-NO2 and [13C18]OA-NO2, respectively (Fig. 4B).
Stability of OA-NO2—OA-NO2 was found to be fully stable for >3 months when stored at -80 °C in MeOH (data not shown). However, some decay was observed in MeOH at 37 °C, showing 10% decay after 1 month (Fig. 5). In phosphate buffer, OA-NO2 decayed much faster, with 40% loss after 2 h. In both solvent environments, LNO2 was much less stable than OA-NO2, with this attributed to the greater reactivity of the bisallylic bond arrangement in LNO2.
Characterization and Quantitation of Endogenous OA-NO2 by ESI MS/MS—Using the gradient HPLC elution protocol described under "Materials and Methods," synthetic OA-NO2 regioisomers eluted from the reverse-phase column as two partially overlapping peaks (Fig. 6). The HPLC elution profiles for synthetic OA-NO2 and [13C18]OA-NO2 were identical (Fig. 6A, left panels). Concurrent product ion analysis of the overlapping peaks showed spectra consistent with OA-NO2-derived species (Fig. 6A, right panels), with major fragments identified in TABLE TWO. Using these same parameters and machine settings, lipid extracts of packed red cells and plasma were analyzed (Fig. 6B). The product ion spectra for the OA-NO2 present in red cells and plasma were identical to those obtained from synthetic OA-NO2, revealing that OA-NO2 is endogenously present in healthy human blood. Interestingly, the HPLC elution profiles for plasma- and blood-derived OA-NO2 acquired during qualitative analysis show single peaks rather than overlapping species as seen for the synthetic standard, suggesting the possibility that only one regioisomer is present in vivo. The peaks in the elution profiles for both urine and plasma have the same retention times as the second peak of the synthetic standard.
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1.2 fmol on column (data not shown). Blood samples obtained from 10 healthy human volunteers (5 female, 5 male, ages ranging from 24 to 53) revealed free OA-NO2 in red cells (i.e. OA-NO2 not esterified to glycerophospholipids or neutral lipids) to be 59 ± 11 pmol/ml packed cells (TABLE THREE). Total free and esterified OA-NO2, the amount present in saponified samples, was 214 ± 76 pmol/ml packed cells. Thus, 75% of OA-NO2 in red cells is esterified to complex lipids (9). In plasma, the free and esterified OA-NO2 concentrations were 619 ± 52 and 302 ± 369 nM, respectively, and thus are more abundant than linoleic acid nitration products (9). Control studies revealed that the extraction and analysis conditions do not induce OA-NO2 formation. Present data also show that saponification reactions induce loss of fatty acid nitro derivatives (data not shown), suggesting that current quantitative results may be an underestimation of actual pool sizes of esterified fatty acid nitroalkene adducts.
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In Vitro Nitration of Oleic Acid to OA-NO2—The in vivo detection of nitrated mono-unsaturated fatty acids raised question as to how these derivatives may be formed in vivo. To gain insight into potential mechanisms of formation of OA-NO2, in vitro reactions were performed to determine whether free radical or alternative mechanisms can generate this nitrated fatty acid species (Fig. 8). Treatment of oleic acid with MPO, H2O2 and yielded OA-NO2 with an HPLC elution profile identical to synthetic OA-NO2. Additionally, nitrohydroxy adducts were observed. Oleic acid treated under mild acidic conditions (pH 4.0) in the presence of also generated OA-NO2 with the same physical characteristics as OA-NO2 prepared by nitrosenylation. Relatively less nitrohydroxy adducts were generated as compared with the amount generated by MPO. Finally, treatment of oleic acid with ONOO- resulted in significant formation of OA-NO2 and nitrohydroxy adducts.
Activation of PPARs by OA-NO2—Recently, LNO2 was identified as an endogenous PPAR ligand (17). Considering the even greater levels of OA-NO2 detected in vivo, OA-NO2 was compared with LNO2 as a PPAR, PPAR, and PPAR ligand. CV-1 cells were transiently co-transfected with a plasmid containing the luciferase gene under regulation by three PPREs in concert with PPAR, PPAR, or PPAR expression plasmids. Dose-dependent activation by OA-NO2 was observed for all PPARs (Fig. 9A), with PPAR showing the greatest response (significant activation at 100 nM). PPAR and PPAR showed significant activation at 300 nM OA-NO2. Nitrated oleic acid was consistently more potent than LNO2 in the activation of PPAR. A concentration of 1 µM OA-NO2 typically induced the same degree of reporter gene expression as 3 µM LNO2 and 1 µM rosiglitazone, with these activities partially inhibited by the PPAR antagonist GW9662 (Fig. 9B). Native fatty acids did not activate PPARs at these concentrations (data not shown). The greater potency of OA-NO2 as a PPAR agonist, compared with LNO2, motivated evaluation of the relative stability of these molecules. Current data indicate that LNO2 decays in aqueous milieu to generate products that do not activate PPARs (17, 20). Compared with LNO2, OA-NO2 is relatively stable in aqueous conditions with only minimal decay occurring after 2 h (Fig. 5).
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ligand were further assessed by evaluating its impact on adipocyte differentiation, as PPAR-dependent gene expression plays an essential role in the development of adipose tissue (28, 33). 3T3-L1 preadipocytes were treated with OA-NO2 (3 µM), LNO2 (3 µM), and negative controls for 2 weeks (Fig. 10A). Adipocyte differentiation was assessed both morphologically and via oil red O staining, which indicated the accumulation of intracellular lipids. Vehicle, oleic acid and linoleic acid did not induce adipogenesis. In contrast, OA-NO2 (3 µM) and LNO2 (3 µM) induced 60 and 30% of 3T3-L1 preadiopcyte differentiation, respectively. Rosiglitazone, a synthetic PPAR ligand, also induced PPAR-dependent preadiopcyte differentiation (Ref. 17 and data not shown). OA-NO2 and rosiglitazone-induced pre-adipocyte differentiation resulted in expression of specific adipocyte markers (PPAR2 and aP2); oleic acid had no effect on these gene products (Fig. 10B). PPAR ligands also play a central role in glucose uptake and metabolism, with agonists widely used as insulin-sensitizing drugs. Consistent with its potent PPAR ligand activity, OA-NO2 induced an increase in the deoxyglucose uptake for the differentiated adipocytes (Fig. 11A). This effect of OA-NO2 (1 µM) was almost paralleled by higher concentrations of LNO2 (3 µM). The increased adipocyte glucose uptake, induced by nitrated fatty acids and the positive control rosiglitazone, was partially inhibited by GW9662 (Fig. 11B). In aggregate, these observations reveal that OA-NO2 manifests well characterized PPAR-dependent signaling actions. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The reactions of .NO and its redox-derived products with lipids are multifaceted. Model studies of photochemical air pollutant-induced lipid oxidation reveal that exceedingly high concentrations of nitrogen dioxide (.NO2) induce both oxidation and nitration of fatty acids in phosphatidylcholine liposomes and fatty acid methyl ester preparations (37-39). Subsequently, reaction systems were designed to explore the interactions of endogenous .NO and .NO-derived species with fatty acids, including the superoxide reaction product ONOO- and the nitrite acidification product nitrous acid (HNO2). These model studies of the inflammatory reactions of .NO with fatty acids supported that (a) .NO mediates potent inhibition of autocatalytic radical chain propagation reactions of lipid peroxidation (40, 41) and (b) .NO-derived species produce both nitrated and oxidized derivatives of unsaturated fatty acids (3, 42). One product of these reaction pathways, LNO2, is present at 500 nM concentration in healthy human red cells and plasma and serves as a ligand for the PPAR nuclear lipid receptor family (9, 17). This insight, coupled with the fact that oleic acid is the most abundant unsaturated fatty acid in living organisms, motivated the present search for other potential endogenous nitrated fatty acid derivatives that might translate tissue redox signaling reactions.
The structure of OA-NO2 (Fig. 1) was defined on the basis of the synthetic rationale and NMR analysis (Fig. 3). Proton and 13C NMR spectra indicate that synthetic OA-NO2 is comprised of two regioisomers, 9- and 10-nitro-9-cis-octadecenoic acids, with no trans-isomers apparent. Peaks characteristic of the nitroalkene and olefinic carbons in the 13C spectrum appear as doublets that are equal in intensity, indicating an equivalent distribution between regioisomers. HPLC ESI MS/MS further characterized synthetic OA-NO2. The combined fragmentation pattern of OA-NO2 regioisomers was obtained by CID, which provided a "molecular fingerprint" used to identify OA-NO2 in biological samples (Fig. 6). ESI MS/MS analysis of lipid extracts derived from plasma and red cells yielded spectra with identical HPLC retention times and major product ions, confirming that OA-NO2 exists endogenously. It is not possible from MS analysis, however, to determine the cis/trans conformation of OA-NO2 regioisomers. Quantitative analysis of plasma and red cells showed that OA-NO2 is present in the vasculature at net concentrations 50% greater than LNO2 (TABLE THREE). The combined concentrations of free and esterified OA-NO2 and LNO2 are well above 1 µM. Multiple in vitro studies support that this is a concentration range capable of eliciting robust cell signaling responses.
The NO2 functional groups of OA-NO2 and LNO2 are located on olefinic carbons. This configuration imparts a unique chemical reactivity that enables the release of .NO during aqueous decay of nitroalkenes via a modified Nef reaction (20). Furthermore, the -carbon proximal to the alkenyl NO2 group is strongly electrophilic and reacts with H2O via a Michael addition-like mechanism to generate nitrohydroxy adducts (Figs. 2 and 7). Nitrohydroxyarachidonic acid species have been detected in bovine cardiac muscle (43), and nitrohydroxylinoleic acid has been identified in lipid extracts obtained from hypercholesterolemic and post-prandial human plasma, suggesting that this is a ubiquitous derivative (44). The present identification of a wide spectrum of nitrated fatty acids and corresponding nitrohydroxy fatty acid derivatives in human plasma and urine reveals that nitration reactions occur with all unsaturated fatty acids (Figs. 2 and 7). The hydroxyl moiety of nitrohydroxy fatty acid derivatives destabilizes the adjacent carbon-carbon bond, facilitating heterolytic scission reactions that generate predictable fragments during CID (Fig. 7). Present data indicate that nitrohydroxy adducts of LNO2 and OA-NO2 are not ligands for PPAR (Ref. 17 and data not shown).
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). The oxidation of by heme peroxidases, such as myeloperoxidase, is also a significant source of inflammatory .NO2 production (49, 50). These alkene nitration mechanisms yield nitrated fatty acids that are structurally similar or identical to the OA-NO2 and nitrohydroxy adducts detected clinically (Fig. 8). Nitration by a free radical mechanism might suggest that all olefinic carbons within a fatty acid would be susceptible nitration targets, with the additional likelihood of double bond rearrangement and conjugation. The discovery of OA-NO2 lends critical perspective to this issue, because monounsaturated fatty acids are less susceptible but still capable of oxidation reactions (51). In view of the present structural data regarding nitroalkene positional isomer distribution, alternative fatty acid nitration mechanisms may also occur. For example, nitration by an ionic addition reaction (e.g. nitronium ion, 
) can generate singly nitrated fatty acids with no double bond-rearrangement (26). Since readily reacts with H2O, this species may require localized catalysis (e.g. reaction of ONOO- with transition metals) to serve as a biologically relevant nitrating species. Finally, data in Fig. 8 indicate that acidic nitration reactions occur with both mono- and polyunsaturated fatty acids to yield non-conjugated nitroalkene derivatives of polyunsaturated fatty acids. This precept is also supported by acidified and .NO2-mediated fatty acid methyl ester oxidation and nitration profiles (39, 48, 52).
Of relevance to mechanisms underlying fatty acid nitration in vivo, the nitrohydroxy adducts of 
9 unsaturated fatty acids examined in the present study (18:1, 18:2, and 18:3) all yield a predominant CID fragment of m/z 171 (Fig. 7). This mass is consistent with 9-oxo-nonanoic acid, a CID fragment generated with standards when the NO2 group is located at the 10-carbon and the hydroxyl moiety at the 9-carbon. There are several interpretations of these data. First, the differences in relative intensities of the CID products may be due to differential fragmentation efficiencies. Indeed, the m/z 171 product generated from the C-10 adduct is a 9-oxo-nonanoic anion, whereas the C-9 product (m/z 202) is 9-nitro-nonanoic anion. An alternative interpretation is that the C-10 nitrohydroxy adduct is more predominant, suggesting that either strict steric control or enzymatic mechanisms regulate the stereospecificity of biological fatty acid nitration. The nitration of 9 unsaturated fatty acids to C-10 nitroalkene derivatives, with retention of double bond arrangement, supports that stereospecific enzymatic reactions may mediate fatty acid nitration. It is also possible that nitrated fatty acids are made bioavailable from dietary sources consisting of stereospecific fatty acid nitroalkene derivatives. Further studies are currently under way to address this issue.
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(17), a nuclear hormone receptor that binds lipophilic ligands and induces DNA binding of the transcription factor complex at DR1-type motifs in the promoter sites of target genes. Downstream effects of PPAR activation include modulation of metabolic and cellular differentiation genes, regulation of inflammatory responses, adipogenesis, and glucose homeostasis (18, 19). In the vasculature, PPAR is expressed in monocytes, macrophages, smooth muscle cells, and endothelium (53) and plays a central role in regulating the expression of genes related to lipid trafficking, cell proliferation, and inflammatory signaling. Herein we show that OA-NO2 also serves as a PPAR, -, and - ligand that exceeds the potency of LNO2 and rivals the potency of synthetic PPAR ligands such as fibrates and thiazo-lidinediones (Figs. 9, 10, 11). The greater potency of OA-NO2 as a PPAR ligand relative to LNO2 is either due to increased aqueous stability (Fig. 5), increased receptor affinity, or both.
The combined blood concentrations of OA-NO2 and LNO2 in healthy humans exceeds 1 µM (TABLE THREE); thus, they are present at concentrations capable of modulating inflammatory cell function and activation of PPAR receptors. Endogenous blood concentrations of nitroalkenes also far exceed those of previously proposed endogenous PPAR ligands (17). These data thus have broad implications for the .NO and redox signaling reactions that play a crucial role in dysregulated cell growth and differentiation, metabolic syndrome, atherosclerosis, diabetes, and a variety of inflammatory conditions, all clinical pathologies that include a significant contribution from PPAR-regulated cell signaling mechanisms (54).
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FOOTNOTES |
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0.05; Student's t test). B, nitrated oleic acid was more potent than LNO2 in the activation of PPAR
