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Bermudez, M. Lipid Droplets, Phospholipase A2, Arachidonic Acid, and Atherosclerosis. Encyclopedia. Available online: https://encyclopedia.pub/entry/18513 (accessed on 18 December 2025).
Bermudez M. Lipid Droplets, Phospholipase A2, Arachidonic Acid, and Atherosclerosis. Encyclopedia. Available at: https://encyclopedia.pub/entry/18513. Accessed December 18, 2025.
Bermudez, Miguel. "Lipid Droplets, Phospholipase A2, Arachidonic Acid, and Atherosclerosis" Encyclopedia, https://encyclopedia.pub/entry/18513 (accessed December 18, 2025).
Bermudez, M. (2022, January 19). Lipid Droplets, Phospholipase A2, Arachidonic Acid, and Atherosclerosis. In Encyclopedia. https://encyclopedia.pub/entry/18513
Bermudez, Miguel. "Lipid Droplets, Phospholipase A2, Arachidonic Acid, and Atherosclerosis." Encyclopedia. Web. 19 January, 2022.
Lipid Droplets, Phospholipase A2, Arachidonic Acid, and Atherosclerosis
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This entry is adapted from the peer-reviewed paper 10.3390/biomedicines9121891

Lipid droplets, classically regarded as static storage organelles, are currently considered as dynamic structures involved in key processes of lipid metabolism, cellular homeostasis and signaling. Studies on the inflammatory state of atherosclerotic plaques suggest that circulating monocytes interact with products released by endothelial cells and may acquire a foamy phenotype before crossing the endothelial barrier and differentiating into macrophages. One such compound released in significant amounts into the bloodstream is arachidonic acid, the common precursor of eicosanoids, and a potent inducer of neutral lipid synthesis and lipid droplet formation in circulating monocytes. Members of the family of phospholipase A2, which hydrolyze the fatty acid present at the sn-2 position of phospholipids, have recently emerged as key controllers of lipid droplet homeostasis, regulating their formation and the availability of fatty acids for lipid mediator production.

lipid droplet phospholipase A2 arachidonic acid atherosclerosis

1. Lipid Droplet Biogenesis. General Aspects

Mammalian cells accumulate excess neutral lipids, i.e., triacylglycerol (TG) and cholesterol esters (CE), in cytoplasmic organelles known as lipid droplets (LD). While LDs are present in almost all known cell types, their number, size and composition vary greatly from cell to cell and even within a given cell type. LDs are spherical particles composed of a phospholipid monolayer which encases a core made up mainly of TG and CE [1][2][3]. A large number of unique phospholipid molecular species has been identified in the LD monolayer. Most of these belong to the choline- and ethanolamine-containing glycerophospholipid classes (abbreviated as PC and PE, respectively), which represent approx. 64% and 24% of the total. Minor amounts of other classes such as phosphatidylinositol (PI), phosphatidylserine (PS), phosphatidic acid (PA), sphingomyelin, and lysophospholipids have also been identified [1][2][3].
The phospholipid monolayer is decorated with a variety of proteins that play numerous roles, ranging from regulating lipid mobilization from and to the organelle, to serving to support and stabilize the organelle structure itself. Of particular importance is the PAT family of proteins, which includes perilipin-1 (PLIN1), adipophilin (ADPR, PLIN2) and TIP47 (Tail-Interacting protein of 47 Kda, PLIN3) [4]. PLIN1 contributes to the growth and stabilization of LDs [5]. PLIN2 is involved in adipocyte differentiation and appears to be important for proper hydrolysis of the LDs [6]. The functions of PLIN3 are related to the regulation of lipolysis in skeletal muscle cells [7]. PLIN4 has been implicated in stabilizing the LD by interacting directly with TG moieties when their phospholipid coverage is limited [8]. PLIN5 appears to be involved in protective roles against mitochondrial damage and endoplasmic reticulum stress in liver [9].
In addition to the PAT proteins, numerous enzymes related to lipid metabolism have also been found on the surface of LDs; these include adipose triacylglycerol lipase (ATGL) [10], hormone-sensitive lipase [11], cytidine triphosphate:phosphocholine cytidylyltransferase [12][13], lysophosphatidylcholine acyltransferases [14], long-chain acyl-CoA synthetase 3 [15], lipin-1 [16][17], cyclooxygenase [18], and group IVA cytosolic phospholipase A2 (cPLA2α) [19][20], among others. Some of these proteins also participate in signaling [1][21][22][23][24]. Thus, the presence of such a varied number of lipid-metabolizing enzymes attests to the role of LDs as intracellular hubs where lipid signaling enzymes dock and interact to activate select pathways [1][21][22][23][24].
Although significant progress has been made in recent years, the mechanism of LD biogenesis is yet to be fully established. Formation of LDs involves a large number of steps and is carried out in the endoplasmic reticulum (ER), where the amphipathic monolayer of LDs originates [25][26] (Figure 1).
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Figure 1. Stages of LD synthesis at the cytosolic face of smooth ER membranes. The various enzymes of lipid metabolism participating in each stage are highlighted in red. Note the essential role of cPLA2α in favoring the induction of positive membrane curvature. TG and CE inside the LD are represented by brown and blue dots, respectively. Lysophospholipids at the membrane are highlighted as pink dots, phosphatidic acid as green dots. PL, phospholipid; DGAT, diacylglycerol:acyl-CoA acyl transferase; ACAT, acyl-CoA:cholesterol acyl transferase; PLD, phospholipase D; LPCAT, lysophosphatidylcholine:acyl-CoA acyl transferase.
The first step in LD formation is the synthesis of TG and CE, the major neutral lipids that compose the core of the LD. Different enzymes located in the ER are involved in their synthesis, namely diacylglycerol acyltransferases (DGAT) for TG, and acyl-CoA:cholesterol acyltransferases (ACAT) for CE. These enzymes may be activated by different stimuli including excess free fatty acids present in the medium, cell activation via surface receptors, or ER stress [27]. The free fatty acids that are used for neutral lipid synthesis may arise from multiple sources, both exogenous (lipoproteins from blood plasma), and endogenous (destruction of intracellular membranes or stimulation of the de novo fatty acid biosynthesis) [27].
A widely held assumption to explain the formation of LDs posits that the newly formed TG and CE accumulate in the ER. Some studies demonstrated that up to 3 mol% of TG and 5 mol% of CE can be accommodated before the LD breaks off to the cytosol, while other studies have suggested even greater accumulation, nearing 5–10% of both TG and CE. In either case, a highly regulated rate of phospholipid to neutral lipid must be maintained at all instances to support the shape and biophysical properties of the nascent organelle [28][29].
In order to grow, nascent LDs need a positive local membrane curvature while still in the ER membrane (Figure 1). This is achieved by the accumulation of lysophospholipids particularly lysoPA, lysoPC and lysoPE which, having a wedge-shaped conformation, help the LD to grow and counteract the neutral or slightly negative curvature produced by PC and PE, respectively [3][25][28][29]. Since lysophospholipids are generated by phospholipase A2s (PLA2), this kind of enzymes, in particular cPLA2α, are key regulators of the initial stages of LD formation [24][30][31][32][33]. Of note, cPLA2α manifests a marked selectivity for hydrolyzing phospholipids containing arachidonic acid at the sn-2 position [34], which makes the activation of this enzyme a key regulatory event for the synthesis of arachidonate-derived bioactive eicosanoids [35][36][37]. LDs have repeatedly been suggested to constitute a prominent eicosanoid-synthesizing site within the cells [38][39]; thus, it appears that cPLA2α may simultaneously serve two key functions of LD biology, i.e., to provide lysophospholipids to allow for continued growth of the organelle and, at the same time, to provide free AA substrate for the synthesis of bioactive lipid mediators in situ. This is further discussed in Section 7 below.
Once a positive curve has been generated at the LD baseline, cells are now able to store more TG and CE in between the phospholipid monolayers. As a result of such enhanced accumulation of neutral lipids, a simultaneous increase in the amount of PLs is necessary to maintain the biophysical properties of the monolayer of the nascent LD. This process requires the coordinate action of enzymes from the phospholipid biosynthetic and acyl chain remodeling pathways (i.e., phospholipid:acyl-CoA acyltransferases), the latter of which contribute to remove the lysophospholipids that accumulated in the membrane as a result of PLA2 action. As the LD emerges from the ER, a neutral curvature is established that stabilizes and protects the hydrophobic core from lipolysis [3][12][29].
In the latter stages of LD biogenesis, a negative curvature is generated at the connection sites with the ER to allow for the complete cleavage and release of the newly-formed LD to the cytosol. This effect is made possible by the accumulated presence of PA at the connection sites, which promotes a strong negative curvature [3][40]. PA is thought to be produced by phospholipase D enzymes [41][42] and removed, as needed, by lipin-1 [16][17]. Along the whole process described above, recruitment of a wide range of accessory proteins to the LD continuously takes place, such as the aforementioned proteins of the PAT family, which help stabilize the structure and assist in the recruitment of additional proteins and lipids [1][3][28][29][43].

2. Arachidonic Acid, a Compound Released in Atherosclerotic Lesions

Arachidonic acid (5,8,11,14-eicosatetraenoic acid, 20:4n-6, AA) is a member of the n-6 family of polyunsaturated fatty acids and can be obtained directly from the diet or synthesized from linoleic acid (18:2n-6) through the successive actions of Δ6-desaturase, elongase and Δ5-desaturase. Although produced mainly in the liver, practically all cells in the body are endowed with the machinery to produce AA from linoleic acid [44].
AA is the common precursor of the eicosanoids, a family of lipid mediators with key roles in physiology and especially in pathophysiological situations involving inflammatory reactions [36] (Figure 2). The potent biological activity of the eicosanoids requires the cells to exert a tight control on AA levels in a way that the availability of the fatty acid in free form is often a limiting factor for eicosanoid biosynthesis [45][46]. Of note, when present at sufficiently high concentrations, AA may be significantly converted to its two-carbon elongation product, adrenic acid (7,10,13,16-docosatetraenoic acid, 22:4n-6) [47][48][49], which can also be oxygenated to a variety of bioactive products [50][51][52].
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Figure 2. Pathways for the oxidative metabolism of AA. A variety of eicosanoids can be produced through four major pathways. These are: (i) the cyclooxygenase (COX) pathway, yielding prostaglandins and thromboxane; (ii) the lipoxygenase (LOX) pathway, yielding leukotrienes, hydroxyeicosatetraenoic acids (HETEs), hydroperoxyeicosatetraenoic acids (HPETEs), eoxins and leukotrienes; (iii) the cytochrome P450 (CYP450) pathway, yielding epoxyeicosatrienoic acids (EETs) and dihydroxyeicosatrienoic acids (DHETEs); and (iv) non-enzymatic oxidation reactions (Non-Enz), yielding isoprostanes and HETEs.
It has long been recognized that free AA is liberated in significant amounts into the bloodstream during the early stages of atherosclerosis [53]. Endothelial cells activated by oxidized LDLs and other stimuli constitute a major source of free AA [54][55][56]. In addition, the action of sPLA2s released from a variety of cells directly on the LDL-bound phospholipids constitutes another important source of AA released into the bloodstream [57][58][59][60]. Finally, platelets recruited to the activated endothelium also contribute to increase the availability of free AA [53] (Figure 3).
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Figure 3. Free AA production in atherosclerotic foci. Damaged endothelium releases substantial amounts of free AA which can be taken up by circulating monocytes and promote their change to a foamy phenotype. Platelets recruited to the activated endothelium also contribute to generating free AA. The sPLA2 action on circulating LDLs can also provide significant amounts of free AA.
Increased levels of free AA in the vicinity of atherosclerotic lesions may contribute greatly to LD formation by circulating monocytes, thereby transforming them into foamy cells and hence, into pro-atherogenic monocytes. This view has been experimentally supported by data demonstrating that free AA, at pathophysiologically relevant concentrations [61], is a strong inducer of LD formation in human peripheral blood monocytes [30]. Free AA appears to serve two different roles; on the one hand it serves as a substrate for the synthesis of TG but, interestingly, not CE. Conversely, AA activates intracellular signaling via p38 and JNK-mediated phosphorylation cascades that enhance neutral lipid synthesis and also activate cPLA2α, which in turn is required to regulate the biogenesis of the LD [30].
LD production by monocytes exposed to AA proceeds the same in the presence of a number of cyclooxygenase or lipoxygenase inhibitors [30]. Moreover, removal of the oxidized impurities from the commercial fatty acid (hydroxy, hydroperoxy and oxo derivatives of AA generated spontaneously) used in the above studies yielded a fraction of pure free AA that completely recapitulated LD formation in monocytes, while the oxidized AA fraction did not [62]. These results demonstrate that it is free AA itself and not an oxygenated product that is responsible for inducing the conversion of monocytes into foamy cells.
Molecular analyses of the lipid composition of the LDs produced by AA in foamy monocytes has shown that the neutral lipid fractions of these cells are enriched with an uncommon positional isomer of palmitoleic acid, namely cis-7-hexadecenoic acid, (16:1n-9) [63]. 16:1n-9 exhibits significant anti-inflammatory activity both in vivo and vitro which is comparable to that of n-3 fatty acids, and clearly distinguishable from that of palmitoleic acid [63][64]. Further, 16:1n-9 can be mobilized from phospholipids in activated phagocytic cells to form novel lipid species such as 16:1n-9-containing PI molecules and esters of 16:1n-9 with various hydroxyfatty acids [65]. These two kinds of compounds have been shown to possess growth-factor-like properties [66] and anti-diabetic/anti-inflammatory activities [67], respectively. The selective accumulation in the neutral lipid fraction of phagocytic cells of an uncommon fatty acid such as 16:1n-9 may reveal an early phenotypic change that could provide a biomarker of proatherogenicity, and a potential target for pharmacological intervention in the first stages of cardiovascular disease. Intriguingly, the 16:1n-9 fatty acid has also been recently identified in the TG fraction of some cancer cells [68], suggesting perhaps a wider biological role.

3. Glycerophospholipid Hydrolysis as a Major Pathway for the Mobilization of AA

The production of lipid mediators is intrinsically linked to the availability of polyunsaturated fatty acid (PUFA) precursors necessary for their synthesis. This depends on the activities of numerous enzymes and proteins that regulate the uptake, transport, storage, hydrolysis, remodeling and trafficking of PUFA among the different cellular and extracellular lipid pools. The major metabolic route supplying free PUFA for lipid mediator synthesis is that regulated by PLA2s, because these enzymes can directly access the major cellular reservoir of readily mobilizable fatty acid, i.e., the sn-2 position of membrane glycerophospholipids, mainly PC, PE, and PI [35][36][37]. It is relevant to indicate, however, that despite the overwhelming quantitative importance of PLA2s to overall PUFA release, there are other minor routes not involving PLA2 which can play important roles under limited, tightly controlled conditions. These include monoacylglycerol lipases acting on endocannabinoids [69][70], acid lipases acting on lysosomal lipids [71], and triacylglycerol lipase (ATGL) acting on TG [72][73].
The PLA2 enzymes typically involved in cellular signaling leading to lipid mediator production have been classically categorized into three major families, namely the Ca2+-dependent cytosolic enzymes (cPLA2), the Ca2+-independent enzymes (iPLA2), and the secreted enzymes (sPLA2). A number of excellent comprehensive reviews covering the classification, characteristics and activation properties of the more than 30 members of the PLA2 superfamily have recently been published, and the reader is kindly referred to these for specific details [34][74][75][76][77][78][79][80][81][82][83][84].
The group IVA PLA2, or cPLA2α, is widely recognized as the key enzyme effecting the AA release because of its unique preference for AA-containing phospholipid substrates and its activation properties, which place it at the center of a number of key signaling pathways involving phosphorylation cascades and/or intracellular Ca2+ movements [34][76][77]. In accordance, studies using cPLA2α-deficient mice have confirmed that this enzyme is essential for stimulus-induced eicosanoid production in practically all cells and tissues [34][74][75][76][77]. A myriad of stimuli, acting on surface receptors, are able to trigger the translocation activation of cPLA2α from the cytosol to a number of intracellular membranes, including the LD monolayer [19][20]. This allows positioning of the enzyme in the vicinity of cyclooxygenases and lipoxygenases for efficient supply of the free fatty acid for eicosanoid formation [85][86].
The group VIA calcium-independent PLA2, frequently referred to as iPLA2β, is another important enzyme for lipid mediator production which, unlike cPLA2α, does not manifest overt specificity for any particular fatty acid, being able to efficiently hydrolyze all kinds of phospholipid substrates [87]. Studies in macrophages have suggested that cPLA2α and iPLA2β preferentially act on different membrane phospholipid subsets, the former cleaving AA-containing phospholipids, and the latter liberating other fatty acids, such as adrenic acid and palmitoleic acid [47][48][88]. These in vivo preferences suggest that the activity of each PLA2 in stimulated cells can also be limited by the nature of the stimulus, the subcellular localization of the enzyme, and the accessibility to a given phospholipid pool [89][90].
The secreted PLA2s (sPLA2) constitute the third family of PLA2 enzymes involved in lipid mediator production [79][91]. The sPLA2s are secreted by a variety of cells, particularly those of the innate immune system, and act mainly on extracellular substrates such as lipoproteins, microparticles, bacteria and viruses, and the outer plasma membrane of mammalian cells. In some cases, they can also be incorporated back to the cells that first released them or to neighboring cells, and act on different intracellular membrane locations to regulate innate immune responses [92][93][94]. sPLA2 enzymes often act in concert with cPLA2α to assist and/o amplify the cPLA2α regulated response leading to PUFA mobilization [95][96][97]. However, under some circumstances, some sPLA2s, especially those of groups IID, III, and X, elicit significant production of PUFAs and associated mediators by acting on their own on different extracellular targets, and showing certain degree of selectivity for hydrolysis of PUFA over other monounsaturated/saturated fatty acids [98][99][100][101].
While less generally recognized, free AA levels can also be significantly increased in cells if the reacylation of lysophospholipids is inhibited [37][45][77][102][103][104]. This is because AA is an intermediate of a reacylation/deacylation cycle, the so-called Lands cycle, where the fatty acid is hydrolyzed from phospholipids by PLA2s and reincorporated back by CoA-dependent acyltrasferases [37][45]. In resting cells, reacylation reactions dominate and, as a result, free AA levels in unstimulated cells are kept at very low levels. In stimulated cells, activation of cPLA2α makes deacylation to dominate over reacylation, which results in the net accumulation of free AA. Nevertheless, AA reacylation under activation conditions is still significant, as demonstrated by the finding that a large amount of the AA initially liberated by cPLA2α is returned back to phospholipids. Hence, blockade of the CoA-dependent acyltransferases involved in phospholipid AA reacylation can result in greatly elevated levels of the free AA which is then available for eicosanoid synthesis [45]. The substrate specificity of the acyltransferases involved in AA reacylation is the reason why this fatty acid is overwhelmingly incorporated in the position sn-2 of phospholipids [77].
A third layer of complexity in the regulation of free AA availability stems from the fact that, once incorporated into phospholipids, the AA does not remain in the phospholipid molecular species that initially incorporated it but moves between different phospholipid species by a series of CoA-independent transacylation reactions [37][104][105]. These reactions are key for the cells to maintain the appropriate distribution of AA within the various cellular pools so that, depending on stimulation conditions, they are accessible to the relevant PLA2 [89][90]. This is an important aspect for the regulation of eicosanoid synthesis, as the quantity and distribution of eicosanoids produced under a given condition also depends on the composition and cellular localization of the phospholipid pool where the AA-hydrolyzing PLA2 primarily acts [89][90].
These transacylation reactions are catalyzed by CoA-independent transacylase (CoA-IT), an enzyme that primarily transfers AA moieties from PC (diacyl species) directly to PE (both diacyl and plasmalogens species), circumventing the need for CoA or ATP [37][104][105]. The sequence of CoA-IT is still unknown which has made it difficult to advance our knowledge on the cellular regulation of phospholipid transacylation. Recent studies have provided suggestive evidence that the CoA-IT-mediated reaction is primarily catalyzed by a well described PLA2 enzyme, the group IVC phospholipase A2γ (cPLA2γ) [89][105]. Unlike its homolog cPLA2α, cPLA2γ is a calcium-independent enzyme and does not manifest clear selectivity for AA residues [106]. Since the CoA-IT reaction involving AA-containing phospholipids appears to be critically involved in the inflammatory response of macrophages to certain stimuli [90][107], it is conceivable that other enzymes in addition to cPLA2γ may also serve a role as CoA-IT in cells. This is currently an area of active research [89][90][105][106][107][108].

References

  1. Olzmann, J.A.; Carvalho, P. Dynamics and Functions of Lipid Droplets. Nat. Rev. Mol. Cell Biol. 2019, 20, 137–155.
  2. Grillitsch, K.; Connerth, M.; Köfeler, H.; Arrey, T.N.; Rietschel, B.; Wagner, B.; Karas, M.; Daum, G. Lipid Particles/Droplets of the Yeast Saccharomyces Cerevisiae Revisited: Lipidome Meets Proteome. Biochim. Biophys. Acta 2011, 1811, 1165–1176.
  3. Thiam, A.R.; Farese, R.V., Jr.; Walther, T.C. The Biophysics and Cell Biology of Lipid Droplets. Nat. Rev. Mol. Cell Biol. 2013, 14, 775–786.
  4. Brasaemle, D.L. The Perilipin Family of Structural Lipid Droplet Proteins: Stabilization of Lipid Droplets and Control of Lipolysis. J. Lipid Res. 2007, 48, 2547–2559.
  5. Sun, Z.; Gong, J.; Wu, H.; Xu, W.; Wu, L.; Xu, D.; Gao, J.; Wu, J.W.; Yang, H.; Yang, M.; et al. Perilipin1 Promotes Unilocular Lipid Droplet Formation through the Activation of Fsp27 in Adipocytes. Nat. Commun. 2013, 4, 1594.
  6. Mardani, I.; Dalen, K.I.; Drevinge, C.; Miljanovic, A.; Ståhlman, M.; Klevstig, M.; Täng, M.S.; Fogelstrand, P.; Levin, M.; Ekstrand, M.; et al. Plin2-Deficiency Reduces Lipophagy and Results in Increased Lipid Accumulation in the Heart. Sci. Rep. 2019, 9, 6909.
  7. MacPherson, R.E.K.; Vandenboom, R.; Roy, B.D.; Peters, S.J. Skeletal Muscle PLIN3 and PLIN5 Are Serine Phosphorylated at Rest and Following Lipolysis during Adrenergic or Contractile Stimulation. Physiol. Rep. 2013, 1, e00084.
  8. Čopič, A.; Antoine-Bally, S.; Giménez-Andrés, M.; La Torre Garay, C.; Antonny, B.; Manni, M.M.; Pagnotta, S.; Guihot, J.; Jackson, C.L. A Giant Amphipathic Helix from a Perilipin That Is Adapted for Coating Lipid Droplets. Nat. Commun. 2018, 9, 1332.
  9. Sanchez, P.B.M.; Krizanac, M.; Weiskirchen, R.; Asimakopoulos, A. Understanding the Role of Perilipin 5 in Non-Alcoholic Fatty Liver Disease and Its Role in Hepatocellular Carcinoma: A Review of Novel Insights. Int. J. Mol. Sci. 2021, 22, 5284.
  10. Lass, A.; Zimmermann, R.; Oberer, M.; Zechner, R. Lipolysis-A Highly Regulated Multi-Enzyme Complex Mediates the Catabolism of Cellular Fat Stores. Prog. Lipid Res. 2011, 50, 14–27.
  11. Shen, W.J.; Patel, S.; Miyoshi, H.; Greenberg, A.S.; Kraemer, F.B. Functional Interaction of Hormone-Sensitive Lipase and Perilipin in Lipolysis. J. Lipid Res. 2009, 50, 2306–2313.
  12. Krahmer, N.; Guo, Y.; Wilfling, F.; Hilger, M.; Lingrell, S.; Heger, K.; Newman, H.W.; Schmidt-Supprian, M.; Vance, D.E.; Mann, M.; et al. Phosphatidylcholine Synthesis for Lipid Droplet Expansion Is Mediated by Localized Activation of CTP:Phosphocholine Cytidylyltransferase. Cell Metab. 2011, 14, 504–515.
  13. Aitchison, A.J.; Arsenault, D.J.; Ridgway, N.D. Nuclear-Localized CTP: Phosphocholine Cytidylyltransferase α Regulates Phosphatidylcholine Synthesis Required for Lipid Droplet Biogenesis. Mol. Biol. Cell 2015, 26, 2927–2938.
  14. Moessinger, C.; Kuerschner, L.; Spandl, J.; Shevchenko, A.; Thiele, C. Human Lysophosphatidylcholine Acyltransferases 1 and 2 Are Located in Lipid Droplets Where They Catalyze the Formation of Phosphatidylcholine. J. Biol. Chem. 2011, 286, 21330–21339.
  15. Poppelreuther, M.; Rudolph, B.; Du, C.; Grossmann, R.; Becker, M.; Thiele, C.; Ehehalt, R.; Füllekrug, J. The N-Terminal Region of Acyl-CoA Synthetase 3 Is Essential for Both the Localization on Lipid Droplets and the Function in Fatty Acid Uptake. J. Lipid Res. 2012, 53, 888–900.
  16. Valdearcos, M.; Esquinas, E.; Meana, C.; Gil-de-Gómez, L.; Guijas, C.; Balsinde, J.; Balboa, M.A. Subcellular Localization and Role of Lipin-1 in Human Macrophages. J. Immunol. 2011, 186, 6004–6013.
  17. Sembongi, H.; Miranda, M.; Han, G.S.; Fakas, S.; Grimsey, N.; Vendrell, J.; Carman, G.M.; Siniossoglou, S. Distinct Roles of the Phosphatidate Phosphatases Lipin 1 and 2 during Adipogenesis and Lipid Droplet Biogenesis in 3T3-L1 Cells. J. Biol. Chem. 2013, 288, 34502–34513.
  18. Melo, R.C.N.; Weller, P.F. Unraveling the Complexity of Lipid Body Organelles in Human Eosinophils. J. Leukoc. Biol. 2014, 96, 703–712.
  19. Yu, W.; Bozza, P.T.; Tzizik, D.M.; Gray, J.P.; Cassara, J.; Dvorak, A.M.; Weller, P.F. Co-Compartmentalization of MAP Kinases and Cytosolic Phospholipase A2 at Cytoplasmic Arachidonate-Rich Lipid Bodies. Am. J. Pathol. 1998, 152, 759–769.
  20. Wooten, R.E.; Willingham, M.C.; Daniel, L.W.; Leslie, C.C.; Rogers, L.C.; Sergeant, S.; O’Flaherty, J.T. Novel Translocation Responses of Cytosolic Phospholipase A2α Fluorescent Proteins. Biochim. Biophys. Acta 2008, 1783, 1544–1550.
  21. Jarc, E.; Petan, T. A Twist of FATe: Lipid Droplets and Inflammatory Lipid Mediators. Biochimie 2020, 169, 69–87.
  22. Pérez-Chacón, G.; Astudillo, A.M.; Ruipérez, V.; Balboa, M.A.; Balsinde, J. Signaling Role for Lysophosphatidylcholine Acyltransferase 3 in Receptor-Regulated Arachidonic Acid Reacylation Reactions in Human Monocytes. J. Immunol. 2010, 184, 1071–1078.
  23. Arrese, E.L.; Saudale, F.Z.; Soulages, J.L. Lipid Droplets as Signaling Platforms Linking Metabolic and Cellular Functions. Lipid Insights 2014, 7, 7–16.
  24. Guijas, C.; Rodríguez, J.P.; Rubio, J.M.; Balboa, M.A.; Balsinde, J. Phospholipase A2 Regulation of Lipid Droplet Formation. Biochim. Biophys. Acta 2014, 1841, 1661–1671.
  25. Wilfling, F.; Wang, H.; Haas, J.T.; Krahmer, N.; Gould, T.J.; Uchida, A.; Cheng, J.X.; Graham, M.; Christiano, R.; Fröhlich, F.; et al. Triacylglycerol Synthesis Enzymes Mediate Lipid Droplet Growth by Relocalizing from the ER to Lipid Droplets. Dev. Cell 2013, 24, 384–399.
  26. Jackson, C.L. Lipid Droplet Biogenesis. Curr. Opin. Cell Biol. 2018, 59, 88–96.
  27. Coleman, R.A.; Lee, D.P. Enzymes of Triacylglycerol Synthesis and Their Regulation. Prog. Lipid Res. 2004, 43, 134–176.
  28. Thiam, A.R.; Ikonen, E. Lipid Droplet Nucleation. Trends Cell Biol. 2021, 31, 108–118.
  29. Pol, A.; Gross, S.P.; Parton, R.G. Biogenesis of the Multifunctional Lipid Droplet: Lipids, Proteins, and Sites. J. Cell Biol. 2014, 204, 635–646.
  30. Guijas, C.; Pérez-Chacón, G.; Astudillo, A.M.; Rubio, J.M.; Gil-de-Gómez, L.; Balboa, M.A.; Balsinde, J. Simultaneous Activation of p38 and JNK by Arachidonic Acid Stimulates the Cytosolic Phospholipase A2-Dependent Synthesis of Lipid Droplets in Human Monocytes. J. Lipid Res. 2012, 53, 2343–2354.
  31. Gubern, A.; Casas, J.; Barceló-Torns, M.; Barneda, D.; de la Rosa, X.; Masgrau, R.; Picatoste, F.; Balsinde, J.; Balboa, M.A.; Claro, E. Group IVA Phospholipase A2 Is Necessary for the Biogenesis of Lipid Droplets. J. Biol. Chem. 2008, 283, 27369–27382.
  32. Gubern, A.; Barceló-Torns, M.; Casas, J.; Barneda, D.; Masgrau, R.; Picatoste, F.; Balsinde, J.; Balboa, M.A.; Claro, E. Lipid Droplet Biogenesis Induced by Stress Involves Triacylglycerol Synthesis that Depends on Group VIA Phospholipase A2. J. Biol. Chem. 2009, 284, 5697–5708.
  33. Gubern, A.; Barceló-Torns, M.; Barneda, D.; López, J.M.; Masgrau, R.; Picatoste, F.; Chalfant, C.E.; Balsinde, J.; Balboa, M.A.; Claro, E. JNK and Ceramide Kinase Govern the Biogenesis of Lipid Droplets through Activation of Group IVA Phospholipase A2. J. Biol. Chem. 2009, 284, 32359–32369.
  34. Leslie, C.C. Cytosolic Phospholipase A2: Physiological Function and Role in Disease. J. Lipid Res. 2015, 56, 1386–1402.
  35. Balsinde, J.; Winstead, M.V.; Dennis, E.A. Phospholipase A2 Regulation of Arachidonic Acid Mobilization. FEBS Lett. 2002, 531, 2–6.
  36. Dennis, E.A.; Norris, P.C. Eicosanoid Storm in Infection and Inflammation. Nat. Rev. Immunol. 2015, 15, 511–523.
  37. Astudillo, A.M.; Balboa, M.A.; Balsinde, J. Selectivity of Phospholipid Hydrolysis by Phospholipase A2 Enzymes in Activated Cells Leading to Polyunsaturated Fatty Acid Mobilization. Biochim. Biophys. Acta 2019, 1864, 772–783.
  38. Melo, R.C.N.; Weller, P. Lipid Droplets in Leukocytes: Organelles Linked to Inflammatory Responses. Exp. Cell. Res. 2016, 340, 193–197.
  39. Bozza, P.T.; Bakker-Abreu, I.; Navarro-Xavier, R.A.; Bandeira-Melo, C. Lipid Body Function in Eicosanoid Synthesis: An Update. Prostaglandins Leukot. Essent. Fatty Acids 2011, 85, 205–213.
  40. Kooijman, E.E.; Chupin, V.; Fuller, N.L.; Kozlov, M.M.; de Kruijff, B.; Burger, K.N.; Rand, P.R. Spontaneous Curvature of Phosphatidic Acid and Lysophosphatidic Acid. Biochemistry 2005, 44, 2097–2102.
  41. Andersson, L.; Boström, P.; Ericson, J.; Rutberg, M.; Magnusson, B.; Marchesan, D.; Ruiz, M.; Asp, L.; Huang, P.; Frohman, M.A.; et al. PLD1 and ERK2 Regulate Cytosolic Lipid Droplet Formation. J. Cell Sci. 2006, 119, 2246–2257.
  42. Balsinde, J.; Diez, E.; Fernández, B.; Mollinedo, F. Biochemical Characterization of Phospholipase D Activity from Human Neutrophils. Eur. J. Biochem. 1989, 186, 717–724.
  43. Tan, J.S.; Seow, C.J.; Goh, V.J.; Silver, D.L. Recent Advances in Understanding Proteins Involved in Lipid Droplet Formation, Growth and Fusion. J. Genet. Genom. 2014, 41, 251–259.
  44. Sprecher, H. Metabolism of Highly Unsaturated N-3 and N-6 Fatty Acids. Biochim. Biophys. Acta 2000, 1486, 219–231.
  45. Pérez-Chacón, G.; Astudillo, A.M.; Balgoma, D.; Balboa, M.A.; Balsinde, J. Control of Free Arachidonic Acid Levels by Phospholipases A2 and Lysophospholipid Acyltransferases. Biochim. Biophys. Acta 2009, 1791, 1103–1113.
  46. Astudillo, A.M.; Balgoma, D.; Balboa, M.A.; Balsinde, J. Dynamics of Arachidonic Acid Mobilization by Inflammatory Cells. Biochim. Biophys. Acta 2012, 1821, 249–256.
  47. Guijas, C.; Astudillo, A.M.; Gil-de-Gómez, L.; Rubio, J.M.; Balboa, M.A.; Balsinde, J. Phospholipid Sources for Adrenic Acid Mobilization in RAW 264.7 Macrophages: Comparison with Arachidonic Acid. Biochim. Biophys. Acta 2012, 1821, 1386–1393.
  48. Monge, P.; Garrido, A.; Rubio, J.M.; Magrioti, V.; Kokotos, G.; Balboa, M.A.; Balsinde, J. The Contribution of Cytosolic Group IVA and Calcium-Independent Group VIA Phospholipase A2s to Adrenic Acid Mobilization in Murine Macrophages. Biomolecules 2020, 10, 542.
  49. Brouwers, H.; Jónasdóttir, H.S.; Kuipers, M.E.; Kwekkeboom, J.C.; Auger, J.L.; González-Torres, M.; López-Vicario, C.; Clària, J.; Freysdottir, J.; Hardardottir, I.; et al. Anti-Inflammatory and Proresolving Effects of the Omega-6 Polyunsaturated Fatty Acid Adrenic Acid. J. Immunol. 2020, 205, 2840–2849.
  50. Harkewicz, R.; Fahy, E.; Andreyev, A.; Dennis, E.A. Arachidonate-Derived Dihomoprostaglandin Production Observed in Endotoxin-Stimulated Macrophage-like Cells. J. Biol. Chem. 2007, 282, 2899–2910.
  51. Sprecher, H.; Van Rollins, M.; Sun, F.; Wyche, A.; Needleman, P. Dihomo-Prostaglandins and -Thromboxane. A Prostaglandin Family from Adrenic Acid That May Be Preferentially Synthesized in the Kidney. J. Biol. Chem. 1982, 257, 3912–3918.
  52. Kopf, P.G.; Zhang, D.X.; Gauthier, K.M.; Nithipatikom, K.; Yi, X.Y.; Falck, J.R.; Campbell, W.B. Adrenic Acid Metabolites as Endogenous Endothelium-Derived and Zona Glomerulosa-Derived Hyperpolarizing Factors. Hypertension 2010, 55, 547–554.
  53. Ross, R. Atherosclerosis—an Inflammatory Disease. N. Engl. J. Med. 1999, 340, 115–126.
  54. Wong, J.T.; Tran, K.; Pierce, G.N.; Chan, A.C.; O., K.; Choy, P.C. Lysophosphatidylcholine Stimulates the Release of Arachidonic Acid in Human Endothelial Cells. J. Biol. Chem. 1998, 273, 6830–6836.
  55. Bogatcheva, N.V.; Sergeeva, M.G.; Dudek, S.M.; Verin, A.D. Arachidonic Acid Cascade in Endothelial Pathobiology. Microvasc. Res. 2005, 69, 107–127.
  56. Badimon, L.; Vilahur, G.; Rocca, B.; Patrono, C. The Key Contribution of Platelet and Vascular Arachidonic Acid Metabolism to the Pathophysiology of Atherothrombosis. Cardiovasc. Res. 2021, 117, 2001–2015.
  57. Hanasaki, K.; Yamada, K.; Yamamoto, S.; Ishimoto, Y.; Saiga, A.; Ono, T.; Ikeda, M.; Notoya, M.; Kamitani, S.; Arita, H. Potent Modification of Low Density Lipoprotein by Group X Secretory Phospholipase A2 Is Linked to Macrophage Foam Cell Formation. J. Biol. Chem. 2002, 277, 29116–29124.
  58. Balsinde, J.; Balboa, M.A.; Yedgar, S.; Dennis, E.A. Group V Phospholipase A2-Mediated Oleic Acid Mobilization in Lipopolysaccharide-Stimulated P388D1 Macrophages. J. Biol. Chem. 2000, 275, 4783–4786.
  59. Sato, H.; Kato, R.; Isogai, Y.; Saka, G.; Ohtsuki, M.; Taketomi, Y.; Yamamoto, K.; Tsutsumi, K.; Yamada, J.; Masuda, S.; et al. Analyses of Group III Secreted Phospholipase A2 Transgenic Mice Reveal Potential Participation of This Enzyme in Plasma Lipoprotein Modification, Macrophage Foam Cell Formation, and Atherosclerosis. J. Biol. Chem. 2008, 283, 23483–33497.
  60. Karabina, S.A.; Brochériou, I.; Le Naour, G.; Agrapart, M.; Durand, H.; Gelb, M.; Lambeau, G.; Ninio, E. Atherogenic Properties of LDL Particles Modified by Human Group X Secreted Phospholipase A2 on Human Endothelial Cell Function. FASEB J. 2006, 20, 2547–2549.
  61. Tallima, H.; El Ridi, R. Arachidonic Acid: Physiological Roles and Potential Health Benefits–A Review. J. Adv. Res. 2018, 11, 33–41.
  62. Guijas, C.; Bermúdez, M.A.; Meana, C.; Astudillo, A.M.; Pereira, L.; Fernández-Caballero, L.; Balboa, M.A.; Balsinde, J. Neutral Lipids Are Not a Source of Arachidonic Acid for Lipid Mediator Signaling in Human Foamy Monocytes. Cells 2019, 8, 941.
  63. Guijas, C.; Meana, C.; Astudillo, A.M.; Balboa, M.A.; Balsinde, J. Foamy Monocytes Are Enriched in Cis-7-Hexadecenoic Fatty Acid (16:1n-9), a Possible Biomarker for Early Detection of Cardiovascular Disease. Cell Chem. Biol. 2016, 23, 689–699.
  64. Astudillo, A.M.; Meana, C.; Guijas, C.; Pereira, L.; Lebrero, R.; Balboa, M.A.; Balsinde, J. Occurrence and Biological Activity of Palmitoleic Acid Isomers in Phagocytic Cells. J. Lipid Res. 2018, 59, 237–249.
  65. Astudillo, A.M.; Meana, C.; Bermúdez, M.A.; Pérez-Encabo, A.; Balboa, M.A.; Balsinde, J. Release of Anti-Inflammatory Palmitoleic Acid and Its Positional Isomers by Mouse Peritoneal Macrophages. Biomedicines 2020, 8, 480.
  66. Scanferlato, R.; Bortolotti, M.; Sansone, A.; Chatgilialoglu, C.; Polito, L.; De Spirito, M.; Maulucci, G.; Bolognesi, A.; Ferreri, C. Hexadecenoic Fatty Acid Positional Isomers and De Novo PUFA Synthesis in Colon Cancer Cells. Int. J. Mol. Sci. 2019, 20, 832.
  67. Aryal, P.; Syed, I.; Lee, J.; Patel, R.; Nelson, A.T.; Siegel, D.; Saghatelian, A.; Kahn, B.A. Distinct Biological Activities of Isomers from Several Families of Branched Fatty Acid Esters of Hydroxy Fatty Acids (FAHFAs). J. Lipid Res. 2021, 62, 100108.
  68. Young, R.S.E.; Bowman, A.P.; Williams, E.D.; Tousignant, K.D.; Bidgood, C.L.; Narreddula, V.R.; Gupta, R.; Marshall, D.L.; Poad, B.L.J.; Nelson, C.C.; et al. Apocryphal FADS2 Activity Promotes Fatty Acid Diversification in Cancer. Cell Rep. 2021, 34, 108738.
  69. Nomura, D.K.; Morrison, B.E.; Blankman, J.L.; Long, J.Z.; Kinsey, S.G.; Marcondes, M.C.; Ward, A.M.; Hahn, Y.K.; Lichtman, A.H.; Conti, B.; et al. Endocannabinoid Hydrolysis Generates Brain Prostaglandins That Promote Neuroinflammation. Science 2011, 334, 809–813.
  70. Grabner, G.F.; Eichmann, T.O.; Wagner, B.; Gao, Y.; Farzi, A.; Taschler, U.; Radner, F.P.; Schweiger, M.; Lass, A.; Holzer, P.; et al. Deletion of Monoglyceride Lipase in Astrocytes Attenuates Lipopolysaccharide-Induced Neuroinflammation. J. Biol. Chem. 2016, 291, 913–923.
  71. Schlager, S.; Vujic, N.; Korbelius, M.; Duta-Mare, M.; Dorow, J.; Leopold, C.; Rainer, S.; Wegscheider, M.; Reicher, H.; Ceglarek, U.; et al. Lysosomal Lipid Hydrolysis Provides Substrates for Lipid Mediator Synthesis in Murine Macrophages. Oncotarget 2017, 8, 40037–40051.
  72. Dichlberger, A.; Schlager, S.; Maaninka, K.; Schneider, W.J.; Kovanen, P.T. Adipose Triglyceride Lipase Regulates Eicosanoid Production in Activated Human Mast Cells. J. Lipid Res. 2014, 55, 2471–2478.
  73. Schlager, S.; Goeritzer, M.; Jandl, K.; Frei, R.; Vujic, N.; Kolb, D.; Strohmaier, H.; Dorow, J.; Eichmann, T.O.; Rosenberger, A.; et al. Adipose Triglyceride Lipase Acts on Neutrophil Lipid Droplets to Regulate Substrate Availability for Lipid Mediator Synthesis. J. Leukoc. Biol. 2015, 98, 837–850.
  74. 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. Chem. Rev. 2011, 111, 6130–6185.
  75. Murakami, M. Novel Functions of Phospholipase A2s: Overview. Biochim. Biophys. Acta 2019, 1864, 763–765.
  76. Mouchlis, V.D.; Dennis, E.A. Phospholipase A2 Catalysis and Lipid Mediator Lipidomics. Biochim. Biophys. Acta 2019, 1864, 766–771.
  77. Kita, Y.; Shindou, H.; Shimizu, T. Cytosolic Phospholipase A2 and Lysophospholipid Acyltransferases. Biochim. Biophys. Acta 2019, 1864, 838–845.
  78. Turk, J.; White, T.D.; Nelson, A.J.; Lei, X.; Ramanadham, S. iPLA2β and Its Role in Male Fertility, Neurological Disorders, Metabolic Disorders, and Inflammation. Biochim. Biophys. Acta 2019, 1864, 846–860.
  79. Murakami, M.; Sato, H.; Miki, Y.; Yamamoto, K.; Taketomi, Y. A New Era of Secreted Phospholipase A2. J. Lipid Res. 2015, 56, 1248–1261.
  80. Murakami, M.; Sato, H.; Taketomi, Y. Updating Phospholipase A2 Biology. Biomolecules 2020, 10, 1457.
  81. White, T.D.; Almutairi, A.; Tusing, Y.G.; Lei, X.; Ramanadham, S. The Impact of the Ca2+-Independent Phospholipase A2β on Immune Cells. Biomolecules 2021, 11, 577.
  82. Dabral, D.; van den Bogaart, G. The Roles of Phospholipase A2 in Phagocytes. Front. Cell Dev. Biol. 2021, 9, 673502.
  83. Peng, Z.; Chang, Y.; Fan, J.; Ji, W.; Su, C. Phospholipase A2 Superfamily in Cancer. Cancer Lett. 2021, 497, 165–177.
  84. Sun, G.Y.; Geng, X.; Teng, T.; Yang, B.; Appenteng, M.K.; Greenlief, C.M. Dynamic Role of Phospholipases A2 in Health and Diseases in the Central Nervous System. Cells 2021, 10, 2963.
  85. Pindado, J.; Balsinde, J.; Balboa, M.A. TLR3-Dependent Induction of Nitric Oxide Synthase in RAW 264.7 Macrophage-like Cells Via a Cytosolic Phospholipase A2/Cyclooxygenase-2 Pathway. J. Immunol. 2007, 179, 4821–4828.
  86. Astudillo, A.M.; Rodríguez, J.P.; Guijas, C.; Rubio, J.M.; Balboa, M.A.; Balsinde, J. Choline Glycerophospholipid-Derived Prostaglandins Attenuate TNFα Gene Expression in Macrophages Via a cPLA2α/COX-1 Pathway. Cells 2021, 10, 447.
  87. Mouchlis, V.D.; Chen, Y.; McCammon, J.A.; Dennis, E.A. Membrane Allostery and Unique Hydrophobic Sites Promote Enzyme Substrate Specificity. J. Am. Chem. Soc. 2018, 140, 3285–3291.
  88. Gil-de-Gómez, L.; Astudillo, A.M.; Guijas, C.; Magrioti, V.; Kokotos, G.; Balboa, M.A.; Balsinde, J. Cytosolic Group IVA and Calcium-Independent Group VIA Phospholipase A2s Act on Distinct Phospholipid Pools in Zymosan-Stimulated Mouse Peritoneal Macrophages. J. Immunol. 2014, 192, 752–762.
  89. Lebrero, P.; Astudillo, A.M.; Rubio, J.M.; Fernández-Caballero, J.; Kokotos, G.; Balboa, M.A.; Balsinde, J. Cellular Plasmalogen Content Does Not Influence Arachidonic Acid Levels or Distribution in Macrophages: A Role for Cytosolic Phospholipase A2γ in Phospholipid Remodeling. Cells 2019, 8, 799.
  90. Gil-de-Gómez, L.; Monge, P.; Rodríguez, J.P.; Astudillo, A.M.; Balboa, M.A.; Balsinde, J. Phospholipid Arachidonic acid Remodeling during Phagocytosis in Mouse Peritoneal Macrophages. Biomedicines 2020, 8, 274.
  91. Murakami, M.; Lambeau, G. Emerging Roles of Secreted Phospholipase A2 Enzymes: An Update. Biochimie 2013, 95, 43–50.
  92. Samuchiwal, S.K.; Balestrieri, B. Harmful and Protective Roles of Group V Phospholipase A2: Current Perspectives and Future Directions. Biochim. Biophys. Acta 2019, 1864, 819–826.
  93. Balboa, M.A.; Shirai, Y.; Gaietta, G.; Ellisman, M.H.; Balsinde, J.; Dennis, E.A. Localization of Group V Phospholipase A2 in Caveolin-Enriched Granules in Activated P388D1 Macrophage-like Cells. J. Biol. Chem. 2003, 278, 48059–48065.
  94. Bingham, C.O.; Fijneman, R.J.; Friend, D.S.; Goddeau, R.P.; Rogers, R.A.; Austen, K.F.; Arm, J.P. Low Molecular Weight Group IIA and Group V Phospholipase A2 Enzymes Have Different Intracellular Locations in Mouse Bone Marrow-Derived Mast Cells. J. Biol. Chem. 1999, 274, 31476–31484.
  95. Ruipérez, V.; Astudillo, M.A.; Balboa, M.A.; Balsinde, J. Coordinate Regulation of TLR-Mediated Arachidonic Acid Mobilization in Macrophages by Group IVA and Group V Phospholipase A2s. J. Immunol. 2009, 182, 3877–3883.
  96. Balboa, M.A.; Pérez, R.; Balsinde, J. Amplification Mechanisms of Inflammation: Paracrine Stimulation of Arachidonic Acid Mobilization by Secreted Phospholipase A2 Is Regulated by Cytosolic Phospholipase A2-Derived Hydroperoxyeicosatetraenoic Acid. J. Immunol. 2003, 171, 989–994.
  97. Kikawada, E.; Bonventre, J.V.; Arm, J.P. Group V Secretory PLA2 Regulates TLR2-Dependent Eicosanoid Generation in Mouse Mast Cells through Amplification of ERK and cPLA2α Activation. Blood 2007, 110, 561–567.
  98. Murakami, M.; Miki, Y.; Sato, H.; Murase, R.; Taketomi, Y.; Yamamoto, K. Group IID, IIE, IIF and III Secreted Phospholipase A2s. Biochim. Biophys. Acta 2019, 1864, 803–818.
  99. Miki, Y.; Yamamoto, K.; Taketomi, Y.; Sato, H.; Shimo, K.; Kobayashi, T.; Ishikawa, Y.; Ishii, T.; Nakanishi, H.; Ikeda, K.; et al. Lymphoid Tissue Phospholipase A2 Group IID Resolves Contact Hypersensitivity by Driving Antiinflammatory Lipid Mediators. J. Exp. Med. 2013, 210, 1217–1234.
  100. Ait-Oufella, H.; Herbin, O.; Lahoute, C.; Coatrieux, C.; Loyer, X.; Joffre, J.; Laurans, L.; Ramkhelawon, B.; Blanc-Brude, O.; Karabina, S.; et al. Group X Secreted Phospholipase A2 Limits the Development of Atherosclerosis in LDL Receptor-Null Mice. Arterioscler. Thromb. Vasc. Biol. 2013, 33, 466–473.
  101. Murase, R.; Sato, H.; Yamamoto, K.; Ushida, A.; Nishito, Y.; Ikeda, K.; Kobayashi, T.; Yamamoto, T.; Taketomi, Y.; Murakami, M. Group X Secreted Phospholipase A2 Releases ω3 Polyunsaturated Fatty Acids, Suppresses Colitis, and Promotes Sperm Fertility. J. Biol. Chem. 2016, 291, 6895–6911.
  102. Murphy, R.C.; Folco, G. Lysophospholipid Acyltransferases and Leukotriene Biosynthesis: Intersection of the Lands Cycle and the Arachidonate PI Cycle. J. Lipid Res. 2019, 60, 219–226.
  103. Patton-Vogt, J.; de Kroon, A.I.P.M. Phospholipid Turnover and Acyl Chain Remodeling in the Yeast ER. Biochim. Biophys. Acta 2020, 1865, 158462.
  104. Yamashita, A.; Hayashi, Y.; Nemoto-Sasaki, Y.; Ito, M.; Oka, S.; Tanikawa, T.; Waku, K.; Sugiura, T. Acyltransferases and Transacylases That Determine the Fatty Acid Composition of Glycerolipids and the Metabolism of Bioactive Lipid Mediators in Mammalian Cells and Model Organisms. Prog. Lipid Res. 2014, 53, 18–81.
  105. Yamashita, A.; Hayashi, Y.; Matsumoto, N.; Nemoto-Sasaki, Y.; Koizumi, T.; Inagaki, Y.; Oka, S.; Tanikawa, T.; Sugiura, T. Coenzyme-A-Independent Transacylation System; Possible Involvement of Phospholipase A2 in Transacylation. Biology 2017, 6, 23.
  106. Ghosh, M.; Tucker, D.E.; Burchett, S.A.; Leslie, C.C. Properties of the Group IV Phospholipase A2 Family. Prog. Lipid Res. 2006, 45, 487–510.
  107. Gil-de-Gómez, L.; Astudillo, A.M.; Lebrero, P.; Balboa, M.A.; Balsinde, J. Essential Role for Ethanolamine Plasmalogen Hydrolysis in Bacterial Lipopolysaccharide Priming of Macrophages for Enhanced Arachidonic Acid Release. Front. Immunol. 2017, 8, 1251.
  108. Rubio, J.M.; Astudillo, A.M.; Casas, J.; Balboa, M.A.; Balsinde, J. Regulation of Phagocytosis in Macrophages by Membrane Ethanolamine Plasmalogens. Front. Immunol. 2018, 9, 1723.
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