The Contribution of Cytosolic Group IVA and Calcium-Independent Group VIA Phospholipase A2s to Adrenic Acid Mobilization in Murine Macrophages

biomolecules Article The Contribution of Cytosolic Group IVA and Calcium-Independent Group VIA Phospholipase A2s to Adrenic Acid Mobilization in Murine Macrophages Patricia Monge 1,2, Alvaro Garrido 1,2, Julio M. Rubio 1,2, Victoria Magrioti 3, George Kokotos 3, María A. Balboa 1,2 and Jesús Balsinde 1,2,* 1Instituto de Biología y Genética Molecular, Consejo Superior de Investigaciones Científicas (CSIC), Universidad de Valladolid, 47003 Valladolid, Spain; [email protected] (P.M.); [email protected] (A.G.); [email protected] (J.M.R.); [email protected] (M.A.B.) 2Centro de Investigación Biomédica en Red de Diabetes y Enfermedades Metabólicas Asociadas (CIBERDEM), 28029 Madrid, Spain 3Department of Chemistry, National and Kapodistrian University of Athens, Panepistimiopolis, 15771 Athens, Greece; [email protected] (V.M.); [email protected] (G.K.) *Correspondence: [email protected]; Tel.: +34-983-423-062 Received: 16 March 2020; Accepted: 1 April 2020; Published: 3 April 2020   Abstract: Adrenic acid (AA), the 2-carbon elongation product of arachidonic acid, is present at significant levels in membrane phospholipids of mouse peritoneal macrophages. Despite its abundance and structural similarity to arachidonic acid, very little is known about the molecular mechanisms governing adrenic acid mobilization in cells of the innate immune system. This contrasts with the wide availability of data on arachidonic acid mobilization. In this work, we used mass-spectrometry-based lipidomic procedures to define the profiles of macrophage phospholipids that contain adrenic acid and their behavior during receptor activation. We identified the phospholipid sources from which adrenic acid is mobilized, and compared the data with arachidonic acid mobilization. Taking advantage of the use of selective inhibitors, we also showed that cytosolic group IVA phospholipase A 2 is involved in the release of both adrenic and arachidonic acids. Importantly, calcium independent group VIA phospholipase A 2 spared arachidonate-containing phospholipids and hydrolyzed only those that contain adrenic acid. These results identify separate mechanisms for regulating the utilization of adrenic and arachidonic acids, and suggest that the two fatty acids may serve non-redundant functions in cells. Keywords: adrenic acid; arachidonic acid; mass spectrometry; lipid signaling; inflammation; phospholipase A2; monocytes/macrophages 1. Introduction Arachidonic acid (cis-5,8,11,14-eicosatetraenoic acid, AA), a fatty acid of the n-6 series, is the major polyunsaturated fatty acid present in cells of the innate immunity [ 1 ]. Cleavage of AA-containing membrane phospholipids by phospholipase A 2 (PLA 2 ) enzymes during activation results in substantial release of free AA. The free fatty acid is then metabolized by cyclooxygenases, lipoxygenases, or cytochrome P-450 enzymes into numerous oxygenated metabolites, collectively called the eicosanoids, which have key roles in inflammation [ 2 – 6 ]. The eicosanoids participate in immune regulation by influencing the activation of innate immune phagocytic cells at various levels, including differentiation and migration, phagocytic capacity, and cytokine production [7–10]. Innate immune cells also contain significant quantities of another n-6 fatty acid, adrenic acid (cis-7,10,13,16-docosatetraenoic acid, AdA), which is the 2-carbon elongation product of AA [ 1 ] Biomolecules 2020,10, 542; doi:10.3390/biom10040542 www.mdpi.com/journal/biomolecules Biomolecules 2020,10, 542 2 of 16 (Scheme 1). Similar to AA, AdA can also be metabolized via cyclooxygenase, lipoxygenase, and cytochrome P-450 pathways [ 11 – 14 ], giving rise to a number of oxygenated metabolites. Interestingly, a recent study using a mouse model of steatohepatitis reported elevated levels of free AdA in plasma and liver [ 15 ]. These data raised the possibility that alterations in the homeostatic mechanisms regulating AdA levels may be related to pathophysiological states [ 15 ]. Regarding the role of AdA in innate immune cells, it has been shown that the fatty acid is able to dampen inflammation by blocking leukotriene B 4 formation by neutrophils and enhancing phagocytosis by macrophages [ 16 ]. These effects are consistent with a pro-resolving mediator function for AdA in osteoarthritis [16]. Biomolecules 2020, 10, 2 of 16 Innate immune cells also contain significant quantities of another n-6 fatty acid, adrenic acid (cis- 7,10,13,16-docosatetraenoic acid, AdA), which is the 2-carbon elongation product of AA [1] (Scheme 1). Similar to AA, AdA can also be metabolized via cyclooxygenase, lipoxygenase, and cytochrome P-450 pathways [11–14], giving rise to a number of oxygenated metabolites. Interestingly, a recent study using a mouse model of steatohepatitis reported elevated levels of free AdA in plasma and liver [15]. These data raised the possibility that alterations in the homeostatic mechanisms regulating AdA levels may be related to pathophysiological states [15]. Regarding the role of AdA in innate immune cells, it has been shown that the fatty acid is able to dampen inflammation by blocking leukotriene B4 formation by neutrophils and enhancing phagocytosis by macrophages [16]. These effects are consistent with a pro-resolving mediator function for AdA in osteoarthritis [16]. Scheme 1. Structure of arachidonic acid (AA) and adrenic acid (AdA). The positions of the carbons at which double bonds occur are numbered. Note that all double bonds are in the cis configuration. Recent work from our laboratory has taken advantage of mass-spectrometry-based lipidomic approaches to define at a molecular level the mechanisms regulating AA mobilization from phagocytic cells responding to stimuli of the innate immune response [17–27]. Our work has highlighted not only the importance of multiple PLA2 enzymes in the process, but also of the mechanisms regulating the reacylation of the liberated fatty acid back into phospholipids and the remodeling that places the AA in the appropriate cellular lipid pools. These findings have raised the intriguing possibility that not all AA pools may be reachable by the PLA2 form involved in its mobilization. For instance, release of lipoxygenase products by ionophore-activated human neutrophils [28] and zymosan-stimulated mouse peritoneal macrophages [22] appears to be associated with AA mobilization from choline-containing phospholipids (PC), not ethanolaminecontaining phospholipids (PE) or phosphatidylinositol (PI). Thus, depending on stimulation conditions, the cellular distribution of AA between various phospholipid locations may also limit eicosanoid synthesis [3,26]. In contrast to AA studies, very little is still known about the mechanisms governing cellular AdA availability. In this study we have analyzed the regulatory features of AdA mobilization in activated macrophages. Our work provides an in-depth examination of AdA homeostasis under pathophysiologically relevant conditions, and suggests that both cytosolic group IVA PLA2 (cPLA2α) and calcium-independent group VIA PLA2 (iPLA2-VIA) participate in the process. AdA mobilization shares regulatory features with AA mobilization with regard to cPLA2α involvement, but seems to be a more complex process, as it involves participation of a second PLA2 that is apparently not involved in AA release (i.e., iPLA2-VIA). In addition, our study also suggests that supplementation with AdA does not reduce AA utilization by the cells. Thus, the two fatty acids may play nonredundant biological roles that could be exploited to design selective strategies to control the production of AdA-derived products at the level of their precursor fatty acid. 2. Materials and Methods 2.1. Reagents Cell culture medium was from Molecular Probes-Invitrogen (Carlsbad, CA, USA). Organic solvents (Optima® LC/MS grade) were from Fisher Scientific (Madrid, Spain). Lipid standards were from Avanti (Alabaster, AL, USA) or Cayman (Ann Arbor, MI, USA). Silicagel G thin-layer Scheme 1. Structure of arachidonic acid (AA) and adrenic acid (AdA). The positions of the carbons at which double bonds occur are numbered. Note that all double bonds are in the cis configuration. Recent work from our laboratory has taken advantage of mass-spectrometry-based lipidomic approaches to define at a molecular level the mechanisms regulating AA mobilization from phagocytic cells responding to stimuli of the innate immune response [ 17 – 27 ]. Our work has highlighted not only the importance of multiple PLA 2 enzymes in the process, but also of the mechanisms regulating the reacylation of the liberated fatty acid back into phospholipids and the remodeling that places the AA in the appropriate cellular lipid pools. These findings have raised the intriguing possibility that not all AA pools may be reachable by the PLA 2 form involved in its mobilization. For instance, release of lipoxygenase products by ionophore-activated human neutrophils [ 28 ] and zymosan-stimulated mouse peritoneal macrophages [ 22 ] appears to be associated with AA mobilization from choline-containing phospholipids (PC), not ethanolamine-containing phospholipids (PE) or phosphatidylinositol (PI). Thus, depending on stimulation conditions, the cellular distribution of AA between various phospholipid locations may also limit eicosanoid synthesis [3,26]. In contrast to AA studies, very little is still known about the mechanisms governing cellular AdA availability. In this study we have analyzed the regulatory features of AdA mobilization in activated macrophages. Our work provides an in-depth examination of AdA homeostasis under pathophysiologically relevant conditions, and suggests that both cytosolic group IVA PLA 2 (cPLA 2α ) and calcium-independent group VIA PLA 2 (iPLA 2 -VIA) participate in the process. AdA mobilization shares regulatory features with AA mobilization with regard to cPLA 2α involvement, but seems to be a more complex process, as it involves participation of a second PLA 2 that is apparently not involved in AA release (i.e., iPLA 2 -VIA). In addition, our study also suggests that supplementation with AdA does not reduce AA utilization by the cells. Thus, the two fatty acids may play non-redundant biological roles that could be exploited to design selective strategies to control the production of AdA-derived products at the level of their precursor fatty acid. 2. Materials and Methods 2.1. Reagents Cell culture medium was from Molecular Probes-Invitrogen (Carlsbad, CA, USA). Organic solvents (Optima ® LC/MS grade) were from Fisher Scientific (Madrid, Spain). Lipid standards were from Avanti (Alabaster, AL, USA) or Cayman (Ann Arbor, MI, USA). Silicagel G thin-layer chromatography plates were from Macherey-Nagel (Düren, Germany). The cPLA 2α inhibitor pyrrophenone [ 29 ] was synthesized and provided by Dr. Alfonso P é rez (Department of Organic Chemistry, University of Biomolecules 2020,10, 542 3 of 16 Valladolid, Valladolid, Spain). The iPLA 2 -VIA inhibitors FKGK18 and GK436 were synthesized in the Kokotos laboratory [30,31]. All other reagents were from Sigma-Aldrich (Madrid, Spain). 2.2. Cell Culture Mouse peritoneal macrophages from Swiss mice (University of Valladolid Animal House, 10–12 weeks old) were obtained by peritoneal lavage using 5 mL cold phosphate-buffered saline and cultured in RPMI 1640 medium with 10% (v/v) fetal bovine serum, 100 U/mL penicillin, and 100 µ g/mL streptomycin, as described elsewhere [ 32 , 33 ]. All procedures involving animals were undertaken under the supervision of the Institutional Committee of Animal Care and Usage of the University of Valladolid (No. 7406000), in accordance with the guidelines established by the Spanish Ministry of Agriculture, Food, and Environment and the European Union. Cells were placed in serum-free medium for 1 h before addition of stimuli or inhibitors. Afterward, they were challenged by the stimuli for the time indicated. Zymosan was prepared exactly as described [ 32 , 33 ]. Only zymosan batches that demonstrated no measurable endogenous PLA 2 activity, as measured by in vitro assay under different conditions [ 34 – 37 ], were used in this study. Cell protein content was quantified according to Bradford [ 38 ] using a commercial kit (BioRad Protein Assay, Bio-Rad, Hercules, CA, USA). 2.3. Gas Chromatography/Mass Spectrometry (GC/MS) Analyses Total lipids from approximately 10 7 cells were extracted according to Bligh and Dyer [ 39 ], and the following internal standards were added: 10 nmol of 1,2-diheptadecanoyl-sn-glycero-3-phosphocholine, 10 nmol of 1,2,3-triheptadecanoylglycerol, and 30 nmol of cholesteryl tridecanoate. Phospholipids were separated from neutral lipids by thin-layer chromatography, using n-hexane/diethyl ether/acetic acid (70:30:1, v/v/v) as the mobile phase [ 40 ]. Phospholipid classes were separated twice with chloroform/methanol/28% (w/w) ammonium hydroxide (60:37.5:4, v/v/v) as the mobile phase, using plates impregnated with boric acid [ 41 ]. The bands corresponding to the different lipid classes were scraped offfrom the plate, and fatty acid methyl esters were obtained from the various lipid fractions by transmethylation with 0.5 M KOH in methanol for 60 min at 37 ◦ C [ 42 – 45 ]. Analyses were carried out using an Agilent 7890A gas chromatograph coupled to an Agilent 5975C mass-selective detector operated in an electron impact mode (EI, 70 eV). The apparatus was also equipped with an Agilent 7693 autosampler and an Agilent DB23 column (60 m length × 0.25 mm internal diameter × 0.15 µ M film thickness) (Agilent Technologies, Santa Clara, CA, USA). Data analysis was carried out with the Agilent G1701EA MSD Productivity Chemstation software, revision E.02.00 [42–45]. 2.4. Liquid Chromatography/Mass Spectrometry (LC/MS) Analyses of Phospholipids Lipids were extracted according to Bligh and Dyer [ 39 ], and the following internal standards were added: 20 pmol each of 1,2-dimyristoyl-sn-glycero-3-phosphoglycerol, 1,2-dilauroyl-sn-glycero-3-phosphoethanolamine, 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine, 1,2-di- heptadecanoyl-sn-glycero-3-phosphoethanolamine, 1,2-dimyristoyl-sn-glycero-3-phosphoserine, 1,2-dimyristoyl-sn-glycero-3-phosphate, 1,2-dipentadecanoyl-sn-glycero-3-phosphocholine, 1,2-di- heptadecanoyl-sn-glycero-3-phosphocholine, and 1,2-dinonadecanoyl-sn-glycero-phosphocholine. The samples were re-dissolved in hexanes/2-propanol/water (42:56:2, v/v/v) and injected into a Thermo Scientific Dionex Ultimate 3000 high-performance liquid chromatograph equipped with an Ultimate HPG-3400SD standard binary pump and an Ultimate ACC-3000 autosampler column compartment (Waltham, MA, USA). Separation was carried using a FORTIS HILIC column (150 × 3 mm, 3 µ m particle size) (Fortis Technologies, Neston, UK). The mobile phase consisted of a gradient of solvent A (hexanes/isopropanol 30:40, v/v) and solvent B (hexanes/isopropanol/20 mM ammonium acetate in water, 30:40:7, v/v/v). The gradient started at 75% A from 0 to 5 min, then decreased from 75% A to 40% A at 15 min, from 40% A to 5% A at 20 min, was held at 5% until 40 min, and then increased to 75% at 41 min. Then, the column was re-equilibrated holding 75% A for an additional 14-min period before Biomolecules 2020,10, 542 4 of 16 the next sample injection [ 46 ]. The flow rate through the column was fixed at 0.4 mL/min. The liquid chromatography system was coupled online to an Sciex QTRAP 4500 Mass Spectrometer equipped with a Turbo V ion source and a TurbolonSpray probe for electrospray ionization (AB Sciex, Framingham, MA, USA). Source parameters were set as follows: ion spray voltage, − 4500 V; curtain gas, 30 psi; nebulizer gas, 50 psi; desolvation gas, 60 psi; temperature, 425 ◦ C. Phospholipid species were analyzed in scheduled multiple reaction monitoring mode with negative ionization, detecting in Q3 the m/zof either 303.2 or 331.2, corresponding to AA and AdA, respectively, as [M-H] − . Compound parameters were fixed as follows: declustering potential; − 45 V (choline glycerophospholipids), − 60 V (ethanolamine glycerophospholipids) − 30 V (phosphatidylinositol), − 50 V (phosphatidylserine), − 60 V (phosphatidic acid), − 50 V (phosphatidylglycerol); collision energy: − 50 V (choline glycerophospholipids), − 40 V (ethanolamine glycerophospholipids), − 60 V (phosphatidylinositol), − 50 V (phosphatidylserine), − 45 V (phosphatidic acid), − 45 V (phosphatidylglycerol); entrance potential, − 10 V; and collision cell exit potential, − 8 V. All glycerophospholipids were detected as [M-H] − , ions except choline glycerophospholipids, which were detected as [M +CH 3 COO] − ions. Quantification was carried out by integrating the chromatographic peaks of each species and comparing these with the peak area of the internal standard that corresponded to each class. 3. Results 3.1. Adrenic Acid and Arachidonic Acid Contents of Murine Peritoneal Macrophages Lipid extracts from mouse peritoneal macrophages were analyzed for fatty acid content by GC/MS. Total AA content was 69.9 ± 4.2 nmol/mg cell protein (mean values ± standard error of the mean, n=5), while AdA content was 15.1 ± 1.2 nmol/mg cell protein (mean values ± standard error of the mean, n=5). Both AA and AdA were found almost exclusively in phospholipids. The distribution of AA and AdA between phospholipid classes is shown in Figure 1. Despite the difference in mass between AA and AdA, their distribution between phospholipid classes was remarkably similar, with the majority of both fatty acids being found in ethanolamine glycerophospholipids (PE), followed by choline glycerophospholipids (PC). Minor amounts of both fatty acids were found in phosphatidylinositol (PI) and phosphatidylserine (PS). Biomolecules 2020, 10, 4 of 16 acetate in water, 30:40:7, v/v/v). The gradient started at 75% A from 0 to 5 min, then decreased from 75% A to 40% A at 15 min, from 40% A to 5% A at 20 min, was held at 5% until 40 min, and then increased to 75% at 41 min. Then, the column was re-equilibrated holding 75% A for an additional 14-min period before the next sample injection [46]. The flow rate through the column was fixed at 0.4 mL/min. The liquid chromatography system was coupled online to an Sciex QTRAP 4500 Mass Spectrometer equipped with a Turbo V ion source and a TurbolonSpray probe for electrospray ionization (AB Sciex, Framingham, MA, USA). Source parameters were set as follows: ion spray voltage, -4500 V; curtain gas, 30 psi; nebulizer gas, 50 psi; desolvation gas, 60 psi; temperature, 425 °C. Phospholipid species were analyzed in scheduled multiple reaction monitoring mode with negative ionization, detecting in Q3 the m/z of either 303.2 or 331.2, corresponding to AA and AdA, respectively, as [M-H]−. Compound parameters were fixed as follows: declustering potential; −45 V (choline glycerophospholipids), −60 V (ethanolamine glycerophospholipids) −30 V (phosphatidylinositol), −50 V (phosphatidylserine), −60 V (phosphatidic acid), −50 V (phosphatidylglycerol); collision energy: −50 V (choline glycerophospholipids), −40 V (ethanolamine glycerophospholipids), −60 V (phosphatidylinositol), −50 V (phosphatidylserine), −45 V (phosphatidic acid), −45 V (phosphatidylglycerol); entrance potential, −10 V; and collision cell exit potential, −8 V. All glycerophospholipids were detected as [M-H]−, ions except choline glycerophospholipids, which were detected as [M + CH3COO]− ions. Quantification was carried out by integrating the chromatographic peaks of each species and comparing these with the peak area of the internal standard that corresponded to each class. 3. Results 3.1. Adrenic Acid and Arachidonic Acid Contents of Murine Peritoneal Macrophages Lipid extracts from mouse peritoneal macrophages were analyzed for fatty acid content by GC/MS. Total AA content was 69.9 ± 4.2 nmol/mg cell protein (mean values ± standard error of the mean, n = 5), while AdA content was 15.1 ± 1.2 nmol/mg cell protein (mean values ± standard error of the mean, n = 5). Both AA and AdA were found almost exclusively in phospholipids. The distribution of AA and AdA between phospholipid classes is shown in Figure 1. Despite the difference in mass between AA and AdA, their distribution between phospholipid classes was remarkably similar, with the majority of both fatty acids being found in ethanolamine glycerophospholipids (PE), followed by choline glycerophospholipids (PC). Minor amounts of both fatty acids were found in phosphatidylinositol (PI) and phosphatidylserine (PS). PC PE PI PS AdA Content (nmol/mg protein) 0 2 4 6 8 10 PC PE PI PS AA Content (nmol/mg protein) 0 8 16 24 32 40 BA Figure 1. Distribution of AA and AdA between phospholipid classes. The various phospholipid classes were separated by thin-layer chromatography. The distribution of AA (A) and AdA (B) between choline glycerophospholipids (PC), ethanolamine glycerophospholipids (PE), phosphatidylinositol (PI), and phosphatidylserine (PS) was determined by gas chromatography/mass spectrometry (GC/MS) after converting the phospholipid-bound fatty acids into methyl esters. Results are shown as means ± standard error of the mean (n = 3). Figure 1. Distribution of AA and AdA between phospholipid classes. The various phospholipid classes were separated by thin-layer chromatography. The distribution of AA ( A ) and AdA ( B ) between choline glycerophospholipids (PC), ethanolamine glycerophospholipids (PE), phosphatidylinositol (PI), and phosphatidylserine (PS) was determined by gas chromatography/mass spectrometry (GC/MS) after converting the phospholipid-bound fatty acids into methyl esters. Results are shown as means ±standard error of the mean (n=3). Figure 2shows the distribution of AA- and AdA-containing phospholipid molecular species, as analyzed by liquid chromatography coupled to tandem mass spectrometry (LC/MS). In agreement Biomolecules 2020,10, 542 5 of 16 with previous estimates [ 21 , 22 ], multiple AA-containing species were detected, with the alkenylacyl and diacyl ethanolamine phospholipid species PE(P-16:0/20:4), PE(P-18:0/20:4), and PE(18:0/20:4) predominating, followed by the diacyl choline phospholipid species PC(16:0/20:4) and PC(18:0/20:4), and the unique inositol phospholipid species PI(18:0/20:4) Biomolecules 2020, 10, 5 of 16 Figure 2 shows the distribution of AA- and AdA-containing phospholipid molecular species, as analyzed by liquid chromatography coupled to tandem mass spectrometry (LC/MS). In agreement with previous estimates [21,22], multiple AA-containing species were detected, with the alkenylacyl and diacyl ethanolamine phospholipid species PE(P-16:0/20:4), PE(P-18:0/20:4), and PE(18:0/20:4) predominating, followed by the diacyl choline phospholipid species PC(16:0/20:4) and PC(18:0/20:4), and the unique inositol phospholipid species PI(18:0/20:4) Figure 2. AA and AdA-containing phospholipid molecular species in peritoneal macrophages The profiles of AA- (A) or AdA- (B) containing PC (red), PE (green), PI (yellow), and PS (pink) species in peritoneal macrophages were determined by liquid chromatography/mass spectrometry (LC/MS). Fatty chains within the different phospholipid species are designated by their numbers of carbons and double bonds. A designation of O- before the first fatty chain indicates that the sn-1 position is ether linked, whereas a P- designation indicates a plasmalogen form (sn-1 vinyl ether linkage) [47]. Phospholipids containing two ester bonds have no designation. Results are shown as mean values ± standard error of the mean (n = 3). Figure 2. AA and AdA-containing phospholipid molecular species in peritoneal macrophages The profiles of AA- ( A ) or AdA- ( B ) containing PC (red), PE (green), PI (yellow), and PS (pink) species in peritoneal macrophages were determined by liquid chromatography/mass spectrometry (LC/MS). Fatty chains within the different phospholipid species are designated by their numbers of carbons and double bonds. A designation of O- before the first fatty chain indicates that the sn-1 position is ether linked, whereas a P- designation indicates a plasmalogen form (sn-1 vinyl ether linkage) [ 47 ]. Phospholipids containing two ester bonds have no designation. Results are shown as mean values ± standard error of the mean (n=3). Biomolecules 2020,10, 542 6 of 16 Regarding AdA-containing species, the alkenyl acyl and diacyl ethanolamine phospholipid species PE(P-16:0/22:4), PE(P-18:0/22:4), and PE(18:0/22:4)], and the diacyl choline phospholipid species PC(16:0/22:4) and PC(18:0/22:4) also constituted the major cellular AdA reservoirs. Strikingly, the inositol phospholipid species PI(18:0/22:4) was not as prevalent as its AA equivalent, PI(18:0/20:4), was among AA-containing phospholipids. This may suggest that the acyl-CoA acyltransferase using lysoPI as the acceptor [48] shows selectivity for AA over AdA as a substrate. Macrophage stimulation with yeast-derived zymosan markedly decreased the cellular AA content in PC and PI. Despite PE being the major AA-containing class, AA losses from PE did not reach statistical significance (Figure 3A). It should be noted in this regard that during receptor stimulation, AA is known to be transferred from AA-containing PC (1-acyl species) to PE (plasmalogen species) by CoA-independent transacylase; hence, the decline in the amount of AA-containing PE during cellular stimulation may be greatly reduced [22,24,49]. Regarding AdA, decreases in its cellular content were also observed after zymosan stimulation. However the pattern clearly differed in that PC was the only phospholipid class that contributed to AdA mobilization; AdA reductions from PE and PI did not reach statistical significance (Figure 3B). Biomolecules 2020, 10, 6 of 16 Regarding AdA-containing species, the alkenyl acyl and diacyl ethanolamine phospholipid species PE(P-16:0/22:4), PE(P-18:0/22:4), and PE(18:0/22:4)], and the diacyl choline phospholipid species PC(16:0/22:4) and PC(18:0/22:4) also constituted the major cellular AdA reservoirs. Strikingly, the inositol phospholipid species PI(18:0/22:4) was not as prevalent as its AA equivalent, PI(18:0/20:4), was among AA-containing phospholipids. This may suggest that the acyl-CoA acyltransferase using lysoPI as the acceptor [48] shows selectivity for AA over AdA as a substrate. Macrophage stimulation with yeast-derived zymosan markedly decreased the cellular AA content in PC and PI. Despite PE being the major AA-containing class, AA losses from PE did not reach statistical significance (Figure 3A). It should be noted in this regard that during receptor stimulation, AA is known to be transferred from AA-containing PC (1-acyl species) to PE (plasmalogen species) by CoA-independent transacylase; hence, the decline in the amount of AA- containing PE during cellular stimulation may be greatly reduced [22,24,49]. Regarding AdA, decreases in its cellular content were also observed after zymosan stimulation. However the pattern clearly differed in that PC was the only phospholipid class that contributed to AdA mobilization; AdA reductions from PE and PI did not reach statistical significance (Figure 3B). Figure 3. AA and AdA mobilization in zymosan-stimulated macrophages. The cells were unstimulated (colored bars) or stimulated (open bars) with 1 mg/mL zymosan for 1 h. Afterward, total content of AA (A) or AdA (B) in various phospholipid classes (see x-axis) was measured by GC/MS. To allow for direct comparison between AA and AdA, the fatty acid contents in phospholipid classes in unstimulated cells are given as 100%. Actual mass values can be obtained from Figure 1. Results are shown as mean values ± standard error of the mean. (n = 3). Note: * p < 0.05, significantly different from the corresponding phospholipid class. 3.2. Defining the Role of Various PLA2 Forms in Stimulus-Induced AA and AdA Mobilization The differential involvement of phospholipid classes in AA and AdA mobilization suggests the existence of separate mechanisms for regulating the availability of each fatty acid. To characterize such mechanisms, we took advantage of the use of selective inhibitors of the two major intracellular PLA2 enzymes potentially effecting the fatty acid release in activated peritoneal macrophages, namely group IVA Ca2+-dependent cytosolic PLA2 (cPLA2α) and group VIA Ca2+-independent PLA2 (iPLA2-VIA) [22,50]. The use of selective chemical inhibitors to address the role of intracellular PLA2s during cell activation has some advantages over other widely used methods, such as small interfering RNA or cells from knockout mice. With chemical inhibitors, inhibition develops rapidly, which reduces the impact of unspecific effects that could occur over time. Also, no compensatory mechanisms take place that might obscure the interpretation of results [22]. In addition, chemical inhibitors directly target PLA2 effects that depend on enzymatic activity, without affecting noncatalytic functions of the enzyme [51]. The inhibitors used in this work are the most potent and selective inhibitors currently available to block cPLA2α and iPLA2-VIA in cells. Pyrrophenone potently and selectively inhibits cPLA2α activity using a number of in vitro assays without detectable effects on other PLA2 activities, blocks AA release in mammalian cells in the 0.01–1 µM range [29,52,53], and has been shown to be effective in experimental models of disease involving cPLA2α Figure 3. AA and AdA mobilization in zymosan-stimulated macrophages. The cells were unstimulated (colored bars) or stimulated (open bars) with 1 mg/mL zymosan for 1 h. Afterward, total content of AA ( A ) or AdA ( B ) in various phospholipid classes (see x-axis) was measured by GC/MS. To allow for direct comparison between AA and AdA, the fatty acid contents in phospholipid classes in unstimulated cells are given as 100%. Actual mass values can be obtained from Figure 1. Results are shown as mean values ± standard error of the mean. (n=3). Note: * p<0.05, significantly different from the corresponding phospholipid class. 3.2. Defining the Role of Various PLA2Forms in Stimulus-Induced AA and AdA Mobilization The differential involvement of phospholipid classes in AA and AdA mobilization suggests the existence of separate mechanisms for regulating the availability of each fatty acid. To characterize such mechanisms, we took advantage of the use of selective inhibitors of the two major intracellular PLA 2 enzymes potentially effecting the fatty acid release in activated peritoneal macrophages, namely group IVA Ca 2+ -dependent cytosolic PLA 2 (cPLA 2α ) and group VIA Ca 2+ -independent PLA 2 (iPLA 2 -VIA) [ 22 , 50 ]. The use of selective chemical inhibitors to address the role of intracellular PLA 2 s during cell activation has some advantages over other widely used methods, such as small interfering RNA or cells from knockout mice. With chemical inhibitors, inhibition develops rapidly, which reduces the impact of unspecific effects that could occur over time. Also, no compensatory mechanisms take place that might obscure the interpretation of results [ 22 ]. In addition, chemical inhibitors directly target PLA 2 effects that depend on enzymatic activity, without affecting noncatalytic functions of the enzyme [ 51 ]. The inhibitors used in this work are the most potent and selective inhibitors currently available to block cPLA 2α and iPLA 2 -VIA in cells. Pyrrophenone potently and selectively inhibits cPLA 2α activity using a number of in vitro assays without detectable effects on other PLA 2 activities, blocks AA release in mammalian cells in the 0.01–1 µ M range [ 29 , 52 , 53 ], and has been shown to be effective in experimental Biomolecules 2020,10, 542 7 of 16 models of disease involving cPLA 2α [ 54 , 55 ]. Fluoroketone FKGK18, a selective iPLA 2 -VIA inhibitor, is at least 200-fold more potent for inhibiting iPLA 2 -VIA than cPLA 2α [ 30 ], and is useful for characterizing iPLA 2 -VIA-mediated functions in vivo [ 31 , 56 , 57 ]. β -Lactone GK436, another iPLA 2 -VIA inhibitor, has been found to be at least 1000-fold more potent for iPLA 2 -VIA than for cPLA 2α [ 31 ]. For comparative purposes with data from the bibliography [ 22 , 58 – 64 ], we also used bromoenol lactone (BEL) to inhibit iPLA 2 -VIA. BEL inhibits calcium-independent PLA 2 s and exerts little or no effect on Ca 2+ -dependent enzymes [65,66], albeit it may exhibit off-target effects depending on cell type [67–69]. In agreement with previous observations [ 22 , 24 ], pyrrophenone was found to almost completely inhibit zymosan-stimulated AA mobilization from phospholipids (Figure 4A). Importantly, pyrrophenone also partly inhibited AdA mobilization (Figure 4B). Conversely, none of the iPLA 2 -VIA inhibitors tested exerted appreciable effects on AA mobilization while significantly affecting AdA mobilization (Figure 4). The simultaneous addition of inhibitors of both cPLA 2α and iPLA 2 -VIA resulted in complete inhibition of AdA mobilization (Figure 4B). These results suggest that unlike AA, AdA mobilization involves the action of both cPLA2αand iPLA2-VIA. Biomolecules 2020, 10, 7 of 16 [54,55]. Fluoroketone FKGK18, a selective iPLA2-VIA inhibitor, is at least 200-fold more potent for inhibiting iPLA2-VIA than cPLA2α [30], and is useful for characterizing iPLA2-VIA-mediated functions in vivo [31,56,57]. β-Lactone GK436, another iPLA2-VIA inhibitor, has been found to be at least 1000-fold more potent for iPLA2-VIA than for cPLA2α [31]. For comparative purposes with data from the bibliography [22,58–64], we also used bromoenol lactone (BEL) to inhibit iPLA2-VIA. BEL inhibits calcium-independent PLA2s and exerts little or no effect on Ca2+-dependent enzymes [65,66], albeit it may exhibit off-target effects depending on cell type [67–69]. In agreement with previous observations [22,24], pyrrophenone was found to almost completely inhibit zymosan-stimulated AA mobilization from phospholipids (Figure 4A). Importantly, pyrrophenone also partly inhibited AdA mobilization (Figure 4B). Conversely, none of the iPLA2- VIA inhibitors tested exerted appreciable effects on AA mobilization while significantly affecting AdA mobilization (Figure 4). The simultaneous addition of inhibitors of both cPLA2α and iPLA2-VIA resulted in complete inhibition of AdA mobilization (Figure 4B). These results suggest that unlike AA, AdA mobilization involves the action of both cPLA2α and iPLA2-VIA. Unstim No Inhib. Pyrr FKGK18 GK436 BEL AA Release (nmol/mg protein) 0 5 10 15 20 Unstim No inhib. Pyrr FKGK18 GK436 BEL Pyrr + FKGK18 AdA Release (nmol/mg protein) 0 1 2 3 4 5 * * A B ** * * Figure 4. Release of AA and AdA by stimulated macrophages. The cells were either unstimulated (open bars) or stimulated by 1 mg/mL zymosan for 1 h (colored bars) in the absence (no inhibition) or presence of the following inhibitors: 2 µM pyrrophenone (Pyrr), 10 µM FKGK18, 5 µM GK436, 10 µM BEL, or 2 µM pyrrophenone plus 10 µM FKGK18 (see x-axis). Afterward, total content of AA (A) or AdA (B) was measured by GC/MS. The fatty acid release was calculated by subtracting the amount of phospholipid-bound AA or AdA in stimulated cells from that in unstimulated cells. Results are shown as mean values ± standard error of the mean. (n = 3). Note: * p < 0.05, significantly different from zymosan-stimulated cells in the absence of inhibitors. 3.3. Studies with AdA-Enriched Cells Given the structural similarities between AA and AdA, it could be envisioned that in some instances, AdA competes with AA for incorporation into phospholipids, which might result in reduced amounts of “mobilizable” AA within phospholipids, and hence reduced formation of proinflammatory eicosanoids [16,70,71]. To address this possibility, we incubated the macrophages with exogenous AdA (10 µM, 20 h), which resulted in the cells avidly incorporating the fatty acid into cellular phospholipids. These conditions led to a 2–3-fold increase in the amount of AdA esterified into phospholipids compared with untreated cells (33.2 ± 4.6 nmol per mg protein; mean ± standard error of the mean, n = 4). Analysis by GC/MS of the fatty acid content of the AdA-enriched cells revealed that the fatty acids typically displaced within phospholipids by AdA upon AdA supplementation were oleic acid (18:1n-9) and the n-6 series members linoleic (18:2n-6) and dihomoγ-linolenic (20:3n-6) acids (Figure 5). Remarkably, little AA displacement occurred after AdA Figure 4. Release of AA and AdA by stimulated macrophages. The cells were either unstimulated (open bars) or stimulated by 1 mg/mL zymosan for 1 h (colored bars) in the absence (no inhibition) or presence of the following inhibitors: 2 µ M pyrrophenone (Pyrr), 10 µ M FKGK18, 5 µ M GK436, 10 µ M BEL, or 2 µ M pyrrophenone plus 10 µ M FKGK18 (see x-axis). Afterward, total content of AA ( A ) or AdA ( B ) was measured by GC/MS. The fatty acid release was calculated by subtracting the amount of phospholipid-bound AA or AdA in stimulated cells from that in unstimulated cells. Results are shown as mean values ± standard error of the mean. (n =3). Note: * p<0.05, significantly different from zymosan-stimulated cells in the absence of inhibitors. 3.3. Studies with AdA-Enriched Cells Given the structural similarities between AA and AdA, it could be envisioned that in some instances, AdA competes with AA for incorporation into phospholipids, which might result in reduced amounts of “mobilizable” AA within phospholipids, and hence reduced formation of pro-inflammatory eicosanoids [ 16 , 70 , 71 ]. To address this possibility, we incubated the macrophages with exogenous AdA (10 µ M, 20 h), which resulted in the cells avidly incorporating the fatty acid into cellular phospholipids. These conditions led to a 2–3-fold increase in the amount of AdA esterified into phospholipids compared with untreated cells (33.2 ± 4.6 nmol per mg protein; mean ± standard error of the mean, n=4). Analysis by GC/MS of the fatty acid content of the AdA-enriched cells revealed that the fatty acids typically displaced within phospholipids by AdA upon AdA supplementation were oleic acid (18:1n-9) and the n-6 series members linoleic (18:2n-6) and dihomoγ -linolenic (20:3n-6) acids Biomolecules 2020,10, 542 8 of 16 (Figure 5). Remarkably, little AA displacement occurred after AdA supplementation. Analysis of AA and AdA mobilization in zymosan-stimulated cells indicated that the extent of the AA response did not substantially differ from that observed from cells not treated with AdA, while AdA mobilization expectedly increased (Figure 6). Biomolecules 2020, 10, 8 of 16 supplementation. Analysis of AA and AdA mobilization in zymosan-stimulated cells indicated that the extent of the AA response did not substantially differ from that observed from cells not treated with AdA, while AdA mobilization expectedly increased (Figure 6). Figure 5. Fatty acid composition of murine peritoneal macrophages. The cells were either untreated (orange bars) or treated with exogenous AdA for 20 h (gray bars). Afterward, the total fatty acid profiles in cells were determined by GC/MS after converting the fatty acid glyceryl esters into fatty acid methyl esters. The fatty acids are designated by their number of carbon atoms, and after a colon, their number of double bonds. To differentiate isomers, the n − x (n minus x) nomenclature is used, where n is the number of carbons of a given fatty acid and x is an integer that gives the position of the last double bond of the molecule when subtracted from n. The data are expressed as mean values ± standard error of the mean of three individual replicates. Note: * p < 0.05, significantly different from incubations in the absence of AdA. Untreated AdA-treated AA Content (nmol/mg protein) 0 20 40 60 80 * * A Untreated AdA-treated AdA Content (nmol/mg protein) 0 10 20 30 40 B * * Figure 6. AA and AdA mobilization in zymosan-stimulated macrophages. The cells were either untreated or treated with exogenous AdA for 20 h (see x-axis). Afterward, the cells were unstimulated (colored bars) or stimulated (open bars) with 1 mg/mL zymosan for 1 h, and cellular AA content (A) or AdA content (B) was determined by GC/MS. Results are shown as mean values ± standard error of the mean (n = 3). Note: * p < 0.05, significantly different from unstimulated cells. Analysis of AA-derived metabolites produced by zymosan-stimulated cells did not appreciably change whether the cells had previously been treated with AdA or not, while the levels of the only AdA metabolite detected at significant levels in the stimulated macrophages, dihomoprostaglandin Figure 5. Fatty acid composition of murine peritoneal macrophages. The cells were either untreated (orange bars) or treated with exogenous AdA for 20 h (gray bars). Afterward, the total fatty acid profiles in cells were determined by GC/MS after converting the fatty acid glyceryl esters into fatty acid methyl esters. The fatty acids are designated by their number of carbon atoms, and after a colon, their number of double bonds. To differentiate isomers, the n − x (n minus x) nomenclature is used, where n is the number of carbons of a given fatty acid and x is an integer that gives the position of the last double bond of the molecule when subtracted from n. The data are expressed as mean values ± standard error of the mean of three individual replicates. Note: * p<0.05, significantly different from incubations in the absence of AdA. Biomolecules 2020, 10, 8 of 16 supplementation. Analysis of AA and AdA mobilization in zymosan-stimulated cells indicated that the extent of the AA response did not substantially differ from that observed from cells not treated with AdA, while AdA mobilization expectedly increased (Figure 6). Figure 5. Fatty acid composition of murine peritoneal macrophages. The cells were either untreated (orange bars) or treated with exogenous AdA for 20 h (gray bars). Afterward, the total fatty acid profiles in cells were determined by GC/MS after converting the fatty acid glyceryl esters into fatty acid methyl esters. The fatty acids are designated by their number of carbon atoms, and after a colon, their number of double bonds. To differentiate isomers, the n − x (n minus x) nomenclature is used, where n is the number of carbons of a given fatty acid and x is an integer that gives the position of the last double bond of the molecule when subtracted from n. The data are expressed as mean values ± standard error of the mean of three individual replicates. Note: * p < 0.05, significantly different from incubations in the absence of AdA. Untreated AdA-treated AA Content (nmol/mg protein) 0 20 40 60 80 * * A Untreated AdA-treated AdA Content (nmol/mg protein) 0 10 20 30 40 B * * Figure 6. AA and AdA mobilization in zymosan-stimulated macrophages. The cells were either untreated or treated with exogenous AdA for 20 h (see x-axis). Afterward, the cells were unstimulated (colored bars) or stimulated (open bars) with 1 mg/mL zymosan for 1 h, and cellular AA content (A) or AdA content (B) was determined by GC/MS. Results are shown as mean values ± standard error of the mean (n = 3). Note: * p < 0.05, significantly different from unstimulated cells. Analysis of AA-derived metabolites produced by zymosan-stimulated cells did not appreciably change whether the cells had previously been treated with AdA or not, while the levels of the only AdA metabolite detected at significant levels in the stimulated macrophages, dihomoprostaglandin Figure 6. AA and AdA mobilization in zymosan-stimulated macrophages. The cells were either untreated or treated with exogenous AdA for 20 h (see x-axis). Afterward, the cells were unstimulated (colored bars) or stimulated (open bars) with 1 mg/mL zymosan for 1 h, and cellular AA content ( A ) or AdA content ( B ) was determined by GC/MS. Results are shown as mean values ± standard error of the mean (n=3). Note: * p<0.05, significantly different from unstimulated cells. Biomolecules 2020,10, 542 9 of 16 Analysis of AA-derived metabolites produced by zymosan-stimulated cells did not appreciably change whether the cells had previously been treated with AdA or not, while the levels of the only AdA metabolite detected at significant levels in the stimulated macrophages, dihomoprostaglandin E 2 , increased in the AdA-treated cells (Figure 7). Collectively, these results suggest that in zymosan-activated macrophages, AdA does not influence AA metabolism leading to eicosanoid production. Biomolecules 2020, 10, 9 of 16 E2, increased in the AdA-treated cells (Figure 7). Collectively, these results suggest that in zymosanactivated macrophages, AdA does not influence AA metabolism leading to eicosanoid production. Figure 7. Eicosanoid production by stimulated macrophages. The cells were either untreated (orange bars) or treated with exogenous AdA for 20 h (gray bars). Afterward, the cells were stimulated by 1 mg/mL zymosan for 1 h, and eicosanoid content in the supernatants was analyzed by LC/MS. The data are expressed as mean values ± standard error of the mean of three individual replicates. Note: * p < 0.05, significantly different from incubations in the absence of AdA treatment. PGE2, prostaglandin E2; TXB2, thromboxane B2; 6k-PGF1a, 6-keto prostaglandin F1α (the stable product of prostaglandin I2); 11-HETE, 11-hydroxyeicosatetraenoic acid; 5-HETE, 5-hydroxyeicosatetraenoic acid; 15-HETE, 15-hydroxyeicosatetraenoic acid; 15k-PGF2a, 15-ketoprostaglandin F2α; 12-HHT, 12- hydroxyheptadecatrienoic acid; 5,15-diHETE, 5,15-dihydroxyeicosatetraenoic acid; 11,12-DHET, 11,12-dihydroxyeicosatrienoic acid; dh-PGE2, dihomoprostaglandin E2. 4. Discussion AdA is the 2-carbon elongation product of AA. Similar to AA, AdA can be mobilized from membrane phospholipids to serve as a substrate for the production of oxidized metabolites with a diverse array of biological functions [11–14,72]. We show in this work that AdA is present at significant quantities in the membrane phospholipids of murine peritoneal macrophages. The AdA- to-AA ratio in macrophages is 20–25%, which is similar to that found in other cells [13,14]. Stimulation of the macrophages with yeast-derived zymosan results in the cells releasing significant amounts of AdA, part of which is metabolized to dihomoprostaglandin E2. While macrophages appear to utilize AA for lipid mediator synthesis more efficiently than AdA, both qualitatively and quantitatively, our results support the concept that neither fatty acid competes with the other. Further, the data suggest that the processes leading to the mobilization and metabolism of both fatty acids are independently regulated. In support of these observations, enriching the cells with exogenous AdA has little effect on endogenous AA levels. AA release and metabolism in these AdA-enriched cells proceed in essentially the same way as in cells displaying normal levels of AdA. Consistent with these observations, studies by others have highlighted that under certain settings, some AdA-derived metabolites may display higher potency than their AA-derived counterparts, suggesting that they may serve non-redundant functions on their own [13,14,72–75]. Collectively, the results suggest that separate and selective mechanisms may exist for regulating AA and AdA utilization in macrophages. Figure 7. Eicosanoid production by stimulated macrophages. The cells were either untreated (orange bars) or treated with exogenous AdA for 20 h (gray bars). Afterward, the cells were stimulated by 1 mg/mL zymosan for 1 h, and eicosanoid content in the supernatants was analyzed by LC/MS. The data are expressed as mean values ± standard error of the mean of three individual replicates. Note: * p<0.05, significantly different from incubations in the absence of AdA treatment. PGE2, prostaglandin E 2 ; TXB2, thromboxane B 2 ; 6k-PGF1a, 6-keto prostaglandin F 1α (the stable product of prostaglandin I 2 ); 11-HETE, 11-hydroxyeicosatetraenoic acid; 5-HETE, 5-hydroxyeicosatetraenoic acid; 15-HETE, 15-hydroxyeicosatetraenoic acid; 15k-PGF2a, 15-ketoprostaglandin F 2α ; 12-HHT, 12-hydroxyheptadecatrienoic acid; 5,15-diHETE, 5,15-dihydroxyeicosatetraenoic acid; 11,12-DHET, 11,12-dihydroxyeicosatrienoic acid; dh-PGE2, dihomoprostaglandin E2. 4. Discussion AdA is the 2-carbon elongation product of AA. Similar to AA, AdA can be mobilized from membrane phospholipids to serve as a substrate for the production of oxidized metabolites with a diverse array of biological functions [ 11 – 14 , 72 ]. We show in this work that AdA is present at significant quantities in the membrane phospholipids of murine peritoneal macrophages. The AdA-to-AA ratio in macrophages is 20–25%, which is similar to that found in other cells [ 13 , 14 ]. Stimulation of the macrophages with yeast-derived zymosan results in the cells releasing significant amounts of AdA, part of which is metabolized to dihomoprostaglandin E 2 . While macrophages appear to utilize AA for lipid mediator synthesis more efficiently than AdA, both qualitatively and quantitatively, our results support the concept that neither fatty acid competes with the other. Further, the data suggest that the processes leading to the mobilization and metabolism of both fatty acids are independently regulated. In support of these observations, enriching the cells with exogenous AdA has little effect on endogenous AA levels. AA release and metabolism in these AdA-enriched cells proceed in essentially Biomolecules 2020,10, 542 16 of 16 77. 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