# Patent

## Title
Nitrated lipids and methods of making and using thereof

## Abstract

Described herein are nitrated lipids and methods of making and using the nitrated lipids.

## Background
CROSS REFERENCE TO RELATED APPLICATIONS
This application is a continuation of U.S. patent application Ser. No. 13/646,985, filed Oct. 8, 2012, which is a continuation of U.S. patent application Ser. No. 12/797,460, filed Jun. 9, 2010, which is a division of U.S. patent application Ser. No. 11/568,377, filed Oct. 26, 2006, which claims priority to U.S. Patent Application Ser. No. 60/566,005, filed Apr. 28, 2004, the entire disclosures of which are hereby incorporated by reference in their entireties for all purposes.


ACKNOWLEDGEMENTS
This invention was made with government support under Grant Numbers RO1HL58115 and RO1HL64937 awarded by the National Institutes of Health. The government has certain rights in the invention.


BACKGROUND
Nitric oxide (.NO) is an endogenously generated, lipophilic signaling molecule that maintains vascular homeostasis via stimulation of soluble guanylate cyclase (1). In addition to mediating vascular relaxation, .NO potently modulates oxygen radical reactions, inflammatory cell function, post-translational protein modification and regulation of gene expression (2-5). There are multiple pathways whereby .NO-derived species can mediate the oxidation and nitration of biomolecules such as unsaturated fatty acids. Nitric oxide reacts at diffusion-limited rates with superoxide (O2.−, k=1.9×1010 M−1 sec−1) to yield peroxynitrite (ONOO−) and its conjugate acid, peroxynitritrous acid (ONOOH), the latter of which undergoes homolytic scission to nitrogen dioxide (.NO2) and hydroxyl radical (.OH) (2, 6). Also, biological conditions favor the reaction of ONOO− with CO2, yielding nitrosoperoxycarbonate (ONOOCO2−; k=3×104 M−1 sec−1), which rapidly yields .NO2 and carbonate (.CO3−) radicals via homolysis, or rearrangement to NO3− and CO2 (7). During inflammation, neutrophil myeloperoxidase and heme proteins such as myoglobin and cytochrome c catalyze H2O2-dependent oxidation of nitrite (NO2−) to .NO2, resulting in biomolecule oxidation and nitration that is influenced by the spatial distribution of catalytic heme proteins (8-11). Finally, even though the rate of reaction of NO with O2 is slow, (k=2×106 M−2 sec−1) the small molecular radius, uncharged nature and lipophilicity of .NO and O2 facilitate their diffusion and concentration in membranes and lipoproteins up to 20-fold (12-14). This “molecular lens” effect induced by .NO and O2 solvation in hydrophobic cell compartments accelerates the reaction of .NO with O2 to yield N2O3 and N2O4. As a result of these various reactions, a rich spectrum of primary and secondary reactions yield products capable of concerted oxidation, nitrosation and nitration of target molecules.
Multiple mechanisms can account for the nitration of fatty acids by .NO2 (15-20). During both basal cell signaling and tissue inflammatory conditions, .NO2 generated by the aforementioned reactions can react with membrane and lipoprotein lipids. Environmental sources also yield .NO2 as a product of photochemical air pollution and tobacco smoke. In both in vivo and in vitro systems, .NO2 has been shown to initiate auto-oxidation of polyunsaturated fatty acids via hydrogen abstraction from the bis-allylic carbon to form nitrous acid and a resonance-stabilized allylic radical (21). Depending on the radical environment, the lipid radical species can react with molecular oxygen to form a peroxyl radical. During inflammation or ischemia, when O2 levels are lower, lipid radicals can react to an even greater extent with .NO2 to generate multiple nitration products including singly nitrated, nitrohydroxy- and dinitro-fatty acid adducts (18, 19, 21). These products can be generated via either hydrogen abstraction or direct addition of .NO2 across the double bond. Hydrogen abstraction causes a rearrangement of the double bonds to form a conjugated diene; however, the addition of .NO2 maintains a methylene-interrupted diene configuration to yield singly nitrated polyunsaturated fatty acids (18). This arrangement is similar to nitration products generated by the nitronium ion (NO2+), which can be produced by ONOO− reaction with heme proteins or via secondary products of CO2 reaction with ONOO− (20).
Reaction of polyunsaturated fatty acids with acidified nitrite (HNO2) generates a complex mixture of products similar to those formed by direct reaction with .NO2, including the formation of singly nitrated products that maintain the bis-allylic bond arrangement (18, 19). The acidification of NO2− creates a labile species, HNO2, which is in equilibrium with secondary products, including N2O3, .NO and .NO2, all of which can participate in nitration reactions. The relevance of this pathway as a mechanism of fatty acid nitration is exemplified by physiological and pathological conditions wherein NO2− is exposed to low pH (e.g., <pH 4.0). This may conceivably occur in the gastric compartment, following endosomal or phagolysosomal acidification or in tissues following-post ischemic reperfusion.
Nitrated linoleic acid (LNO2) displays robust cell signaling activities that (at present) are anti-inflammatory in nature (20, 22-25). Synthetic LNO2 inhibits human platelet function via cAMP-dependent mechanisms (26) and inhibits neutrophil O2.− generation, calcium influx, elastase release, CD11b expression and degranulation via non-cAMP, non-cGMP-dependent mechanisms (27). LNO2 also induces vessel relaxation in part via cGMP-dependent mechanisms (22, 28). In aggregate, these data, derived from a synthetic fatty acid adduct, infer that LNO2 species represent a novel class of lipid-derived signaling mediators. To date, a gap in the clinical detection and structural characterization of nitrated fatty acids has limited defining LNO2 derivatives as biologically-relevant lipid signaling mediators that converge .NO and oxygenated lipid signaling pathways.
Therefore, it would be advantageous to produce nitrated lipids in substantially pure form so that their cell signaling activities can be characterized and their purified derivatives can be used to treat various diseases. Described herein are nitrated lipids and methods for producing nitrated lipids in pure form. Also described herein are methods for using the nitrated lipids to treat various diseases.
SUMMARY
Described herein are nitrated lipids and methods of making and using the nitrated lipids. The advantages of the invention will be set forth in part in the description which follows, and in part will be obvious from the description, or may be learned by practice of the aspects described below. The advantages described below will be realized and attained by means of the elements and combinations particularly pointed out in the appended claims. It is to be understood that both the foregoing general description and the following detailed description are exemplary and explanatory only and are not restrictive.



BRIEF DESCRIPTION OF THE DRAWINGS
The accompanying drawings, which are incorporated in and constitute a part of this specification, illustrate several embodiments of the invention and together with the description, serve to explain the principles of the invention.
FIG. 1 shows a reaction scheme for producing nitrated lipids.
FIG. 2 shows the separation of C-10 and C-12 nitrated linoleic acid from a crude reaction mixture by thin layer chromatography.
FIG. 3 shows the extinction coefficient (A) and ultraviolet light absorption spectrum (B) of nitrated linoleic acid.
FIG. 4 shows the characterization of nitrated linoleic acid by GC mass spectrometry.
FIG. 5 shows the HPLC resolution of individual positional isomers of nitro derivatives of linoleic acid present in a) synthetic preparations, b) red cells and c) plasma; and in concert with this exemplification of positional isomer resolution is the structural characterization of individual nitrated linoleic acid positional isomers by electrospray ionization triple quadrupole mass spectrometry.
FIG. 6 shows the standard curve used for quantitative analysis of red blood cell and plasma nitrated linoleic acid content.
FIG. 7 shows that LNO2 is a potent PPAR ligand. (A) CV-1 cells, transiently co-transfected with different nuclear receptor ligand binding domains fused to the Gal4 DNA binding domain and the luciferase reporter gene under the control of four Gal4 DNA binding elements, were incubated with vehicle (methanol) or LNO2 (3 μM, 2 hr, n=4). (A, inset) Dose-response of LNO2-dependent PPARγ ligand binding domain activation (n=4). (B) Dose-response of LNO2-dependent PPARγ, α and δ activation (n=4). The luciferase reporter gene was under the control of three PPAR response elements (C) Response of CV-1 cells transfected with PPARγ and a luciferase reporter construct under the control of PPRE following exposure to LNO2 and other reported PPARγ ligands (1 and 3 μM each of ciglitazone, 15-deoxy-Δ12,14-PGJ2, 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphocholine (LPA 16:0), 1-oleoyl-2-hydroxy-sn-glycero-3-phosphocholine (LPA 18:1), 1-O-hexadecyl-2-azelaoyl-sn-glycero-3-phosphocholine (AzPC), 1-palmitoyl-2-azelaoyl-sn-glycero-3-phosphocholine (azPC ester), Δ9,11-conjugated linoleic acid (CLA-1) and Δ10,12-conjugated linoleic acid (CLA-2), with (n=3 to 5). “Vector” indicates empty vector (C, inset) Using the same reporter construct, the dose response of PPARγ activation by LNO2, rosiglitazone, 15-deoxy-Δ12,14-PGJ2 and linoleic acid was measured (n=3). All values are expressed as mean±SD. (*) represents significantly different (P<0.05) from vehicle control using Student's t test. All experiments were repeated at least three times.
FIG. 8 shows the characterization of the PPARγ ligand activity of LNO2. (A) Using CV-1 cells cotransfected with PPARγ and PPRE-controlled luciferase expression plasmids, the activation of PPARγ by LNO2 was evaluated in the absence or presence of PPARγ-specific antagonist GW9662 added 1 hr prior to LNO2 addition or upon co-addition of the RXR receptor co-activating ligand 9-cis-retinoic acid (n=3). PPARγ activation by LNO2 was inhibited in a dose-dependent manner by GW9662 and was enhanced in the presence of the coactivator 9-cis-retinoic acid. (B) The action of LNO2 as a PPARγ ligand was compared with LNO2-derived decay products. Effective LNO2 concentrations after selected decay periods were measured by LC-MS with electrospray ionization using [13C]LNO2 as internal standard (9). PPARγ activation was assessed via PPRE reporter analysis in CV-1 cells. (n=3) (c) Potential PPARγ ligand activity of LNO2 decay products was measured via PPRE reporter analysis (n=4) (D) Competition of LNO2, linoleate and unlabeled Rosiglitazone for PPARγ-bound [3H] Rosiglitazone. For (A-C), all values are expressed as mean±SD. (*) represents significantly different (P<0.05) from vehicle control, and (#) represents significantly different from LNO2 alone, using Student's t test. All experiments were repeated at least three times.
FIG. 9 shows that LNO2 induces CD36 expression in macrophages and adipogenesis of 3T3-L1 preadipocytes. (A) Mouse RAW264.7 macrophages at  ̃90% confluence were cultured in DMEM with 1% FBS for 16 hours and then treated with various stimuli for 16 hours as indicated. The PPARγ-specific antagonist GW9662 was added 1 h prior to the treatment. The cell lysate was immunoblotted with anti-CD36 and anti-β-actin antibodies. (B,C) Two days after reaching confluence, 3T3-L1 preadipocytes were cultured for 14 days and stained using Oil red O as previously (18) (B) or treated with various stimuli as indicated and the cell lysate was immunoblotted with anti-PPARγ, anti-aP2 and anti-β-actin antibodies (C). (D) LNO2 increases [3H]-2-deoxy-D-glucose uptake in 3T3-L1 adipocytes. Left panel: The dose-dependent effects of LNO2 on [3H]-2-deoxy-D-glucose uptake in 3T3-L1 adipocytes. Right panel: PPARγ-specific antagonist GW9662 was added 1 h before the treatment. [3H]-2-deoxy-D-glucose uptake assay was performed as described in supplemental methods. All experiments were repeated at least three times. Values are expressed as mean±SD (n=6). Statistical analysis was done by using Student's t test (*p<0.05 vs vehicle control; #p<0.05 vs GW9662 untreated groups). Vehicle (Veh); Rosiglitazone (Rosi); 15-deoxy-PGJ2 (15-d-PGJ2); linoleic acid (LA).
FIG. 10 shows nitrated oleic acid (OA-NO2). Two regioisomers of OA-NO2 were synthesized by nitrosenylation of oleic acid and purified as described in Experimental Procedures, generating 9- and 10-nitro-9-cis-octadecenoic acids.
FIG. 11 shows nitrated fatty acid species in plasma and urine. Potential nitroalkene products were evaluated in plasma and urine. Fatty acids were extracted from clinical samples and analyzed by ESI-MS/MS as described in Methods. Nitrated fatty acid adducts (—NO2) and their nitrohydroxy counterparts (L(OH)—NO2) were detected using the multiple reaction monitoring (MRM) scan mode (Table 3) and are presented as base to peak HPLC elution profiles with maximum ion intensity given on the left axis. Six fatty acids were monitored: oleic acid (18:1), linoleic acid (18:2), linolenic acid (18:3), arachidonic acid (20:4) eicosapentaenoic acid (20:5) and docosahexaenoic acid (22:6). In plasma and urine, all of the nitrated fatty acids and their Michael addition products with H2O (nitrohydroxy adducts) appear in the HPLC elution profiles.
FIG. 12 shows 1H and 13C NMR spectrometry of synthetic nitro-oleate (OA-NO2). Proton (A) and 13C (B) NMR spectrometry confirmed the structure of synthetic OA-NO2. Identified protons and carbons are indicated for each regioisomer; downfield shifts are presented in ppm. 13C NMR spectrometry indicates that synthetic OA-NO2 is a mixture of two regioisomers, with most carbon peaks appearing as doublets. The equal height of the doublets suggests an equal molar ratio of the regioisomers. The peaks appearing at 152 ppm and 136 ppm are the carbons a and β to the alkenyl nitro group, respectively.
FIG. 13 shows the spectrophotometric analysis of OA-NO2. (A) 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. An absorbance maximum at 270 nm was identified. (B) Extinction coefficients for OA-NO2 and [13C]OA-NO2 were determined by plotting absorbance (λ270) vs. concentration, resulting in calculated values of ε=8.22 and 8.23 cm−1 mM−1, respectively.
FIG. 14 shows the GC-MS analysis of synthetic OA-NO2. (A) Methyl esters of the two synthetic OA-NO2 regioisomers were generated as described in Methods and analyzed by EI GC MS/MS. The mixture was resolved using a 30 m fused silica column and detected by total ion monitoring. The upper chromatogram shows partial resolution of the two regioisomers. Product ion analysis of each peak (B) revealed that the first and second eluting peaks (37.41 and 37.59 min, respectively) each has a unique identifying ion: m/z 168 (peak 1) and m/z 156 (peak 2).
FIG. 15 shows the identification and characterization of synthetic and blood OA-NO2 by HPLC ESI MS/MS. (A, left panels) OA-NO2 and [13C]OA-NO2 were characterized by HPLC-ESI MS/MS. Nitrated oleic acid species were separated by HPLC and detected by acquiring MRM transitions consistent with the loss of the alkenyl nitro group [M−HNO2]−, m/z 326/279 and m/z 344/297 for OA-NO2 and [13C]OA-NO2, respectively. (A, right panels) Concurrent to MRM detection, product ion analysis was performed to generate identifying fragmentation patterns also used to characterize in vivo OA-NO2. The predominant product ions generated by collision-induced dissociation are identified in Table 3. (B) Total lipid extracts were prepared from packed red cell and plasma fractions of venous blood and directly analyzed by mass spectrometry.
FIG. 16 shows the product ion analysis of fatty acid nitrohydroxy-adducts in urine. The presence of nitrohydroxy fatty acids in urine was confirmed by product ion analysis run concomitant to MRM detection. Structures of possible adducts are presented along with their diagnostic fragments and product ion spectra for (A) 18:1(OH)—NO2, (B) 18:2(OH)—NO2 and (C) 18:3(OH)—NO2. The 10-nitro regioisomer of 18:1(OH)—NO2 is present in urine, as evidenced by the intense peak corresponding to m/z 171; also present are fragments consistent with the 9-nitro regioisomer (m/z 202), loss of a nitro group (m/z 297) and water (m/z 326). 18:2(OH)—NO2 also shows a predominant m/z 171 fragment, again consistent with an oxidation product of LNO2 nitrated at the 10-carbon (B). Diagnostic fragments for the three other potential regioisomers were not apparent. Finally, multiple regioisomers of 18:3(OH)—NO2 are present (C).
FIG. 17 shows that OA-NO2 is a PPARγ agonist. (A) CV-1 cells transiently co-transfected with a plasmid containing the luciferase gene under the control of three tandem PPRE (PPRE×3 TK-Luciferase) and hPPARγ, hPPARα or hPPARδ expression plasmids showed all three PPARs were activated by OA-NO2, with the relative activation of PPARγ>PPARδ>PPARα. All values are expressed as mean±SD (n=3). PPARγ activation was significantly different from vehicle at 100 nM OA-NO2, whereas PPARα and PPARγ activation were significantly different from vehicle at 300 nM and 1 μM OA-NO2, respectively (P≦0.05; Student's t test). (B) Nitrated oleic acid appears to be more potent than LNO2 in the activation of PPARγ, with 1 μM OA-NO2 inducing similar activity as 3 μM LNO2 versus control (P≦0.05; Student's t test). Activation of PPARγ was partially yet significantly blocked using the PPARγ antagonist GW9662 (P≦0.05; Student's t test). (C) Equimolar concentrations of OA-NO2 and LNO2 (3 μM) were incubated in 100 mM phosphate buffer. After 2 hr, only 25% of the initial OA-NO2 had degraded;  ̃80% of LNO2 degrades in the same time period, indicating a great aqueous stability of OA-NO2. All values are expressed as mean±SD (n>3).
FIG. 18 shows that OA-NO2 induces adipogenesis in 3T3 L1 preadipocytes. PPARγ plays an essential role in the differentiation of adipocytes. 3T3-L1 preadipocytes were treated with OA-NO2, LNO2, Rosiglitazone and controls (oleic acid, linoleic acid and DMSO) for two weeks. (A) Adipocyte differentiation was assessed both morphologically and via oil red O staining, which reveals the accumulation of intracellular lipids. Vehicle, oleic acid and linoleic acid did not induce adipogenesis, while OA-NO2 induced  ̃60% of 3T3-L1 preadiopcyte differentiation; LNO2 induced  ̃30%, reflecting the greater potency of OA-NO2. As the positive control, Rosiglitazone also induced PPARγ-dependent adipogenesis. (B) OA-NO2 and Rosiglitazone-induced preadipocyte differentiation resulted in the expression of adipocyte-specific markers (PPARγ2 and aP2), an event not detected for oleic acid.
FIG. 19 shows that OA-NO2 induces [3H]-2-deoxy-D-glucose uptake in differentiated 3T3 L1 adipocytes. (A) PPARγ ligands induce glucose uptake in adipose tissue. To further define the functional significance of OA-NO2 as a PPARγ ligand, 3T3-L1 preadipocytes were differentiated to adipocytes and treated with OA-NO2 or LNO2 for two days prior to addition of [3H]-2-deoxy-D-glucose. OA-NO2 induced significant increases in glucose uptake; these effects were paralleled by LNO2 (P≦0.05; Student's t test). (B) The increases in glucose uptake induced by nitrated lipids and the positive control Rosiglitazone were significantly inhibited by the PPARγ-specific antagonist GW9662 (P≦0.05; Student's t test). All values are expressed as mean±SD (n=3).
FIG. 20 shows EPR and UV/visible spectroscopic detection of .NO release by LNO2. (A) EPR spectral analysis of cPTIO (200 μM, red line) reduction to cPTI (black line) by LNO2 (300 μM) decay during a 60 min decay period. (B) Differential spectra of oxymyoglobin (20 μM) oxidation by LNO2 (200 μM). Spectra were repetitively recorded at 5 min intervals and show the decrease in the 580 nm and 543 nm maxima (characteristic of the α and β visible band absorbance of the oxymyoglobin) and the increase in 630 and 503 nm maxima characteristic of metmyoglobin. (C) .NO release rate detected by oxymyoglobin (20 μM) oxidation in the presence of different concentrations of LNO2. Values expressed as mean±SD of 2 independent experiments repeated four times. (D) UV spectra of LNO2 taken every 10 min, revealing loss of the characteristic absorbance of the NO2 group at 268 nm and the formation of a new chromophore at 320 nm. (A, B and D) Spectra are representative of 3 independent experiments.
FIG. 21 shows nitrite formation during LNO2 decay. The time-dependent formation of NO2− during LNO2 (initial concentration 200 μM) decomposition was measured in parallel with oxymyoglobin (20 μM) oxidation. Nitrite formation was measured in the absence of oxymyoglobin. Values expressed as mean±SD of 3 independent experiments repeated three times.
FIG. 22 shows the pH dependency of .NO formation from LNO2. The rate of .NO formation from 80 μM LNO2 (detected using EPR spectroscopic measurement of cPTIO (80 μM) reduction) was determined in buffers with different pHs.
FIG. 23 shows chemiluminescent detection of .NO release by LNO2. A) LNO2 (5 and 10 mM) was incubated in a capped vial under aerobic conditions for 3 min and the gas phase injected into an O3 chemiluminescence detector. Additionally, known concentrations of DEA-NONOate (nM) (in 10 mM NaOH) were added to a capped vial containing 0.5 M HCl and the gas phase was injected into the chemiluminescence detector. B) Phosphate buffer (50 mM phosphate pH 7.4 containing 10 μM DTPA) was illuminated with a xenon arc lamp. NO formation was examined by O3-based chemiluminescence after the injection of MeOH (20 μl), LNO2 (4 nmol in 20 μl MeOH, two additions made before and after sodium nitrite addition) and sodium nitrite (4 nmol in 20 μl phosphate buffer). C) Blood was obtained by cardiac puncture of LPS-treated rats, red cells removed by centrifugation and plasma samples treated as noted in Experimental Procedures. The following conditions were studied in panel C: (1) I3− alone; (2) I3− plus sulfanilamide; (3) I3− plus sulfanilamide and HgCl2, with 3.5 nmol LNO2 treated with I3− plus sulfanilamide and HgCl2, as for the corresponding plasma sample. Derived .NO was measured by .NO chemiluminescence analysis. Traces are representative from three different experiments.
FIG. 24 shows spectroscopic (EPR, UV and visible) analysis of micellar inhibition of LNO2 decomposition and .NO release. In panels A-D, closed diamonds represent conditions containing OTG and open diamonds represent OG. A) .NO release from LNO2 (80 μM) in the presence of different OTG and OG concentrations after 60 min, as measured by EPR detection of cPTIO (80 μM) conversion to cPTI. The extent of cPTIO (80 μM) conversion to cPTI by known concentrations of proli-NONOate was utilized to calculate yields of NO. B) NO release rate from LNO2 (130 μM) in the presence of different concentrations of OG and OTG, as measured by oxidation of oxymyoglobin (20 μM) to metmyoglobin. An extinction coefficient of 14.4 mM−1 cm−1 was used to calculate yields of .NO. C) Initial decomposition rates of LNO2 (37 μM), measured at 268 nm, in the presence of different concentrations of OTG and OG. D) Same as C, but rates of LNO2 decomposition product formation at 320 nm were measured. Values are expressed as mean±SD of at least 3 independent experiments repeated three or four times.
FIG. 25 shows micellar and phosphatidylcholine-cholesterol liposome inhibition of LNO2 decomposition and .NO release. A) LNO2 (200 μM) decomposition was measured in the absence and presence of 15 mg/ml OTG by mass spectrometry. B) Calculation of partition coefficient of LNO2 into OTG and OG micelles from data shown in FIG. 25D). C) LNO2 (80 μM)-dependent NO formation was measured by cPTIO (200 μM) reduction to cPTI at different times in the presence of increasing liposome concentration (0-5 mg/ml).
FIG. 26 shows formation of L(OH)NO2 from LNO2. A) LNO2 was incubated in the presence (dotted line) or absence (solid line) of 15 mg/ml OTG, lipids were extracted and analyzed by ESI MS/MS. The presence of OTG inhibited LNO2 decay as indicated by the MRM transition m/z 324/277 (A) and the formation of species with transitions m/z 342/171 and 342/295, which correspond to 9-hydroxy-10-nitro-12-octadecaenoic acid specifically (B), and all L(OH)NO2 regioisomers (C), respectively. In the absence of OTG, increased L(OH)NO2 yields were formed. D) Structures of possible nitrohydroxy adducts are presented along with their diagnostic fragments. E) Product ion spectra of L(OH)NO2 showed two predominant ions consistent with expected fragments shown in (D), m/z 171 (9-hydroxy-10-nitro-12-octadecaenoic acid) and m/z 211 (12-hydroxy-13-nitro-9-octadecaenoic acid
FIG. 27 shows mass spectrometric detection of LNO2 decay products. LNO2 (500 μM) was incubated in aqueous buffer phosphate buffer (100 mM phosphate pH 7.4 containing 100 μM DTPA) for 0, 45 and 240 min (A-C, respectively). Decay products were CHCl3-extracted and analyzed by direct ESI MS/MS. Products were detected in the negative ion mode. The 293 m/z ion corresponds to an expected Nef reaction product, a conjugated ketone; m/z 342 is consistent with the mass of vicinal nitrohydroxy linoleic acid; and m/z 340 and 356 represent the hydroxy and peroxy derivatives of LNO2, respectively.
FIG. 28 shows Scheme 1, Hydrophobic regulation of LNO2 decomposition and NO release in lipid bilayers and micelles. The partitioning of LNO2 into different cell compartments is in part governed by its partition coefficient (K ̃1500). LNO2 may also be stabilized and placed in “reserve”, in terms of attenuating NO-mediated cell signaling capabilities, by esterification into complex lipids of membranes or lipoproteins. Alternatively, LNO2 derivatives of complex lipids can be formed by direct nitration of esterified unsaturated fatty acids. During inflammatory conditions or in response to other stimuli, LNO2 may be released from complex lipids by A2-type phospholipases or esterases; thus mobilizing “free” LNO2 that can in turn diffuse to exert receptor-dependent signaling actions or undergo decay reactions to release .NO.
FIG. 29 shows Scheme 2, possible mechanisms for NO formation by LNO2. (Stage 1) Due to the strong electrophilic nature of the carbon adjacent to the nitroalkene and the acidity of its bound hydrogen, the vicinal nitrohydroxy fatty acid derivative is in equilibrium with the nitroalkene. (Stage 2) The mechanism of .NO release from LNO2 can result from the formation of a nitroso intermediate formed during aqueous LNO2 decay. This nitroso intermediate is expected to have an especially weak C—N bond, easily forming .NO and a radical stabilized by conjugation with the alkene and stabilized by the OH group, a moiety known to stabilize adjacent radicals.
FIG. 30 shows that nitroalkenes potently activate p-JNK and p-c-Jun protein kinases by stimulating their phosphorylation. The activation of these cell signaling mediators will profoundly impact on cell inflammatory responses, proliferation and differentiation. This example shows human lung epithelial cell responses of p-JNK and p-c-Jun.
FIG. 31 shows that nitroalkenes activate the ERK MAPK pathway in human lung epithelial cells, as shown by a dramatic increase in ERK phosphorylation (e.g., activation) and the phosphorylation of its downstream target signaling protein, pELK. The activation of these cell signaling mediators will profoundly impact on cell inflammatory responses, proliferation and differentiation. This example shows human lung epithelial cell responses of p-JNK and p-c-Jun.
FIG. 32 shows that nitrolinoleate (LNO2), and not the control fatty acid linoleate (LA), inhibits activity of NF-kB pathways as indicated by a) luciferase-linked NFkB-response element reporter assay in response to the inflammatory mediator TNFa and b) direct analysis of the degradation f the NFkB inhibitor protein, IkB in response to the inflammatory mediator E. coli LPS.
FIG. 33 shows the mass spectra of nitro/hydroxy fatty acid.
FIG. 34 shows the mass spectrum of the Michael addition product between nitro linoleate and glutathione.



DETAILED DESCRIPTION
Before the present compounds, compositions, and/or methods are disclosed and described, it is to be understood that the aspects described below are not limited to specific compounds, synthetic methods, or uses as such may, of course, vary. It is also to be understood that the terminology used herein is for the purpose of describing particular aspects only and is not intended to be limiting.
In this specification and in the claims that follow, reference will be made to a number of terms that shall be defined to have the following meanings:
It must be noted that, as used in the specification and the appended claims, the singular forms “a,” “an” and “the” include plural referents unless the context clearly dictates otherwise. Thus, for example, reference to “a pharmaceutical carrier” includes mixtures of two or more such carriers, and the like.
“Optional” or “optionally” means that the subsequently described event or circumstance can or cannot occur, and that the description includes instances where the event or circumstance occurs and instances where it does not. For example, the phrase “optionally substituted lower alkyl” means that the lower alkyl group can or can not be substituted and that the description includes both unsubstituted lower alkyl and lower alkyl where there is substitution.
Ranges may be expressed herein as from “about” one particular value, and/or to “about” another particular value. When such a range is expressed, another aspect includes from the one particular value and/or to the other particular value. Similarly, when values are expressed as approximations, by use of the antecedent “about,” it will be understood that the particular value forms another aspect. It will be further understood that the endpoints of each of the ranges are significant both in relation to the other endpoint, and independently of the other endpoint.
References in the specification and concluding claims to parts by weight, of a particular element or component in a composition or article, denotes the weight relationship between the element or component and any other elements or components in the composition or article for which a part by weight is expressed. Thus, in a compound containing 2 parts by weight of component X and 5 parts by weight component Y, X and Y are present at a weight ratio of 2:5, and are present in such ratio regardless of whether additional components are contained in the compound.
A weight percent of a component, unless specifically stated to the contrary, is based on the total weight of the formulation or composition in which the component is included.
Variables such as R1-R16 used throughout the application are the same variables as previously defined unless stated to the contrary.
By “subject” is meant an individual. The subject can be a mammal such as a primate or a human. The term “subject” can include domesticated animals including, but not limited to, cats, dogs, etc., livestock (e.g., cattle, horses, pigs, sheep, goats, etc.), and laboratory animals (e.g., mouse, rabbit, rat, guinea pig, etc.).
By “contacting” is meant an instance of exposure by close physical contact of at least one substance to another substance. For example, contacting can include contacting a substance, such as a pharmacologic agent, with a cell. A cell can be contacted with a test compound, for example, a nitrated lipid, by adding the agent to the culture medium (by continuous infusion, by bolus delivery, or by changing the medium to a medium that contains the agent) or by adding the agent to the extracellular fluid in vivo (by local delivery, systemic delivery, intravenous injection, bolus delivery, or continuous infusion). The duration of contact with a cell or group of cells is determined by the time the test compound is present at physiologically effective levels or at presumed physiologically effective levels in the medium or extracellular fluid bathing the cell.
“Treatment” or “treating” means to administer a composition to a subject or a system with an undesired condition (e.g., inflammation) or at risk for the condition. The condition can include a disease or a predisposition to a disease. The effect of the administration of the composition to the subject can have the effect of but is not limited to reducing or preventing the symptoms of the condition, a reduction in the severity of the condition, or the complete ablation of the condition.
By “effective amount” is meant a therapeutic amount needed to achieve the desired result or results, e.g., increasing the expression of a gene, inhibiting Ca+2 mobilization in a cell, inhibiting degranulation or CD11b expression in a neutrophil, etc.
Herein, “inhibition” or “suppression” means to reduce activity as compared to a control. It is understood that inhibition or suppression can mean a slight reduction in activity to the complete ablation of all activity. An “inhibitor” or “suppressor” can be anything that reduces the targeted activity.
Herein, “induce” means initiating a desired response or result that was not present prior to the induction step. The term “potentiate” means sustaining a desired response at the same level prior to the potentiating step or increasing the desired response over a period of time.
The term “alkyl group” as used herein is a branched or unbranched saturated hydrocarbon group of 1 to 24 carbon atoms, such as methyl, ethyl, n-propyl, isopropyl, n-butyl, isobutyl, t-butyl, pentyl, hexyl, heptyl, octyl, decyl, tetradecyl, hexadecyl, eicosyl, tetracosyl and the like. A “lower alkyl” group is an alkyl group containing from one to six carbon atoms.
The term “alkenyl group” is defined as a branched or unbranched hydrocarbon group of 2 to 24 carbon atoms and structural formula containing at least one carbon-carbon double bond.
The term “alkynyl group” is defined as a branched or unbranched hydrocarbon group of 2 to 24 carbon atoms and a structural formula containing at least one carbon-carbon triple bond.
The term “ester” is represented by the formula —OC(O)R, where R can be an alkyl, alkenyl, or group described above.
R1-R16 can, independently, possess two or more of the groups listed above. For example, if R1 is a straight chain alkyl group, one of the hydrogen atoms of the alkyl group can be substituted with an ester group. Depending upon the groups that are selected, a first group may be incorporated within second group or, alternatively, the first group may be pendant (i.e., attached) to the second group. For example, with the phrase “an alkyl group comprising an ester group,” the ester group may be incorporated within the backbone of alkyl group. Alternatively, the ester can be attached the backbone of the alkyl group. The nature of the group(s) that is (are) selected will determine if the first group is embedded or attached to the second group.
Disclosed are compounds, compositions, and components that can be used for, can be used in conjunction with, can be used in preparation for, or are products of the disclosed methods and compositions. These and other materials are disclosed herein, and it is understood that when combinations, subsets, interactions, groups, etc. of these materials are disclosed that while specific reference of each various individual and collective combinations and permutation of these compounds may not be explicitly disclosed, each is specifically contemplated and described herein. For example, if a number of different nucleosides and polymeric substrates are disclosed and discussed, each and every combination and permutation of the nucleoside and the polymeric substrate are specifically contemplated unless specifically indicated to the contrary. Thus, if a class of molecules A, B, and C are disclosed as well as a class of molecules D, E, and F and an example of a combination molecule, A-D is disclosed, then even if each is not individually recited, each is individually and collectively contemplated. Thus, in this example, each of the combinations A-E, A-F, B-D, B-E, B-F, C-D, C-E, and C-F are specifically contemplated and should be considered disclosed from disclosure of A, B, and C; D, E, and F; and the example combination A-D. Likewise, any subset or combination of these is also specifically contemplated and disclosed. Thus, for example, the sub-group of A-E, B-F, and C-E are specifically contemplated and should be considered disclosed from disclosure of A, B, and C; D, E, and F; and the example combination A-D. This concept applies to all aspects of this disclosure including, but not limited to, steps in methods of making and using the disclosed compositions. Thus, if there are a variety of additional steps that can be performed it is understood that each of these additional steps can be performed with any specific embodiment or combination of embodiments of the disclosed methods, and that each such combination is specifically contemplated and should be considered disclosed.
I. Nitrated Lipids
In one aspect, the nitrated lipids described herein are lipids comprising at least one nitro group (NO2) covalently bonded to the lipid, wherein the nitrated lipid is substantially pure. The term “substantially pure” as defined herein is a nitrated lipid that exists predominantly as one species. In certain aspects, nitration of a lipid can produce two or more nitration products. For example, the lipid can be nitrated one or more times at different positions on the lipid. These are referred to as positional isomers. Additionally, if the lipid contains a carbon-carbon double bond, the stereochemistry about the carbon-carbon double bond can also vary. These are referred to as stereoisomers. The nitrated lipids described herein are substantially one compound (positional and stereoisomer). In one aspect, the nitrated lipid is 90%, 92%, 94%, 96%, 98%, 99%, 99.5%, or 100% one compound.
In one aspect, the nitrated lipids possess at least one allylic or vinyl nitro group. The phrase “allylic nitro group” has the general formula —C═C—C—(NO2). The phrase “vinyl nitro group” has the general formula —C═C—(NO2). In one aspect, the nitrated lipid possesses only one allylic nitro group. In another aspect, the nitrated lipid possesses only one vinyl nitro group. In another aspect, the nitrated lipid possesses one or more allylic nitro groups and/or one or more vinyl nitro groups.
Lipids known in the art can be nitrated using the techniques described herein to produce nitrated lipids. In general, lipids useful for producing the nitrated lipid include, but are not limited to, fats and fat derived materials. In one aspect, the nitrated lipid can include, but is not limited to, a nitrated fatty acid or ester thereof, a nitrated fatty alcohol, or a nitrated sterol. In another aspect, the nitrated lipid can be a nitrated complex lipid. Examples of complex lipids include, but are not limited to, glycerolipids (e.g., compounds having a glycerol backbone including, but not limited to, phospholipids, glycolipids, monoglycerides, diglycerides, triglycerides) or cholesterol (e.g., cholesterols having fatty acids attached to it such as cholesterol linoleate). In one aspect, the nitrated lipid comprises a fatty acid having at least one ester linkage [—O—C═O(R)], ether group (C—O—R) or vinyl ether group (C—O—C═C—R). Examples of lipids having at least one ether group or vinyl ether group that can be nitrated are depicted below in A and B, respectively.




wherein

    
    
        R14 comprises C16-C22 alkyl, C16-C22 alkenyl, or C16-C22 alkynyl;
        R15 comprises C1-C20 alkyl, C1-C20 alkenyl, or C1-C20 alkynyl; and
        R1 comprises C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl.
    
    


In one aspect, the nitrated lipid is composed of a fatty acid having at least one carbon-carbon double bond. In one aspect, the nitrated lipid can be a nitrated fatty acid such as, for example, 14:1, 16:1, 18:1 (oleic acid), 18:2 (linoleic acid), 18:3 (linolenic acid), 20:4 (arachidonic acid), 22:6, or docosahexanoic acid, where the first number indicates the carbon chain length of the fatty acid, and the second number indicates the number of carbon-carbon double bonds present in the fatty acid.
In one aspect, the nitrated lipid can have the formula I




wherein

    
    
        R1 comprises C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl;
        R2, R3, R7, and R8 comprise, independently, hydrogen, NO2, OH, or OOH;
        R4 comprises C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl;
        wherein R4 comprises a terminal COOR6 group, wherein R6 comprises hydrogen, C1-C24 alkyl, or a pharmaceutically acceptable counterion, wherein R4 optionally comprises one or more NO2, OH, or OOH groups;
        R5 comprises hydrogen or R4 and R5 collectively forms ═C(R9)(R10), wherein R9 comprises C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl, wherein R9 comprises a terminal COOR6 group, wherein R9 optionally comprises one or more NO2, OH, or OOH groups;
        R10 comprises hydrogen, NO2, OH, or OOH; and
        n is from 1 to 24;

wherein the nitrated lipid comprises at least one NO2 group,

wherein the nitrated lipid is substantially pure. In this aspect, the stereochemistry about the carbon-carbon double bond is substantially cis (or Z) or substantially trans (or E).

    
    


In another aspect, the nitrated lipid can have the formula II




wherein R3 is trans or cis to R8, and R7 is trans or cis to R10,

wherein the nitrated lipid is substantially pure. In this aspect, the stereochemistry about the carbon-carbon double bond is substantially cis (or Z) or substantially trans (or E). In one aspect, when the nitrated lipid has the formula II, R1 comprises a C4-C10 alkyl group, R2, R3, R8, and R10 are hydrogen, R7 is NO2, and R9 comprises a C6-C12 alkyl group. This class of nitrated lipids is depicted below, where the nitrated lipid has one vinyl nitro group.





In a further aspect, when the nitrated lipid has the formula II, R3 and R8 are cis (Z) to one another, and R7 and R10 are cis (Z) to one another. In another aspect, the nitrated lipid is 10-nitro-9-cis,12-cis-octadecadienoic acid, which is depicted in FIG. 4C.
In another aspect, when the nitrated lipid has the formula II, R1 comprises a C4-C10 alkyl group, R2, R3, R7, and R10 are hydrogen, R8 is NO2, and R9 comprises a C6-C12 alkyl group. This class of nitrated lipids is depicted below, where the nitrated lipid has one vinyl nitro group.




In a further aspect, R3 and R8 are cis (Z) to one another and R7 and R10 are cis (Z) to one another. In another aspect, the nitrated lipid is 12-nitro-9-cis,12-cis-octadecadienoic acid, which is depicted in FIG. 4C.
In another aspect, the nitrated lipid has the formula III




wherein

    
    
        R1 comprises C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl;
        R2 and R12 comprise, independently, hydrogen, NO2, OH, or OOH; and
        R13 comprises C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl, wherein R13 comprises a terminal COOR6 group, wherein R6 comprises hydrogen, C1-C24 alkyl, or a pharmaceutically acceptable counterion, wherein R13 optionally comprises one or more NO2, OH, or OOH groups;

wherein the compound comprises at least one NO2 group,

wherein the nitrated lipid is substantially pure.

    
    


In another aspect, when the nitrated lipid has the formula III, R1 comprises a C4-C10 alkyl group, R2 is hydrogen, R12 is NO2, and R13 comprises a C6-C12 alkyl group. This class of nitrated lipids is depicted below, where the nitrated lipid has one allylic nitro group.




In this aspect, R1 comprises a C4-C10 alkyl group, R2 is NO2, R12 is hydrogen, and R13 comprises a C6-C12 alkyl group. In one aspect, the nitrated lipid is 9-nitro-10,12-cis-octadecadienoic acid; 9-nitro-10,12-trans-octadecadienoic acid; 13-nitro-10,12-cis-octadecadienoic acid; or 13-nitro-10,12-trans-octadecadienoic acid.

Methods for preparing the nitrated lipids described herein are described below and in the Examples section.
II. Synthesis of Nitrated Lipids
Described herein are methods for preparing nitrated lipids. In one aspect, the method comprises
(a) reacting an unsaturated lipid with a mercuric salt, a selenium compound, and a nitrating compound to produce a first intermediate, and
(b) reacting the first intermediate with an oxidant.
Any of the lipids described above can be used to produce the nitrated lipids described herein. In one aspect, any unsaturated lipid having at least one carbon-carbon double bond can be used in this aspect to produce nitrated lipids.
In one aspect, step (a) can be performed in situ without isolation of the first intermediate. In another embodiment, the first intermediate can be trapped prior to step (b). The mercuric salt, a selenium compound, and a nitrating compound can be added in any order to the unsaturated lipid.
The selenium compound is any compound that is capable of reacting or interacting with the unsaturated group present in the lipid. In one aspect, when the unsaturated lipid possesses a carbon-carbon double bond, the selenium compound can form a three-membered ring intermediate, which is depicted, for example, in FIG. 1. In one aspect, the selenium compound possesses a leaving group capable of reacting with the three ring intermediate. Examples of leaving groups include, but are not limited to, halides (e.g., F, Cl, Br, I) and esters (e.g., acetate). Not wishing to be bound by theory, it is believed that the leaving group reacts with the three-membered ring intermediate to open the ring to produce a selenium-leaving group intermediate. One example of this intermediate is depicted in FIG. 1, where Br is the leaving group. Examples of selenium compounds useful herein include, but are not limited to, PhSeBr, PhSeCl, PhSeO2CCF3, PhSeO2H, or PhSeCN.
The mercuric salt used in the methods described herein can be any mercuric salt known in the art. Not wishing to be bound by theory, it is believed that the mercuric salt facilitates the formation of the selenium three-membered ring intermediate. In one aspect, the mercuric salt comprises HgCl2, Hg(NO3)2, or Hg(OAc)2.
The nitrating compound is any compound that provides a source of NO2− ions in solution. Not wishing to be bound by theory, it is believed that NO2− reacts with the selenium intermediate formed upon the reaction between the unsaturated lipid and selenium compound by displacing the leaving group present in the intermediate. FIG. 1 depicts one aspect of this, where NO2− displaces Br− by nucleophilic attack to produce a selenium/nitro intermediate. In one aspect, the nitrating compound can be any nitrite salt. In another aspect, the nitrating compound comprises NaNO2 or AgNO2.
The relative amounts of unsaturated lipid, selenium compound, mercuric salt, and nitrating compound can vary depending upon the specific reagents that are selected and reaction conditions. In one aspect, an equimolar amount or slight excess thereof of selenium compound, mercuric salt, and nitrating compound relative to the unsaturated lipid can be used. Step (a) is generally performed in a solvent, such as, for example, a polar or unpolar organic solvent. Examples of solvents useful herein include, but are not limited to, nitriles, ethers, esters, alkanes, alcohols, or combinations thereof. In one aspect, the solvent used in step (a) can be THF/acetonitrile. Step (a) can be performed at various temperatures depending upon the starting materials that are selected. In one aspect, the step (a) can be performed at room temperature.
By varying the reaction conditions in step (a), it is possible to increase the overall yield of the nitrated lipids as well as reduce the number of different types of nitrated lipids. In one aspect, the step (a) can be performed under anaerobic conditions. In another aspect, step (a) can be performed under anhydrous conditions. In a further aspect, step (a) is performed under anaerobic and anhydrous conditions.
After step (a), the intermediate that is produced is reacted with an oxidant to convert the intermediate to the nitrated lipid (step (b)). Not wishing to be bound by theory, the oxidant oxidizes the selenium compound to produce a selenium-oxo group, which rearranges to produce the nitrated lipid. One aspect of this mechanism is depicted in FIG. 1, whereupon oxidation, the selenium-oxo group rearranges to produce PhSeOH and a nitrated lipid possessing a vinyl nitro group. In one aspect, the oxidant comprises H2O2 or an organic hydroperoxide. The amount of oxidant will vary depending upon the selection of the oxidant and reaction conditions. In one aspect, a 2-fold, 3-fold, 5-fold, 7-fold, 9-fold, 10-fold, 15-fold, or 20-fold molar amount of oxidant is used compared to the molar amount of unsaturated lipid. In one aspect, step (b) is performed in an organic solvent at reduced temperature.
In one aspect, the unsaturated lipid comprises oleic acid, linoleic acid, linolenic acid, arachidonic acid, eicosapentaenoic acid, or docosahexanoic acid, the mercuric salt comprises HgCl2, the selenium compound comprises PhSeBr, the nitrating compound comprises NaNO2, and the oxidant comprises H2O2. In a further aspect, step (a) is performed under anaerobic and anhydrous conditions.
After step (b), one nitrated lipid or mixture of two or more nitrated lipid positional or stereoisomers may be present depending upon reagents and starting materials that are selected. In one aspect, if two or more nitrated lipids are present, each of the nitrated lipids can be separated to produce substantially pure nitrated lipid. In one aspect, the mixture of two or more nitrated lipids can be separated by chromatography. For example liquid chromatography, thin layer chromatography or column chromatography using silicic acid, silical gel or other adsorbents useful for lipid separations, can be used to separate the nitrated lipids. The solvent system used in this aspect will vary depending upon the type and number of nitrated lipids to be separated and can be determined by one of ordinary skill in the art.
Also described herein are methods for stabilizing nitrated lipids, comprising placing the nitrated lipid in a hydrophobic medium. The nitrated lipids are generally stable oils and can be stored indefinitely in organic solvents under anaerobic and anhydrous conditions and reduced temperature. Alternatively, the salts of the nitrated lipids are stable and can be further processed into a formulation. Alternatively, the more stable nitrohydroxy derivative can be formed under alkaline conditions, which at more neutral pH reversibly yields the parent nitrated fatty acid. Alternatively, the nitrated lipids can be placed in detergent emulsions or liposomes, which can be later administered to a subject.
III. Nitro/Hydroxy Lipids and Synthesis Thereof
In one aspect, described herein are lipids comprising at least one nitro group and at least one hydroxyl group, wherein the compound is substantially pure. In one aspect, the nitro group and the hydroxyl group are on adjacent carbon atoms. In this aspect, the lipid contains the fragment HO—C—C—NO2. In another aspect, the nitro/hydroxy lipid has the formula X or XI




wherein R1 comprises a C1-C24 alkyl group, a C1-C24 alkenyl group, or C1-C24 alkynyl group, and R9 comprises a terminal COOR6 group, wherein R6 comprises hydrogen, C1-C24 alkyl, or a pharmaceutically acceptable counterion, and p is from 1 to 12. In a further aspect, the nitro/hydroxy lipid has the formula XII or XIII





wherein R1 comprises a C1-C24 alkyl group, a C1-C24 alkenyl group, or C1-C24 alkynyl group, and R9 comprises a terminal COOR6 group, wherein R6 comprises hydrogen, C1-C24 alkyl, or a pharmaceutically acceptable counterion, and m is from 0 to 12.

Any of the nitrated lipids can be converted to the corresponding nitro/hydroxyl compound. In one aspect, the nitrated lipid is placed in an aqueous base. Not wishing to be bound by theory, it is believed that the nitrated lipid undergoes a Michael addition reaction with water followed by deprotonation to produce the nitro/hydroxyl lipid. This mechanism is depicted below.




The nitro/hydroxyl lipid can be subsequently isolated by solvent extraction followed by purification using techniques known in the art (e.g., HPLC or thin layer chromatography).

IV. Methods of Use

Delivery
As used throughout, administration of any of the nitrated lipids described herein can occur in conjunction with other therapeutic agents. Thus, the nitrated lipids can be administered alone or in combination with one or more therapeutic agents. For example, a subject can be treated with a nitrated lipid alone, or in combination with chemotherapeutic agents, antibodies, antivirals, steroidal and non-steroidal anti-inflammatories, conventional immunotherapeutic agents, cytokines, chemokines, and/or growth factors. Combinations may be administered either concomitantly (e.g., as an admixture), separately but simultaneously (e.g., via separate intravenous lines into the same subject), or sequentially (e.g., one of the compounds or agents is given first followed by the second). Thus, the term “combination” or “combined” is used to refer to either concomitant, simultaneous, or sequential administration of two or more agents. Furthermore, two or more nitrated lipids described herein can be administered to a subject concomitantly, simultaneously, or sequentially.
The nitrated lipids can be administered in a number of ways depending on whether local or systemic treatment is desired, and on the area to be treated. Administration may be topically (including opthamalically, vaginally, rectally, intranasally), orally, by inhalation, or parenterally, for example by intravenous drip, subcutaneous, intraperitoneal or intramuscular injection. The disclosed compounds can be administered intravenously, intraperitoneally, intramuscularly, subcutaneously, intracavity, transdermally, intratracheally, extracorporeally, or topically (e.g., topical intranasal administration or administration by inhalant). As used herein, “topical intranasal administration” means delivery of the compositions into the nose and nasal passages through one or both of the nares and can comprise delivery by a spraying mechanism or droplet mechanism, or through aerosolization of the nucleic acid or vector. The latter can be effective when a large number of subjects are to be treated simultaneously. Administration of the compositions by inhalant can be through the nose or mouth via delivery by a spraying or droplet mechanism. Delivery can also be directly to any area of the respiratory system (e.g., lungs) via intubation.
Parenteral administration of the composition, if used, is generally characterized by injection. Injectables can be prepared in conventional forms, either as liquid solutions or suspensions, solid forms suitable for solution of suspension in liquid prior to injection, or as emulsions. A more recently revised approach for parenteral administration involves use of a slow release or sustained release system such that a constant dosage is maintained. See, e.g., U.S. Pat. No. 3,610,795, which is incorporated by reference herein in its entirety for the methods taught.
The compositions may be in solution or in suspension (for example, incorporated into microparticles, liposomes, or cells). These compositions may be targeted to a particular cell type via antibodies, receptors, or receptor ligands. The following references are examples of the use of this technology to target specific proteins to given tissue (Senter, et al., Bioconjugate Chem., 2:447-451, (1991); Bagshawe, K. D., Br. J. Cancer, 60:275-281, (1989); Bagshawe, et al., Br. J. Cancer, 58:700-703, (1988); Senter, et al., Bioconjugate Chem., 4:3-9, (1993); Battelli, et al., Cancer Immunol. Immunother., 35:421-425, (1992); Pietersz and McKenzie, Immunolog. Reviews, 129:57-80, (1992); and Roffler, et al., Biochem. Pharmacol, 42:2062-2065, (1991)). Vehicles such as “stealth” and other antibody conjugated liposomes (including lipid mediated drug targeting to colonic carcinoma), receptor mediated targeting of DNA through cell specific ligands, lymphocyte directed tumor targeting, and highly specific therapeutic retroviral targeting of murine glioma cells in vivo. In general, receptors are involved in pathways of endocytosis, either constitutive or ligand induced. These receptors cluster in clathrin-coated pits, enter the cell via clathrin-coated vesicles, pass through an acidified endosome in which the receptors are sorted, and then either recycle to the cell surface, become stored intracellularly, or are degraded in lysosomes. The internalization pathways serve a variety of functions, such as nutrient uptake, removal of activated proteins, clearance of macromolecules, opportunistic entry of viruses and toxins, dissociation and degradation of ligand, and receptor-level regulation. Many receptors follow more than one intracellular pathway, depending on the cell type, receptor concentration, type of ligand, ligand valency, and ligand concentration. Molecular and cellular mechanisms of receptor-mediated endocytosis has been reviewed (Brown and Greene, DNA and Cell Biology 10:6, 399-409 (1991)).
The exact amount of the compositions required will vary from subject to subject, depending on the species, age, weight and general condition of the subject, the severity of the disorder being treated, the particular nucleic acid to be targeted, its mode of administration and the like. Thus, it is not possible to specify an exact amount for every composition. However, an appropriate amount can be determined by one of ordinary skill in the art using only routine experimentation given the teachings herein. In one aspect, the amount of nitrated lipid that is administered can be from 1 nM to 1 mM, 10 nM to 1 mM, 20 nM to 1 mM, 50 nM to 1 mM, 100 nM to 1 mM, 200 nM to 1 mM, 300 nM to 1 mM, or 500 nM to 1 mM. The time at which the nitrated lipids can be administered will also vary depending upon the subject, the disorder, mode of administration, etc. The nitrated lipid can be administered to the subject prior to the onset of inflammation or during a time when the subject is experiencing inflammation. In one aspect, the nitrated lipid can be administered within 24 hours, 20 hours, 16 hours, 12 hours, 8 hours, 4 hours, 2 hours, 1 hour, or 30 minutes before inflammation occurs or 10 hours, 20 hours, 30 hours, 40 hours, 60 hours, 80 hours, 100 hours, or 120 hours after the onset of the inflammation.
Pharmaceutically Acceptable Carriers
The nitrated lipids can be used therapeutically in combination with a pharmaceutically acceptable carrier. Pharmaceutical carriers are known to those skilled in the art. These most typically would be standard carriers for administration of drugs to humans, including solutions such as sterile water, saline, and buffered solutions at physiological pH. The compositions can be administered intramuscularly or subcutaneously. Other compounds will be administered according to standard procedures used by those skilled in the art.
Pharmaceutical compositions may include carriers, thickeners, diluents, solvents, buffers, preservatives, surface active agents and the like in addition to the molecule of choice. Pharmaceutical compositions may also include one or more active ingredients such as antimicrobial agents, anti-inflammatory agents, anesthetics, and the like.
Preparations for parenteral administration include sterile aqueous or non-aqueous solutions, suspensions, and emulsions. Examples of non-aqueous solvents are propylene glycol, polyethylene glycol, vegetable oils such as olive oil, and injectable organic esters such as ethyl oleate. Aqueous carriers include water, alcoholic/aqueous solutions, emulsions or suspensions, including saline and buffered media. Parenteral vehicles include sodium chloride solution, Ringer's dextrose, dextrose and sodium chloride, lactated Ringer's, or fixed oils. Intravenous vehicles include fluid and nutrient replenishers, electrolyte replenishers (such as those based on Ringer's dextrose), and the like. Preservatives and other additives may also be present such as, for example, antimicrobials, anti-oxidants, chelating agents, and inert gases and the like.
Formulations for topical administration may include ointments, lotions, creams, gels, drops, suppositories, sprays, liquids and powders. Conventional pharmaceutical carriers, aqueous, powder or oily bases, thickeners and the like may be necessary or desirable.
Compositions for oral administration include powders or granules, suspensions or solutions in water or non-aqueous media, capsules, sachets, or tablets. Thickeners, flavorings, diluents, emulsifiers, dispersing aids or binders may be desirable.
Some of the nitrated lipids may potentially be administered as a pharmaceutically acceptable acid- or base-addition salt, formed by reaction with inorganic acids such as hydrochloric acid, hydrobromic acid, perchloric acid, nitric acid, thiocyanic acid, sulfuric acid, and phosphoric acid, and organic acids such as formic acid, acetic acid, propionic acid, glycolic acid, lactic acid, pyruvic acid, oxalic acid, malonic acid, succinic acid, maleic acid, and fumaric acid, or by reaction with an inorganic base such as sodium hydroxide, ammonium hydroxide, potassium hydroxide, and organic bases such as mono-, di-, trialkyl and aryl amines and substituted ethanolamines. In another aspect, the nitrated lipid is in the form of the sodium or potassium salt. In another embodiment, the nitrated lipids can be converted to the corresponding pharmaceutically-acceptable ester such as, for example, the methyl ester.
Therapeutic Uses
The methods described herein contemplate the use of single or mixtures of two or more nitrated or nitro/hydroxyl lipids. In one aspect, disclosed are methods for reducing or preventing inflammation in a subject with inflammation or at risk for inflammation, comprising administering an effective amount of any of the nitrated lipids described herein, wherein the nitrated lipid reduces or prevents the inflammation in the subject. Examples of inflammation include, but are not limited to, pulmonary inflammation, vascular inflammation, renal inflammation, inflammation of the central nervous system, hepatic inflammation, or splanchnic inflammation. The inflammation can be associated with an inflammatory disease including, but not limited to, systemic lupus erythematosus, Hashimoto's disease, rheumatoid arthritis, graft-versus-host disease, Sjögren's syndrome, pernicious anemia, Addison disease, scleroderma, Goodpasture's syndrome, Crohn's disease, autoimmune hemolytic anemia, myasthenia gravis, multiple sclerosis, Alzheimer's disease, amyotrophic lateral sclerosis, Basedow's disease, thrombopenia purpura, insulin-dependent diabetes mellitus, allergy; asthma, inflammatory bowel disease, cancer, ulcerative colitis, scleroderma, cardiomyopathy, atherosclerosis, hypertension, sickle cell disease, or respiratory distress syndrome of neonate and adults. In another aspect, the inflammation can be caused by an organ transplantation, respiratory distress, ventilator induced lung injury, ischemia reperfusion, hemorrhagic shock, or sepsis. In one aspect, when the pulmonary inflammation is caused by respiratory distress or sepsis, the nitrated lipids can reduce or prevent the accumulation of alveolar fluid in a subject.
In one aspect, the nitrated lipids described herein can modulate the expression or activity of one or more nucleic acids that encode one or more inflammatory-related or cell signaling polypeptides. The term “modulate” is defined herein as the ability of the nitrated lipid to decrease or increase the expression or activity relative to a control. The “control” can be either the amount of expression or activity in the absence of a nitrated lipid, or in the presence of solvent or non-nitrated parent lipid used for the synthesis of the nitrated lipid derivative. Alternatively, the “control” can be the amount of expression or activity before or after the period use (i.e., administration of the nitrated lipid). The term “inflammatory-related polyepetide” is defined herein as any polypeptide that can induce or potentiate an inflammatory response. Examples of genes that can be modulated by the nitrated lipids described herein and inflammatory-related polypeptides include, but are not limited to, prostaglandin H synthase-2, gamma glutamyl cysteine synthase, low density lipoprotein receptor, vascular endothelial growth factor, tocopherol binding protein, a heat shock protein (e.g., 10, 40, 60, 70, 90 kD), prostaglandin receptor EP4, a protein tyrosine phosphatase, a Ca, Na, or K ATPase, a G-protein signaling regulator (e.g., 24 kD), a vasoactive intestinal peptide, a guanine nucleotide exchange factor, a bone morphogenetic protein, an aromatic-inducible cytochrome P450s, an ADP-ribosyltransferase, endothelin-1, a vascular cell adhesion molecule-1, an intercellular adhesion molecule-1, a tumor necrosis factor alpha receptor, an interleukin 1 beta receptor, a transforming growth factor beta receptor, an advanced glycation endproduct-specific receptor, a cAMP phosphodiesterase, a cGMP phosphodiesterase, a cyclin-dependent kinase, cathepsins B and D, a connective tissue growth factor, actin, myosin, a tubulin gene, a c-src tyrosine kinase, an insulin growth factor binding protein, a cysteine-rich angiogenic inducer 61, thrombospondin-1, a cadherin-associated protein beta, heme oxygenase-1, one or more of the genes listed in Table 5, one more of the genes listed in Table 6, one or more of the genes listed in Table 7, or a combination thereof. In one aspect, the nitrated lipid can modulate the nucleic acid 1.5 fold, 2 fold, 3-fold, 5-fold, 10-fold, 20-fold, 30-fold, 40-fold, 50-fold, 75-fold, 100-fold, 125-fold, 150-fold, or greater.
The nitrated lipids described herein can be used to mediate receptors in a cell in order to reduce or prevent inflammation in a subject. In one aspect, described herein are methods for inducing or potentiating peroxisome proliferator activated receptor (PPAR) activity, comprising contacting a cell comprising at least one PPAR receptor with one or more nitrated lipids under conditions that allow the compound to induce or potentiate the activity of the PPAR receptor. The PPAR receptor can be α, δ, or γ. Alternatively, other lipid receptors that nitrated lipids can bind to, be transported by, and mediate include the family of fatty acid binding proteins, G protein-coupled receptors and cis-retinoic acid binding protein. Not wishing to be bound by theory, it is believed that activation of cell receptors such as, for example, PPAR, can modulate the tissue expression and activity of inflammatory-related genes. A number of cell-types can be contacted with the nitrated lipids described herein in order to reduce or prevent inflammation in a subject. Examples of such cells include, but are not limited to the constituent cells of lung, airways, nasal passages, eyes, auditory system, liver, spleen, kidney, intestine, colon, genito-urinary tract, heart, brain, spinal cord, muscle, bone, connective tissue, blood and reticuloendothelial system and nervous tissue.
In another aspect, the nitrated lipids described herein can induce the expression or potentiate the activity of an inflammatory-related polypeptide when a cell comprising at least one nucleic acid that encodes the inflammatory-related polypeptide is contacted with a nitrated lipid under conditions that allow the compound to induce the expression or potentiate the activity of the inflammatory-related polypeptide. In one aspect, the following expressions or activities can be induced or potentiated with the nitrated lipids described herein:

    1. Modification of downstream signaling regulated by small G-proteins.
    2. Increased phosphorylation and activation of c-Jun N-terminal kinase.
    3. Increased the phosphorylation and activation of the transcription factor c-Jun.
    4. Modification of activator protein-1 binding and activator protein-1 mediated gene expression
    5. Increased phosphorylation and activation of extracellular-signal regulated kinase.
    6. Increased phosphorylation and activation of the transcription factor Elk-1.
    7. Modification of binding to serum response element (SRE) and SRE mediated gene expression.
    8. Increased synthesis of the transcription factor c-Fos.
    9. Affect the nuclear translocation of the transcription factor Nrf-2 (Nuclear factor erythroid 2 related factor 2).
    10. Modification of the electrophilic response element (also known as antioxidant response element) binding and the electrophilic response element mediated gene expression.
    11. Modification of p38 mitogen activated protein kinase activation.
    12. Modification of the nuclear translocation of the transcription factor p65.
    13. Modification of nuclear factor-kappa B binding and nuclear factor kappa B mediated
    gene expression.


In other aspects, any of the nitrated lipids described herein can reduce a cell's response to an inflammatory stimulus. In one aspect, the cell can be a neutrophil, monocyte, or macrophage. For example, the nitrated lipids can inhibit neutrophil, monocyte, or macrophage degranulation (e.g., azurophilic) or release of hydrolases and proteases following degranulation, neutrophil, monocyte, or macrophage O2.− formation, expression of CD11b expression in a neutrophil, monocyte, or macrophage, and fMLP-induced Ca+2 influx in a neutrophil, monocyte, or macrophage upon contact of the neutrophil, monocyte, or macrophage with the nitrated lipid. In these aspects, the neutrophil, monocyte, or macrophage can be contacted with the nitrated lipid in vivo, in vitro, or ex vivo.
In one aspect, described herein are methods for regulating the activity of a protein kinase signaling pathway, comprising reacting a kinase with a nitro lipid described herein. In another aspect, the nitrated lipids described herein can regulate the activity of a thiol-dependent enzyme in a cell by contacting the cell with a nitrated lipid of the present invention. The cell can be contacted with the nitrated lipid, in vivo, in vitro, or ex vivo. For example, nitroalkenes inhibit enzymes that depend on thiols as catalytic residues. Not wishing to be bound by theory, it is believed that the thiol can react with the nitrated lipid via a Michael addition reaction. Via this property, nitroalkenes also serve a potent stimuli of thiol modification-dependent protein kinases and their downstream cell signaling pathways. In this regard, a variety of cell protein kinases are activated by nitroalkenes and the associated thiol-dependent phosphoprotein phosphatases can also be inhibited by thiol alkylation. In one aspect, any thiol-dependent structural, cell signaling and catalytic protein can be regulated by the nitrated fatty acid compounds described herein.
In another aspect, the nitrated lipids described herein can control protein trafficking in cells by serving as a thiol-attached hydrophobic membrane trafficking agent for proteins to which they attach. Thus, the alkylation of proteins by membrane-avid nitrated fatty acids will facilitate the localization of proteins containing hydrophobic nitrated fatty acid-amino acid adducts to cytosol, plasma membrane and organelle (nucleus, endoplasmaic reticulum, mitochondrial, golgi, secretory vesicles) membranes.
In another aspect, the nitrated lipids described herein can inhibit platelet function in a subject upon administration of the nitrated lipid to the subject. In one aspect, the nitrated lipids can inhibit thrombin- or other stimuli-induced platelet aggregation by attenuating cAMP-dependent Ca+2 mobilization and activation of the phosphorylation of vasodilator-stimulated phosphoprotein (VASP).
In another aspect, the nitrated lipids described herein can be used to induce or potentiate tissue repair in a subject suffering from inflammation. Not wishing to be bound by theory, it is believed that the nitrated lipids can down regulate events that result in inflammation or the impairment of vascular function and blood flow.
In another aspect, the nitrated lipids described herein can promote satiety in a subject upon administration of the nitrated lipid to the subject.
In another aspect, the nitrated lipids described herein can treat cancer upon administering an effective amount of the nitrated lipid to the subject. Not wishing to be bound by theory, it is believed that the nitrated lipids directly stimulate tumor cell killing and potentiate the killing of tumor cells by standard chemotherapeutic drugs. The cell necrosis and apoptosis-inducing activity is believed to be the result of nitrated lipid/PPAR ligand activity (e.g., PPAR activation), as well as by stimulating other cell signaling pathways noted above that mediate cell growth, cell differentiation and death signaling pathways.
In another aspect, the nitrated lipids set forth herein can act as nitric oxide (NO) donors, as described in the Examples. Therefore, the nitrated lipids described herein can be used to administer NO to a subject and/or treat a NO-related condition in a subject. These conditions include, but are not limited to, atherosclerosis, myocardial infarction, peripheral vascular disease, coronary artery diseases, heart failure, stroke, essential hypertension, diabetes mellitus, pre-eclampsia, erectile dysfunction, impotence, diabetic nephropathy, inflammatory glomerular diseases, acute renal failure, chronic renal failure, inflammation, bacterial infection, septic shock, respiratory distress syndromes, arthritis, cancer, impetigo, epidermolysis bullosa, eczema, neurodermatitis, psoriasis, pruritis, erythema, hidradenitis suppurativa warts, diaper rash and jock itch.
In another aspect, described herein are methods for detecting inflammation in a subject, comprising (a) measuring the amount of a nitrated lipid present in the subject and (b) comparing the amount of nitrated lipid in the subject to the amount of nitrated lipid present in a subject that is not experiencing any inflammation. In one aspect, patients with cardiovascular disease have increased amounts of nitrated fatty acid. Based on the presence of nitrated lipids, the detection of increased levels of nitrated fatty acids in blood, tissues and bodily fluids can serve to diagnose the occurrence, progression and/or resolution of the inflammatory process.
EXAMPLES
The following examples are put forth so as to provide those of ordinary skill in the art with a complete disclosure and description of how the compounds, compositions, and methods described and claimed herein are made and evaluated, and are intended to be purely exemplary and are not intended to limit the scope of what the inventors regard as their invention. Efforts have been made to ensure accuracy with respect to numbers (e.g., amounts, temperature, etc.) but some errors and deviations should be accounted for. Unless indicated otherwise, parts are parts by weight, temperature is in ° C. or is at ambient temperature, and pressure is at or near atmospheric. There are numerous variations and combinations of reaction conditions, e.g., component concentrations, desired solvents, solvent mixtures, temperatures, pressures and other reaction ranges and conditions that can be used to optimize the product purity and yield obtained from the described process. Only reasonable and routine experimentation will be required to optimize such process conditions.
Example 1
Materials
Linoleic acid was purchased from Nu-Check Prep (Elysian, Minn.). Phenylselenium bromide, HgCl2, NaNO2, anhydrous tetrahydrofuran, N,N-diisopropylethylamine (99.5%) and acetonitrile were obtained from Sigma/Aldrich (St Louis, Mo.). Silica gel HF thin layer chromatography (TLC) plates (250 μm) were from Analtech (Newark, Del.). Pentafluorobenzyl bromide and methanolic BF3 was from Pierce (Rockford, Ill.). Solvents used in synthesis were HPLC grade or better and were purchased from Fisher Scientific (Fairlawn, N.J.). Solvents used for mass spectrometric analyses from Burdick and Jackson (Muskigon, Mich.). [13C]Linoleic acid was from Spectra Stable Isotopes (Columbia, Md.) and [15N]NaNO2 was from Cambridge Isotope Laboratories, Inc. (Andover, Mass.). [14N]LNO2, [13C]LNO2 and [15]LNO2 positional isomers were synthesized as described previously for nitrated fatty acids.
LNO2 Synthesis.
Linoleic acid/HgCl2/phenylselenium bromide/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 linoleic acid. Care was taken to use anhydrous solvents, dry glassware and reagents that had been dried in vacuo over phosphorus pentoxide. 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 then allowed to rest in an ice bath for 20 min followed by a gradual warming to room temp (45 min). The product mixture was extracted with equal parts saturated NaCl and diethyl ether, the organic phase collected, the solvent removed in vacuo and the lipid products were resolvated in CH2Cl2/CH3OH (4:1, v/v). A mixture of LNO2 positional isomers were initially separated from the product mixture 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). Regions of silica containing LNO2 were scraped, extracted (29) and stored in CH3OH under argon at −80° C. Under these conditions, purified nitrated linoleic acid is stable for >3 months. Large scale purification of the individual positional isomers was performed by preparative HPLC using a 250×21.2 mm C18 Phenomenex Luna column (5 μm particle size). Lipids were eluted from the column using a gradient solvent system consisting of A (H2O containing 0.1% NH4OH) and B (CNCH3 containing 0.1% H2O) under the following conditions: 20-80% B (linear increase, 45 min), 80% B (2 min), 20% B (5 min). Fractions were collected as positional isomers eluted, the solvent removed in vacuo and the lipids were stored in CH3OH under argon at −80° C. Stable isotopes of LNO2, specifically [13C]LNO2 and [15]LNO2, were synthesized as above, except that [13C]linoleic acid or [15N]NaNO2 were substituted in the synthetic scheme. FIG. 1 depicts a synthetic scheme for producing nitrated lipids. The process described above significantly increased the purity and yield of fatty acid allylic nitration products, facilitating structural resolution of specific LNO2 positional isomers and future cell signaling studies (22, 32, 33). Preparative TLC permitted the initial resolution of nitrated fatty acids from starting materials and oxidized linoleic acid species (FIG. 2). The principal C-10 and C-12 positional isomers have Rf values of 0.45 and 0.50, respectively. The changes in synthetic approaches, use of anhydrous reagents and the execution of nitrosenylation reaction steps under a nitrogen atmosphere resulted in an increase in yield of nitrated linoleic acid products from 4% to 56%.
In an alternate aspect, nitrated fatty acids can be separated and purified from oils such as, for example, fish oil, soy bean oil, and olive oil. For example, silica gel and silicic acid columns can be used to fractionate the nitro-fatty acids, followed by preparative TLC or HPLC to separate the different nitrated fatty acids. Nitro-oleate is the predominant (>90%) marine and plant-nitroalkene.
Spectral Analysis of LNO2.
Initial concentrations of synthetic LNO2 preparations were measured by chemiluminescent nitrogen analysis (Antek Instruments, Houston, Tex.) using caffeine as a standard. This data was utilized to determine dilution concentrations for subsequent spectral analysis. The extinction coefficients (ε) for LNO2 and the isotopic derivatives [13C]LNO2 and [15]LNO2 were measured using a UV-VIS spectrophotometer (Shimadzu, Japan) set to measure absorbance at 329 nm, the absorbance maximum specific to LNO2 as compared to linoleic acid. Absorbance values for increasing concentrations of LNO2, [15]LNO2 or [13C]LNO2 in MeOH containing 20 mM NaOH were plotted against concentration to calculate slope/extinction coefficient.
Nitrated linoleic acid displays a characteristic absorption profile and maximum, permitting determination of an extinction coefficient and providing a facile method for measuring concentrations of synthetic LNO2. This species displays a unique absorbance maximum at 329 nm, compared with linoleic acid (FIG. 3A). Plotting absorbance versus concentration profiles for each of the synthetic LNO2 preparations reported herein—[14N]LNO2, [15]LNO2 and [13C]LNO2—generated identical extinction coefficients for all LNO2 derivatives: ε=10.1 cm−1 M−1 (FIG. 3B).
Red Blood Cell and Plasma Lipid Isolation and Extraction.
Peripheral blood from healthy human volunteers was collected by venipuncture in heparinized tubes, centrifuged (1200×g; 10 min) and plasma isolated from red cell pellets from which the buffy coat was removed. Crude lipid extracts were prepared from packed red cells and plasma by the method of Bligh and Dyer (29) and analyzed by mass spectrometry. Care was taken to avoid acidification during all steps of plasma fractionation and lipid extraction to prevent artifactual lipid nitration due to the presence of endogenous NO2−. Extracts from red cells were analyzed by mass spectrometry; however, lipid extracts from plasma were first fractionated by TLC to separate LNO2 from the bulk of neutral lipids present in plasma, minimizing ionization dampening during LNO2 analysis by mass spectrometry. TLC plates were developed twice in a solvent system consisting of hexane:ether:acetic acid (70:30:1, v/v), and regions of silica containing LNO2, identified by comparing to the migration of synthetic standards, were scraped and extracted (29). To measure the esterified LNO2 content in red cell membranes and plasma lipoproteins, lipid extracts were first hydrolyzed (30), fractionated by TLC and analyzed by mass spectrometry.
In FIG. 6A, [13C]LNO2 was added to lipid extractions as an internal standard to quantitate the free and esterified LNO2 content of red cells and plasma obtained from healthy human volunteers. The mean age of the 5 female and 5 male subjects was 34 yr. The endogenous LNO2 isomers co-eluted with the added 13C-labeled LNO2 internal standard, with the internal standard differentiated from endogenous LNO2 by monitoring its unique m/z 342/295 MRM transition. In FIG. 6B, the internal standard curve for LNO2 reveals linear detector responses over five orders of magnitude. The limit of quantitation (LOQ) for LNO2, as defined by ten times the standard deviation of the noise, is approximately 0.3 fmol ( ̃100 fg) injected on column.
Analysis of Synthetic LNO2 Methyl and Pentafluorobenzyl Esters by Gas Chromatography Mass Spectrometry.
Methyl ester-derivatives of synthetic LNO2 isomers were analyzed by gas chromatography mass spectrometry (GC-MS) in both positive and negative ion modes. Electron-impact (EI) ionization was used to identify and characterize the fragmentation pattern of the two main positional isomers of LNO2 methyl esters. Methyl esters were prepared by drying LNO2 under a stream of nitrogen and redissolving in methanolic BF3 (14% BF3, 86% CH3OH); with the methylation reaction proceeding for 8 min at 60° C. Methyl esters were then extracted with hexane, washed twice with saturated saline, redissolved in undecane and analyzed by electron-impact GC-MS. EI GC-MS was performed using a Saturn 2000 mass spectrometer coupled with a Varian 3800 gas chromatograph. Samples were ionized by electron impact at +70 eV. Methyl ester-derivatized LNO2 isomers were resolved by GC using a CP-7420 capillary column (0.25 mm ID, 100 m fused silica, Varian, Palo Alto, Calif.) with the following temperature gradient: 60° C. (2 min); 60 to 120° C. at 20° C./min, held for 2 min; 120 to 270° C. at 20° C./min, held for 20 min. Helium was used as the carrier gas.
Due to the low sensitivity of positive ion GC-MS to LNO2, negative ion chemical ionization (NICI) was used to characterize pentafluorobenzyl (PFB) esters of synthetic LNO2 and detect LNO2 species in vivo. PFB esters of synthetic LNO2, red cell and plasma lipids were prepared (29), with biological lipids first partially purified by TLC as previously noted. Lipids were then subjected to PFB esterification and analysis by negative ion chemical ionization GC-MS using a Hewlett Packard 5890 GC (Palo Alto, Calif.) coupled to a single quadrupole Hewlett Packard MS using a 30 m CP-Sil 8CB-MS column (5% phenyl, 95% dimethylpolysiloxane; Varian, Palo Alto, Calif.) (31). The following temperature gradient was used to resolve the positional isomers of LNO2: 165 to 190° C. at 10° C./min; 190 to 270° C. at 2° C./min; and 270 to 290° C. at 5° C./min. Helium and methane were used as carrier and reagent gases, respectively. The detection of LNO2 was conducted by total ion count monitoring of [M−PFB]−, i.e., m/z 324 ([12C14N]LNO2), 325 ([12C, 15N]) and 342 ([13C, 14N]LNO2).
Initial characterization of positional isomers of synthetic LNO2 was via EI GC-MS on LNO2 methyl esters. Total ion count monitoring of the derivatized parent ion LNO2 revealed 2 dominant peaks at 38.75 and 39 min when resolved using a 100 m column (FIG. 4A). Product ion analysis of each peak (FIG. 4C) generated fragmentation patterns similar to previously reported for acidic nitration of ethyl linoleate (18). The first and second peaks correspond to linoleic acid that is nitrated on the 12- and the 10-carbon, respectively (referred to as C12 and C10 isomers). These two isomers are identified by the unique daughter ions m/z 250 and m/z 282 (specific to C12), and m/z 196 (specific to C10). Analysis of [15]LNO2 revealed fragmentation patterns with these identifying ions shifted by m/z=+1, further affirming fragments containing the nitro group (FIG. 4C). As a control for potential BF3-dependent artifactual products, anhydrous methanolic sulfuric acid (1%) esterification was performed to further verify the formation of the same respective LNO2 methyl ester positional isomers.
Structural analysis of synthetic LNO2 isomers by EI GC MS yielded important isomeric structural information and identification; but lacks sensitivity for the detection, structural characterization and quantification of LNO2 derivatives present in biological samples. Thus, these analyses were performed by NICI GC-MS on LNO2 species that had been derivatized to PFB esters (FIG. 4B). Chromatographic separation of both synthetic LNO2 and TLC-separated red cell lipid extracts showed two dominant peaks, C12 and C10, and two minor peaks ascribed to C13 and C9 positional isomers of LNO2; with approximately 95% of total peak area accounted for by C12 and C10 isomers. To confirm that the identities of the LNO2—PFB derivatives were the same as those identified as LNO2-methyl esters, the LNO2—PFB derivatives were run on EI positive ionization mode and identified by formation of characteristic fragmentation pattern (data not shown). Initial detection and quantitation of endogenous levels of LNO2 was performed using NICI GC-MS. [13C]LNO2 was added during the monophase of lipid extractions as an internal standard to correct for losses due to TLC and derivatization. The requirement for TLC separation to enrich fractions containing LNO2, the additional requirement for derivatization and on-column thermal decomposition creating limitations for analyzing nitrohydroxy- and nitrohydroperoxy fatty acids all combined to yield inconsistent quantitation of LNO2 levels in biological samples upon analysis by GC-MS. Consequently, we developed a LC-MS/MS based method to more reliably characterize and quantitate LNO2 in biological samples.
Analysis of LNO2 Positional Isomers by Electrospray Ionization Triple Quadrupole Mass Spectrometry.
Qualitative analysis of nitrated linoleic acid positional isomers by electrospray ionization mass spectrometry was performed using an Applied Biosystems/MDS Sciex 4000 Q TrapTM, a hybrid triple quadrupole-linear ion trap mass spectrometer. To separate and characterize the two major LNO2 positional isomers, synthetic standards and lipid extracts from biological samples were resolved by reverse-phase high performance liquid chromatography (HPLC) using a 150×2 mm C18 Phenomenex Luna column (3 μm particle size). Lipids were eluted using a gradient solvent system consisting of A (H2O containing 0.1% NH4OH) and B (CNCH3 containing 0.1% H2O) under the following conditions: 20-70% B (linear increase, 20 min), 70-95% B (2 min), 95% B (8 min), 95-20% B (1 min) and 20% B (5 min). Using these gradient conditions, two major and two minor LNO2 positional isomers were separated with baseline resolution. The resolved positional isomers of LNO2 were detected by mass spectrometry using a multiple reaction monitoring (MRM) scan mode by reporting molecules that undergo an m/z 324/277 mass transition. This transition, consistent with the loss of HNO2 ([M−(HNO2)—H]−), is common for all mono-nitrated positional isomers of linoleic acid. Concurrent with MRM, enhanced product ion analysis (EPI) was performed to generate characteristic and identifying fragmentation patterns of the eluting species with a precursor mass of m/z 324. In some analyses, specific fragments appearing in the EPI spectra were further fragmented in the ion trap to generate a MS3 spectrum to verify structural elucidation.
The most sensitive and selective technique available for analyte quantitation is triple quadrupole mass spectrometry detecting in the MRM. An HPLC separation strategy that baseline-resolved individual LNO2 positional isomers permitted the MS-based quantitation of LNO2 positional isomers, following chromatography of red cell and plasma lipid extracts. By monitoring the m/z 324/277 mass transition of the linoleate nitro derivative and the fatty acid parent molecule, respectively (FIG. 5A-1). To initially identify individual LNO2 positional isomers resolved by HPLC separation, the two major species were collected (peaks 1 and 2, FIG. 6B), derivatized to PFB-esters and analyzed by GC-MS. From GC-MS analysis, the first species eluting upon HPLC separation was identified as the C12 isomer and the second the C10 nitro derivative. To further characterize synthetic LNO2 isomeric structures, the linear ion trap mode of the hybrid mass spectrometer was used to perform enhanced product ion analysis on eluting peaks to generate a characteristic, identifying fragmentation pattern for each LNO2 positional isomer (FIGS. 5B-1, C-1). The ion trap enabled the concentration and thus detection of otherwise minor fragment ions, with MS/MS analysis revealing the unique fragments m/z 168 and 228 for the C10 isomer, as well as the fragments common to all LNO2 isomers (m/z 233, 244, 277, 293 and 306, FIG. 5C-1). The m/z 228 ion was further fragmented in the ion trap using the MS3 scanning mode, generating a fragment with m/z 46, indicating the loss of a nitro group and assigning the nitro group to the 9-10 double bond. Using [15]LNO2, the C10 isomer generated a fragment of m/z 229 that undergoes further fragmentation to m/z 47 (not shown). FIG. 5B-1 presents the characteristic fragmentation pattern obtained from MS/MS of the C12 isomer. The unique fragments to C12-LNO2, m/z 196 and 157, and the common fragments (m/z 233, 244, 277, 293 and 306) are present and consistent with a nitro group located at the 12-carbon of linoleic acid. In aggregate, from the synthetic rationale and GC MS and HPLC-ESI MS/MS data, the C10 and the C12 isomers are identified as 10-nitro-9-cis,12-cis-octadecadienoic acid and 12-nitro-9-cis,12-cis-octadecadienoic acid, respectively.
HPLC-ESI MS/MS was used to characterize and quantitate LNO2 species present in healthy human blood, specifically red cells and plasma (FIG. 5). The MRM elution profiles for red cell and plasma lipid extracts were identical to those obtained from the synthetic standards (FIGS. 5B-2, C-2 and 5B-3, C-3). All characteristic fragments found in the EPI spectra of the synthetic C10 and C12 LNO2 isomers were present in the resolved biological extracts, further affirming that red cells and plasma contain LNO2 positional isomers identical to synthetic standards.
Control Studies Relevant to Processing and Analysis-Induced Lipid Nitration.
In the presence of nitrite and acidic conditions, unsaturated fatty acids undergo nitration to products that reflect the chromatographic and MS characteristics of nitrosenylation-derived synthetic LNO2 isomers (18-20). Thus, acidic conditions were either avoided or documented for potential influence on product yield and nature. Importantly, lipid extractions were routinely conducted in the presence of pH 7.4 aqueous buffers. Comparison of LNO2 content in preparations extracted under neutral or acidic conditions revealed that there was no difference in LNO2 yield or LNO2 isomer distribution only in the complete absence of NO2−. Acidic lipid extraction conditions (pH<4.0) in the presence of exogenously-added biological concentrations of NO2− (10-500 μM) induced additional linoleate nitration, as indicated by nitration of 13C-linoleic acid added to pure linoleate or biological lipid extracts. When lipid extracts are separated by TLC prior to MS analysis, effective resolution of LNO2 from native and oxidized fatty acids requires the use of a 1% acetic acid-containing solvent system. It was observed that when NO2− was present in the TLC chromatographic solvent at concentrations not expected from biological analyses (>1 mM), LNO2 was formed de novo by acid-catalyzed nitration reactions. Since the initial solvent extraction of lipids from synthetic mixtures or biological materials resulted in removal of >95% of all adventitious NO2−, confidence exists that artifactual fatty acid nitration reactions were not occurring during acidic HPLC or TLC resolution. With regard to biological sample analysis, this latter precept was affirmed by a) adding 13C-labeled linoleic acid prior to red cell and plasma lipid extraction, purification and MS analysis and b) observing no formation of 13C-labeled nitrated linoleic acid derivatives.
Detection and Quantitation of LNO2 in Human Red Blood Cells.
Quantitation of LNO2 in biological samples was performed as above, with modification. During the monophase stage of lipid extractions (29), a known quantity of [13C]LNO2 was added as internal standard to correct for loss during extraction and TLC. The gradient elution profile was changed so that all LNO2 positional isomers co-eluted. Lipids were eluted from the same column used for structural characterization using a gradient solvent system consisting of A (H2O containing 0.1% NH4OH) and B (CNCH3 containing 0.1% H2O) under the following conditions: 80-90% B (2 min), 90% B (3 min), 90-80% B (1 min) and 80% B (2 min). Two MRM transitions were monitored: m/z 324/277, for LNO2 isomers, and m/z 342/295, a transition consistent with the loss of HNO2 for the 13C-labeled internal standard. The areas under each peak were integrated, the ratio of analyte to internal standard areas was determined and amounts of LNO2 were quantitated using Analyst 1.4 quantitation software (Applied Biosystems, Foster City, Calif.) by fitting the data to an internal standard curve. Adjustable mass spectrometer settings for both qualitative and quantitative analyses were as follows: CUR 10; IS −4500; TEM 450; GS1 50; GS2 60; CAD 2 (8 when EPI experiments were concurrently performed); DP −50; and EP −10. House-generated zero grade air was used as the source gas and nitrogen was used as the curtain and collision gases.
To quantitate net LNO2 species present in red cells and plasma, HPLC gradient conditions were changed (see Experimental Methods) so that all positional isomers co-eluted (FIG. 6). MRM transitions for the combined LNO2 and [13C]LNO2 (m/z 324 and 342, respectively) were monitored, with analytes eluting at 2 min. Nitrated linoleic acid concentration was a function of the ratio of analyte to internal standard peak areas using an internal standard curve that is linear over five orders of magnitude. Blood samples obtained from ten healthy human volunteers (5 female, 5 male, ages ranging from 22 to 45) revealed free LNO2 in red cells (i.e., LNO2 not esterified to glycerophospholipids or neutral lipids) to be 49.6±16.6 pmol/ml packed cells. Total free and esterified LNO2, the amount present in saponified samples, was 249±104 pmol/ml packed cells. Thus, approximately 75% of LNO2 in red cells exists as esterified fatty acids. In plasma, free and total (free plus esterified) LNO2 was 78.9±34.5 pmol/ml plasma 629±242 nM, respectively, with free LNO2 representing 85% of total (Table 1). These values indicate that LNO2 species are quantitatively the greatest pool of bioactive oxides of nitrogen in the vascular compartment (34-37).
Throughout the Background and Example 1, several publications have been referenced. These publications are listed in the Reference List following Example 1. The disclosures of these publications in their entireties are hereby incorporated by reference into this application in order to more fully describe the compounds, compositions and methods described herein.







TABLE 1







Biologically active nitrogen oxide derivatives in human blood -


comparison with net C10- and C12-nitro derivatives of linoleic acid.


Venous blood was obtained from healthy human volunteers, centrifuged


(1200 x g; 10 min) and plasma isolated from red cell pellets from which


the buffy coat was removed. Total lipid extracts were prepared from


packed red cells and plasma (29) and analyzed by mass spectrometry as


described in Experimental Methods. Total LNO2 (free plus esterified)


was determined in lipid extracts following saponification. Free and total


LNO2 was quantitated by fitting analyte to internal standard area


ratios obtained by mass spectrometry to an internal standard curve.


Data are expressed as mean ± SD (n = 10; 5 female, 5 male).










Species
Compartment
Concentration (nM)
Reference





NO2−
Plasma
205 ± 21 
(35, 36)


RSNO
Plasma
7.2 ± 1.1
(35, 36)



Plasma


LNO2
Free
80 ± 34



Esterified
550 ± 274



Total
630 ± 240


Hb-NO
Blood
>50
(37)


Hb-SNO
Blood
0-150
(34)


LNO2
Packed red cells



Free
25 ± 8 



Esterified
199 ± 60 



Total
224 ± 52 


LNO2
Whole Blood*
477 ± 128





*assuming a 40% hematocrit






REFERENCES


    1. Arnold, W. P., Mittal, C. K., Katsuki, S. & Murad, F. (1977) Proc. Natl. Acad. Sci. U.S.A 74, 3203-3207.
    2. Beckman, J. S., Beckman, T. W., Chen, J., Marshall, P. A. & Freeman, B. A. (1990) Proc. Natl. Acad. Sci. U.S.A 87, 1620-1624.
    3. Nathan, C. (1992) Federation of American Societies for Experimental Biology Journal 6, 3051-3064.
    4. Jourd'heuil, D., Miranda, K. M., Kim, S. M., Espey, M. G., Vodovotz, Y., Laroux, S., Mai, C. T., Miles, A. M., Grisham, M. B. & Wink, D. A. (1999) Arch. Biochem. Biophys. 365, 92-100.
    5. Rubbo, H., Darley-Usmar, V. & Freeman, B. A. (1996) Chem. Res. Toxicol. 9, 809-820.
    6. Kissner, R., Nauser, T., Bugnon, P., Lye, P. G. & Koppenol, W. H. (1997) Chem. Res. Toxicol. 10, 1285-1292.
    7. Radi, R., Denicola, A. & Freeman, B. A. (1999) Methods Enzymol. 301, 353-367.
    8. Baldus, S., Eiserich, J. P., Mani, A., Castro, L., Figueroa, M., Chumley, P., Ma, W., Tousson, A., White, C. R., Bullard, D. C. et al. (2001) J. Clin. Invest 108, 1759-1770.
    9. Castro, L., Eiserich, J. P., Sweeney, S., Radi, R. & Freeman, B. A. (2004) Arch. Biochem. Biophys. 421, 99-107.
    10. Eiserich, J. P., Baldus, S., Brennan, M. L., Ma, W., Zhang, C., Tousson, A., Castro, L., Lusis, A. J., Nauseef, W. M., White, C. R. et al. (2002) Science 296, 2391-2394.
    11. Grisham, M. B. (1985) J. Free Radic. Biol. Med. 1, 227-232.
    12. Liu, X., Miller, M. J., Joshi, M. S., Thomas, D. D. & Lancaster, J. R., Jr. (1998) Proc. Natl. Acad. Sci. U.S.A 95, 2175-2179.
    13. Subczynski, W. K., Lomnicka, M. & Hyde, J. S. (1996) Free Radic. Res. 24, 343-349.
    14. Thomas, D. D., Liu, X., Kantrow, S. P. & Lancaster, J. R., Jr. (2001) Proc. Natl. Acad. Sci. U.S.A 98, 355-360.
    15. Baldus, S., Castro, L., Eiserich, J. P. & Freeman, B. A. (2001) Am. J. Respir. Crit Care Med. 163, 308-310.
    16. Gallon, A. A. & Pryor, W. A. (1993) Lipids 28, 125-133.
    17. Gallon, A. A. & Pryor, W. A. (1994) Lipids 29, 171-176.
    18. Napolitano, A., Camera, E., Picardo, M. & d'Ischia, M. (2000) J. Org. Chem. 65, 4853-4860.
    19. Napolitano, A., Crescenzi, O., Camera, E., Giudicianni, I., Picardo, M. & d'Ischia, M. (2004) Tetrahedron 58, 5061-5067.
    20. O'Donnell, V. B., Eiserich, J. P., Chumley, P. H., Jablonsky, M. J., Krishna, N. R., Kirk, M., Barnes, S., Darley-Usmar, V. M. & Freeman, B. A. (1999) Chem. Res. Toxicol. 12, 83-92.
    21. Pryor, W. A., Lightsey, J. W. & Church, D. F. (2004) Journal of the american chemical society 104, 6685-6692.
    22. Lim, D. G., Sweeney, S., Bloodsworth, A., White, C. R., Chumley, P. H., Krishna, N. R., Schopfer, F., O'Donnell, V. B., Eiserich, J. P. & Freeman, B. A. (2002) Proc. Natl. Acad. Sci. U.S.A 99, 15941-15946.
    23. Coles, B., Bloodsworth, A., Clark, S. R., Lewis, M. J., Cross, A. R., Freeman, B. A. & O'Donnell, V. B. (2002) Circ. Res. 91, 375-381.
    24. O'Donnell, V. B. & Freeman, B. A. (2001) Circ. Res. 88, 12-21. \
    25. O'Donnell, V. B., Chumley, P. H., Hogg, N., Bloodsworth, A., Darley-Usmar, V. M. & Freeman, B. A. (1997) Biochemistry 36, 15216-15223.
    26. Coles, B., Bloodsworth, A., Eiserich, J. P., Coffey, M. J., McLoughlin, R. M., Giddings, J. C., Lewis, M. J., Haslam, R. J., Freeman, B. A. & O'Donnell, V. B. (2002) J. Biol. Chem. 277, 5832-5840.
    27. Coles, B., Bloodsworth, A., Clark, S. R., Lewis, M. J., Cross, A. R., Freeman, B. A. & O'Donnell, V. B. (2002) Circ. Res. 91, 375-381.
    28. Balazy, M., Iesaki, T., Park, J. L., Jiang, H., Kaminski, P. M. & Wolin, M. S. (2001) J. Pharmacol. Exp. Ther. 299, 611-619.
    29. Bligh, E. G. & Dyer, W. L. (1959) Can. J. Biochem. Physiol. 37, 911-917.
    30. Nakamura, T., Bratton, D. L. & Murphy, R. C. (1997) J. Mass Spectrom. 32, 888-896.
    31. Wheelen, P., Zirrolli, J. A. & Murphy, R. C. (1994) J. Am. Soc. Mass Spectrom. 6, 40-51.
    32. d'Ischia, M., Rega, N. & Barone, V. (1999) Tetrahedron 55, 9297-9308.
    33. Ranu, B. C. & Chakraborty, R. (1991) Tetrahedron Letters 32, 3579-3582.
    34. Gladwin, M. T., Shelhamer, J. H., Schechter, A. N., Pease-Fye, M. E., Waclawiw, M. A., Panza, J. A., Ognibene, F. P. & Cannon, R. O., III (2000) Proc. Natl. Acad. Sci. U.S.A 97, 11482-11487.
    35. Rassaf, T., Bryan, N. S., Kelm, M. & Feelisch, M. (2002) Free Radic. Biol. Med. 33, 1590-1596.
    36. Rassaf, T., Bryan, N. S., Maloney, R. E., Specian, V., Kelm, M., Kalyanaraman, B., Rodriguez, J. & Feelisch, M. (2003) Nat. Med. 9, 481-482.
    37. Cosby, K., Partovi, K. S., Crawford, J. H., Patel, R. P., Reiter, C. D., Martyr, S., Yang, B. K., Waclawiw, M. A., Zalos, G., Xu, X. et al. (2003) Nat. Med. 9, 1498-1505.
    38. Zelinsky, N. D. & Rosanoff, M. A. (1912) Z. Physikal Chem. 78, 629-633.
    39. Finlayson-Pitts, B. J., Sweetman, L. L. & Weissbart, B. (1987) Toxicol. Appl. Pharmacol. 89, 438-448.
    40. Hogg, N., Kalyanaraman, B., Joseph, J., Struck, A. & Parthasarathy, S. (1993) FEBS Lett. 334, 170-174.
    41. Rubbo, H., Radi, R., Trujillo, M., Telleri, R., Kalyanaraman, B., Barnes, S., Kirk, M. & Freeman, B. A. (1994) J. Biol. Chem. 269, 26066-26075.
    42. Rubbo, H., Parthasarathy, S., Barnes, S., Kirk, M., Kalyanaraman, B. & Freeman, B. A. (1995) Arch. Biochem. Biophys. 324, 15-25.
    43. Foster, M. W., McMahon, T. J. & Stamler, J. S. (2003) Trends Mol. Med. 9, 160-168.
    44. Schopfer, F. J., Baker, P. R. & Freeman, B. A. (2003) Trends Biochem. Sci. 28, 646-654.
    45. Coles, B., Bloodsworth, A., Eiserich, J. P., Coffey, M. J., McLoughlin, R. M., Giddings, J. C., Lewis, M. J., Haslam, R. J., Freeman, B. A. & O'Donnell, V. B. (2002) J. Biol. Chem. 277, 5832-5840.
    46. Lima, E. S., Di Mascio, P., Rubbo, H. & Abdalla, D. S. (2002) Biochemistry 41, 10717-10722.
    47. Lima, E. S., Di Mascio, P. & Abdalla, D. S. (2003) J. Lipid Res. 44, 1660-1666.
    48. Knowles, M. E., McWeeny, D. J., Couchman, L. & Thorogood, M. (1974) Nature 247, 288-289.
    49. Hayama, T., Tomoda, S., Takeuchi, Y. & Nomura, Y. (1982) Tetrahedron Letters 23, 4733-4734.


Example 2
Allylic nitro derivatives of linoleic acid (nitrolinoleic acid, LNO2) are formed via nitric oxide-dependent oxidative inflammatory reactions and are found at concentrations of  ̃500 nM in the blood of healthy individuals. It will be shown that nitrolinoleic acid (LNO2) is a potent endogenous ligand for peroxisome proliferator-activated receptor γ (PPAR γ; Ki ̃133 nM) that acts within physiological concentration ranges. This nuclear hormone receptor (PPARγ) regulates glucose homeostasis, lipid metabolism and inflammation. PPARγ ligand activity is specific for LNO2 and not mediated by LNO2 decay products, NO donors, linoleic acid or oxidized linoleic acid. LNO2 is a significantly more robust PPARγ ligand than other reported endogenous PPARγ ligands, including lysophosphatidic acid (LPA 16:0 and LPA 18:1), 15-deoxy-Δ12,14-PGJ2, conjugated linoleic acid and azelaoyl-PC. LNO2 activation of PPAR γ via CV-1 cell luciferase reporter gene expression analysis revealed a ligand activity that rivals or exceeds synthetic PPARγ agonists such as Rosiglitazone and Ciglitazone, is co-activated by 9 cis-retinoic acid and is inhibited by the PPAR γ antagonist GW9662. LNO2 induces PPARγ-dependent macrophage CD-36 expression, adipocyte differentiation and glucose uptake also at a potency rivaling thiazolidinediones. These observations reveal that nitric oxide (.NO)-mediated cell signaling reactions can be transduced by fatty acid nitration products and PPAR-dependent gene expression.
The reaction of nitric oxide (.NO) with tissue free radical and oxidative intermediates yields secondary oxides of nitrogen that mediate oxidation, nitration and nitrosation reactions (1, 2). Of present relevance, the reaction of .NO and .NO-derived species with oxidizing unsaturated fatty acids is kinetically rapid and exerts a multifaceted impact on cell redox and signaling reactions. Nitric oxide readily out-competes lipophilic antioxidants for the scavenging of lipid radicals, resulting in the inhibition of peroxyl radical-mediated chain propagation reactions (3). Both the catalytic activity and gene expression of eicosanoid biosynthetic enzymes are also regulated by NO, affirming a strong linkage between NO and fatty acid oxygenation product synthesis and signaling (4, 5). Consistent with this latter precept, fatty acid nitration products generated by NO-derived species inhibit multiple aspects of inflammatory cell function, indicating that nitrated fatty acids are both byproducts and mediators of redox signaling reactions (6-8).
Recently, the structural characterization and quantitation of nitrolinoleic acid (LNO2) in human red cells and plasma revealed this unsaturated fatty acid derivative to be the most abundant bioactive oxide of nitrogen in the vasculature. Net blood levels of  ̃80 and 550 nM free and esterified LNO2, respectively, were measured in healthy humans (9). The observation that NO-dependent oxidative inflammatory reactions yields allylic nitro derivatives of unsaturated fatty acids displaying cGMP-independent cell signaling properties (5) led to the identification of a receptor that can transduce LNO2 signaling. Affymetrix oligonucleotide microarray analysis of cRNA prepared from methanol (vehicle), linoleic acid (LA)- and LNO2-treated human aortic smooth muscle cells indicated that LNO2 specifically and potently regulated the expression of key inflammatory, cell proliferation and cell differentiation-related proteins. Multiple PPARγ target genes were significantly regulated, suggesting that LNO2 serves as an endogenous PPARγ ligand.
PPARγ is a nuclear hormone receptor that binds lipophilic ligands. Downstream effects of PPARγ activation include modulation of metabolic and cellular differentiation genes and regulation of inflammatory responses (e.g., integrin expression, lipid transport by monocytes), adipogenesis and glucose homeostasis (10, 11). In the vasculature, PPARγ is expressed in monocytes, macrophages, smooth muscle cells and endothelium (12) and plays a central role in regulating the expression of genes related to lipid trafficking, cell proliferation and inflammatory signaling(13). While synthetic thiazolidinediones (TZDs) such as Rosiglitazone and Ciglitazone are appreciated to be the most potent PPARγ ligands yet described, considerable interest and debate remains focused on the identity of endogenous PPARγ ligands because of therapeutic potential and their intrinsic value in understanding cell signaling. At present, tissue and plasma levels of putative PPAR ligands are frequently not precisely defined and when so, are found in concentrations sometimes orders of magnitude lower than those required to activate specific α, γ or δ PPAR subtypes (14-16). As set forth herein, the allylic nitro derivatives of fatty acids are robust endogenous PPARγ ligands that act within physiological concentration ranges to modulate key PPARγ-regulated signaling events including adipogenesis, adipocyte glucose homeostasis and CD36 expression in macrophages.
Materials.
LNO2 and [13C]LNO2 were synthesized and purified using linoleic acid (NuCheckPrep, Elysian, Minn.) and [13C]linoleic acid (Spectra Stables Isotopes, Columbia, Md.) subjected to nitroselenylation as previously described (9). LNO2 concentrations were quantified spectroscopically and by chemiluminescent nitrogen analysis (Antec Instruments, Houston, Tex.) using caffeine as a standard (9). Anti-CD36 antibody (kindly provided by Dr. de Beer at University of Kentucky Medical Center, Lexington, Ky.); anti-PPARγ and anti-β-actin antibodies were from Santa Cruz (Santa Cruz, Calif.); and anti-aP2 antibody was from Chemicon International Inc. (Temecula, Calif.). Horseradish peroxidase-linked goat anti-rabbit IgG and Coomasie Blue were from Pierce (Rockford, Ill.). [3H]Rosiglitazone was from American Radiolabeled Chemical, Inc. (St. Louis, Mo.). [3H]2-Deoxy-D-glucose was from Sigma (St Louis, Mich.). Scintil-safe PlusTM 50% was from Fisher Scientific (Pittsburgh, Pa.). Rosiglitazone, Ciglitazone, 15-deoxy-Δ12,14-PGJ2, Conjugated Linoleic Acid (CLA1, CLA2) and GW9662 were from Cayman Chemical (Ann Arbor, Mich.). 1-palmitoyl-2-hydroxy-sn-glycero-3-Phosphate (16:0 LPA), 1-O-9-(Z)-octadecenyl-2-hydroxy-sn-glycero-3-phosphate (18:1 LPA), 1-O-hexadecyl-2-azelaoyl-sn-glycero-3-phosphocholine (azPC), 1-Palmitoy-2-Azelaoyl-sn-Glycero-3-Phosphocholine (azPC Ester) were from Avanti Polar Lipids, Inc. (Alabaster, Ala.).
Cell Transient Transfection Assay.
CV-1 cells from ATCC (Manassas, Va.) were grown to  ̃85% confluence in DMEM/F12 supplemented with 10% FBS, 1% penicillin-streptomycin. Then, cells were transiently co-transfected with a plasmid containing the luciferase gene under regulation by four Gal4 DNA binding elements (UASG×4 TK-Luciferase, a gift from Dr. Ronald M. Evans), in concert with plasmids containing the ligand binding domain for the different nuclear receptors fused to the Gal4 DNA binding domain. For assessing full-length PPAR receptors, CV-1 cells were transiently co-transfected with a plasmid containing the luciferase gene under the control of three tandem PPAR response elements (PPRE) (PPRE×3 TK-Luciferase) and hPPARγ, hPPARα or hPPARδ expression plasmids, respectively. In all cases, fluorescence protein (GFP) expression plasmid was co-transfected as the control for the transfection efficiency. Twenty-four hours after the transfection, cells were cultured for another 24 hours in Optium-MEM (Invitrogen, Carlsbad, Calif.). Then, cells were treated with different compounds as indicated in figures for 24 hours in Optium-MEM. Reporter luciferase assay kits from Promega (Madison, Wis.) were used to measure the luciferase activity according to the manufacturer's instructions with a luminometer (Victor II, Perking Elmer). Luciferase activity was normalized by GFP units. Each condition was performed at least in triplicates in each experiment. All experiments were repeated at least three times.
PPARγ Competition Binding Assay.
Human PPARγ1 cDNA was inserted into pGEX from Amersham Biosciences Corp (Piscataway, N.J.) containing the gene encoding glutathione S-transferase (GST). GST-PPARγ protein induction and receptor binding was assessed as previously described (17).
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 postconfluent preadipocytes (designated day 0) were cultured in DMEM containing 10% FBS plus 3 μM LNO2 for 14 days. The medium was changed every two days. Rosiglitazone (3 μM) and linoleic acid (10 μM) were used as the positive and negative control, respectively. The differentiated adipocytes were stained by Oil red 0 as previously described (18).
[3H]-2-Deoxy-D-glucose Uptake Assay in Differentiated 3T3-L1 Adipocyte.
[3H]-2-Deoxy-D-glucoseuptake assay as previously described (19) 3T3-L1 preadipocytes were grown in 24-well tissue culture plates, 2-day postconfluent preadipocytes were treated by 10 μg/ml insulin (Sigma), 1 μM dexamethasone (Sigma), and 0.5 mM 3-isobutyl-1-methylxanthine (Sigma) in DMEM containing 10% FBS for two days, then cells were kept in 10 μg/ml insulin also in DMEM containing 10% FBS for 6 days (changed medium every three days). Eight days after induction of adipogenesis, test compounds in DMEM containing 10% FBS were added for an additional 2 days (changed medium every day) and PPARγ-specific antagonist GW9662 were pretreated 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. The treated cells were rinsed carefully three times with cold PBS, lysed overnight in 0.8N NaOH (0.4 ml/well), and neutralized with 13.3 μl of 12N HCl. Lysate (360 μl) was added to 4 ml Scinti-safe PlusTM 50% in a scintillation vial, and the vials were mixed and counted.
RNA and Protein Preparation and Analysis.
RNA and protein expression levels were analyzed by quantitative real-time PCR and Western blot analysis as previously described.25 LNO2 decay. LNO2 decay was induced by incubating 3 μM LNO2 in medium+serum at 37° C. At different times, samples were removed and analyzed for bioactivty or for LNO2 content by Bligh and Dyer extraction in the presence of 1 μM [13C]LNO2 as an internal standard. Non-decayed LNO2 was quantified via triple quadrupole mass spectrometric analysis (Applied Biosystems/MDS Sciex, Thornhill, Ontario, Canada) as previously reported (9).
To characterize LNO2 as a potential ligand for a lipid-binding nuclear receptor [e.g., PPARα, PPARγ, PPARγ, androgen receptor (AR), glucocorticoid receptor (GR), mineralocorticoid receptor (MR), progesterone receptor (PR) and retinoic X receptor α (RXRα)], CV-1 reporter cells were co-transfected with plasmids containing the ligand binding domain for these nuclear receptors fused to the Gal4 DNA binding domain and the luciferase gene under regulation of four Gal4 DNA binding elements. LNO2 (1 μM) induced significant activation of PPARγ (620%), PPARα (325%) and PPARγ (221%), with no impact on AR, GR, MR, PR or RXRα receptor activation (FIG. 7A). To further explore PPAR activation by LNO2, CV-1 cells were transiently co-transfected with a plasmid containing the luciferase gene under three PPAR response elements (PPRE) in concert with PPARγ, PPARα or PPARγ expression plasmids. Dose-dependent activation by LNO2 was observed for all PPARs by LNO2 (FIG. 7B), with PPARγ showing the greatest response at clinically-relevant concentrations of LNO2.
PPARγ activation by LNO2 rivaled that induced by Ciglitazone and Rosiglitazone and exceeded that of 15-deoxy-Δ12,14-PGJ2 (20, 21), which only occurred at concentrations 3 orders of magnitude greater than found clinically (FIG. 7C and inset). Several other reported endogenous PPARγ activators added in equimolar concentration with LNO2 [linoleic acid (LA), conjugated linoleic acid (CLA1, CLA2), lysophosphatidic acid (LPA-16:0, LPA-18:1), azPC (azelaoylPC) and azPC ester; 1 and 3 μM], displayed no significant activation of PPARγ reporter gene expression when compared with vehicle control (FIG. 7C) (22-24) LNO2-mediated PPARγ activation was inhibited by the PPARγ-specific antagonist GW9662 and enhanced  ̃180% by co-addition of the RXRα agonist 9-cis-retinoic acid, which facilitates PPRE promoter activation via heterodimerization of activated RXRα with PPARγ (FIG. 8A).
LNO2 slowly undergoes decay reactions in aqueous solution, displaying a 30-60 min half-life (FIG. 8B) and yielding .NO and an array of oxidation products. The activation of PPARγ paralleled the presence and concentration of the LNO2 parent molecule, as determined by concomitant receptor activation analysis and mass spectrometric quantitation of residual LNO2 in reaction mixtures undergoing different degrees of aqueous decomposition prior to addition to CV-1 cells (FIG. 8B). Affirmation that activation of PPARγ is LNO2-specific, rather than a consequence of LNO2 decay products, was established by treatment of PPARγ-transfected CV-1 cells with .NO donors and oxidized linoleic acid derivatives. Additionally, the fatty acid oxidation products 9- and 13-oxoODE are reported endogenous PPARγ stimuli (25-27). 13-oxoODE had no effect on PPARγ-dependent reporter gene expression (FIG. 8C). Only high and non-physiological concentrations of 9-oxoODE, and the concerted addition of high concentrations of 13-oxoODE and S-nitrosoglutathione or spermine-NONOate, resulted in modest PPARγ activation (FIG. 8C). The .NO donors added individually did not activate PPARγ, even at high concentrations, eliminating the possibility that PPARγ-mediated signaling by LNO2 is due to .NO derivatives released during LNO2 decay (FIG. 8C).
Competitive PPARγ binding analysis quantified the displacement of [3H]Rosiglitazone by unlabeled Rosiglitazone, LNO2 and linoleic acid. The calculated binding affinity (Ki) for Rosiglitazone was 53 nM, consistent with reported values (FIG. 8D, 40-50 nM (28)). LNO2 displayed an estimated Ki of 133 nM and linoleic acid an estimated Ki of >1,000 nM (previously reported as 1.7 to 17 μM (28)). This reveals that the Ki for LNO2 displacement of [3H]Rosiglitazone from PPARγ is comparable to this highly avid ligand.
The PPARγ agonist actions of LNO2 were also examined in a biological context, using cell models noted for well-established PPARγ-dependent functions. The scavenger receptor CD36 is expressed in diverse cell types, including platelets, adipocytes and macrophages. In macrophages, CD36 is a receptor for oxidized LDL, with expression positively regulated by PPARγ (29). Treatment of mouse RAW264.7 macrophages with LNO2 induced greater CD36 receptor protein expression than an equivalent Rosiglitazone concentration, a response partially inhibitable by the PPARγ-specific antagonist GW9662 (FIG. 9A). Moreover, quantitative real-time PCR revealed that LNO2-dependent PPARγ transactivation induced a dose-dependent increase in CD36 mRNA expression in these macrophages.
PPARγ plays an essential role in the differentiation of adipocytes (30, 31). In support of this precept, selective disruption of PPARγ results in impaired development of adipose tissue (18, 32). To define if LNO2 induces PPARγ-activated adipogenesis, 3T3-L1 preadipocytes were treated with LNO2, Rosiglitazone or linoleic acid for two weeks. Adipocyte differentiation was assessed both morphologically and via oil red O staining, which reveals the accumulation of intracellular lipids. Vehicle and linoleic acid did not affect differentiation, while LNO2 induced >30% of 3T3-L1 preadiopcyte differentiation (FIG. 9B). Rosiglitazone treatment affirmed a positive PPARγ-dependent response. LNO2 and Rosiglitazone-induced preadipocyte differentiation also resulted in expression of specific adipocyte markers (PPARγ2 and aP2), an event not detected for linoleic acid (FIG. 9C). PPARγ ligands play a central role in glucose metabolism, with TZDs widely used as insulin-sensitizing drugs. Addition of LNO2 (1-10 μM) to differentiated adipocytes induced a dose-dependent increase in glucose uptake (FIG. 9D, Left panel). The impact of LNO2 on glucose uptake was greater than that observed for equimolar 15-deoxy-PGJ2, equivalent to Rosiglitazone and similarly inhibited by the PPARγ-specific antagonist GW9662 (FIG. 9D, Right panel). In aggregate, these observations establish that LNO2 induces well-characterized PPARγ-dependent signaling actions towards macrophage CD36 expression and adipogenesis.
The identification of bona fide high affinity endogenous PPARγ ligands has been a provocative issue that, when resolved, will advance our understanding of endogenous PPARγ modulation and can reveal new means for intervention in diverse metabolic disorders and disease processes. This will also shed light on the broader contributions of this nuclear hormone receptor family to both the maintenance of tissue homeostasis and the regulation of cell and organ dysfunction. Of relevance, the generation of PPAR-activating intermediates from complex lipids often requires hydrolysis by phospholipase A2 (PLA2), with the identity of phospholipid sn-2 position fatty acid derivatives that might serve as PPAR agonists still forthcoming (33, 34). Thus, the fact that  ̃80% of LNO2 present in a variety of tissue compartments is esterified encourages the notion that this pool of high affinity PPAR ligand activity will be mobilized upon the PLA2 activation that occurs during inflammation to regulate the formation of cell signaling molecules.
Comparison of PPARγ-dependent gene expression induced by LNO2 with that of TZDs and putative fatty acid- and phospholipid-derived PPARγ ligands further affirmed the robust activity of LNO2 as a PPARγ ligand. While a number of endogenous lipophilic species are proposed as PPARγ ligands, their intrinsically low binding affinities and in vivo concentrations do not support a capability to serve as physiologically-relevant signaling mediators. Presently-reported endogenous PPARγ agonists include free fatty acids, components of oxidized plasma lipoproteins (9- and 13-oxoODE, azPC), conjugated linoleic acid derivatives (CLA1 and CLA2), products of phospholipase hydrolysis of complex lipids (LPA), platelet activating factor (PAF) and eicosanoid derivatives such as the dehydration product of PGD2, 15-deoxy-Δ12,14-PGJ2 (14, 22, 23). Herein, minimal or no activation of PPARγ reporter gene expression by 1-3 μM concentrations of these putative ligands was observed, in contrast to the dose-dependent PPAR activation by LNO2 that, for PPARγ, was significant at clinically-relevant concentrations as low as 100 nM. A dilemma exists in that some putative endogenous PPARγ agonists have only been generated by aggressive in vitro oxidizing conditions (e.g., Cu-mediated LDL oxidation) and have not been clinically quantified or detected (for instance, azPC, CLA). Other lipid derivatives proposed as PPAR ligands are present in <100 nM tissue concentrations, orders of magnitude below their binding affinities (1-15 μM) and are not expected to result in significant receptor occupancy and activation in vivo. This latter category includes free fatty acids, eicosanoids, 9- and 13-oxoODE, PAF and 15-deoxy-Δ12,14-PGJ2 (14). For example, while LPA is a notable PPARγ ligand of relevance to vascular, inflammatory and cell proliferative diseases the plasma concentration of LPA is well below 100 nM (35, 36) and its binding affinity for PPARγ has not been established. Also, the in vivo downstream vascular signaling actions of LPA are inconsistent with its proposed PPARγ ligand activity (24, 37-39). Future studies using mice genetically deficient for PPARγ and NOS isoforms should assist in defining the mechanisms of formation and endogenous PPAR ligand activity of nitrated fatty acids.
In summary, LNO2 is a high affinity ligand for PPARs, especially PPARγ, that activates both reporter constructs and cells at physiological concentrations. Fatty acid nitration products, generated by NO-dependent reactions, are thus expected to display broad cell signaling capabilities as endogenous nuclear receptor-dependent paracrine signaling molecules with a potency that rivals TZDs (FIG. 7C). Present data reveals that LNO2 mediates cell differentiation in adipocytes, CD36 expression in macrophages, and inflammatory-related signaling events in endothelium (e.g., inhibition of VCAM-1 expression and function, not shown) with an important contribution from PPARγ-dependent mechanisms (FIG. 9A-D). Allylic nitro derivatives of fatty acids thus represent a unique class of receptor-dependent cell differentiation, metabolic and anti-inflammatory signaling molecules that serve to converge NO and oxygenated lipid redox signaling pathways.
Throughout Example 2, several publications have been referenced. These publications are listed in the Reference List for Example 2. The disclosures of these publications in their entireties are hereby incorporated by reference into this application in order to more fully describe the compounds, compositions and methods described herein.
REFERENCE LIST FOR EXAMPLE 2


    1. Schopfer, F. J., Baker, P. R. & Freeman, B. A. (2003) Trends Biochem. Sci. 28, 646-654.
    2. Hogg, N. (2002) Annu. Rev. Pharmacol. Toxicol. 42, 585-600.
    3. Rubbo, H., Radi, R., Anselmi, D., Kirk, M., Barnes, S., Butler, J., Eiserich, J. P. & Freeman, B. A. (2000) J. Biol. Chem. 275, 10812-10818.
    4. Marnett, L. J., Wright, T. L., Crews, B. C., Tannenbaum, S. R. & Morrow, J. D. (2000) J. Biol. Chem. 275, 13427-13430.
    5. O'Donnell, V. B. & Freeman, B. A. (2001) Circ. Res. 88, 12-21.
    6. Coles, B., Bloodsworth, A., Eiserich, J. P., Coffey, M. J., McLoughlin, R. M., Giddings, J. C., Lewis, M. J., Haslam, R. J., Freeman, B. A. & O'Donnell, V. B. (2002) J. Biol. Chem. 277, 5832-5840.
    7. Coles, B., Bloodsworth, A., Clark, S. R., Lewis, M. J., Cross, A. R., Freeman, B. A. & O'Donnell, V. B. (2002) Circ. Res. 91, 375-381.
    8. Rubbo, H., Radi, R., Trujillo, M., Telleri, R., Kalyanaraman, B., Barnes, S., Kirk, M. & Freeman, B. A. (1994) J. Biol. Chem. 269, 26066-26075.
    9. Baker, P. R., Schopfer, F. J., Sweeney, S. & Freeman, B. A. (2004) Proc. Natl. Acad. Sci. U.S.A 101, 11577-11582.
    10. Lee, C. H. & Evans, R. M. (2002) Trends Endocrinol. Metab. 13, 331-335.
    11. Marx, N., Duez, H., Fruchart, J. C. & Staels, B. (2004) Circ. Res. 94, 1168-1178.
    12. Wang, N., Verna, L., Chen, N. G., Chen, J., Li, H., Forman, B. M. & Stemerman, M. B. (2002) J. Biol. Chem. 277, 34176-34181.
    13. Chen, Y. E., Fu, M., Zhang, J., Zhu, X., Lin, Y., Akinbami, M. A. & Song, Q. (2003) Vitam. Horm. 66, 157-188.
    14. Bell-Parikh, L. C., Ide, T., Lawson, J. A., McNamara, P., Reilly, M. & FitzGerald, G. A. (2003) J. Clin. Invest. 112, 945-955.
    15. Rosen, E. D. & Spiegelman, B. M. (2001) J Biol. Chem. 276, 37731-37734.
    16. Tzameli, I., Fang, H., Ollero, M., Shi, H., Hamm, J. K., Kievit, P., Hollenberg, A. N. & Flier, J. S. (2004) J. Biol. Chem. 
    17. Fu, J., Gaetani, S., Oveisi, F., Lo, V. J., Serrano, A., Rodriguez, D. F., Rosengarth, A., Luecke, H., Di Giacomo, B., Tarzia, G. et al. (2003) Nature 425, 90-93.
    18. Zhang, J., Fu, M., Cui, T., Xiong, C., Xu, K., Zhong, W., Xiao, Y., Floyd, D., Liang, J., Li, E. et al. (2004) Proc. Natl. Acad. Sci. U.S.A 101, 10703-10708.
    19. Mukherjee, R., Hoener, P. A., Jow, L., Bilakovics, J., Klausing, K., Mais, D. E., Faulkner, A., Croston, G. E. & Paterniti, J. R., Jr. (2000) Mol. Endocrinol. 14, 1425-1433.
    20. Forman, B. M., Tontonoz, P., Chen, J., Brun, R. P., Spiegelman, B. M. & Evans, R. M. (1995) Cell 83, 803-812.
    21. Kliewer, S. A., Lenhard, J. M., Willson, T. M., Patel, I., Morris, D. C. & Lehmann, J. M. (1995) Cell 83, 813-819.
    22. Davies, S. S., Pontsler, A. V., Marathe, G. K., Harrison, K. A., Murphy, R. C., Hinshaw, J. C., Prestwich, G. D., Hilaire, A. S., Prescott, S. M., Zimmerman, G. A. et al. (2001) J. Biol. Chem. 276, 16015-16023.
    23. McIntyre, T. M., Pontsler, A. V., Silva, A. R., St Hilaire, A., Xu, Y., Hinshaw, J. C., Zimmerman, G. A., Hama, K., Aoki, J., Arai, H. et al. (2003) Proc. Natl. Acad. Sci. U.S.A 100, 131-136.
    24. Zhang, C., Baker, D. L., Yasuda, S., Makarova, N., Balazs, L., Johnson, L. R., Marathe, G. K., McIntyre, T. M., Xu, Y., Prestwich, G. D. et al. (2004) J. Exp. Med. 199, 763-774.
    25. Kliewer, S. A., Sundseth, S. S., Jones, S. A., Brown, P. J., Wisely, G. B., Koble, C. S., Devchand, P., Wahli, W., Willson, T. M., Lenhard, J. M. et al. (1997) Proc. Natl. Acad. Sci. U.S.A 94, 4318-4323.
    26. Nagy, L., Tontonoz, P., Alvarez, J. G., Chen, H. & Evans, R. M. (1998) Cell 93, 229-240.
    27. Von Knethen, A. & Brune, B. (2002) J. Immunol. 169, 2619-2626.
    28. Ferry, G., Bruneau, V., Beauverger, P., Goussard, M., Rodriguez, M., Lamamy, V., Dromaint, S., Canet, E., Galizzi, J. P. & Boutin, J. A. (2001) Eur. J. Pharmacol. 417, 77-89.
    29. Chawla, A., Barak, Y., Nagy, L., Liao, D., Tontonoz, P. & Evans, R. M. (2001) Nat. Med. 7, 48-52.
    30. Tontonoz, P., Hu, E., Graves, R. A., Budavari, A. I. & Spiegelman, B. M. (1994) Genes Dev. 8, 1224-1234.
    31. Tontonoz, P., Hu, E. & Spiegelman, B. M. (1994) Cell 79, 1147-1156.
    32. Evans, R. M., Barish, G. D. & Wang, Y. X. (2004) Nat. Med. 10, 355-361.
    33. Delerive, P., Furman, C., Teissier, E., Fruchart, J., Duriez, P. & Staels, B. (2000) FEBS Lett. 471, 34-38.
    34. Pawliczak, R., Han, C., Huang, X. L., Demetris, A. J., Shelhamer, J. H. & Wu, T. (2002) J. Biol. Chem. 277, 33153-33163.
    35. Saulnier-Blache, J. S., Girard, A., Simon, M. F., Lafontan, M. & Valet, P. (2000) J. Lipid Res. 41, 1947-1951.
    36. Xu, Y., Shen, Z., Wiper, D. W., Wu, M., Morton, R. E., Elson, P., Kennedy, A. W., Belinson, J., Markman, M. & Casey, G. (1998) JAMA 280, 719-723.
    37. Chen, Z., Ishibashi, S., Perrey, S., Osuga, J., Gotoda, T., Kitamine, T., Tamura, Y., Okazaki, H., Yahagi, N., Iizuka, Y. et al. (2001) Arterioscler. Thromb. Vasc. Biol. 21, 372-377.
    38. Claudel, T., Leibowitz, M. D., Fievet, C., Tailleux, A., Wagner, B., Repa, J. J., Torpier, G., Lobaccaro, J. M., Paterniti, J. R., Mangelsdorf, D. J. et al. (2001) Proc. Natl. Acad. Sci. U.S.A 98, 2610-2615.
    39. Collins, A. R., Meehan, W. P., Kintscher, U., Jackson, S., Wakino, S., Noh, G., Palinski, W., Hsueh, W. A. & Law, R. E. (2001) Arterioscler. Thromb. Vasc. Biol. 21, 365-371.


Example 3
The synthesis, structural characterization, clinical quantitation and cell signaling activity of nitrated oleic acid (OA-NO2) are reported. Analysis of plasma and urine also revealed the presence of additional nitrated fatty acids, including nitrated linolenic, arachidonic, eicosapentaenoic and docosahexaenoic acids and multiple nitrohydroxy derivatives, which reveal the ubiquity of nitrated fatty acid derivatives in humans. Two nitroalkene derivatives of oleic acid were synthesized (9- and 10-nitro-9-cis-octadecenoic acid), structurally characterized and compared with nitrated fatty acids present in plasma, red cells and urine of healthy humans. Based on HPLC elution and mass spectrometric characteristics, these two regioisomers of OA-NO2 were identified in clinical samples. Using 13C isotope dilution, OA-NO2 was quantitated, with plasma free and esterified levels of 619±52 and 302±369 nM, respectively. In red blood cells, free and esterified OA-NO2 was 59±11 and 155±65 nM, respectively. Assuming a 40% hematocrit, OA-NO2 levels are  ̃50% greater than that of nitrated linoleic acid; with combined free and esterified levels of these two nitroalkene derivatives exceeding 1 μM. OA-NO2 potently activated peroxisome proliferator activated receptor-γ and induced PPAR γ-dependent adipogenesis and deoxyglucose uptake in 3T3 L1 preadipocytes. These data reveal that multiple nitrated fatty acids comprise a class of NO and fatty acid-derived signaling molecules.
The oxidation of unsaturated fatty acids converts lipids, otherwise serving as cell metabolic precursors and structural components, into potent signaling molecules including prostaglandins, leukotrienes, isoprostanes, hydroxy- and hydroperoxy-eicosatetraenoates and platelet-activating factor. This process, either enzymatic or auto-oxidative, orchestrates immune responses, neurotransmission and the regulation of cell growth. For example, prostaglandins are cyclooxygenase-derived lipid mediators that signal via receptor-ligand interactions to regulate inflammatory responses, vascular function, initiation of parturition, cell survival and angiogenesis (1). In contrast, the various isoprostane products of arachidonic acid auto-oxidation exert vasoconstrictive and pro-inflammatory signaling actions via both receptor-dependent and -independent mechanisms (2). A common element of these diverse lipid signaling actions is that nitric oxide (.NO) and other reactive nitrogen species significantly impact lipid mediator formation and bioactivities.
The ability of .NO and .NO-derived reactive species to oxidize, nitrosate and nitrate biomolecules suggests that .NO might also influence the synthesis and reactions of bioactive lipids (3-5). Interactions between .NO and lipid oxidation pathways are multifaceted and interdependent. For example, .NO regulates both the activity and expression of prostaglandin H synthase (6). Conversely, leukotriene products of lipoxygenases induce nitric oxide synthase-2 expression to increase .NO production (7). Furthermore, the autocatalytic chain propagation reactions of lipid peroxyl radicals during membrane and lipoprotein oxidation are potently inhibited by .NO (8). Of relevance, reactions between .NO-derived species and lipid oxidation intermediates yield nitrated fatty acids. Recently, the nitroalkene derivative of linoleic acid (LNO2) has been 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 (NO2−), nitrosothiols (RSNO) and heme-nitrosyl complexes, LNO2 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 (PPAR) (17), a class of nuclear hormone receptors that modulates the expression of metabolic and cellular differentiation and inflammatory-related genes (18,19).
The identification of LNO2 as an endogenous PPARγ ligand that acts within physiologically-relevant concentrations motivated a search for other nitrated lipids that might serve related signaling actions. Herein, fatty acid nitroalkene products in plasma and urine are abundant and ubiquitous are reported. Of the total fatty acid content in red cells, linoleic acid and oleic acid comprise  ̃8% and  ̃18%, respectively (20). Thus, due to its prevalence and structural simplicity, oleic acid was evaluated as a potential candidate for nitration. Herein, reported is the synthesis, structural characterization and cell signaling activity of 9- and 10-nitro-9-cis-octadecaenoic acids (nitrated oleic acid; OA-NO2; FIG. 10) and that OA-NO2 regioisomers are present in human blood at levels exceeding those of LNO2. Furthermore, OA-NO2 activates PPARγ with a greater potency than LNO2. These data reveal that nitrated unsaturated fatty acids represent a novel class of lipid-derived, receptor-dependent signaling mediators.
Methods
Materials.
9-Octadecenoic acid (oleic acid) and its respective methyl ester, methyl-9-octadecenoate was purchased from Nu-Check Prep (Elysian, Minn.). LNO2 and [13C]LNO2 were synthesized as previously described (9,12); OA-NO2 and [13C]OA-NO2 were synthesized as described in below. Phenylselenium bromide, HgCl2, NaNO2, anhydrous tetrahydrofuran (THF), CH3CN, CDCl3, insulin, dexamethasone and 3-isobutyl-1-methylxanthine were obtained from Sigma/Aldrich (St Louis, Mo.). Silica gel G and HF thin layer chromatography plates (250 and 2000 μm) were from Analtech (Newark, Del.). Methanolic BF3, horseradish peroxidase-linked goat anti-rabbit IgG and Coomasie Blue were from Pierce (Rockford, Ill.). Synthetic solvents were of HPLC grade or better from Fisher Scientific (Fairlawn, N.J.). Solvents used for extractions and mass spectrometric analyses were from Burdick and Jackson (Muskegon, Mich.). [13C]Oleic acid and [13C]linoleic acid were purchased from Cambridge Isotope Laboratories, Inc. (Andover, Mass.). Anti-PPARγ and anti-β-actin antibodies were from Santa Cruz (Santa Cruz, Calif.); anti-aP2 antibody was from Chemicon International Inc. (Temecula, Calif.).
Synthesis of OA-NO2.
Oleic acid and [13C]oleic acid were nitrated as described (9,12), with modifications. Briefly, 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 ten-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 (21). Based on this synthetic rationale, two regioisomers are generated: 9- and 10-nitro-9-cis-octadecenoic acids (generically termed OA-NO2). Thin layer chromatography does not resolve the two isomers. [13C]OA-NO2 was synthesized using [13C]oleic acid as a reactant. Stock concentrations of OA-NO2 isomers were quantitated by chemiluminescent nitrogen analysis (Antek Instruments, Houston, Tex.), using caffeine as a standard. All standards were diluted in methanol, aliquoted and stored under argon gas at −80° C. Under these conditions, OA-NO2 isomers remain stable for >3 months.
OA-NO2 Spectrophotometric Characterization.
OA-NO2 stock solution concentrations were initially determined by chemiluminescent nitrogen analysis. These data 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 (ε) for OA-NO2 and the isotopic derivative [13C]OA-NO2 were measured (λ270) using a UV-VIS spectrophotometer (Shimadzu, Japan). Absorbance values for increasing concentrations of OA-NO2 and [13C]OA-NO2 were plotted against concentration to calculate ε.
NMR spectrometric analysis of OA-NO2—
1H and 13C NMR spectra were measured using a Varian NOVA 300 and 500 MHz NMR and recorded in CDCl3. Chemical shifts are in δ 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).
Gas Chromatography Mass Spectrometric Characterization of OA-NO2.
Methyl esters of purified synthetic OA-NO2 regioisomers were analyzed by electron impact ionization gas chromatography mass spectrometry (EI GC-MS). Fatty acid methyl esters of OA-NO2 were synthesized, extracted with hexane, washed twice with saturated saline, redissolved in undecane and analyzed by GC-MS using a Saturn 2000 Tandem Mass Spectrometer coupled to a Varian 3800 Gas Chromatograph. Samples were ionized by electron impact at +70 eV. The methyl ester-derivatized OA-NO2 regioisomers were resolved using a 30 m capillary column (5% phenyl 95% dimethylpolysiloxane; CP-Sil 8CB-MS, Varian) with the following temperature gradient: 80° C. (2 min); 80 to 170° C. at 20° C./min; 170 to 240° C. at 2° C./min and 240 to 280° C. at 5° C./min. Helium was used as a carrier gas.
Structural Characterization of OA-NO2 by Electrospray Ionization Triple Quadrupole Mass Spectrometry (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 separation was developed using a 150×2 mm C18 Phenomenex Luna column (3 μm particle size). Lipids were 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 to 65% B (10 min); 65 to 95% B (1 min; hold for 3 min) and 95 to 20% B (1 min; hold for 3 min). Using these gradient conditions, OA-NO2 elutes after LNO2 positional isomers. OA-NO2 was detected using a multiple reaction monitoring (MRM) scan mode by reporting molecules that undergo a m/z 326/279 mass transition, which is 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. EPI analysis utilizes the trap functionality of the triple quadrupole to “concentrate” fragment ions to enhance sensitivity. 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 healthy human volunteers was collected by venipuncture into heparinized tubes (UAB Institutional Review Board-approved protocol #X040311001). Blood was centrifuged (1200×g; 10 min), the buffy coat removed and erythrocytes were isolated. Lipid extracts were prepared from red cells and plasma (21) and directly analyzed by mass spectrometry. 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 #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 hr fasting; urine was collected from the first void of the day. During the monophase stage of the lipid extraction (21), [13C]OA-NO2 was added as internal standard to correct for losses due to extraction. Nitrated fatty acids were then analyzed by HPLC ESI MS/MS. Lipids were eluted from the HPLC column using an isocratic solvent system consisting of CH3CN:H2O:NH4OH (85:15:0.1, v/v), resulting in the co-elution of the two OA-NO2 regioisomers. During quantitative analyses, two MRM transitions were monitored: m/z 326/279 (OA-NO2) and m/z 344/297 ([13C]OA-NO2), transitions consistent with the loss of the nitro group from the respective precursor ions. The areas under each peak were integrated, the ratio of analyte to internal standard areas was determined and levels of OA-NO2 were quantitated using Analyst 1.4 quantitation software (Applied Biosystems/MDS Sciex) by fitting the data to an internal standard curve. Data are expressed as mean±std dev (n=10; 5 female and 5 male).
Qualitative Analysis of Nitro- and Nitrohydroxy-Adducts of Fatty Acids.
Using HPLC ESI-MS/MS, blood and urine samples were evaluated for the presence of allylic nitro derivatives other than LNO2 and OA-NO2. HPLC separations were performed similarly to those used to characterize OA-NO2, with some modifications. Alternative MRM transitions were used to detect other potential nitroalkene derivatives. Based on what appears to be a common fragmentation product of nitrated fatty acids (i.e., loss of the nitro group; [M−HNO2]−), theoretical MRM transitions were determined for nitrated linolenic (18:3—NO2), arachidonic (20:4—NO2) and docosahexaenoic acids (22:6—NO2). MRM transitions for nitrohydroxy adducts were also monitored: 18:1(OH)—NO2; 18:2(OH)—NO2; 18:3(OH)—NO2; 20:4(OH)—NO2 and 22:6(OH)—NO2.
LNO2 and OA-NO2 Decay.
The relative rates of LNO2 and OA-NO2 decay in aqueous solution were determined by incubating 3 μM LNO2 and OA-NO2 in phosphate buffer (100 mM, pH 7.4, 37° C.). During the two hour incubation, aliquots were removed and analyzed for LNO2 and OA-NO2 content. The aliquots were extracted as described (21) and 1 μM [13C]LNO2 was added during the monophase stage of the extraction procedure as an internal standard. Non-decayed LNO2 and OA-NO2 were quantitated via HPLC ESI-MS/MS as described above.
PPAR Transient Transfection Assay.
CV-1 cells from the ATCC (Manassas, Va.) were grown to  ̃85% confluence in DMEM/F12 supplemented with 10% FBS, 1% penicillin-streptomycin. Twelve hours before transfection, the media was removed and antibiotic-free media was applied. Cells were transiently co-transfected with a plasmid containing the luciferase gene under the control of three tandem PPAR response elements (PPRE) (PPRE×3 TK-Luciferase) and PPARγ, PPARα or PPARδ expression plasmids, respectively. In all cases, fluorescence protein (GFP) expression plasmid was co-transfected as the control for transfection efficiency. Twenty-four hours after transfection, cells the returned to OptiMEM (Invitrogen, Carlsbad, Calif.) for 24 hr and then treated as indicated for another 24 hr. Reporter luciferase assay kits from Promega (Madison, Wis.) were used to measure the luciferase activity according to the manufacturer's instructions with a luminometer (Victor II, Perkin-Elmer). Luciferase activity was normalized by GFP units. Each condition was performed in triplicate in 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 two 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 previously (22).
[3H]-2-Deoxy-D-Glucose Uptake Assay in Differentiated 3T3-L1 Adipocytes.
[3H]-2-deoxy-D-glucose uptake was analyzed as previously (23). 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 two days, then cells were maintained in 10 μg/ml insulin in DMEM containing 10% FBS for 6 days (medium was changed every three 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 PPARγ-specific antagonist GW9662 was added 1 hr before other additions. After two rinses with serum-free DMEM, cells were incubated for 3 hr 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 μl of 12 N HCl and 360 μl of lysate was added into 4 ml Scinti-safe PlusTM 50% for radioactivity determination by liquid scintillation counting.
Results
Detection and Identification of Nitrated PUFA.
The discovery that LNO2 is present in vivo motivated a search for additional endogenous nitrated fatty acids that may also act as lipid signaling molecules. To survey plasma and urine for other nitrated fatty acids, lipid extracts from healthy human blood donors were analyzed by HPLC ESI MS/MS in the multiple reaction monitoring (MRM) scan mode. MRM transitions were calculated for the nitro- and nitrohydroxy-adducts of six fatty acids, as shown in Table 2, and were used to detect nitro- and nitrohydroxy-adducts present in plasma and urine lipid extracts using an isocratic HPLC elution methodology (FIG. 11). Gradient elution methods using MRM and EPI scan modes were used for structural confirmation (not shown). Due to the lack of appropriate stable isotope internal standards for all derivatives, data are presented as base-peak spectra and are qualitative evidence that these species exist in vivo. Mass spectrometric analysis revealed that nitrated adducts of all monitored unsaturated fatty acids are present in blood and urine, which include the Michael-like addition products of these species with H2O detected as nitrohydroxy adducts. Owing to its predominant abundance and structural simplicity, oleic acid was synthesized as a standard to specifically quantitate endogenous OA-NO2 content and signaling activity.







TABLE 2







Multiple reaction monitoring (MRM) transitions


for fatty acid nitroalkene derivatives


MRM values for nitroalkene and nitrohydroxy-adducts


of fatty acids were based on the common loss of the


nitro group that occurs during collision-induced


dissociation of nitrated fatty acids.











Carbons:dou-
Nitro adduct
Nitrohydroxy adduct


Fatty Acid
ble bonds
(—NO2)
(L(OH)—NO2)





Oleic
18:1
326/279
344/297


Linoleic
18:2
324/277
342/295


Linolenic
18:3
322/275
340/293


Arachidonic
20:4
348/301
366/319


Eicosapentaenoic
20:5
346/299
364/317


Docosahexaenoic
22:6
372/325
390/343









Synthesis and Purification of OA-NO2.

Nitration of oleic acid by nitrosenylation will yield two potential regioisomers of OA-NO2 (FIG. 10). Upon purification, analytical TLC, GC and LC-mass spectrometry of synthetic OA-NO2 indicated no contamination by oleic acid nor oxidized products (not shown).
NMR Analysis of OA-NO2.
The structure of synthetic OA-NO2 (a 1:1 mixture of C9- and C10 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): δ 11.1 (br s, 1H); 7.06 (dd, 1H, J=7.8 Hz); 3.75 (t, 2H, J=6.7 Hz); 2.55 (t, 2H, J=7.6 Hz); 2.36 (q, 2H, J=7.6 Hz); 2.33 (m, 2H); 2.20 (q, 2H, J=7.3 Hz); 1.85 (m, 2H); 1.61 (m, 4H); 1.47 (m, 4H); 1.32-1.25 (m, 8H); 0.87 (dt, 3H, J=7.0 Hz). The 1H spectrum and proposed assignments of diagnostic peaks are presented in FIG. 12A: 11.1 (COOH); 7.06 (C9 or C10, alkene proton, each a triplet from coupling to neighboring methylene —CH2, with regioisomers superimposed on each other, appearing on one NMR spectrometer as a doublet of triplets and on the other as a quartet, which is really a superimposed pair of triplets); 3.75 (C8 or C11, allylic methylene neighboring nitro group; nitroalkene more electron-withdrawing than carbonyl); 2.55 (C2 methylene neighboring carbonyl); 2.20 (C8 or C11, allylic methylene); 1.85 (C7 or C12, methylene next to nitro-allyl position); 1.61 (C3 methylene); 0.87 (C18 terminal methyl, superimposed regioisomers resulting in doublet of triplets). There is no indication of oleic acid or synthetic intermediates and no trans-isomers of OA-NO2 in the proton spectrum. The local environment of each proton in the 9- and 10-nitrated cis-isomers is not distinguishable, so each signal is indicative of relative positions in the alkene region, and absolute positions in the extreme portions of the molecule (i.e., the terminal methyl and acid groups). The aliphatic regions are too similar for exact assignments.
Further structural characterization was performed by 13C NMR. From the spectrum, 30 total peaks were observed: δ180.1, 180.0; 152.2, 152.0; 136.8, 136.4; 68.1; 34.2; 32.0; 29.6; 29.5; 29.5; 29.5; 29.4; 29.4; 29.3; 29.2; 29.1; 28.8; 28.7; 28.3; 28.2; 28.1; 28.1; 26.6; 26.6; 25.8; 24.8; 22.9; 14.3. The proposed assignment of diagnostic peaks is presented in FIG. 12B: 180.1, 180.0 (C1 carbonyl); 152.2, 152.0 (C9 or C10, alkene attached to NO2); 136.7, 136.4 (C9 or C10, alkene); 68.2 (C8 or C11, CH2 allylic to NO2); 34.2 (C2, CH2 neighboring carbonyl); 32.1 (C8 or C11, allylic distal to NO2); 14.3 (C18, terminal CH3). The total integrated signal is less than the total number of carbons for the two isomers (i.e., 30 vs. 36); thus, some peaks are identifiable doublets from isomers (e.g., carbonyl peaks), whereas others are indistinguishable and appear as a singlet (e.g., the methyl peak).
Spectral Characterization of OA-NO2.
The spectrum of OA-NO2 was acquired in phosphate buffer in the presence of the iron chelator DTPA (FIG. 13A). An absorbance maximum at 270 nm was identified that is ascribed to electron absorption by the NO2 group. Extinction coefficients for OA-NO2 and [13C]OA-NO2 were determined by plotting absorbance (λ270) vs. concentration, giving m=AU·cm−1·mM−1 and a calculated ε=8.22 and 8.23 cm−1 mM−1, for OA-NO2 and [13C]OA-NO2, respectively (FIG. 13B).
Characterization of OA-NO2 by GC-MS/MS.
Capillary columns used for gas chromatography have high resolving capacity and can separate regio- and diasteroisomers. Thus, initial mass spectrometric characterization of OA-NO2 regioisomer methyl esters was performed by GC-MS (FIG. 14). Filtering total ion count (TIC) data to show ions with m/z 342 ([M+H]+) and 306 (source fragment of m/z 342; [M−2H2O]+) reveals two peaks appearing at 37.4-37.7 min (FIG. 14A, upper panel). Product ion analysis of these peaks generated an identifying spectrum for each species (FIG. 14B). Ions with m/z 324 and 306 represent the loss of one and two H2O from the parent compound. The unique product ions m/z 168 and 156 for the first and second eluting peaks, respectively, were chosen as identifiers of each regioisomer. Filtering the MS/MS TIC for these product ions generated chromatograms that show the individual regioisomers of OA-NO2 (FIG. 14A, middle and lower panels).
Characterization and Quantitation of OA-NO2 by ESI-MS/MS.
Using the gradient HPLC protocol described in Methods, synthetic OA-NO2 regioisomers eluted from the reverse phase column as two overlapping peaks (FIG. 15). The HPLC elution profiles for synthetic OA-NO2 and [13C]OA-NO2 were virtually identical (FIG. 15A, left panels). Concurrent product ion analysis of the overlapping peaks showed spectra consistent with OA-NO2-derived species (FIG. 15A, right panels), with major fragments identified in Table 3. Using these same parameters and machine settings, lipid extracts of packed red cells and plasma were analyzed (FIG. 15B). The product ion spectra for OA-NO2 of red cells and plasma are identical to those obtained from synthetic OA-NO2, revealing that OA-NO2 is endogenously present in blood. Interestingly, the HPLC elution profiles for plasma- and blood-derived OA-NO2 show single peaks rather than overlapping species, suggesting only one regio-isomer is present in vivo.







TABLE 3







Collision-induced dissociation fragments of


nitroalkene fatty acid derivatives


Nitroalkene derivatives of fatty acids were analyzed by


electrospray-ionization tandem mass spectrometry. Product


ion spectra from synthetic standards were obtained in the


negative ion mode as described in Experimental Procedures.


Major fragments generated for each standard are listed below.










Mass/





charge


(m/z)
OA-NO2
[13C]OA-NO2
LNO2





344
—
[M-H]
—


326
[M-H]
[M-H2O]
—


324
—
—
[M-H2O]


308
[M-H2O]
[M-2H2O]
—


306
—
—
[M-H2O]


295
—
[M-HNO2]
—


293
—
—
[M-HNO]


279
[M-HNO2]
—
—


277
—
—
[M-HNO2]


264
—
[M-(2H2O + CO2)]
—


246
[M-(2H2O + CO2)]
—
—


244
—
—
[M-2H2O + CO2)]









To quantitate OA-NO2 content in red cells and plasma, lipid extracts were separated using an isocratic HPLC elution protocol wherein analytes co-elute at 2 min; MRM transitions for OA-NO2 and [13C]OA-NO2 were monitored (not shown). The concentration of OA-NO2 in biological samples was determined from the ratio of analyte to internal standard peak areas using an internal standard curve that is linear over four orders of magnitude. The limit of quantitation (LOQ; determined as ten times the standard deviation of the noise) was calculated to be  ̃1.2 fmol on column (not shown). Blood samples obtained from ten healthy human volunteers (5 female, 5 male, ages ranging from 24 to 51) 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 4). 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 exists as esterified fatty acids (9). In plasma, the free and esterified OA-NO2 concentrations were 619±52 and 302±369 nM, respectively, and were observed to be more abundant than linoleic acid nitration products (9).







TABLE 4







Nitrated Oleic Acid in human blood a comparison


with nitrated linoleic acid


Venous blood was obtained from healthy human volunteers and


centrifuged, and plasma and red cells were extracted and prepared


for mass spectrometric analysis as described in Experimental Procedures.


During sample preparation, [13C]OA-NO2 was added as an internal


standard to correct for losses. OA-NO2 was quantitated by fitting analyte


to internal standard area ratios obtained by MS to an internal standard


curve. Concentration values for LNO2 in the vascular compartment


were obtained from ref. 9. Data are expressed as mean ± std dev


(n − 10; 5 female and 5 male)










Compartment
Fraction
[OA-NO2] (nM)
[LNO2] (nM)





Plasma
Free
619 ± 52 
79 ± 35



Esterified
302 ± 369
550 ± 275



Total
921 ± 421
630 ± 240


Packed red cells
Free
59 ± 11
50 ± 17



Esterified
155 ± 65 
199 ± 121



Total
214 ± 76 
249 ± 104


Whole Blood*
Total
 639 ± 366*
 477 ± 128*





*Assuming a 40% hematocrit






Characterization of Nitrohydroxy Allylic Derivatives.

The initial survey to detect nitrated fatty acids indicated that nitrohydroxy allylic derivatives are also present in plasma and urine (FIG. 11). Their presence was confirmed by product ion analysis run concomitantly with MRM detection (FIG. 16). Structures of possible adducts are presented with diagnostic fragments and product ion spectra for 18:1(OH)—NO2, 18:2(OH)—NO2 and 18:3(OH)—NO2. Both the 9- and 10-nitro regioisomers of 18:1(OH)—NO2 are present in urine (A) and plasma (not shown), as evidenced by the intense peaks corresponding to m/z 171 and 202 (A). Also present are fragments consistent with the loss of a nitro group and water (m/z 297 and 326, respectively). The product ion spectrum obtained from 18:2(OH)—NO2 shows a predominant fragment (m/z 171), consistent with an oxidation product of LNO2 nitrated at the 10-carbon (B). Diagnostic fragments for the three other potential regioisomers were not apparent. Finally, multiple regioisomers of 18:3(OH)—NO2 are present (C). For all three Δ-9 fatty acids monitored, the predominant fragment generated during collision-induced dissociation (CID) is m/z 171.
Activation of PPARs by OA-NO2.
Recently, LNO2 has been identified as an endogenous PPAR ligand (17). Considering the high levels of OA-NO2 detected in vivo, OA-NO2 was evaluated and compared with LNO2 as a potential ligand for PPARα, PPARγ and PPARδ. CV-1 cells were transiently co-transfected with a plasmid containing the luciferase gene under regulation by three PPAR response elements (PPRE) in concert with PPARγ, PPARα or PPARδ expression plasmids. Dose-dependent activation by OA-NO2 was observed for all PPARs, (FIG. 17A), with PPARγ showing the greatest response (significant activation at 100 nM); PPARα and PPARδ showed significant activation at  ̃300 nM OA-NO2. Nitrated oleic acid was consistently more potent than LNO2 in the activation of PPARγ, with 1 μM OA-NO2 typically inducing the same degree of reporter gene expression as 3 μM LNO2 or 1 μM Rosiglitazone; these increases were partially inhibited by the PPARγ antagonist GW9662 (FIG. 17B). Native fatty acids did not activate PPARs at these concentrations (not shown). The greater potency of OA-NO2 as a PPARγ agonist compared to LNO2 motivated evaluation of the relative stability of these molecules, because LNO2 decays in aqueous milieu to generate products that do not activate PPARs (17,24). Compared with LNO2, OA-NO2 is relatively stable, with only minimal decay occurring after 2 hr; a time when  ̃80% LNO2 decay was noted (FIG. 17C).
The signaling actions of OA-NO2 as a PPARγ ligand was assessed by evaluating its impact on adipocyte differentiation, as PPARγ-dependent gene expression plays an essential role in the development of adipose tissue (25,26). 3T3-L1 preadipocytes were treated with OA-NO2 (1 μM), LNO2 (3 μM) and negative controls for two weeks (FIG. 18A). Previously, LNO2 was shown to induce pre-adipocyte differentiation to the same extent as Rosiglitazone (17). Adipocyte differentiation was assessed both morphologically and via oil red O staining, which revealed the accumulation of intracellular lipids. Vehicle, oleic acid and linoleic acid did not induce adipogenesis. In contrast, OA-NO2 and LNO2 induced >30% of 3T3-L1 preadiopcyte differentiation. Rosiglitazone, a synthetic PPARγ ligand, also induced PPARγ-dependent preadiopcyte differentiation. OA-NO2- and Rosiglitazone-induced pre-adipocyte differentiation resulted in expression of specific adipocyte markers (PPARγ2 and aP2); oleic acid had no effect on these gene products (FIG. 18B). PPARγ ligands also play a central role in glucose uptake and metabolism, with agonists widely used as insulin-sensitizing drugs. Consistent with its potent PPARγ ligand activity, OA-NO2 induced an increase in differentiated adipocyte glucose uptake (FIG. 19A). This effect of OA-NO2 was 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. 19B). In aggregate, these observations reveal that OA-NO2 manifests well-characterized PPARγ-dependent signaling actions.
DISCUSSION
The nitration of hydrocarbons has long been recognized (27). Following the more recent discovery of vascular cell signaling actions of oxides of nitrogen (1,28), it is now also appreciated that .NO-derived species mediate oxidation, nitrosation, nitrosylation and nitration reactions of protein, DNA and unsaturated fatty acids (29). These reactions frequently yield stable products that influence target molecule structure and function to either a) translate the signaling actions of .NO or b) mediate pathogenic responses when occurring in “excess”.
The reactions of .NO and its redox-derived products with lipids is multifaceted. Model studies of photochemical air pollutant-induced lipid oxidation revealed that exceedingly high concentrations of nitrogen dioxide (.NO2) could induce nitration of fatty acids in phosphatidylcholine liposomes and fatty acid methyl ester preparations (30-32). Subsequently, reaction systems designed to model the interactions of endogenous .NO and .NO-derived species [e.g., peroxynitrite (ONOO−) and nitrous acid (HNO2)] with fatty acids showed that a) .NO mediates potent inhibition of autocatalytic radical chain propagation reactions of lipid peroxidation (33,34) and b) .NO-derived species produce both nitrated and oxidized derivatives of unsaturated fatty acids (3,35). 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 mammals and plants, 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. 10) was defined on the basis of the synthetic rationale and NMR analysis (FIG. 12). Proton and 13C NMR spectra indicate that the 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 nitro-allylic and allylic carbons in the 13C spectrum both appear as doublets that are equal in intensity, indicating an equivalent distribution between regioisomers. EI GC MS confirmed the presence of 9- and 10-nitro-9-cis-octadecenoic acids by mass and differential retention times (FIG. 14). HPLC ESI MS/MS was used to further characterize synthetic OA-NO2. The combined fragmentation pattern of OA-NO2 regioisomers was obtained by CID, which provided a “molecular fingerprint” that was used to identify OA-NO2 in biological samples (FIG. 15). 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; however, it is not possible from MS analysis to determine the cis/trans conformation of regioisomers. Quantitative analysis of plasma and red cells revealed that OA-NO2 is present in the vasculature at net concentrations  ̃50% greater than LNO2 (Table 4). Combined, esterified and free OA-NO2 and LNO2 are well above 1 μM, a concentration range capable of eliciting cell signaling responses.
The nitro 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 these nitroalkene derivatives via a modified Nef reaction (24). Furthermore, the α-carbon proximal to the alkenyl nitro group is strongly electrophilic, which readily reacts with H2O via a Michael addition mechanism to generate nitrohydroxy adducts (FIGS. 11 and 16). Nitrohydroxy-arachidonic acid species have previously been detected in bovine cardiac muscle (36), and nitrohydroxy-linoleic acid has been identified in lipid extracts obtained from hypercholesterolemic and post-prandial human plasma, suggesting that this is a ubiquitous derivative (37). 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. 11, and 16). The hydroxyl moiety of nitrohydroxy-fatty acids destabilizes the adjacent carbon-carbon bond, resulting in heterolytic scission reactions that a) generate predictable fragments during CID (FIG. 16) and b) render potentially unique cell signaling actions to fatty acid nitrohydroxy derivatives. Present data indicates, however, that nitrohydroxy adducts of LNO2 and OA-NO2 are not avid ligands for PPARγ (FIG. 17C).
Multiple mechanisms can support the basal and inflammatory nitration of fatty acids by .NO-derived species, including .NO2-initiated auto-oxidation of polyunsaturated fatty acids via hydrogen abstraction from the bis-allylic carbon and nitration by acidified NO2− (31,38-42). Of relevance to both basal and inflammatory cell signaling, .NO2 can be derived from multiple reactions. This includes the homolytic scission of both peroxynitrous acid (ONOOH) and nitrosoperoxocarbonate (ONOOCO2−), as well as the oxidation of NO2− by heme peroxidases (43,44). Present data supports that all of these alkenyl nitration mechanisms can yield nitrated fatty acids that are structurally similar or identical to the OA-NO2 and LNO2 detected clinically. 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 to free radical-mediated hydrogen abstraction reactions. In view of the data presented herein, alternative fatty acid nitration mechanisms may thus also be viable. For example, nitration by an ionic addition reaction (e.g., nitronium ion, NO2+) can generate singly nitrated fatty acids with no double bond-rearrangement. Since NO2+ readily reacts with H2O, this species may require stabilization in the hydrophobic milieu of the membrane bilayer or localized catalysis (e.g., reaction of ONOO— with transition metals) to serve as a biologically-relevant nitrating species. Finally, radical addition reactions may also occur with mono- and polyunsaturated fatty acids to yield non-conjugated nitroalkene derivatives of polyunsaturated fatty acids, as revealed by studies of acidified NO2− and .NO2-mediated fatty acid methyl ester oxidation and nitration profiles (32,41).
Of important relevance to mechanisms underlying fatty acid nitration in vivo, the nitrohydroxy adducts of Δ9 unsaturated fatty acids examined in the present study (18:1, 18:2 and 18:3) all have a predominant fragment of m/z 171 (FIG. 16). This mass is consistent with 9-oxo-nonanoic acid, a fragment generated with authentic standards when the nitro group is located at the 10-carbon and the hydroxy moiety at the 9-carbon (data not shown). This suggests either strict steric control or enzymatic mechanisms regulating the stereospecificity of biological fatty acid nitration. The nitration of Δ9 unsaturated fatty acids to C10 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 that give rise to specific fatty acid nitroalkene derivatives.
Designation of nitroalkene derivatives as a class of signaling molecules is contingent upon ascribing specific bioactivities to multiple members within the class at clinically-relevant concentrations. Nitrolinoleate has been observed to inhibit neutrophil and platelet function via cGMP-independent, cAMP-mediated mechanisms (10-12). Also, aqueous decay of LNO2 yields .NO, a reaction that is facilitated by translocation of LNO2 from a hydrophilic to hydrophobic microenvironment, which in turn induces cGMP-dependent vessel relaxation (12,24). Recently, LNO2 has also been reported to serve as a robust ligand for PPARγ (17), a nuclear hormone receptor that binds lipophilic ligands to induce DNA binding of the transcription factor complex at DR1-type motifs in the promoter sites of target genes. Downstream effects of PPARγ activation include modulation of metabolic and cellular differentiation genes and regulation of inflammatory responses, adipogenesis and glucose homeostasis (45,46). In the vasculature, PPARγ is expressed in monocytes, macrophages, smooth muscle cells and endothelium (47) 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 PPARγ, α and δ ligand that rivals or exceeds the potency of LNO2 and synthetic PPAR ligands such as fibrates and thiazolidinediones (FIGS. 17-19). The increased potency of OA-NO2 as a PPARγ ligand is either due to increased aqueous stability relative to LNO2 (FIG. 17C) or increased receptor affinity. The combined blood concentrations of OA-NO2 and LNO2 in healthy humans exceeds 1 μM (Table 4), concentrations well-within the range needed to activate PPAR receptors and exceeding those of previously-proposed endogenous PPARγ ligands. These observations have broad implications for the NO and redox signaling reactions that play a crucial role in dysregulated cell growth and differentiation, metabolic syndrome, atherosclerosis and diabetes—all clinical pathologies that also include a significant contribution from PPAR-regulated cell signaling mechanisms (48).
The regulation of inflammation by inhibiting eicosanoid synthesis is a well-established and prevalent target of anti-inflammatory drug strategies. Much less well-understood are the concerted cell signaling mechanisms by which inflammation is favorably resolved in vivo. While the integrated in vivo tissue signaling activities of nitrated fatty acids remain to be defined, studies to date indicate that these pluripotent signaling mediators generally manifest salutary metabolic and anti-inflammatory actions (10-12,17). The capability of redox-derived lipid signaling molecules to mediate the resolution of inflammation is a relatively new concept, with lipoxins representing one new class of lipid mediators that may also act in this manner (49). Endogenous concentrations of OA-NO2 and LNO2 are abundant and are increased by oxidative inflammatory reactions. Thus, nitrated fatty acids are expected to play both receptor-dependent (via PPAR ligand activity) and cyclic nucleotide-mediated roles in transducing the redox signaling actions of oxygen and .NO, thereby regulating organ function, cell differentiation, cell metabolism and systemic inflammatory responses.
Throughout Example 3, several publications have been referenced. These publications are listed in the Reference List for Example 3. The disclosures of these publications in their entireties are hereby incorporated by reference into this application in order to more fully describe the compounds, compositions and methods described herein.
REFERENCE LIST FOR EXAMPLE 3
Reference List


    1. Smith, W. L. (1992) Am. J Physiol 263, F181-F191
    2. Montuschi, P., Barnes, P. J., and Roberts, L. J. (2004) FASEB J 18, 1791-1800
    3. Rubbo, H., Radi, R., Trujillo, M., Telleri, R., Kalyanaraman, B., Barnes, S., Kirk, M., and Freeman, B. A. (1994) J. Biol. Chem. 269, 26066-26075
    4. Schopfer, F. J., Baker, P. R., and Freeman, B. A. (2003) Trends Biochem. Sci. 28, 646-654
    5. Marshall, H. E., Merchant, K., and Stamler, J. S. (2000) FASEB J 14, 1889-1900
    6. Vidwans, A. S., Uliasz, T. F., Hewett, J. A., and Hewett, S. J. (2001) Biochemistry 40, 11533-11542
    7. Larfars, G., Lantoine, F., Devynck, M. A., Palmblad, J., and Gyllenhammar, H. (1999) Blood 93, 1399-405
    8. Rubbo, H., Radi, R., Anselmi, D., Kirk, M., Barnes, S., Butler, J., Eiserich, J. P., and Freeman, B. A. (2000) J. Biol. Chem. 275, 10812-10818
    9. Baker, P. R., Schopfer, F. J., Sweeney, S., and Freeman, B. A. (2004) Proc. Natl. Acad. Sci. U.S.A 101, 11577-11582
    10. Coles, B., Bloodsworth, A., Eiserich, J. P., Coffey, M. J., McLoughlin, R. M., Giddings, J. C., Lewis, M. J., Haslam, R. J., Freeman, B. A., and O'Donnell, V. B. (2002) J. Biol. Chem. 277, 5832-5840
    11. Coles, B., Bloodsworth, A., Clark, S. R., Lewis, M. J., Cross, A. R., Freeman, B. A., and O'Donnell, V. B. (2002) Circ. Res. 91, 375-381
    12. Lim, D. G., Sweeney, S., Bloodsworth, A., White, C. R., Chumley, P. H., Krishna, N. R., Schopfer, F., O'Donnell, V. B., Eiserich, J. P., and Freeman, B. A. (2002) Proc. Natl. Acad. Sci. U.S.A 99, 15941-15946
    13. Gladwin, M. T., Shelhamer, J. H., Schechter, A. N., Pease-Fye, M. E., Waclawiw, M. A., Panza, J. A., Ognibene, F. P., and Cannon, R. O., III (2000) Proc. Natl. Acad. Sci. U.S.A 97, 11482-11487
    14. Rassaf, T., Bryan, N. S., Kelm, M., and Feelisch, M. (2002) Free Radic. Biol. Med. 33, 1590-1596
    15. Rassaf, T., Bryan, N. S., Maloney, R. E., Specian, V., Kelm, M., Kalyanaraman, B., Rodriguez, J., and Feelisch, M. (2003) Nat. Med. 9, 481-482
    16. Cosby, K., Partovi, K. S., Crawford, J. H., Patel, R. P., Reiter, C. D., Martyr, S., Yang, B. K., Waclawiw, M. A., Zalos, G., Xu, X., Huang, K. T., Shields, H., Kim-Shapiro, D. B., Schechter, A. N., Cannon, R. O., III, and Gladwin, M. T. (2003) Nat. Med. 9, 1498-1505
    17. Schopfer, F. J., Lin, Y., Baker, P. R., Cui, T., Garcia-Barrio, M., Zhang, J., Chen, K., Chen, Y. E., and Freeman, B. A. (2005) Proc Natl Acad Sci USA 102, 2340-2345
    18. Lee, C. H. and Evans, R. M. (2002) Trends Endocrinol. Metab 13, 331-335
    19. Marx, N., Duez, H., Fruchart, J. C., and Staels, B. (2004) Circ. Res. 94, 1168-1178
    20. Dodge, J. T. and Phillips, G. B. (1967) J Lipid Res. 8, 667-675
    21. Bligh, E. G. and Dyer, W. L. (1959) Can. J. Biochem. Physiol. 37, 911-917
    22. Zhang, J., Fu, M., Cui, T., Xiong, C., Xu, K., Zhong, W., Xiao, Y., Floyd, D., Liang, J., Li, E., Song, Q., and Chen, Y. E. (2004) Proc. Natl. Acad. Sci. U.S.A 101, 10703-10708
    23. Mukherjee, R., Hoener, P. A., Jow, L., Bilakovics, J., Klausing, K., Mais, D. E., Faulkner, A., Croston, G. E., and Paterniti, J. R., Jr. (2000) Mol. Endocrinol. 14, 1425-1433
    24. Schopfer, F. J., Baker, P. R., Giles, G., Chumley, P., Batthyany, C., Crawford, J., Patel, R. P., Hogg, N., Branchaud, B. P., Lancaster, J. R., Jr., and Freeman, B. A. (2005) J Biol Chem 
    25. Evans, R. M., Barish, G. D., and Wang, Y. X. (2004) Nat. Med. 10, 355-361
    26. Zhang, J., Fu, M., Cui, T., Xiong, C., Xu, K., Zhong, W., Xiao, Y., Floyd, D., Liang, J., Li, E., Song, Q., and Chen, Y. E. (2004) Proc Natl Acad Sci USA 101, 10703-10708
    27. Zelinsky, N. D. and Rosanoff, M. A. (1912) Z. Physikal Chem. 78, 629-633
    28. Ignarro, L. J., Byrns, R. E., Buga, G. M., Wood, K. S., and Chaudhuri, G. (1988) J Pharmacol. Exp. Ther. 244, 181-189
    29. Miranda, K. M., Espey, M. G., Jourd'heuil, D., Grisham, M. B., Fukoto, J. M., Feelisch, M., and Wink, D. A. (2000) The Chemical Biology of Nitric Oxide. In Ignarro, L. J., editor. Nitric Oxide: Biology and Pathobiology, Academic Press, San Diego
    30. Finlayson-Pitts, B. J., Sweetman, L. L., and Weissbart, B. (1987) Toxicol. Appl. Pharmacol. 89, 438-448
    31. Gallon, A. A. and Pryor, W. A. (1993) Lipids 28, 125-133
    32. d'Ischia, M., Rega, N., and Barone, V. (1999) Tetrahedron 55, 9297-9308
    33. O'Donnell, V. B., Chumley, P. H., Hogg, N., Bloodsworth, A., Darley-Usmar, V. M., and Freeman, B. A. (1997) Biochemistry 36, 15216-15223
    34. Hogg, N., Kalyanaraman, B., Joseph, J., Struck, A., and Parthasarathy, S. (1993) FEBS Lett. 334, 170-174
    35. Rubbo, H., Parthasarathy, S., Barnes, S., Kirk, M., Kalyanaraman, B., and Freeman, B. A. (1995) Arch. Biochem. Biophys. 324, 15-25
    36. Balazy, M., Iesaki, T., Park, J. L., Jiang, H., Kaminski, P. M., and Wolin, M. S. (2001) J. Pharmacol. Exp. Ther. 299, 611-619
    37. Lima, E. S., Di Mascio, P., Rubbo, H., and Abdalla, D. S. (2002) Biochemistry 41, 10717-10722
    38. Baldus, S., Castro, L., Eiserich, J. P., and Freeman, B. A. (2001) Am. J. Respir. Crit Care Med. 163, 308-310
    39. Gallon, A. A. and Pryor, W. A. (1994) Lipids 29, 171-176
    40. Napolitano, A., Camera, E., Picardo, M., and d'Ischia, M. (2000) J. Org. Chem. 65, 4853-4860
    41. Napolitano, A., Crescenzi, O., Camera, E., Giudicianni, I., Picardo, M., and d'Ischia, M. (2004) Tetrahedron 58, 5061-5067
    42. O'Donnell, V. B., Eiserich, J. P., Chumley, P. H., Jablonsky, M. J., Krishna, N. R., Kirk, M., Barnes, S., Darley-Usmar, V. M., and Freeman, B. A. (1999) Chem. Res. Toxicol. 12, 83-92
    43. Beckman, J. S., Beckman, T. W., Chen, J., Marshall, P. A., and Freeman, B. A. (1990) Proc. Natl. Acad. Sci. U.S.A 87, 1620-1624
    44. Castro, L., Eiserich, J. P., Sweeney, S., Radi, R., and Freeman, B. A. (2004) Arch. Biochem. Biophys. 421, 99-107
    45. Lee, C. H. and Evans, R. M. (2002) Trends Endocrinol. Metab 13, 331-335
    46. Marx, N., Duez, H., Fruchart, J. C., and Staels, B. (2004) Circ. Res. 94, 1168-1178
    47. Wang, N., Verna, L., Chen, N. G., Chen, J., Li, H., Forman, B. M., and Stemerman, M. B. (2002) J. Biol. Chem. 277, 34176-34181
    48. Li, A. C., Binder, C. J., Gutierrez, A., Brown, K. K., Plotkin, C. R., Pattison, J. W., Valledor, A. F., Davis, R. A., Willson, T. M., Witztum, J. L., Palinski, W., and Glass, C. K. (2004) J Clin. Invest 114, 1564-1576
    49. Levy, B. D., Clish, C. B., Schmidt, B., Gronert, K., and Serhan, C. N. (2001) Nat. Immunol. 2, 612-619


Example 4
The aqueous decay and concomitant release of nitric oxide (.NO) by nitrolinoleic acid (10-nitro-9,12-octadecadienoic acid and 12-nitro-9,12-octadecadienoic acid; LNO2) is reported. Mass spectrometric analysis of reaction products support the Nef reaction as the mechanism accounting for the generation of .NO by the aqueous reactions of fatty acid nitroalkene derivatives. Nitrolinoleic acid is stabilized by aprotic milieu, with LNO2 decay and .NO release strongly inhibited by phosphatidylcholine-cholesterol liposome membranes and detergents when present at levels above their critical micellar concentrations. The release of .NO from LNO2 was induced by the UV photolysis and I3-based ozone chemiluminescence reactions currently being used to quantify putative protein nitrosothiol (RSNO) and N-nitrosamine derivatives. This reactivity of LNO2 complicates the qualitative and quantitative analysis of biological oxides of nitrogen when applying UV photolysis and triiodide (I3−)-based analytical systems in biological preparations typically abundant in nitrated fatty acids. These results reveal that nitroalkene derivatives of linoleic acid are pluripotent signaling mediators that act via not only receptor-dependent mechanisms, but also by transducing the signaling actions of .NO via pathways subject to regulation by the relative distribution of LNO2 to hydrophobic versus aqueous micro environments.
Nitrolinoleic acid (10-nitro-9,12-octadecadienoic acid and 12-nitro-9,12-octadecadienoic acid; abbreviated as LNO2) is present in plasma lipoproteins and red blood cell membranes at concentrations of  ̃500 nM, rendering this species the most quantitatively abundant biologically-active oxide of nitrogen in the human vascular compartment (1). Nitrolinoleic acid is a product of nitric oxide (.NO)-dependent linoleic acid nitration reactions that predominantly occur at the C10 and C12 alkene carbons. The positional isomer distribution of the LNO2 alkenyl nitro group indicates that in vivo fatty acid nitration is a consequence of nucleophilic (nitronium group, NO2+) and/or radical (nitrogen dioxide, .NO2) addition reactions with olefinic carbons.
Recent observations reveal that LNO2 is a pluripotent signaling mediator that acts via both receptor-dependent and -independent pathways. Nitrated fatty acids are specific and high affinity endogenous ligands for peroxisome proliferator-activated receptors (2), and serve to activate receptor-dependent gene expression at physiological concentrations. LNO2 also activates cAMP-dependent protein kinase signaling pathways in neutrophils and platelets, serving to down-regulate the activation of these inflammatory cells (3,4). Finally, LNO2 induces vessel relaxation in an endothelial-independent manner (5). This LNO2-mediated relaxation of phenylephrine-preconstricted aortic rings was 1) a consequence of LNO2-induced stimulation of smooth muscle cell and aortic segment cGMP content, 2) inhibitable by the NO scavenger oxyhemoglobin and 3) ODQ-inhibitable (e.g., guanylate cyclase-dependent). Although these vessel responses to LNO2 suggest .NO as the mediator of guanylate cyclase activation, the identity of the proximal LNO2-derived, cGMP-dependent signaling molecule was not directly identified (5).
Nitric oxide, synthesized by three different nitric oxide synthase isoforms, was first shown to mediate endothelial-dependent relaxation via reaction with the heme iron of guanylate cyclase and subsequent activation of cGMP-dependent protein kinases (6). Subsequent to this discovery, there has been a growing appreciation that the cell signaling actions of .NO are also transduced by secondary products derived from redox reactions of .NO. These redox reactions yield a variety of oxides of nitrogen displaying both unique and overlapping reactivities that can regulate differentiated cell function via both cGMP- and non-cGMP-dependent mechanisms. These products include nitrite (NO2−), .NO2, peroxynitrite (ONOO−), nitrosothiols (RSNO) and dinitrogen trioxide (N2O3). These reactive species serve to transduce the cell signaling actions of .NO by inducing changes in target molecule structure and function via oxidation, nitration or nitrosation reactions (7,8).
The lipophilicity and intrinsic chemical reactivities of .NO facilitate multiple interactions with lipids that impact both cellular redox and .NO signaling reactions. For example, .NO concentrates in membranes and lipoproteins, where it more readily reacts with oxygen to yield oxidizing, nitrosating and nitrating species such as N2O3 and N2O4 (9-11). In these lipophilic compartments, .NO can react with lipid peroxyl radicals (LOO.) at diffusion-limited rates, readily out-competing tocopherols and ascorbate for the scavenging of intermediates that would otherwise propagate lipid oxidation. In this regard, .NO displays an oxidant-protective, anti-inflammatory role (12,13). Of relevance to inflammatory signaling, heme and non-heme-containing peroxidases and oxygenases that catalyze physiologic and pathologic fatty acid oxygenation reactions also catalytically consume .NO during enzyme turnover [e.g., lipoxygenases (14,15), cyclooxygenase (16) and myeloperoxidase (17)]. The reaction of .NO with these enzymatic catalysts and free radical intermediates of fatty acid oxygenation in turn inhibits rates of fatty acid oxygenation product formation. The convergence of .NO and fatty acid oxygenation reactions thus can influence the steady state concentration of both .NO and eicosanoids in a concerted fashion.
Redox reactions of .NO frequently induce the chemical modification of target molecules, including the nitrosylation (addition of .NO) of heme proteins (18), the nitrosation (addition of the nitroso group .NO) of thiol substituents (7) and the nitration (addition of the nitro group NO2) of protein tyrosine residues and DNA bases (8). Herein, the present invention shows that LNO2, a product of .NO-dependent unsaturated fatty acid nitration reactions that is abundant in red cells and plasma, decays in aqueous milieu to release .NO. This generation of .NO by LNO2 is inhibited by aprotic environments, a milieu that concomitantly stabilizes LNO2. Moreover, it is shown that UV photolysis and I3−-based chemiluminescence approaches currently used to quantify .NO derived from protein heme-nitrosyl, RSNO and N-nitrosamine (RNNO) derivatives, also facilitate .NO release from LNO2. This complicates the interpretation of quantitative and qualitative results from the application of these analytical systems in biological preparations. In aggregate, these results reveal that nitroalkene derivatives of fatty acids serve to transduce the signaling actions of .NO via pathways subject to regulation by the relative distribution of LNO2 to hydrophobic versus aqueous microenvironments.
Materials—
Horse heart myoglobin, octyl-β-glucopyranoside (OG) and octyl-thio-β-glucopyranoside (OTG), carboxy 2-phenyl-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO), diethylenetriaminepentaacetate (DTPA), Na2HPO4 and sodium dithionite were from Sigma (St Louis, Mo.). LNO2 and [13C]LNO2 were synthesized and purified as previously described (1). Metmyoglobin was reduced using sodium dithionite, desalted by exclusion chromatography on a Sephadex PD-10 column and further oxygenated by equilibration with 100% oxygen.
Electron Paramagnetic Resonance—
EPR measurements were performed at room temperature using a Bruker Elexsys E-500 spectrometer equipped with an ER049X microwave bridge and an AquaX liquid sample cell. The following instrument setting were used: modulation frequency, 100 kHz; modulation amplitude, 0.05 G; receiver gain, 60 dB; time constant, 1.28 ms; sweep time, 5.24 s; center field, 3510 G; sweep width, 100 G; power, 20 mW; scan parameter, 16 scans.
Spectrophotometry—
The UV spectrum of LNO2 and repetitive scans of LNO2 decay kinetics were collected using a Hitachi UV 2401 PC spectrophotometer. Apparent .NO formation was calculated from extents of oxymyoglobin oxidation in the visible wavelength range (spectrum) and at 580 nm (kinetic mode). Initial oxymyoglobin oxidation was calculated using a UV probe Version 1.10 (ε580 14.4 mM−1 cm−1). Decomposition of the NO2 group was followed at 268 nm and the appearance of oxidized products at 320 nm.
Liposome Preparation—
Reverse phase evaporation liposomes were formed from dipalmitoylphosphatidylcholine (DPPC), cholesterol and stearylamine (4:2:1 mole ratio) following an established procedure (19). Briefly DPPC, cholesterol and stearylamine were dissolved in CHCl3 and sonicated with 10 mM KPi buffer (CHCl3/KPi, 2:1, v/v). The organic solvent was then removed by evaporation under reduced pressure at 45° C. The liposomes were allowed to anneal for 12 h at room temperature then centrifuged and the pellet resuspended in the experimental buffer.
Chemiluminescence and UV Photolysis Analyses—
For direct detection of NO release, LNO2 (75 μM) was incubated directly or with different additions (sulfanilamide, 1.5% w/v in 2 M HCl, 5 min, 25° C., with or without HgCl2, 50 mM) under aerobic conditions in a capped vial for 3 min. The gas phase was then injected into a chemiluminescence detector (ANTEK Instruments, Houston, Tex.). Additionally, known concentrations of DEA-NONOate (in 10 mM NaOH) were added to a capped vial containing 0.5 M HCl. NOx concentration profiles of plasma samples were performed by .NO chemiluminescence analysis. Measurement of putative NO2−, RSNO, and other .NO derivatives present in plasma was performed by using an I3−-based reducing system as previously (20,21). Rats were treated by intraperitoneal injection of 50 mg/kg E. coli LPS and 5 hr later blood was collected in EDTA anticoagulation tubes following cardiac puncture. Following removal of red cells by centrifugation (500 g×10 min), plasma samples were pretreated with either sulfanilamide (final concentration 1.5% w/v in 2 M HCl, 5 min, 25° C.) with or without HgCl2 (50 mM) prior to injecting into the chemiluminescence detector to measure NO2− and HgCl2-resistant NOx derivatives, respectively. For UV photolysis studies, a water-cooled reaction chamber was filled with 1 ml of phosphate buffer (50 mM, pH 7.4 containing 10 μM DTPA) and continuously bubbled with argon. The chamber was illuminated using an ILC PS300-1A xenon arc source (ILC Technology, Sunnyvale, Calif.). Samples were injected into the reaction chamber through an air-tight septum and released .NO was passed to the reaction chamber of a Sievers NOA 280 NO analyzer and detected by chemiluminescence after reaction with ozone (O3).
Mass Spectrometric Analysis—
LNO2 was extracted using the method of Bligh and Dyer (22). During extraction, [13C]LNO2 was added as internal standard and the LNO2 content of samples quantified using LC/MS/MS (1). Qualitative and quantitative analysis of LNO2 by ESI MS/MS was performed using a hybrid triple quadrupole-linear ion trap mass spectrometer (4000 Q trap, Applied Biosystems/MDS Sciex) as described (1). For the detection and characterization of L(OH)NO2, the hydration product of LNO2 generated by a Michael-like addition between H2O and the nitroalkene, LNO2 (3 μM) was incubated at 25° C. for 60 min in 100 mM phosphate buffer containing 100 μM DTPA pH 7.4 and extracted (Bligh and Dyer). L(OH)NO2 was detected using a multiple reaction monitoring (MRM) scan mode by reporting molecules that undergo an m/z 342/295 mass transition. This method selects (m/z 342) in the first quadrupole, consistent with the precursor ion, and following collision-induced dissociation (CID) yields in Q3 a species (m/z 295) consistent with loss of the nitro group ([M−(HNO2)]−). Presence of the nitrohydroxy-adduct was confirmed by product ion analysis of m/z 342. The degradation of LNO2 to secondary products was followed in negative ion mode after chloroform extraction and direct injection into an ion trap mass spectrometer with electrospray ionization (LCQ Deca, ThermoFinnigan).
Characterization of .NO Release from LNO2—
cPTIO is a selective spin trap for .NO (k=104 M−1 s−1, (23), with the product of this reaction, cPTI, displaying a characteristic EPR spectrum. In order to determine if .NO is derived from LNO2, it was incubated at 25° C. for different times in 100 mM phosphate buffer containing 100 μM DTPA, pH 7.4 in the presence of cPTIO. This resulted in a time-dependent decrease of the characteristic five peak cPTIO signal and the appearance of a new signal ascribed to cPTI (FIG. 20A). This release of .NO by LNO2 was concentration-dependent and followed first order decay kinetics for LNO2. Due to limitations of the .NO-cPTIO reaction for quantitating yields of .NO, oxymyoglobin was utilized to measure .NO release rates (24).
LNO2 was incubated with oxymyoglobin in 100 mM phosphate buffer containing 100 μM DTPA, pH 7.4 and .NO-dependent oxymyoglobin oxidation was followed spectrophotometrically. LNO2 oxidized oxymyoglobin in a dose- and time-dependent fashion, yielding metmyoglobin as indicated by the spectral changes depicted in FIG. 20B. The apparent rate constant for .NO release by LNO2, calculated from the oxidation of oxymyoglobin to metmyoglobin, was k=9.67×10−6 s−1 (FIG. 20C). To monitor the concomitant decomposition of the parent LNO2 molecule, its UV spectrum was first analyzed. LNO2 displays a characteristic absorbance spectrum with a peak at 268 nm, ascribed to the π electrons of the NO2 group. During aqueous LNO2 decay, this maximum decreases and a new maximum appears at 320 nm, corresponding to a mixture of oxygen and conjugated diene-containing products not yet fully characterized by mass spectroscopy (FIG. 20D). The decrease in absorbance at 268 nm paralleled .NO release, as detected by both EPR and oxymyoglobin oxidation.
Nitrite Formation During LNO2 Decomposition—
During LNO2-dependent .NO formation, measured via oxymyoglobin oxidation and MS analysis of LNO2 parent molecule loss in aqueous buffers, the stable .NO oxidation product NO2− accumulates with time (FIG. 21). The release of .NO from LNO2 was maximal at pH 7.4 (22), suggesting a role for protonation and deprotonation reactions in NO formation from LNO2.
Chemiluminescence Analysis of LNO2-Derived .NO—
Gas phase O3-mediated chemiluminescence detection of NO is a highly sensitive and specific method for detecting .NO. LNO2 was incubated in capped vials in 100 mM phosphate buffer, 100 μM DTPA, pH 7.4 in air and the gas phase directly injected into the detector. The .NO-dependent chemiluminescence yield was a function of concentration of DEA-NONOate and LNO2 concentrations, studied separately (FIG. 23A). Chemiluminescence was also time-dependent, increasing with time of LNO2 decay prior to gas sampling from vials.
UV photolysis has been used to quantitate RSNO derivatives of proteins and other NO-containing biomolecules (25,26). When LNO2 (4 nmol) was subjected to UV photolysis in concert with .NO chemiluminescence detection, UV light exposure stimulated .NO release from LNO2 (FIG. 23B). The .NO chemiluminescence response to NO2− (4 nmol) added to samples being subjected to UV photolysis and repetitive LNO2 addition was also examined to address the possibility that LNO2-derived NO2− formed during decay reactions might have accounted for some fraction of net chemiluminescent yield; it did not.
Appreciating that nitroalkene derivatives of red cell membrane and plasma fatty acids are present in human blood, whether LNO2-derived .NO has the potential to interfere with the chemiluminescent detection of NO2−, RSNO, RNNO or NO-heme compounds in plasma, when also analyzed via a triiodide (I3−)-based reaction system (21) was examined. Plasma from LPS-treated rats was used to exemplify this reaction system, since LPS treatment of rodents induces a robust elevation in plasma biomolecule NO-adduct levels (27). First, plasma was directly injected into the detector chamber and I3− reagent added, yielding a signal indicative of net plasma NO2−, RSNO, RNNO and NO-heme compounds (FIG. 23C, peak 1). Then, plasma treated with acidic sulfanilamide (which removes NO2−) was injected, giving a peak of lower intensity after I3− reagent addition, indicative of RSNO and putative RNNO derivatives (20,28). Finally, a plasma sample treated with acidic sulfanilamide and HgCl2 was injected, which resulted in an even smaller peak following I3− reagent addition (FIG. 23C, peak 3). This latter peak has been referred to as Hg-resistant RNNO derivatives (20,28). Using this strategy and combination of reagents, LNO2 pretreated with acidic sulfanilamide and HgCl2 also generated NO chemiluminescence for extended periods of time following I3− addition (FIG. 23C, LNO2 inset).
Hydrophobic Stabilization of LNO2—
The observation that LNO2 is stable in organic solvents such as n-octanol, undergoing decay only after solvation in aqueous solutions, led us to analyze rates of .NO formation from LNO2 in the presence of non-ionic detergents. The formation of .NO was followed by EPR (measuring cPTI formation) in the presence of different concentrations of octyl-β-glucopyranoside (OG) and octyl-thio-β-glucopyranoside (OTG). The rate of .NO release was constant and not influenced by these detergents until the critical micellar concentration (CMC) for each was achieved, after which .NO formation was inhibited as the volume of the hydrophobic environment increased (FIG. 24A). Similar results were obtained when measuring .NO formation via conversion of oxymyoglobin to metmyoglobin. Apparent .NO release rates remained constant until OG concentration reached  ̃2.8 mg/ml (CMC=2.77 mg/ml (11)). For OTG, inhibition of LNO2-dependent .NO release occurred at  ̃7 mg/ml (CMC=7.8 mg/ml (11)) (FIG. 24B). To further confirm that LNO2 was protected in lipophilic environments, LNO2 decomposition was followed by UV absorbance at 268 nm and 320 nm (FIGS. 24 C-D). The inhibition of the NO2 group loss at 268 nm was paralleled by inhibition of the formation of oxidation products at 320 nm, similarly paralleling the detergent-induced inhibition of .NO release observed by EPR and oxymyoglobin-based detection. The micellar stabilization of LNO2 was also documented in OTG-containing buffers by MS-based quantification of LNO2 after Bligh and Dyer extraction (FIG. 25A). Assuming rapid partitioning of LNO2 between the aqueous and hydrophobic compartments, and that LNO2 decay occurs only in the aqueous compartment, it can be shown that the rate of reaction v is given by the relationship:




  v
  =
  
    
      k
      ⁡
      
        [
        
          LNO
          2
        
        ]
      
    
    
      1
      +
      
        α
        ⁡
        
          (
          
            K
            -
            1
          
          )
        
      
    
  




where k is the rate constant for aqueous breakdown, a is the fraction of total volume that is the hydrophobic volume, and K (hydrophobic/aqueous concentration ratio) is the partition constant for LNO2. Thus, a plot of 1/v vs. α will yield a linear plot with slope divided by y-axis intercept equal to K-1. FIG. 25B shows this plot for OG and OTG, yielding values for K of 1580 and 1320 respectively.

For evaluating the stability of LNO2 in bilayers rather than micelles, phosphatidylcholine-cholesterol liposomes prepared by reverse phase evaporation were utilized (3:1). This alternative hydrophobic bilayer environment also resulted in a dose-dependent inhibition of the release of .NO from LNO2, as detected by EPR analysis of cPTI formation from cPTIO (FIG. 25C).
The decay of LNO2 in aqueous solutions results in the formation of multiple secondary fatty acid-derived products, as well as NO. One pathway that may be involved in aqueous LNO2 decay is the Michael-like addition reaction with H2O at the α carbon of the nitroalkene moiety. To test this possibility, and the influence of micellar stabilization of LNO2, the formation of nitrohydroxylinoleic acid (L(OH)NO2, m/z 342) was analyzed by MS. LNO2-derived L(OH)NO2 was evident after 60 min incubation in aqueous buffer at pH 7.4, with a concomitant decrease in LNO2 levels (m/z 324). Addition of OTG at a concentration above the CMC significantly decreased extents of L(OH)NO2 formation (FIG. 26A-C). Product ion analysis of L(OH)NO2 generated a pattern of CID product ions that indicate the presence of two predominant regioisomers consistent with the heterolytic scission products of L(OH)NO2, 9-hydroxy-10-nitro-12-octadecaenoic acid (m/z 171) and 12-hydroxy-13-nitro-9-octadecaenoic acid (m/z 211, FIG. 26 D-E).
The release of .NO by LNO2 via a modified Nef reaction mechanism was further supported by detecting an aqueous degradation product with m/z 293 (FIG. 27). This mass to charge ratio is consistent with the formation of a conjugated ketone (Scheme 2 (FIG. 29), Stage 2). Also present in the mass spectrum is a peak for the vicinal nitrohydroxy adduct (m/z 342) and minor peaks corresponding to the hydroxy and peroxy derivatives of LNO2 (m/z 340 and 356, respectively).
Nitrolinoleic acid is a pluripotent signaling molecule that exerts its bioactivity by acting as a high affinity ligand for PPARγ (2), activating protein kinase signaling cascades and, as shown herein, by serving as a hydrophobically-stabilized reserve for .NO. The activation of PPARγ-dependent gene expression by LNO2 requires this ligand to be stabilized and transported as the intact nitroalkene to the nuclear receptor (2). The mechanism(s) involved in protein kinase activation by LNO2 remain unclear, but can include direct ligation of receptors at the plasma membrane and/or covalent modification and activation of signaling mediators via Michael addition reactions. Current data reveals that the signaling actions of LNO2 are multifaceted, with the activation of protein kinases and/or PPAR receptor activation not fully explaining observed cellular responses, such as the stimulation of cGMP-dependent vessel relaxation (5). The observation herein that LNO2 decay yields .NO and that LNO2 is subject to hydrophobic stabilization thus lends additional perspective to our understanding of how compartmentalization will influence the nature of cell signaling reactions mediated by fatty acid nitroalkene derivatives.
A central challenge in detecting .NO generation by relatively slow-releasing compounds (e.g., RSNO and organonitrate derivatives) is the risk of lack of specificity and sensitivity. This is especially the case when concurrent oxygen, heme, lipid, protein and probe-related redox reactions are possible. Quantitative rigor is also always a concern. To circumvent these problems, multiple approaches for the qualitative and quantitative detection of .NO generation by LNO2 were employed herein. The release of .NO by LNO2 was assessed quantitatively by spectrophotometric analysis of oxymyoglobin oxidation. Additional qualitative proof of LNO2-derived NO release came from EPR analysis of .NO-dependent cPTI formation, .NO-dependent chemiluminescence following reaction with O3 and mass spectroscopic detection of anticipated decay products of LNO2. Also, in aqueous solutions and in the absence of alternative reaction pathways, 4 mol NO react with 1 mol O2 to ultimately yield 4 mol NO2−. Thus, formation of NO2− was used as additional evidence for NO formation. The yield of NO2− during LNO2 decay was 3.5 fold lower than predicted from more direct NO measurements based on oxymyoglobin oxidation. Several explanations can account for this apparent discrepancy. First, in the absence of .NO scavengers, .NO rapidly equilibrates with the gas phase, thus decreasing NO available for oxidation to NO2−. Second, .NO reactions with carbonyl, hydroxyl and peroxyl radicals are extremely fast [k> ̃1×1010 M−1 s−1, (29)]. These free radical intermediates are likely formed during LNO2 decomposition, as evidenced by products with mass to charge ratio 340 and 356 (FIG. 27). Thus, products of the reaction of these species with .NO may not contribute to NO2− formation. Overall, multiple independent criteria support the capacity of LNO2 to release of .NO.
The gas-phase chemiluminescence reaction of .NO with O3 is a highly sensitive and specific method for detecting .NO and nitroso-derivatives of biomolecules. One widely utilized analytical strategy relies on the reductive cleavage of NO2− and nitroso-derivatives by I3−. Treatment of samples with acidic sulfanilamide and HgCl2 permits additional discrimination between heme-NO, NO2−, and putative RSNO and RNNO derivatives (20, 21, 25-28). The latter HgCl2-resistant species [proposed as RNNO, (20)] may be best termed XNOx at this juncture, since LNO2 also yields O3 chemiluminescence following reaction with acidified sulfanilamide and HgCl2 prior to injecting into iodine/triiodide mixtures and the detection chamber. These data reveal that a contribution of fatty acid nitroalkene derivatives to the measurement of various tissue biomolecule NO derivatives must additionally be considered. Of additional interest, the UV photolysis approach for NOx detection in biological samples directly stimulates decay of LNO2 to yield .NO. This new insight thus raises significant concern about the accuracy of reported concentrations for .NO-derived species using UV photolysis, since nitrated fatty acids are the most prevalent bioactive oxides of nitrogen yet found in vivo (1). Protein fractionation via solvent extraction (e.g., acetone) prior to analysis of .NO derivatives in biological samples does not eliminate the possibility that nitrated fatty acids are a source of “detectable” or RSNO-like .NO formation by UV photolysis, as LNO2 and other nitroalkenes readily partition into the polar phase of many extraction strategies including those employing acetone.
The observation that LNO2 undergoes decay reactions to yield .NO in aqueous solution initially raised concern regarding how a significant and consistent LNO2 content in plasma and red cells of healthy humans could be detected at near-micromolar concentrations (1). Appreciating that synthetic LNO2 is stable in methanol suggested that the ionic microenvironment in which LNO2 was solvated would significantly modulate stability. To first address the possibility that LNO2 is stabilized by hydrophobic environments reminiscent of membranes and lipoproteins, it was observed that .NO release from LNO2 was inhibited upon LNO2 solvation in n-octanol. Further analysis using non-ionic detergent micelles, wherein the relatively hydrophobic NO2 group of LNO2 is expected to partition into non-polar microenvironments, revealed that LNO2 decomposition and .NO release was inhibited (FIG. 24). Importantly, this occurred at and above the CMC of each detergent and lipid studied. Similar results were obtained using DPPC-phosphatidylcholine-cholesterol (3:1, mol/mol) liposomes, also revealing that LNO2 is readily incorporated into and stabilized by lipid bilayers (FIG. 25C). This stabilizing influence of liposomes, which have a very low CMC, occurred at low hydrophobic phase volumes. These data reveal that LNO2 will be stable in hydrophobic environments and that cell membranes and lipoproteins can serve as an endogenous reserve for LNO2 and its downstream cell signaling capabilities. Indeed,  ̃80% of LNO2 is esterified to complex lipids in blood, including phospholipids derived from red cell membrane lipid bilayers (1). This further suggests that during inflammatory responses, esterases and A2-type phospholipases may hydrolyze and mobilize membrane-stabilized LNO2 for mediating cell signaling actions. This regulated disposition of LNO2 in lipophilic versus aqueous environments thus represents a “hydrophobic switch” that will control the nature of LNO2 signaling activity (Scheme 1 (FIG. 28)).
The mechanisms accounting for NO release from organic nitrites and nitrates are controversial, appear to be multifaceted and remain to be incisively defined. For example, the nitrate ester derivative nitroglycerin (NTG) has been used as a vasodilator for more than a century in the treatment of angina pectoris. Nitroglycerin does not directly decay to yield NO or an .NO-like species that will activate soluble guanylate cyclase (sGC), rather cellular metabolism is required to yield a species capable of NO-like activation of sGC. While several enzymes are identified as competent to mediate the denitration and “bioactivation” of NTG (e.g., xanthine oxidoreductase, cytochrome P450 oxidase and reductase, old yellow protein and mitochondrial aldehyde dehydrogenase-2), detailed insight is lacking as to unified redox chemistry, enzymatic and cellular mechanisms accounting for a) the 3 e-reduction of nitrate to an .NO-like species and b) the attenuated NTG metabolism that occurs during nitrate tolerance (30).
The present report of non-enzymatic release of .NO from endogenous fatty acid nitroalkene derivatives (e.g., LNO2) lends additional perspective to how nitric oxide synthase-dependent .NO signaling can be transduced. It is shown via 3 different analytical approaches that the product of LNO2 decay is unambiguously .NO. Mass spectrometric analysis and LNO2 decay studies reported herein, in concert with previous understanding of the chemical reactivity of nitroalkenes, reveals a viable mechanism for how nitrated fatty acids can serve to transduce tissue .NO signaling capacity (Schemes 1 and 2).
The release of .NO by a vicinal nitrohydroxy arachidonic acid derivative detected in cardiac lipid extracts has been proposed (31). These derivatives induce vasorelaxation of rat aortic rings via possible .NO-dependent activation of guanylate cyclase. The intermediate formation of an analogous hydroxy derivative of nitrolinoleate, L(OH)NO2, is documented herein to occur during LNO2 decay in aqueous milieu (FIG. 27). Fatty acid nitroalkene derivatives appear to be clinically abundant, since both nitro and nitrohydroxy derivatives of all principal unsaturated fatty acids are present in healthy human blood plasma and urine (32). Present results indicate that hydroxy derivatives of fatty acid nitroalkenes represent the accumulation of Michael addition-like reaction products with H2O that are in equilibrium with the parent nitroalkene and are not a direct precursor to .NO release.
A more viable mechanism accounting for .NO release by nitroalkenes is supported by 1) mass spectroscopic detection of expected decay products and 2) the aqueous and pH dependency of this process (FIG. 22), with LNO2 decay and consequent .NO release involving protonation and deprotonation events. The mechanism accounting for .NO release by nitroalkenes is based on the Nef reaction (33,34), a standard reaction of organic nitro derivatives first described in 1894 (35).
The original Nef reaction entails complete deprotonation of an alkyl nitro compound with base to yield the nitroanion, followed by quenching with aqueous acid to cause hydrolysis to the corresponding carbonyl compound and oxides of nitrogen. Most Nef reactions are now performed using additional oxidants or reductants, rather than the simple acid-base chemistry of the original reaction (36-44). There are a few noteworthy points about this proposed mechanism that relate to how .NO can be ultimately produced. The nitrogen-containing product of the original Nef reaction is N2O, a stable oxide of nitrogen that would not be a precursor to .NO under the neutral aqueous conditions used herein to model biologically-relevant LNO2 decay. The initial oxide of nitrogen formed, HNO, is unstable and quickly disproportionates to form N2O as shown in Scheme 2 (FIG. 29) (Stage 2). While HNO (or the NO− anion) might conceivably yield one electron and be oxidized to NO, this is not expected under neutral aqueous conditions. Alternatively, a nitroso intermediate formed during LNO2 decay provides a plausible pathway to yield .NO. This nitroso intermediate is expected to have an especially weak C—N bond, easily forming .NO and a radical stabilized by conjugation with the alkene and stabilized by the OH group, a moiety known to stabilize adjacent radicals.
In Scheme 2 (FIG. 29) (Stage 1) the vicinal nitrohydroxy fatty acid derivative is in equilibrium with the nitroalkene. This is possible for two reasons. First, the nitro group in the vicinal nitrohydroxy fatty acid makes the adjacent hydrogen very acid (pKa  ̃7-8), thus facilitating formation of a significant amount of the nitronate anion at physiological pH. The anion can then release hydroxide, which when neutralized with the proton removed in the first step results in the net loss of neutral water. Second, the fatty acid nitroalkene is a strong electrophile and can readily undergo Michael conjugate addition reaction with the small amounts of hydroxide anion that are always present in aqueous solution under physiological pH conditions, explaining the facile equilibrium of vicinal nitrohydroxy fatty acids with their corresponding nitroalkene derivatives. In Scheme 2 (FIG. 29) (Stage 2), the lipid nitroalkene forms .NO as described above.
These proposed mechanisms for .NO formation from LNO2 provided the testable hypothesis for how nitrated fatty acids can serve as a source of .NO using simple acid/base chemistry with no additional oxidants or reductants. Mass spectrometric detection of expected oxidized fatty acid products and direct detection of .NO formation supported this pathway of nitroalkene decay. This acid/base chemistry could also be employed by as-yet-undescribed enzymes that could catalyze physiologically-significant extents of .NO release from the multiple lipid nitroalkene derivatives now being observed (32).
Therapeutic agents that release .NO are a rapidly expanding area of drug design. Dual-acting nitro and nitroso derivatives of existing drugs have been synthesized and are being studied for efficacy in treating diabetes, metabolic syndrome, hypertension and atherosclerosis. These include .NO-releasing statin derivatives and NO-non-steroidal anti-inflammatory derivatives such as NO-acetacylic acid, NO-ibuprofen and NO-piroxicam. These adducts were devised based on the precept that an .NO donor moiety will augment therapeutic breadth and value. This class of pharmaceuticals are of particular relevance when alterations in endogenous .NO signaling contributes to tissue pathogenesis. In this regard, LNO2 shares similarities with these classes of “chimeric” inflammatory-regulating compounds, as LNO2 is a potent endogenous PPARγ agonist that rivals extents of PPARγ activation induced by similar concentrations of thiazolidinediones (2). Herein, the present invention shows that LNO2 also has the capability to release .NO in a regulated manner. Thus, the potential signaling actions of LNO2 are expected to be pluripotent in nature.
In summary, .NO-mediated oxidative reactions with unsaturated fatty acids yield nitroalkene derivatives. Once formed, nitrated fatty acids are hydrophobically stabilized by lipid bilayers and lipoproteins or alternatively, can be redistributed to aqueous environments to release .NO via a Nef-like reaction. In its native form, LNO2 also activates nuclear PPAR receptor-mediated regulation of gene expression. These combined actions are expected to transduce the salutary inflammatory signaling reactions that have been described for both .NO and LNO2. Because LNO2 production is increased by oxidative inflammatory reactions, this species thus represents an adaptive mediator that regulates potentially pathogenic tissue responses to inflammation.
Throughout Example 4, several publications have been referenced. These publications are listed in the Reference List for Example 4. The disclosures of these publications in their entireties are hereby incorporated by reference into this application in order to more fully describe the compounds, compositions and methods described herein.
REFERENCE LIST FOR EXAMPLE 4


    1. Baker, P. R., Schopfer, F. J., Sweeney, S., and Freeman, B. A. (2004) Proc. Natl. Acad. Sci. U.S.A 101, 11577-11582
    2. Schopfer, F. J., Lin, Y., Baker, P. R., Cui, T., Garcia-Barrio, M., Zhang, J., Chen, K., Chen, Y. E., and Freeman, B. A. (2005) Proc. Natl. Acad. Sci. U.S.A 102, 2340-2345
    3. Coles, B., Bloodsworth, A., Eiserich, J. P., Coffey, M. J., McLoughlin, R. M., Giddings, J. C., Lewis, M. J., Haslam, R. J., Freeman, B. A., and O'Donnell, V. B. (2002) J. Biol. Chem. 277, 5832-5840
    4. Coles, B., Bloodsworth, A., Clark, S. R., Lewis, M. J., Cross, A. R., Freeman, B. A., and O'Donnell, V. B. (2002) Circ. Res. 91, 375-381
    5. Lim, D. G., Sweeney, S., Bloodsworth, A., White, C. R., Chumley, P. H., Krishna, N. R., Schopfer, F., O'Donnell, V. B., Eiserich, J. P., and Freeman, B. A. (2002) Proc. Natl. Acad. Sci. U.S.A 99, 15941-15946
    6. Ignarro, L. J., Byrns, R. E., Buga, G. M., and Wood, K. S. (1987) Circ. Res. 61, 866-879
    7. Hogg, N. (2002) Annu. Rev. Pharmacol. Toxicol. 42, 585-600
    8. Schopfer, F. J., Baker, P. R., and Freeman, B. A. (2003) Trends Biochem. Sci. 28, 646-654
    9. Denicola, A., Souza, J. M., and Radi, R. (1998) Proc. Natl. Acad. Sci. U.S.A 95, 3566-3571
    10. Denicola, A., Batthyany, C., Lissi, E., Freeman, B. A., Rubbo, H., and Radi, R. (2002) J. Biol. Chem. 277, 932-936
    11. Liu, X., Miller, M. J., Joshi, M. S., Thomas, D. D., and Lancaster, J. R., Jr. (1998) Proc. Natl. Acad. Sci. U.S.A 95, 2175-2179
    12. O'Donnell, V. B., Chumley, P. H., Hogg, N., Bloodsworth, A., Darley-Usmar, V. M., and Freeman, B. A. (1997) Biochemistry 36, 15216-15223
    13. Rubbo, H., Radi, R., Anselmi, D., Kirk, M., Barnes, S., Butler, J., Eiserich, J. P., and Freeman, B. A. (2000) J. Biol. Chem. 275, 10812-10818
    14. Coffey, M. J., Natarajan, R., Chumley, P. H., Coles, B., Thimmalapura, P. R., Nowell, M., Kuhn, H., Lewis, M. J., Freeman, B. A., and O'Donnell, V. B. (2001) Proc. Natl. Acad. Sci. U.S.A 98, 8006-8011
    15. O'Donnell, V. B., Taylor, K. B., Parthasarathy, S., Kuhn, H., Koesling, D., Friebe, A., Bloodsworth, A., Darley-Usmar, V. M., and Freeman, B. A. (1999) J. Biol. Chem. 274, 20083-20091
    16. O'Donnell, V. B., Coles, B., Lewis, M. J., Crews, B. C., Marnett, L. J., and Freeman, B. A. (2000) J. Biol. Chem. 275, 38239-38244
    17. Eiserich, J. P., Baldus, S., Brennan, M. L., Ma, W., Zhang, C., Tousson, A., Castro, L., Lusis, A. J., Nauseef, W. M., White, C. R., and Freeman, B. A. (2002) Science 296, 2391-2394
    18. Arnold, W. P., Mittal, C. K., Katsuki, S., and Murad, F. (1977) Proc. Natl. Acad. Sci. U.S.A 74, 3203-3207
    19. Szoka, F., Jr. and Papahadjopoulos, D. (1980) Annu. Rev. Biophys. Bioeng. 9, 467-508
    20. Feelisch, M., Rassaf, T., Mnaimneh, S., Singh, N., Bryan, N. S., Jourd'heuil, D., and Kelm, M. (2002) FASEB J. 16, 1775-1785
    21. Gladwin, M. T., Wang, X., Reiter, C. D., Yang, B. K., Vivas, E. X., Bonaventura, C., and Schechter, A. N. (2002) J. Biol. Chem. 277, 27818-27828
    22. Bligh, E. G. and Dyer, W. L. (1959) Can. J. Biochem. Physiol. 37, 911-917
    23. Akaike, T., Yoshida, M., Miyamoto, Y., Sato, K., Kohno, M., Sasamoto, K., Miyazaki, K., Ueda, S., and Maeda, H. (1993) Biochemistry 32, 827-832
    24. Hogg, N., Singh, R. J., Joseph, J., Neese, F., and Kalyanaraman, B. (1995) Free Radic. Res. 22, 47-56
    25. McMahon, T. J., Moon, R. E., Luschinger, B. P., Carraway, M. S., Stone, A. E., Stolp, B. W., Gow, A. J., Pawloski, J. R., Watke, P., Singel, D. J., Piantadosi, C. A., and Stamler, J. S. (2002) Nat. Med. 8, 711-717
    26. Stamler, J. S., Jaraki, O., Osborne, J., Simon, D. I., Keaney, J., Vita, J., Singel, D., Valeri, C. R., and Loscalzo, J. (1992) Proc. Natl. Acad. Sci. U.S.A 89, 7674-7677
    27. Crawford, J. H., Chacko, B. K., Pruitt, H. M., Piknova, B., Hogg, N., and Patel, R. P. (2004) Blood 104, 1375-1382
    28. Janero, D. R., Bryan, N. S., Saijo, F., Dhawan, V., Schwalb, D. J., Warren, M. C., and Feelisch, M. (2004) Proc. Natl. Acad. Sci. U.S.A 101, 16958-16963
    29. Padmaja, S. and Huie, R. E. (1993) Biochem. Biophys. Res. Commun. 195, 539-544
    30. Thatcher, G. R., Nicolescu, A. C., Bennett, B. M., and Toader, V. (2004) Free Radic. Biol. Med. 37, 1122-1143
    31. Balazy, M., Iesaki, T., Park, J. L., Jiang, H., Kaminski, P. M., and Wolin, M. S. (2001) J. Pharmacol. Exp. Ther. 299, 611-619
    32. Baker, P. R. S., Lin, Y., Schopfer, F. J., Woodcock, S. T., Long, M. H., Batthyany, C., Iles, K. E., Baker, L. M. S., Sweeney, S., Braunchaud, B. P., Chen, Y. E., and Freeman, B. A. (2005) J Biol Chem Submitted,
    33. Pinnick, H. W. (1990) Organic Reactions 38, 655-792
    34. Ballini, R. and Petrini, M. (2004) Tetrahedron 60, 1017-1047
    35. Nef, J. U. (1894) Justis Liebigs Annalan der Chemie 263-342
    36. Van Tamelen, E. E. and Thiede, R. J. (1952) J. Am. Chem. Soc. 74, 2615-2618
    37. Leitch, L. C. (1955) Canadian Journal of Chemistry 33, 400-404
    38. Noland, W. E. (1955) Chemical Reviews 55, 137-155
    39. Hawthorne, M. F. (1957) J. Am. Chem. Soc. 79, 2510-2515
    40. Feuer, H. and Nielsen, A. T. (1962) J. Am. Chem. Soc. 84, 688
    41. Armand, J. (1965) Bulletin de la Societe Chimique de France 3246-3255
    42. Kornblum, N. and Brown, R. A. (1965) J. Am. Chem. Soc. 87, 1742-1747
    43. Cundall, R. B. and Locke, A. W. (1968) Journal of the Chemical Society [section] B: Physical Organic 98-103
    44. Wilson, H. and Lewis, E. S. (1972) J. Am. Chem. Soc. 94, 2283-2285


Example 5
The nitroalkenes of the present invention also display regulatory actions towards the expression of genes related to inflammatory responses, cell growth, cell differentiation, cell signaling, cell death (apoptosis) and metabolism. For example, nitrolinoleate regulates gene expression in human vascular endothelial cells and macrophages. Cells were stimulated with vehicle and linoleic acid (2.5 μM, the latter two treatments as controls) and LNO2 (2.5 mM) for 24 hours. The total RNA was purified with a Qiagen Kit and analyzed with a Whole Human Genome Oligo Array (G4112A, 41K Genes, Agilent). Results showed that nitrolinoleate regulates >5000 genes, with statistically-significant up-regulation of >2300 genes and down-regulation of >3000 genes. In contrast, the native (precursor) fatty acid, linoleic acid, only regulates 14 genes, with an up-regulation of 5 genes and the down-regulation of 9 genes.
Therefore, the present invention also provides for the selective regulation of clinically significant genes by nitrolinoleate. Table 5 lists examples of genes down-regulated by nitrolineate that are associated with cell signaling, growth, differentiation, metabolism, inflammatory responses, migration and apoptosis. Table 5 also includes genes involved in the Jak/STAT signaling pathway, the NF-κB pathway and the P13K/Akt pathway. The genes listed in Table 5 can be involved in one or more processes associated with cell signaling, growth, differentiation, metabolism, inflammatory responses, migration and apoptosis. These genes can also be involved in one or more pathways selected from the group consisting of the Jak/STAT signaling pathway, the NF-κB pathway and the P13K/Akt pathway.







TABLE 5





Selected genes down-regulated by nitrolineate (>1.5 fold, p < 0.05)
















RAC2; ras-related C3 botulinum toxin substrate 2 (rho
2.47


family, small GTP binding protein Rac2)


CDC2; cell division cycle 2, G1 to S and G2 to M
2.75


CDC42; cell division cycle 42 (GTP binding protein,
1.65


25 kDa)


CCND3; cyclin D3
1.51


CCNE1; cyclin E1
1.58


CAMK1; calcium/calmodulin-dependent protein kinase I
2.24


TCF4; transcription factor 4
2.63


PPARGC1B; peroxisome proliferative activated
2.20


receptor, gamma, coactivator 1, beta


RB1; retinoblastoma 1 (including osteosarcoma)
2.06


TNF; tumor necrosis factor (TNF superfamily, member 2)
2.01


TYK2; tyrosine kinase 2
2.00


STAT1; signal transducer and activator of transcription
1.74


1, 91 kDa


CAV3; caveolin 3
1.76


EDARADD; EDAR-associated death domain
1.69


DAPK2; death-associated protein kinase 2
1.73


CASP8AP2; CASP8 associated protein 2
1.52


ITR; intimal thickness-related receptor
1.68









Table 6 lists additional genes that are also down-regulated by nitrolineate and are associated with cell signaling, growth, differentiation, metabolism, inflammatory responses, migration and apoptosis








TABLE 6





Down-regulated genes (>1.5 fold, p < 0.05). Genes associated with cell signaling,


growth, differentiation, metabolism, inflammatory responses, migration and apoptosis




















TGFBI;
9.80
ILK; integrin-linked kinase
2.94
MAPKAPK3; mitogen-activated
2.18


transforming



protein kinase-activated protein


growth factor,



kinase 3


beta-induced,


68 kDa


PLAU;
7.58
EGR2; early growth response
2.65
ADRB2; adrenergic, beta-2-,
2.04


plasminogen

2 (Krox-20 homolog,

receptor, surface


activator,

Drosophila)


urokinase


VASP;
4.85
FDPS; farnesyl diphosphate
2.46
IL11; interleukin 11
2.02


vasodilator-

synthase (farnesyl


stimulated

pyrophosphate synthetase,


phosphoprotein

dimethylallyltranstransferase,




geranyltranstransferase)


AIF1; allograft
3.95
TGFB1; transforming growth
2.46
PIK3CB; phosphoinositide-3-
1.93


inflammatory

factor, beta 1 (Camurati-

kinase, catalytic, beta


factor 1

Engelmann disease)

polypeptide


PTGS1;
3.95
IRS1; insulin receptor
2.40
TIMP2; tissue inhibitor of
1.92


prostaglandin-

substrate 1

metalloproteinase 2


endoperoxide


synthase 1


(prostaglandin


G/H synthase and


cyclooxygenase)


PTGES2;
1.59
PDGFRA; platelet-derived
2.36
TLR2; toll-like receptor 2
1.89


prostaglandin E

growth factor receptor, alpha


synthase 2

polypeptide


E2F2; E2F
3.70
RACGAP1; Rac GTPase
2.35
LDLR; low density lipoprotein
1.77


transcription

activating protein 1

receptor (familial


factor 2



hypercholesterolemia)


NCOR2; nuclear
1.59
NCR1; natural cytotoxicity
2.30
CARD9; caspase recruitment
3.33


receptor co-

triggering receptor 1

domain family, member 9


repressor 2


SRF; serum
1.56
PDGFA; platelet-derived
2.27
FLJ23091; putative NFkB
1.74


response factor

growth factor alpha

activating protein 373


(c-fos serum

polypeptide


response


element-binding


transcription


factor)


BCL2A1;
3.50
TIMP1; tissue inhibitor of
3.62
PDGFC; platelet derived growth
1.72


BCL2-related

metalloproteinase 1

factor C


protein A1

(erythroid potentiating




activity, collagenase




inhibitor)


CARD9;
3.33
TIMP3; tissue inhibitor of
2.27
ARHGEF1; Rho guanine
1.64


caspase

metalloproteinase 3 (Sorsby

nucleotide exchange factor


recruitment

fundus dystrophy,

(GEF) 1


domain family,

pseudoinflammatory)


member 9


FN1; fibronectin 1
3.33
MMP19; matrix
1.58
JUNB; jun B proto-oncogene
1.60




metalloproteinase 19


PPARD;
3.12
CAMK1;
2.24
MYLK; myosin, light
3.13


peroxisome

calcium/calmodulin-

polypeptide kinase


proliferative

dependent protein kinase I


activated


receptor, delta




MAPKAPK3; mitogen-
2.18
ROCK1; Rho-associated, coiled-
1.59




activated protein kinase-

coil containing protein kinase 1




activated protein kinase 3






IGF1; insulin-like growth factor
1.54






1 (somatomedin C)






VEGFB; vascular endothelial
1.51






growth factor B






ECGF1; endothelial cell growth
1.73






factor 1 (platelet-derived)









Table 7 provides gene that are up-regulated by nitrolinoleate. These genes are associated with apoptosis, cell signaling and/or growth.








TABLE 7





Genes up-regulated by nitrolinoleate.


















IL10RA; interleukin 10 receptor,
3.173



alpha



GADD45G; growth arrest and DNA-
3.155



damage-inducible, gamma



IRS2; insulin receptor substrate 2
3.243



HO-1, heme oxygenase-1
4.086



GADD45A; growth arrest and DNA-
2.823



damage-inducible, alpha



CASP1; caspase 1, apoptosis-related
2.572



cysteine protease (interleukin 1, beta,



convertase)



CASP4; caspase 4, apoptosis-related
2.57



cysteine protease



CCNA1; cyclin A1
2.558



JUN; v-jun sarcoma virus 17
2.454



oncogene homolog (avian)



CASP3; caspase 3, apoptosis-related
1.689



cysteine protease



GADD45B; growth arrest and DNA-
1.674



damage-inducible, beta



AATK; apoptosis-associated tyrosine
1.503



kinase



AMID; apoptosis-inducing factor
8.362



(AIF)-homologous mitochondrion-



associated inducer of death










Example 6
The present invention also provides the activation of p-JNK and p-c-Jun protein kinases by nitrated lipids, by stimulating the phosphorylation of these kinases. The activation of these cell signaling mediators allow modulation of cell signaling, growth, differentiation, metabolism, inflammatory responses, migration and apoptosis. FIG. 30 shows human lung epithelial cell responses of p-JNK and p-c-JUN upon administration of nitrolineate. The nitroalkenes of the present invention also activate the ERK MAPK pathway in human lung epithelial cells, as shown by a dramatic increase in ERK phosphorylation (e.g. activation) and the phosphorylation of its downstream target signaling protein, pELK, illustrated in FIG. 31. The nitroalkenes of the present invention also inhibit activity of NF-κB pathway as indicated by a) luciferase-linked NFkB-response element reporter assay in response to the inflammatory mediator TNFα and b) direct analysis of the degradation f the NFkB inhibitor protein, IkB in response to the inflammatory mediator E. coli LPS (FIG. 32).
Example 7
Synthesis and Characterization of Nitro/Hydroxy Fattay Acids
The Michael addition reaction of water to nitroalkenes results in formation of nitro-hydroxy derivative (18:1, 18:2 and 18:3 nitro-hydroxy species shown in FIG. 33). These hydroxylated species are prepared by placing nitroalkenes in basic aqueous conditions, isolation by solvent extraction and purification by HPLC or thin layer chromatography. Fatty acid nitro-hydroxy derivatives can (a) display unique cell signaling activities, (b) represent a more stable “storage form” of the PPAR receptor-avid nitroalkene parent molecule and (c) permit determination of specific positional isomers of nitroalkenes by directing, upon hydroxylation, nucleophilic heterolytic scission between the nitro and hydroxy-bonded fatty acid alkene. Nitro isomer-specific fragments can then be detected by mass spectrometry (FIG. 33).
Example 8
Addition of Glutathione to Nitraed Fatty Acids
In a neutral buffered solution, the thiol and nitrated FA (e.g., nitrated oleate or linoleate) can be combined from equimolar to 10:1 ratios of nitroalkene to thiol. Mass spectrometry shows nitrolinoleate forms a covalent Michael addition reaction product with the tripeptide glutathione (m/z 631.3, LNO2-GSH) (FIG. 34). This adduct can be “fingerprinted” by its source fragmentation in the mass spectrometer to the precursors glutathione (m/z 306.3) and nitrolinoleate (m/z 324.2, LNO2).
Throughout this application, various publications are referenced. The disclosures of these publications in their entireties are hereby incorporated by reference into this application in order to more fully describe the compounds, compositions and methods described herein.
Various modifications and variations can be made to the compounds, compositions and methods described herein. Other aspects of the compounds, compositions and methods described herein will be apparent from consideration of the specification and practice of the compounds, compositions and methods disclosed herein. It is intended that the specification and examples be considered as exemplary.

## Claims

1. A method of treating renal inflammation, sickle cell disease, or pulmonary inflammation in a subject, comprising administering to the subject a compound comprising a lipid having the structure:




wherein:
R1 is selected from the group consisting of C1-C24 alkyl, C1-C24 alkenyl, and C1-C24 alkynyl;
R2, R3, R7, and R8 are each independently selected from the group consisting of hydrogen, NO2, OH, and OOH, wherein at least one of R2, R3, R7, and R8 is NO2;
R4 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkenyl, wherein R4 comprises a terminal COOR6 and R6 is hydrogen, C1-C24 alkyl, or a pharmaceutically acceptable counterion;
R5 is hydrogen or R4 and R5 collectively form ═C(R9)(R10), wherein R9 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl and R10 is hydrogen, NO2, OH, or OOH; and
n is from 1 to 24,

wherein the compound is substantially pure.

2. The method of claim 1, wherein the lipid comprises a fatty acid.

3. The method of claim 1, wherein the lipid is selected from the group consisting of glycolipids, glycerolipids, phospholipids, and cholesterol.

4. The method of claim 1, wherein the lipid is selected from the group consisting of oleic acid (18:1), 22:6, and docosahexanoic acid.

5. The method of claim 1, wherein the lipid comprises 10-nitro-9-cis,12-cis-octadecadienoic acid.

6. The method of claim 1, wherein the lipid is selected from the group consisting of linoleic acid (18:2), linolenic acid (18:3), and cholesterol linoleate.

7. The method of claim 1, wherein the lipid comprises arachidonic acid (20:4).

8. The method of claim 1, the lipid having the formula:




wherein:
R9 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkenyl and includes a terminal COOR6; and
R10 is hydrogen, NO2, OH, or OOH,
wherein at least one of R2, R3, R7, R8, and R10 is NO2.

9. The method of claim 8, wherein R1 is C4-C10 alkyl; R2, R8, and R10 are hydrogen; R7 is NO2; and R9 is C6-C12 alkyl.

10. The method of claim 8, wherein R3 and R8 are cis to one another, and R7 and R10 are cis to one another.

11. The method of claim 1, wherein R1 is C4-C10 alkyl; R2, R3, and R7 are hydrogen; R8 is NO2; and R4 is C6-C12 alkyl.

12. The method of claim 1, wherein the lipid comprises 12-nitro-9-cis,12-cis-octadecadienoic acid.

13. The method of claim 1, wherein R9 is C6-C12 alkyl.

14. The method of claim 1, wherein the lipid is about 90% one compound.

15. The method of claim 1, wherein the lipid is about 100% one compound.

16. The method of claim 1, wherein the lipid is 10-nitro-9-cis-octadecaenoic acid.

17. The method of claim 1, wherein renal inflammation is diabetic nephropathy.

18. The method of claim 1, wherein pulmonary inflammation is essential hypertension.

19. A method of treating renal inflammation comprising administering to a subject in need of treatment an effective amount of a pharmaceutical composition containing a compound of formula:



wherein:
R1 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl;
R2, R3, R7, and R8 are each independently hydrogen, NO2, OH, or OOH, and at least one of R2, R3, R7, and R8 is NO2;
R4 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkenyl, and R4 comprises a terminal COOR6, and R6 is hydrogen, C1-C24 alkyl, or a pharmaceutically acceptable counterion;
R5 is hydrogen or R4 and R5 collectively form ═C(R9)(R10), wherein R9 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl and R10 is hydrogen, NO2, OH, or OOH; and
n is from 1 to 24, wherein the compound is substantially pure; and
a pharmaceutically acceptable carrier, thickener, diluent, solvent, buffer, preservative, or surface active agent.

20. The method of claim 19, wherein the compound is 10-nitro-9-cis-octadecaenoic acid.

21. The method of claim 19, wherein renal inflammation is diabetic nephropathy.

22. A method for treating pulmonary inflammation comprising administering to a patient in need of treatment an effective amount of a pharmaceutical composition containing a compound of formula:



wherein:
R1 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl;
R2, R3, R7, and R8 are each independently hydrogen, NO2, OH, or OOH, and at least one of R2, R3, R7, and R8 is NO2;
R4 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkenyl, and R4 comprises a terminal COOR6, and R6 is hydrogen, C1-C24 alkyl, or a pharmaceutically acceptable counterion;
R5 is hydrogen or R4 and R5 collectively form ═C(R9)(R10), wherein R9 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl and R10 is hydrogen, NO2, OH, or OOH; and
n is from 1 to 24, wherein the compound is substantially pure; and
a pharmaceutically acceptable carrier, thickener, diluent, solvent, buffer, preservative, or surface active agent.

23. The method of claim 22, wherein the compound is 10-nitro-9-cis-octadecaenoic acid.

24. The method of claim 22, wherein the pulmonary inflammation is essential hypertension.

25. A method for treating sickle cell disease comprising administering to a patient in need of treatment an effective amount of a pharmaceutical composition containing a compound of formula:



wherein:
R1 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl;
R2, R3, R7, and R8 are each independently hydrogen, NO2, OH, or OOH, and at least one of R2, R3, R7, and R8 is NO2;
R4 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkenyl, and R4 comprises a terminal COOR6, and R6 is hydrogen, C1-C24 alkyl, or a pharmaceutically acceptable counterion;
R5 is hydrogen or R4 and R5 collectively form ═C(R9)(R10), wherein R9 is C1-C24 alkyl, C1-C24 alkenyl, or C1-C24 alkynyl and R10 is hydrogen, NO2, OH, or OOH; and
n is from 1 to 24, wherein the compound is substantially pure; and
a pharmaceutically acceptable carrier, thickener, diluent, solvent, buffer, preservative, or surface active agent.

26. The method of claim 25, wherein the compound is 10-nitro-9-cis-octadecaenoic acid.