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Quantitative determination of esterified eicosanoids and related oxygenated metabolites after base hydrolysis

Open AccessPublished:October 15, 2018DOI:https://doi.org/10.1194/jlr.D089516
      Eicosanoids and related metabolites (oxylipins) possess potent signaling properties, elicit numerous important physiologic responses, and serve as biomarkers of disease. In addition to their presence in free form, a considerable portion of these bioactive lipids is esterified to complex lipids in cell membranes and plasma lipoproteins. We developed a rapid and sensitive method for the analysis of esterified oxylipins using alkaline hydrolysis to release them followed by ultra-performance LC coupled with mass spectrometric analysis. Detailed evaluation of the data revealed that several oxylipins are susceptible to alkaline-induced degradation. Nevertheless, of the 136 metabolites we examined, 56 were reproducibly recovered after alkaline hydrolysis. We classified those metabolites that were resistant to alkaline-induced degradation and applied this methodology to quantify metabolite levels in a macrophage cell model and in plasma of healthy subjects. After alkaline hydrolysis of lipids, 34 metabolites could be detected and quantified in resting and activated macrophages, and 38 metabolites were recovered from human plasma at levels that were substantially greater than in free form. By carefully selecting internal standards and taking the observed experimental limitations into account, we established a robust method that can be reliably employed for the measurement of esterified oxylipins in biological samples.
      Eicosanoids and related metabolites, sometimes referred to as oxylipins, are a group of structurally diverse metabolites that derive from the oxidation of PUFAs, including arachidonic acid, linoleic acid, α and γ linolenic acid, dihomo γ linolenic acid, eicosapentaenoic acid, and docosahexaenoic acid. They are locally acting bioactive signaling lipids that regulate a diverse set of homeostatic and inflammatory processes (
      • Funk C.D.
      Prostaglandins and leukotrienes: advances in eicosanoid biology.
      ,
      • Astarita G.
      • Kendall A.C.
      • Dennis E.A.
      • Nicolaou A.
      Targeted lipidomic strategies for oxygenated metabolites of polyunsaturated fatty acids.
      ). Given the important regulatory functions in numerous physiological and pathophysiological states, the accurate measurement of eicosanoids and other oxylipins is of great clinical interest and lipidomics is now widely used to screen effectively for potential disease biomarkers (
      • Quehenberger O.
      • Dennis E.A.
      The human plasma lipidome.
      ).
      The biosynthesis of eicosanoids and oxylipins involves the action of multiple enzymes organized into a complex and intertwined lipid-anabolic network (
      • Dennis E.A.
      • Norris P.C.
      Eicosanoid storm in infection and inflammation.
      ). Generally, the enzymatic formation of eicosanoids requires free fatty acids as substrates; thus, the pathway is initiated by the hydrolysis of phospholipids (PLs) by phospholipase A2 upon physiological stimuli (
      • Dennis E.A.
      • Cao J.
      • Hsu Y.H.
      • Magrioti V.
      • Kokotos G.
      Phospholipase A2 enzymes: physical structure, biological function, disease implication, chemical inhibition, and therapeutic intervention.
      ). The hydrolyzed PUFAs are then processed by three enzyme systems: cyclooxygenases (COXs), lipoxygenases (LOXs), and cytochrome P450 enzymes. Each of these enzyme systems produces unique collections of oxygenated metabolites that function either as end-products or as intermediates for a cascade of downstream enzymes. The resulting eicosanoids exhibit diverse biological activities, half-lives, and utilities in regulating many physiologic processes in health and disease, including the immune response, inflammation, and homeostasis (
      • Smith W.L.
      • Urade Y.
      • Jakobsson P.J.
      Enzymes of the cyclooxygenase pathways of prostanoid biosynthesis.
      ,
      • Morisseau C.
      • Hammock B.D.
      Impact of soluble epoxide hydrolase and epoxyeicosanoids on human health.
      ,
      • Lone A.M.
      • Tasken K.
      Proinflammatory and immunoregulatory roles of eicosanoids in T cells.
      ,
      • Serhan C.N.
      Pro-resolving lipid mediators are leads for resolution physiology.
      ,
      • Kuhn H.
      • Banthiya S.
      • van Leyen K.
      Mammalian lipoxygenases and their biological relevance.
      ). Additionally, nonenzymatic processes can produce oxidized PUFA metabolites via free radical reactions giving rise to isoprostanes and other oxidized fatty acids (
      • Roberts L.J.
      • Morrow J.D.
      Measurement of F(2)-isoprostanes as an index of oxidative stress in vivo.
      ).
      Eicosanoids are either secreted and signal through G protein-coupled receptors in an autocrine or paracrine fashion or act intracellularly via various peroxisome proliferator-activating receptors (
      • Dennis E.A.
      • Norris P.C.
      Eicosanoid storm in infection and inflammation.
      ,
      • Ricciotti E.
      • FitzGerald G.A.
      Prostaglandins and inflammation.
      ,
      • Korotkova M.
      • Jakobsson P.J.
      Persisting eicosanoid pathways in rheumatic diseases.
      ). For optimal biological activity, these mediators need to be present in their free nonesterified form. However, a number of studies reported that a portion of eicosanoids are naturally esterified and can also be contained in cell membrane lipids, including PLs, in the form of esters (
      • Brezinski M.E.
      • Serhan C.N.
      Selective incorporation of (15S)-hydroxyeicosatetraenoic acid in phosphatidylinositol of human neutrophils: agonist-induced deacylation and transformation of stored hydroxyeicosanoids.
      ,
      • Bernstrom K.
      • Kayganich K.
      • Murphy R.C.
      • Fitzpatrick F.A.
      Incorporation and distribution of epoxyeicosatrienoic acids into cellular phospholipids.
      ,
      • VanRollins M.
      • Kaduce T.L.
      • Fang X.
      • Knapp H.R.
      • Spector A.A.
      Arachidonic acid diols produced by cytochrome P-450 monooxygenases are incorporated into phospholipids of vascular endothelial cells.
      ). The role of esterified eicosanoids is not clear but they may be signaling molecules in their own right or serve as a cellular reservoir for rapid release upon cell stimulation (
      • Shearer G.C.
      • Newman J.W.
      Impact of circulating esterified eicosanoids and other oxylipins on endothelial function.
      ,
      • Hammond V.J.
      • O'Donnell V.B.
      Esterified eicosanoids: generation, characterization and function.
      ).
      Two potential mechanisms for the formation of eicosanoid-containing PLs have been proposed: i) direct oxidation of PUFAs on the intact PLs; and ii) reacylation of preformed free oxylipins into lysoPLs. COXs require free fatty acid as substrate and show little activity toward PUFAs in intact PLs (
      • Lands W.E.
      • Samuelsson B.
      Phospholipid precursors of prostaglandins.
      ). A number of subsequent studies support the concept that prostaglandins (PGs) are first formed enzymatically and then incorporated into PLs by the sequential actions of long-chain acyl-CoA synthases and lysoPL acyltransferases (
      • Aldrovandi M.
      • Hammond V.J.
      • Podmore H.
      • Hornshaw M.
      • Clark S.R.
      • Marnett L.J.
      • Slatter D.A.
      • Murphy R.C.
      • Collins P.W.
      • O'Donnell V.B.
      Human platelets generate phospholipid-esterified prostaglandins via cyclooxygenase-1 that are inhibited by low dose aspirin supplementation.
      ,
      • Klett E.L.
      • Chen S.
      • Yechoor A.
      • Lih F.B.
      • Coleman R.A.
      Long-chain acyl-CoA synthetase isoforms differ in preferences for eicosanoid species and long-chain fatty acids.
      ). Additionally, preformed fatty acid epoxides, including the regioisomers of epoxyeicosatrienoic acid (EET), are effectively incorporated primarily into the PL fraction of cellular lipids, presumably via CoA-dependent mechanisms (
      • Spector A.A.
      • Fang X.
      • Snyder G.D.
      • Weintraub N.L.
      Epoxyeicosatrienoic acids (EETs): metabolism and biochemical function.
      ).
      In contrast, mammalian 12/15 LOX can act directly on PLs to generate esterified HETE isomers, including esterified 12-HETE and 15-HETE (
      • Brash A.R.
      Lipoxygenases: occurrence, functions, catalysis, and acquisition of substrate.
      ,
      • O'Donnell V.B.
      • Murphy R.C.
      New families of bioactive oxidized phospholipids generated by immune cells: identification and signaling actions.
      ). Similarly, the endocannabinoid 2-arachidonylglycerol is a substrate for COX-2 and is metabolized to PGH2 glycerol ester as effectively as free arachidonic acid (
      • Kozak K.R.
      • Rowlinson S.W.
      • Marnett L.J.
      Oxygenation of the endocannabinoid, 2-arachidonylglycerol, to glyceryl prostaglandins by cyclooxygenase-2.
      ). The final products derived from this direct PL oxygenation pathway include esterified PGs as well as 11-HETE and 15-HETE. PUFAs contained in PLs can also be oxidized by nonenzymatic reactions. Free radical peroxidation reactions observed under conditions of oxidative stress can freely proceed on intact PLs resulting in the formation of isoprostanes (
      • Morrow J.D.
      • Awad J.A.
      • Boss H.J.
      • Blair I.A.
      • Roberts L.J.
      Non-cyclooxygenase-derived prostanoids (F2-isoprostanes) are formed in situ on phospholipids.
      ).
      Previously, we and others applied LC-MS/MS protocols to test whether plasma levels of oxylipins can be used as biomarkers to differentiate the progressive form of nonalcoholic fatty liver disease, termed nonalcoholic steatohepatitis, from the milder form termed nonalcoholic fatty liver. In that study, we identified a panel of nonesterified oxylipins that when used together is able to discriminate nonalcoholic steatohepatitis from nonalcoholic fatty liver with a high degree of certainty (
      • Loomba R.
      • Quehenberger O.
      • Armando A.
      • Dennis E.A.
      Polyunsaturated fatty acid metabolites as novel lipidomic biomarkers for noninvasive diagnosis of nonalcoholic steatohepatitis.
      ). Another study used an approach that included an alkaline hydrolysis step with the aim of measuring the sum total of free and esterified oxylipins (
      • Feldstein A.E.
      • Lopez R.
      • Tamimi T.A.
      • Yerian L.
      • Chung Y.M.
      • Berk M.
      • Zhang R.
      • McIntyre T.M.
      • Hazen S.L.
      Mass spectrometric profiling of oxidized lipid products in human nonalcoholic fatty liver disease and nonalcoholic steatohepatitis.
      ). Of the markers monitored, products derived from free radical-mediated oxidation of linoleic acid were reported to be significantly elevated in nonalcoholic steatohepatitis. These results differed significantly from our findings, but can in part be explained by the difference in the experimental approach, as we measured the free oxylipins present in plasma, not those appearing after alkaline hydrolysis (
      • Loomba R.
      • Quehenberger O.
      • Armando A.
      • Dennis E.A.
      Polyunsaturated fatty acid metabolites as novel lipidomic biomarkers for noninvasive diagnosis of nonalcoholic steatohepatitis.
      ). In order to quantitatively capture the sum total of esterified and free oxylipins, all plasma samples need to be hydrolyzed, which requires strong alkaline conditions to quantitatively release the oxidized PUFAs before analysis. However, neither any specific experimental conditions nor systematic testing of the effect of strong bases on eicosanoid stability were reported in the later study (
      • Feldstein A.E.
      • Lopez R.
      • Tamimi T.A.
      • Yerian L.
      • Chung Y.M.
      • Berk M.
      • Zhang R.
      • McIntyre T.M.
      • Hazen S.L.
      Mass spectrometric profiling of oxidized lipid products in human nonalcoholic fatty liver disease and nonalcoholic steatohepatitis.
      ). In contrast, in the present study, we have specifically determined the stability of the oxylipins under the hydrolysis conditions employed, and compiled a list of metabolites that can be reproducibly measured in biological samples.
      From previous studies in our and other laboratories, we know that eicosanoids and specifically PGs are sensitive to alkaline-induced degradation. The objective of the current study was to develop precise conditions to minimize degradation of lipid metabolites during alkaline treatment and to identify specific eicosanoids and related oxidized PUFAs that are released intact from esterified lipids and which can be quantitatively included in searches for potential biomarkers.

      MATERIALS AND METHODS

      Reagents

      All solvents were ultra-performance LC (UPLC) grade and were purchased from Fisher Scientific (Waltham, MA). All primary standards (PSTDs) for standard curves and deuterated internal standards (ISTDs) were purchased from Cayman Chemicals (Ann Arbor, MI) or Enzo Life Sciences (Farmingdale, NY). Strata-X polymeric reversed phase columns were purchased from Phenomenex (Torrance, CA). Human plasma was purchased from Gemini Bio Products (West Sacramento, CA).

      Cell culture

      RAW 264.7 cells (ATCC) were used in all cell experiments. Briefly, 4x1E5 cells were plated into each well of a 6-well plate and cultured overnight in 2 ml of DMEM containing 10% FBS (Atlanta Biologicals, Flowery Branch, GA) and were grown overnight at 37°C and 5% CO2. For the experiment, the culture medium was exchanged with 2 ml of DMEM without FBS, and the cells were primed with Kdo2-lipid A (Avanti Polar Lipids, Alabaster, AL) at 100 ng/ml for 4 h, then stimulated with ATP (5 mM) for an additional 20 h. At the end of the incubation period, DMEM was removed and cells were harvested into 1 ml of PBS, counted, homogenized, and both cell homogenate and medium fractions were frozen. For analysis, we used 500 μl of the cell homogenates.

      Lipid extraction

      Free eicosanoids.

      For the extraction of free eicosanoids, 50 μl of plasma, 500 μl of cell homogenates, or the PSTD collection consisting of 136 individual standards were spiked with 100 μl of the ISTD mix (1 ng of each of 26 deuterated standards in ethanol) and diluted with Dulbecco's PBS to give a 10% total ethanol concentration (
      • Wang Y.
      • Armando A.M.
      • Quehenberger O.
      • Yan C.
      • Dennis E.A.
      Comprehensive ultra-performance liquid chromatographic separation and mass spectrometric analysis of eicosanoid metabolites in human samples.
      ). Eicosanoids were then isolated by solid phase extraction (SPE) using Strata-X polymeric reversed phase columns. The columns were activated with consecutive washes of 3 ml of 100% methanol and 3 ml of water. The samples were loaded and washed with 3 ml of 10% methanol. Eicosanoids were then eluted with 1 ml of 100% methanol, dried under vacuum, and dissolved in 50 μl of buffer A consisting of water/acetonitrile/acetic acid (60/40/0.02, v/v/v). Samples were immediately analyzed using UPLC-MS/MS. A complete list of all PSTDs used for standard curves and deuterated standards and their assignments for normalization is provided in supplemental Table S1.

      Total eicosanoids.

      To extract total eicosanoids, 50 μl of plasma, 500 μl of cell homogenates, or 50 μl of the PSTDs mix were spiked with ISTDs (in 100 μl of ethanol) and added to a mixture consisting of 100 μg of butylated hydroxytoluene (in 100 μl ethanol), 250 μl methanol, 50 μl KOH (4 M), and water to a final volume of 1 ml (Fig. 1). The mixture was kept for 30 min at 37°C to hydrolyze the esterified eicosanoids. Following hydrolysis, 3.5 ml of glycine-HCl buffer (0.1 mM, pH 4) were added. The free eicosanoids were then isolated by SPE and analyzed according to the protocol for the free eicosanoids, as described above. An unadulterated mix of pure PSTDs and ISTDs that was not subjected to hydrolysis conditions or SPE served as a control to estimate recoveries.
      Figure thumbnail gr1
      Fig. 1Experimental flow chart for the analysis of free and esterified eicosanoids and other oxylipins.

      Separation and quantification of eicosanoids

      Chromatographic separation.

      Separation was performed on an Acquity UPLC system (Waters, Milford, MA), equipped with a RP C18 BEH shield column (2.1 × 100 nm; 1.7 μm; Waters). For the separation of eicosanoids, a binary buffer system was used consisting of buffer A (described above) and buffer B composed of acetonitrile/2-propanol (50/50, v/v). At a flow rate of 0.5 ml/min, buffer A was held at 100% for 1 min followed by a gradient over 3 min to 55% buffer B, then further increased over 1.5 min to 100% buffer B and kept at this level for 0.5 min. The starting conditions were reconstituted in 1 min. The column was kept at 40°C and the sample manager at 4°C. The samples (10 μl) were injected via partial loop injection using needle overfill mode. To minimize carryover, needle washes were carried out between samples.

      MS data acquisition.

      Data were collected on an AB/Sciex 6500 QTRAP hybrid triple quadrupole mass spectrometer (Sciex, Framingham, MA) using negative electrospray and scheduled multiple reaction monitoring (MRM) mode. The source settings were as follows: curtain gas (CUR = 20 psi), nebulizer gas (GS1 = 30 psi), turbo heater gas (GS2 = 20 psi), electrospray voltage (TEM = −4,500 V), source temperature (500°C), and collision gas (CAD = medium).

      Quantitation.

      Eicosanoids were quantified by the stable isotope dilution method. Briefly, identical amounts of ISTDs were added to each sample and to all the PSTDs. Nine point standard curves were generated for each of the 136 PSTDs, ranging from 0.03 ng to 10 ng. To calculate the amount of eicosanoids in a sample, ratios of peak areas between endogenous eicosanoids and matching deuterated internal eicosanoids were calculated. Ratios were converted to absolute amounts by linear regression analysis of standard curves. Currently, we quantify most eicosanoids at low femtomole levels.
      To determine recovery values of the PSTDs under alkaline hydrolysis conditions, MS peak areas were compared before and after the addition of base. All measurements were performed in triplicate or five replicate measurements and the data are reported as averaged values. The coefficient of variance (CV) determines the precision of this quantitation method.

      PL measurements

      The PL measurements were carried out by LC-MS using a multiplex approach as previously described (
      • Baker P.R.
      • Armando A.M.
      • Campbell J.L.
      • Quehenberger O.
      • Dennis E.A.
      Three-dimensional enhanced lipidomics analysis combining UPLC, differential ion mobility spectrometry, and mass spectrometric separation strategies.
      ). Briefly, 50 μl of human plasma were hydrolyzed as described above and extracted according to Bligh and Dyer (
      • Bligh E.G.
      • Dyer W.J.
      A rapid method of total lipid extraction and purification.
      ). As a control, nonhydrolyzed plasma was included. The organic solvent was removed and the lipids were reconstituted in buffer A (isopropanol/hexane/water = 59/40/1, v/v/v, with 10 mM ammonium acetate) and analyzed on a Waters Acquity UPLC-Sciex 6500 QTrap mass spectrometer system. The lipids were separated on a silica column (2.1 × 150 mm; 3 μm; Phenomenex) using a binary solvent gradient from 100% A to 100% B (isopropanol/hexane/water = 50/40/10, v/v/v, with 10 mM ammonium acetate) over 16 min. PL molecular species (432 isobaric species in total) were identified by MS using precursor ion and neutral loss scans in positive and negative modes. For the PL class analysis, the sum total of individual molecular species within each class was used.

      RESULTS AND DISCUSSION

      Quantitation of eicosanoid-related oxylipins in their free nonesterified form

      Eicosanoids are important lipid metabolites that are involved in a number of physiological processes at the cellular level. As with fatty acids, they can exist either in the free form or esterified to complex lipids, such as PLs. The objective of the current study was to develop precise conditions that allow the quantitative measurement of both free and esterified oxylipins. To achieve this, we first examined the recovery of free eicosanoids during the prepurification step prior to LC-MS analysis using a defined set of quantitation standards consisting of 161 authentic metabolites that were mixed at precisely measured concentrations. Included in the standard cocktail were also 26 deuterated analogs that can be used to offset any potential losses during workup. The standard mix was then divided into two aliquots and analyzed by LC-MS with or without SPE prepurification. Figure 2 shows the recovery of a subset of eicosanoids that were selected based on their metabolic pathway. In order to assess the degree of any potential losses during sample preparation and SPE purification, we plotted the raw MS data after SPE without normalization as percent recovery compared with the raw MS data obtained with standards set that did not undergo SPE purification. As can be seen, the recovery of these lipid metabolites in their free nonesterified form and undergoing our standard purification procedure was largely quantitative, ranging between 90% and 100%, even without normalization to ISTDs (Fig. 2). Any potential losses were minimal and could easily be offset with the application of our routine normalization procedure using deuterated eicosanoid analogs as ISTDs. A complete list of the recoveries for all metabolites as well as ISTD assignments, pertinent technical information, and instrument settings are provided in the supplemental Table S1.
      Figure thumbnail gr2
      Fig. 2Recoveries of oxylipin standards in their free form after SPE. A collection of primary oxylipin standards was analyzed with and without SPE purification. Recoveries were determined by comparing MS intensities obtained with standards after undergoing SPE purification with MS intensities of standards without SPE purification that were analyzed in parallel and served as controls. The open bars show non-normalized recoveries (Non-Norm), which were calculated from the raw MS peak areas without normalization to ISTDs; the closed bars show the same data set but normalized to ISTD (Norm). All data are expressed as percent of untreated controls. The mean and SD of triplicate measurements are displayed. Shown is a representative subset of standards (see for the complete data set).

      Optimization of hydrolysis conditions to preserve oxylipin structure

      A number of studies reported that a portion of eicosanoids are naturally esterified and can be contained in membrane lipids in the form of esters. To profile quantitatively all eicosanoids incorporated into the various lipid fractions using an approach that preserves the intact molecule represents an enormous technical challenge. An alternative approach is to release the eicosanoids first by hydrolysis and then measure the metabolites in their free form. Several laboratories have applied alkaline hydrolysis for this purpose; however, the hydrolysis conditions varied considerably as they were often optimized for the analysis of certain subclasses of eicosanoids, including isoprostanes, fatty acid alcohols, ketones, and epoxides (
      • Morrow J.D.
      • Awad J.A.
      • Boss H.J.
      • Blair I.A.
      • Roberts L.J.
      Non-cyclooxygenase-derived prostanoids (F2-isoprostanes) are formed in situ on phospholipids.
      ,
      • Mallat Z.
      • Nakamura T.
      • Ohan J.
      • Leseche G.
      • Tedgui A.
      • Maclouf J.
      • Murphy R.C.
      The relationship of hydroxyeicosatetraenoic acids and F2-isoprostanes to plaque instability in human carotid atherosclerosis.
      ,
      • Newman J.W.
      • Kaysen G.A.
      • Hammock B.D.
      • Shearer G.C.
      Proteinuria increases oxylipid concentrations in VLDL and HDL but not LDL particles in the rat.
      ,
      • Keenan A.H.
      • Pedersen T.L.
      • Fillaus K.
      • Larson M.K.
      • Shearer G.C.
      • Newman J.W.
      Basal omega-3 fatty acid status affects fatty acid and oxylipin responses to high-dose n3-HUFA in healthy volunteers.
      ,
      • Shearer G.C.
      • Harris W.S.
      • Pedersen T.L.
      • Newman J.W.
      Detection of omega-3 oxylipins in human plasma and response to treatment with omega-3 acid ethyl esters.
      ). Considering that many oxylipins are unstable under extreme alkaline or acidic conditions (
      • Stehle R.G.
      • Oesterling T.O.
      Stability of prostaglandin E1 and dinoprostone (prostaglandin E2) under strongly acidic and basic conditions.
      ,
      • Demers L.M.
      • Brennecke S.P.
      • Mountford L.A.
      • Brunt J.D.
      • Turnbull A.C.
      Development and validation of a radioimmunoassay for prostaglandin E2 metabolite levels in plasma.
      ,
      • Younger E.W.
      • Szabo R.M.
      The stability of prostaglandin E1 in dilute physiological solutions at 37 degrees C.
      ), it is important to balance hydrolysis efficiency and structural preservation of the analytes. To achieve this, we explored mild alkaline conditions for their efficacies to hydrolyze oxylipins esterified to complex lipids, including PLs. We established the optimal base concentration at 0.2 M KOH and tested the hydrolysis efficiency at this concentration on human plasma at various temperatures and incubation times. The majority of base-stable metabolites that are generated by enzymes, including the fatty acid epoxides, are contained in PLs (
      • Spector A.A.
      • Fang X.
      • Snyder G.D.
      • Weintraub N.L.
      Epoxyeicosatrienoic acids (EETs): metabolism and biochemical function.
      ). Thus, we focused on the PL fraction to measure hydrolysis efficiency. The mass chromatograms for several PL classes taken before and after base hydrolysis indicated that the mildest condition, 0.2 N KOH at 37°C for 30 min, was sufficient to hydrolyze >95% of the plasma PLs (Fig. 3). There was some remaining sphingomyelin, which contains N-linked fatty acids that are more resistant to hydrolysis, even at 60°C. No lysoPLs were detectable post hydrolysis, which indicates that the PLs were not converted to the lyso moieties and the hydrolysis step effectively released all sn1 and sn2 fatty acids (Fig. 3B). We also subjected some selected oxylipin standards to the same conditions and observed that at 37°C and 30 min, the base-induced destruction of these metabolites was least, as exemplified by the recovery of intact 7-hydroxy-docosahexaenoic acid (Fig. 4). Like the nondeuterated metabolites, the degradation of the deuterated ISTDs increased similarly with increasing temperature and time. As a result, the sensitivity and precision of the analysis decreases proportionally. Our data show that the hydrolysis condition of 0.2 N KOH at 37°C for 30 min provides the optimal balance between hydrolysis efficiency and structural preservation of the analytes. Deviating from these conditions augments metabolite degradation and low abundance metabolites may fall below the lower limit of detection. Furthermore, reproducibility decreases with increasing degradation and, consequently, the precision of the analysis deteriorates.
      Figure thumbnail gr3
      Fig. 3Alkaline hydrolysis efficiency. To determine the hydrolysis efficiency, lipids in human plasma were hydrolyzed with 0.2 N potassium hydroxide for the indicated times and temperatures. Remaining intact PLs (A) and lysoPLs (B) were then measured by LC-MS. Shown are the chromatograms for the PL classes, phosphatidylcholine (PC), phosphatidylethanolamine (PE), and phosphatidylinositol (PI), as well as the respective lyso-lipids. The mildest condition tested (37°C, 30 min) was sufficient to effectively hydrolyze all PLs and lysoPLs. Ether, ether-linked PE.
      Figure thumbnail gr4
      Fig. 4Recovery of 7-hydroxy-docosahexaenoic acid (7-HDoHE) subjected to alkaline conditions. The 7-HDoHE is shown as a representative example to test the stability of related hydroxylated PUFAs. Three conditions were tested: A, no saponification; B, 37°C for 30 min; C, 37°C for 60 min; and D, 60°C for 30 min. The mass spectral response for 7-HDoHE is shown in panel A. The average relative intensity of triplicate measurements is shown in panel B. The untreated control was set at 100%. As can be seen, there was a time- and temperature-dependent decline in the recovery of 7-HDoHE.

      Analysis of eicosanoids after alkaline treatment

      Next, we expanded our stability tests and examined our entire library of PSTDs for their resistance to base-induced degradation (Fig. 5). As shown in Fig. 6 and supplemental Table S2, many of the eicosanoids are susceptible to base-induced degradation, as indicated by the changes in their mass spectral intensities. The data shown represent the average of five replicate measurements performed on a single day. In particular, PGs and leukotrienes (LTs) were virtually undetectable. The exceptions were PGF2a and LTB4, which were resistant to degradation (supplemental Table S2). Additionally, fatty acids containing hydroxy, di-hydroxy, or epoxy groups showed resistance to degradation to various degrees. Even though some degradation may have occurred during the saponification step, when we normalized the spectral data of the nondeuterated eicosanoids to their deuterated analogs, we were able to neutralize the loss for many of the eicosanoids. For example, the nonnormalized “raw” recovery of thromboxane (TX)B2 was about 78%. However, when we normalized the value to its deuterated analog measured under identical conditions, the recovery was calculated to be 100%. Similarly, the normalized recoveries were quantitative for 5-HETE, 13-HODE, and 9,10-EpOME, compared with the nonnormalized recoveries of about 70–80% based on raw mass spectral peak areas (Fig. 6). As a class, the epoxides and diols are quite stable under basic conditions (
      • Newman J.W.
      • Kaysen G.A.
      • Hammock B.D.
      • Shearer G.C.
      Proteinuria increases oxylipid concentrations in VLDL and HDL but not LDL particles in the rat.
      ), and many of them were recovered quantitatively from the standard mix under alkaline hydrolysis conditions (Table 1). For some eicosanoids, normalization to ISTDs did not improve the overall recovery. This applies specifically to the ones with more complex structural elements, including PGs and LTs as well as ketones (supplemental Table S2).
      Figure thumbnail gr5
      Fig. 5MRM chromatogram of oxylipin standards subjected to alkaline hydrolysis conditions. A set of pure oxylipin standards (136 metabolites) was subjected to alkaline hydrolysis conditions, purified by SPE, and analyzed by LC-MS. Shown is the complete MRM chromatogram of all 136 metabolites. Inset: As an example and to demonstrate peak quality, the MRMs for 5-HETE, 9-HODE, and 13-HODE were extracted and shown individually.
      Figure thumbnail gr6
      Fig. 6Recoveries of oxylipins under alkaline hydrolysis conditions. A set of pure oxylipin standards (136 metabolites) was subjected to alkaline hydrolysis conditions, purified by SPE, and analyzed by LC-MS. Recoveries were determined by comparing MS intensities of the standards after alkaline treatment with MS intensities of the untreated standards that were analyzed in parallel and served as controls. The open bars show non-normalized recoveries (Non-Norm), which were calculated from the raw MS peak areas without normalization to ISTDs; the closed bars show the same data set but normalized to ISTD (Norm). All data are expressed as percent of untreated controls. The mean and SD of five replicate measurements are displayed. Shown is a representative subset of standards (see for the complete data set).
      TABLE 1List of eicosanoids and related oxylipins that can be reliably measured after alkaline hydrolysis
      PathwayAnalyteDay 1 Recovery (%)Day 2 Recovery (%)Day 3 Recovery (%)Average Recovery (%)CV (%)
      COX11 HEPE9482818510
      11 HETE9698102988
      12 HHTRE86911029311
      PGF1a1109410410212
      d17 6k PGF1a1179211710814
      6k PGF1a10587999710
      PGF2a1029210510011
      2,3 dinor 11b PGF2a104103941019
      dh PGF2a10992899613
      20oh PGF2a1101079010311
      PGF3a979192936
      TXB11001051011026
      TXB2101110951037
      TXB3981179410311
      LOX5 HETE1009389948
      8 HETE991011061027
      12 HETE100105981007
      Tetranor 12-HETE1149911811311
      15 HETE102106951009
      5 HETrE1161059710610
      8 HETrE105104971017
      15 HETrE105110981059
      9 HODE92104101998
      13 HODE11811610311111
      9 HOTrE981091091069
      13 HOTrE (y)11311610511012
      LTB49110093957
      14 HDoHE979395957
      17 HDoHE107101919910
      7(R) Maresin-110180868812
      CYP9,10 EpOME991051071047
      12,13 EpOME9810098997
      18 HEPE9080838410
      5,15 diHETE108831089915
      19,20 diHDPA11185939715
      8,9 diHETrE10088849012
      11,12 diHETrE1139411911015
      14,15 diHETrE10099859512
      9,10 diHOME107971031036
      12,13 diHOME1071031011035
      8,9 EET9793949615
      11,12 EET10698971005
      14,15 EET104941021018
      14,15 EpETE878593889
      Non-enzyme4 HDoHE10686919412
      7 HDoHE878180829
      8 HDoHE97100909510
      10 HDoHE99107981017
      11 HDoHE10196919512
      13 HDoHE1201109310713
      16 HDoHE1009993976
      20 HDoHE10010294997
      9 HETE1041061011029
      8 iso PGF2a III10391103999
      2,3 dinor 8-iso PGF2a9998919610
      8-iso PGF3a96105969812
      The set of PSTDs was supplemented with ISTDs, subjected to alkaline hydrolysis conditions, and analyzed by LC-MS. An identical set of untreated standards was analyzed in parallel and served as control. The analyses were performed in five replicates on each of three consecutive days. Recoveries after alkaline treatment were calculated by comparing the mass spectral intensities with those of untreated standards and after normalization to ISTDs. Shown is the list of metabolites that were recovered at 80–120% and with a CV of ≤15%. A list of all abbreviations is provided on the website (http://www.lipidmaps.org/data/standards/standards.php?lipidclass=LMFA).
      Most naturally occurring PGs have a considerable potential for hydrolysis, dehydration, or isomerization, depending on their immediate environment (
      • Fitzpatrick F.A.
      • Wynalda M.A.
      Albumin-lipid interactions: prostaglandin stability as a probe for characterizing binding sites on vertebrate albumins.
      ). PGs contain multiple hydroxyl groups and keto groups and a rigid five-member prostane ring. The resulting β-hydroxy ketone system is unstable and readily undergoes dehydration under acidic or basic conditions to A- or B-type PGs (
      • Stehle R.G.
      • Oesterling T.O.
      Stability of prostaglandin E1 and dinoprostone (prostaglandin E2) under strongly acidic and basic conditions.
      ,
      • Stehle R.G.
      Physical chemistry, stability, and handling of prostaglandins E2, F2α, D2, and I2: A critical summary.
      ). Alternatively, PGD and PGE can oxidize to the 9,11-diketones, PGK1, and PGK2, which were generated and increased about 3-fold during alkaline hydrolysis. Furthermore, bicyclo PGE2, a base-catalyzed breakdown product of PGE2 and 13,14-dihydro-15-keto PGE2 (dhk PGE2) was found to be substantially increased after alkaline hydrolysis (supplemental Table S2). These breakdown or conversion products cannot be reliably measured after alkaline hydrolysis and should not be included in quantitative analyses of esterified eicosanoids.

      Precision of the method

      For the method to be applicable to biological samples, it has to be accurate and reliable. For this purpose, we compiled a panel of eicosanoids that were either resistant to base-induced degradation or suffered only minor destruction and were reproducibly recovered. Each metabolite in our library of standards was measured in five replicate measurements on three consecutive days with and without saponification, normalized to ISTDs, and averaged. The recovery after alkaline hydrolysis was calculated and the precision was expressed as the coefficient of variation.
      For practical purposes, we used a cut-off point of 80–120% normalized recovery and a precision (CV) of 15% or less to assemble a list of eicosanoids that can be reproducibly measured (Table 1). Of the 136 eicosanoids in the standard mix that we used for this purpose (supplemental Table S2), 56 metabolites satisfied these criteria and were included in the list. The 3 day average was close to 100% recovery for most of these metabolites. These data indicated that the assay is reproducible and, with careful selection of ISTDs for normalization and neutralization of potential losses, the method is useful for the analysis of selected esterified eicosanoids.

      Application of the method to measure esterified eicosanoids in biological samples

      To establish usefulness, we applied this protocol to the analysis of esterified eicosanoids in biological samples, including RAW cells, a cell model of mouse macrophages (
      • Buczynski M.W.
      • Stephens D.L.
      • Bowers-Gentry R.C.
      • Grkovich A.
      • Deems R.A.
      • Dennis E.A.
      TLR-4 and sustained calcium agonists synergistically produce eicosanoids independent of protein synthesis in RAW264.7 cells.
      ,
      • Dennis E.A.
      • Deems R.A.
      • Harkewicz R.
      • Quehenberger O.
      • Brown H.A.
      • Milne S.B.
      • Myers D.S.
      • Glass C.K.
      • Hardiman G.T.
      • Reichart D.
      • et al.
      A mouse macrophage lipidome.
      ), and human plasma (
      • Quehenberger O.
      • Armando A.M.
      • Brown A.H.
      • Milne S.B.
      • Myers D.S.
      • Merrill A.H.
      • Bandyopadhyay S.
      • Jones K.N.
      • Kelly S.
      • Shaner R.L.
      • et al.
      Lipidomics reveals a remarkable diversity of lipids in human plasma.
      ). For the cell model, mouse RAW macrophages were activated with the Toll-like receptor agonist Kdo2-lipid A, and ATP and eicosanoids were analyzed in their free and esterified form in both stimulated and unstimulated control cells. Considering the limitations outlined in Table 1 and using the algorithm for the identification of stable metabolites, we identified and quantified a number of eicosanoids that were present in their free and esterified form (Table 2). In total, we detected 34 metabolites that met the criteria for inclusion. In the unstimulated control cells, the COX-derived metabolites, 11-HETE and PGF2a, were present only in their free form. In contrast, metabolites formed by cytochrome P450 and nonenzymatic pathways were mainly found in their esterified form. As can be seen, most eicosanoids increased during stimulation. The eicosanoid ratio (esterified vs. free) in the stimulated cells is a reflection of de novo synthesis, hydrolysis by phospholipase A2, and release into the extracellular medium. Of course it is possible that some eicosanoids are tightly bound to receptors or carrier proteins even during SPE purification, but are released upon base hydrolysis and would therefore be included in the “esterified” measure. Additionally, it is well-established that PGs, including PGF2a, are formed and rapidly secreted during stimulation with Toll-like receptor agonists and ATP (
      • Buczynski M.W.
      • Stephens D.L.
      • Bowers-Gentry R.C.
      • Grkovich A.
      • Deems R.A.
      • Dennis E.A.
      TLR-4 and sustained calcium agonists synergistically produce eicosanoids independent of protein synthesis in RAW264.7 cells.
      ). Consequently, there may be a relative increase in the esterified portion due to secretion of metabolites in their free form. Of note, the data contain the cell-associated free eicosanoids and do not include the secreted fraction.
      TABLE 2Total eicosanoids and other oxylipins in stimulated RAW macrophages that satisfy our selection criteria for accurate measurement
      PathwayAnalyteTotal OxylipinsFree OxylipinsEsterified
      StimulatedControlStimulatedControlStimulatedControl
      MeanSDMeanSDMeanSDMeanSD
      pmol/1E6 cellspmol/1E6 cells%%
      COX11 HEPENDND0.180.02NDNDNDNDND100
      11 HETE2.480.430.330.031.840.270.350.06260
      PGF2a1.360.150.290.021.090.110.290.01190
      LOX5 HETE1.85*0.481.250.050.16*0.040.210.059283
      8 HETE0.230.030.420.070.03*0.010.020.008994
      12 HETE0.330.030.180.01NDNDNDND100100
      15 HETE1.070.160.280.030.590.130.210.064525
      5 HETrE0.330.020.190.030.07*0.010.070.018062
      8 HETrE0.14*0.050.110.02NDND0.020.0110083
      15 HETrE0.130.030.030.000.080.010.020.01378
      9 HODE0.660.140.230.040.710.290.190.03016
      13 HODE0.230.010.110.030.180.040.050.011951
      13 HOTrE (y)0.160.040.030.00NDNDNDND100100
      14 HDoHE0.400.040.150.020.140.010.020.016686
      CYP9,10 EpOME0.780.150.940.170.130.020.050.018395
      12,13 EpOME0.680.130.790.120.150.030.050.017794
      18 HEPENDND0.150.02NDNDNDNDND100
      11,12 diHETrE0.070.030.110.02NDNDNDND100100
      19,20 diHDPA0.120.050.160.02NDND0.020.0010089
      9,10 diHOME0.080.010.030.020.050.030.010.004672
      12,13 diHOME0.070.010.040.010.050.020.010.003469
      8,9 EET0.69*0.300.870.14NDND0.030.0210097
      11,12 EET1.560.322.520.28NDNDNDND100100
      14,15 EET2.030.613.660.74NDND0.040.0110099
      14,15 EpETENDND0.320.16NDNDNDNDND100
      Non-enzyme4 HDoHE2.010.152.530.260.160.000.080.019297
      7 HDoHE0.43*0.050.410.06NDNDNDND100100
      8 HDoHE0.930.311.430.36NDND0.10.0010093
      10 HDoHE0.200.040.100.02NDND0.030.0110075
      11 HDoHE0.23*0.040.220.030.06*0.030.040.027683
      13 HDoHE0.630.170.090.020.240.080.060.016236
      16 HDoHE0.250.010.100.020.03*0.010.020.018682
      20 HDoHE0.670.120.390.06NDND0.070.0310081
      9 HETE0.290.050.550.11NDNDNDND100100
      RAW macrophages were stimulated with KdO2-lipid A and ATP and the total eicosanoids were measured in lipid extracts after saponification (Stimulated). As a control, unstimulated cells were processed identically (Control). For comparison, the levels of cell-associated free metabolites were measured in parallel. The mean and SD of triplicate measurements are displayed. The esterified fraction is expressed as percent of the total (Esterified). ND, not detected. All metabolites changed significantly (P < 0.05) upon stimulation, except the ones marked with an asterisk (*). A list of all abbreviations is provided on the website (http://www.lipidmaps.org/data/standards/standards.php?lipidclass=LMFA).
      We next examined esterified eicosanoids in human plasma. In all, we recovered 38 eicosanoids after base hydrolysis (Table 3). For comparison, we also measured these metabolites in their free form as previously reported (
      • Quehenberger O.
      • Dennis E.A.
      The human plasma lipidome.
      ,
      • Quehenberger O.
      • Armando A.M.
      • Brown A.H.
      • Milne S.B.
      • Myers D.S.
      • Merrill A.H.
      • Bandyopadhyay S.
      • Jones K.N.
      • Kelly S.
      • Shaner R.L.
      • et al.
      Lipidomics reveals a remarkable diversity of lipids in human plasma.
      ). The dynamic range for the total eicosanoids spans several orders of magnitude from about 5 to 44,500 pmol/ml and, on average, about 70% of all eicosanoids were esterified. A potential matrix effect adds to the complexity of the measurement. The matrix effect was determined previously by spiking the ISTD mixture into a plasma sample (
      • Wang Y.
      • Armando A.M.
      • Quehenberger O.
      • Yan C.
      • Dennis E.A.
      Comprehensive ultra-performance liquid chromatographic separation and mass spectrometric analysis of eicosanoid metabolites in human samples.
      ). The results showed that the recovery of ISTDs that are resistant to alkaline-induced degradation is greater than 80%, indicating that matrix effects and ion suppression are minimal. The plasma measurements were carried out on 50 μl of plasma of which the equivalent of 10 μl was injected into the UPLC/MS for an analysis. Thus, the lower limits of quantification for eicosanoid measurements in biological material are in the low femtomole range.
      TABLE 3Total eicosanoids and other oxylipins in human plasma that satisfy our selection criteria for accurate measurement
      PathwayAnalyteTotal OxylipinsFree OxylipinsEsterified (%)
      Mean (pmol/ml)SD (pmol/ml)Mean (pmol/ml)SD (pmol/ml)
      COX11 HEPE335.0340.59104.9417.6769
      11 HETE4,692.851,011.692,400.561,112.2149
      PGF2a20.942.1112.492.3840
      LOX5 HETE8,447.45579.754,493.63547.7147
      8 HETE9,495.391,504.951,750.21207.6082
      Tetranor 12-HETE42.494.086.971.4684
      12 HETE10,988.431,631.181,865.15179.1483
      15 HETE7,386.67744.521,883.5144.2875
      5 HETrE143.8627.52103.807.5228
      8 HETrE7,057.361,139.551,317.45130.9681
      15 HETrE1,076.25171.56280.087.3874
      9 HODE17,145.923,441.526,628.18721.6561
      13 HODE14,788.06564.636,065.741,056.6659
      9 HOTrE565.75155.94114.863.4780
      13 HOTrE (y)244.5517.0786.167.6865
      14 HDoHE1,459.56206.03467.66207.0968
      17 HDoHE1,888.64297.49359.2154.1781
      CYP9,10 EpOME19,274.99857.236,024.431,114.2869
      12,13 EpOME23,762.051,515.638,998.901,662.6262
      18 HEPE300.2939.73103.159.8066
      5,15 diHETE57.977.7366.8932.400
      8,9 diHETrE14.373.104.210.6171
      11,12 diHETrE4.891.341.350.4272
      14,15 diHETrE11.910.403.780.6768
      9,10 diHOME27.071.457.221.0373
      12,13 diHOME10.271.176.411.0338
      8,9 EET8,719.221,600.981,308.17179.9085
      14,15 EET90.289.6926.412.8271
      14,15 EpETE6,405.69469.561,871.3320.3071
      Non-enzyme4 HDoHE827.04142.28262.8247.5568
      7 HDoHE314.5944.9276.1615.1276
      8 HDoHE11,437.321,610.951,187.05110.7090
      10 HDoHE64.5510.6823.634.4163
      11 HDoHE1,700.85311.31337.7923.5180
      13 HDoHE670.10108.67176.5460.9974
      16 HDoHE1,046.17144.11257.299.2675
      20 HDoHE1,392.5783.96168.070.5788
      9 HETE44,523.251,009.615,479.52355.1988
      For comparison, the levels of the free metabolites isolated without saponification were measured in parallel. The mean and SD of triplicate measurements are displayed. The esterified fraction is expressed as percent of the total (esterified). A list of all abbreviations is provided on the website (http://www.lipidmaps.org/data/standards/standards.php?lipidclass=LMFA).

      CONCLUSION

      In this study, we assessed the suitability of base hydrolysis to quantify esterified eicosanoids in biological material. We showed that about 75% of all cell-associated eicosanoids were esterified. Globally this percentage remained largely unchanged between control and Kdo2-lipid A/ATP-stimulated cells, even though some selected metabolites increased either in their free or esterified form. In a previous report, we demonstrated the presence of free eicosanoids in human plasma (
      • Quehenberger O.
      • Armando A.M.
      • Brown A.H.
      • Milne S.B.
      • Myers D.S.
      • Merrill A.H.
      • Bandyopadhyay S.
      • Jones K.N.
      • Kelly S.
      • Shaner R.L.
      • et al.
      Lipidomics reveals a remarkable diversity of lipids in human plasma.
      ) and found that the levels are correlated with various inflammatory and metabolic diseases, including nonalcoholic fatty liver disease (
      • Loomba R.
      • Quehenberger O.
      • Armando A.
      • Dennis E.A.
      Polyunsaturated fatty acid metabolites as novel lipidomic biomarkers for noninvasive diagnosis of nonalcoholic steatohepatitis.
      ). In the current study, we expanded our method to include esterified eicosanoids in the analysis. Overall, we fine-tuned the alkaline hydrolysis conditions to minimize metabolite degradation, adjusted the prepurification steps to enhance metabolite recovery, optimized the assignment of ISTDs to compensate for potential losses, and established an algorithm based on recovery and reproducibility to compile a table of metabolites that can be accurately measured. When taking these limitations into consideration, this method can now be successfully applied to accurately measure the sum total, i.e., free and esterified oxylipins, in human plasma and a variety of biological samples.

      Supplementary Material

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