Myeloid-Specific Deficiency of Long-Chain Acyl CoA Synthetase 4 Reduces Inflammation by Remodeling Phospholipids and Reducing Production of Arachidonic Acid–Derived Proinflammatory Lipid Mediators

Tissue-resident macrophages are sentinel cells that survey their environment to detect pathogens or tissue damage (1). Resident peritoneal macrophages (rpMACs) are long-lived cells derived from the yolk sac during development and maintain their populations through self-renewal rather than recruitment from bone marrow–derived monocytes (2). Upon sensing infection or injury, resident macrophages become activated and produce inflammatory signaling molecules that lead to endothelial permeability, swelling, and the recruitment of immune cells (3, 4). Chronologically, neutrophils are typically the first myeloid cell type recruited after tissue injury or infection, followed by monocytes and then macrophages that aid in the clearance of pathogens and dead cells, and help to orchestrate the resolution of inflammation and the repair and remodeling of tissue (5).

An important aspect of inflammatory activation of macrophages is the release of lipid mediators generated by the enzymatic oxidation of polyunsaturated fatty acids (PUFAs). Arachidonic acid (AA; 20:4n-6) is an ω-6 PUFA that is the precursor for the biosynthesis of many proinflammatory lipid mediators. AA can be enzymatically oxidized to generate proinflammatory lipid mediators, such as PGs, thromboxanes, and leukotrienes, as well as several other classes of lipid mediators, all of which have a broad range of biological activities. Each of the lipid mediators can have various, but specific, roles during inflammation. For instance, PGE₂ can cause pain, increased blood flow to the injured area, and ultimately lead to edema (6, 7). The actions of PGE₂ on macrophages during inflammation are multifactorial and include both proinflammatory and anti-inflammatory/immunosuppressive functions. For example, PGE₂ enhances IL-1β production in response to LPS but can also blunt TLR4 expression, enhance suppressor of cytokine 3 signaling, and diminish macrophage phagocytosis (8–11). Leukotriene B₄ (LTB₄) is a potent chemoattractant that recruits new proinflammatory immune cells, such as neutrophils and macrophages (12, 13). Another group of AA-derived lipid mediators, the hydroxyeicosatetraenoic acids (HETEs), has diverse effects on inflammation based on the position and stereochemistry of the hydroxyl group. The murine 12/15-lipoxegenases (12/15-LOX), for example, can generate both 12- and 15-HETE from AA and are thus markers of this lipoxygenase activity (14). The 12-HETE product is proinflammatory and leads to neutrophil aggregation and an itching sensation demonstrating that the position of an individual hydroxyl group is critical for the activity of many of these lipid mediators (15, 16).

The resolution of inflammation is an active process that macrophages contribute to as well, which involves reduction of inflammatory signals and coordinated remodeling of the damaged tissues by clearance of apoptotic immune cell infiltrate and dead cell debris (17, 18). AA can have a crucial role in resolving inflammation when it is sequentially oxidized by murine 12/15-LOX and 5-LOX to generate the specialized proresolving lipid mediators (SPMs) termed lipoxins (LXs) (13). Similar to the proinflammatory lipid mediators, the biological actions of SPMs are mediated by activation of specific receptors that can result in a broad range of effects, such as suppressing inflammatory signaling and neutrophil recruitment, increasing macrophage efferocytosis, and promoting tissue repair, among many of their various functions (19).

Long-chain acyl-CoA synthetases (ACSLs) catalyze the thioesterification of long-chain fatty acids (FAs) and CoAs to form a fatty acyl-CoA. There are five ACSL isoforms (1, 3–6) in mammals that localize to several different subcellular compartments and differ in their preferences for specific FAs (20). Mouse, rat, and human ACSL4 have strong preferences for AA, EPA, and adrenic acid (22:4n-6) (21–24). Human ACSL4 has a very strong preference for highly unsaturated FAs that contain greater than three double bonds, such as AA, EPA, and DHA, with negligible activity toward saturated FAs and monounsaturated FAs (22). Once the fatty acyl-CoA is generated, it can be used in various metabolic pathways, such as synthesis of phospholipids (PLs), triglycerides, cholesterol esters, as well as oxidative phosphorylation to generate ATP (25). Our laboratory previously demonstrated that ACSL4-deficient adipocytes from mice fed a high-fat diet had reduced AA incorporation by ∼30–50% into all PL classes within adipocytes and a reciprocal 25–89% increase in the incorporation of linoleic acid (LA; 18:2) in differing PL species (26). However, deficiency of ACSL4 in adipocytes did not alter the incorporation of any FA species into triglycerides. Furthermore, mouse ACSL4 has also been demonstrated to direct adrenic acid, as well as AA, into PLs, which is necessary for ferroptosis, a nonapoptotic form of programmed cell death (23).

Diet is the predominant source of all PUFAs. AAs can come from the diet directly or, alternatively, dietary LA can be converted into AA through several enzymatic steps within cells. Both exogenous and endogenous FAs can be incorporated into PL through the consecutive actions of ACSLs and acyltransferases. The FA composition of cellular PL is constantly being remodeled through a process of de-acylation by phospholipases and re-esterification by ACSLs and acyltransferases, termed the Lands’ cycle, which enriches PUFA at the sn-2 position of PL as a result of the specificity of the acyltransferases (27–29). ACSL4 has been demonstrated to be involved in the re-esterification of AA during constitutive and stimulated lipid remodeling (30, 31).

To elucidate the role of ACSL4 in PL remodeling and inflammation within macrophages, we isolated rpMACs from mice with or without myeloid-specific deficiency of Acsl4. Macrophage Acsl4 deficiency resulted in dramatic reduction in AAs and other specific PUFAs within PL and decreased production of proinflammatory AA-derived lipid mediators and cytokine expression. Furthermore, neutrophil migration and lipid mediator production were decreased in vivo during LPS-induced peritonitis in myeloid Acsl4-deficient mice, which demonstrated a physiological role of ACSL4 in mediating acute inflammation.

All animal breeding and procedures were approved by the Institutional Animal Care and Use Committee at Tufts University (protocol number H2019-145). Homozygous mice containing two LoxP sites flanking exons 3 and 4 of Acsl4 (floxed Acsl4) were generated at the North Carolina Animal Models Core as previously reported (26). The floxed Acsl4 mice were mated with heterozygous LysM-Cre mice from the Jackson Laboratory (B6.129P2-Lyz2^tm1(cre)Ifo/J), and the male progeny, Acsl4^FL/Y Lyz2^Cre/WT (Acsl4^mKO) or Acsl4^FL/Y Lyz2^WT/WT (Acsl4^Flox), were used for our studies. The mice were maintained on Teklad 2016 chow diet and were euthanized by CO₂ asphyxiation and cervical dislocation when 12–16 wk old, at which point there were no differences in body weights.

After euthanasia, the peritoneal cavity was exposed and 3 mL of PBS containing 0.5% BSA and 2 mM EDTA (MACS buffer) was injected into the peritoneal cavity and massaged for 30 s. The lavage fluid was removed with a 20-gauge needle and centrifuged at 300 × g for 10 min. The pellet was resuspended in MACS buffer, and the macrophage population was purified by negative selection through magnetic columns as per the manufacturer’s protocol (Mouse Macrophage Isolation Kit [Peritoneum]; Miltenyi Biotec), which resulted in a 99.5% pure population of macrophages as demonstrated by FACS analysis of CD11b⁺ F4/80⁺ cells (Supplemental Fig. 1B).

For the culture of rpMACs, cells were resuspended in DMEM, low glucose (11885; Thermo Fisher), at 1 × 10⁶ cells/ml and allowed to adhere to tissue-culture plastic for 45 min at 37°C. For lipid mediator analysis, 1 ml of the cell suspension per well of a 12-well plate was used, and for gene expression analysis, 0.5 ml of the suspension in each well of a 24-well plate was used. After adhesion, cells were washed once with PBS, new DMEM containing either opsonized zymosan (opZym) or LPS was added, and cells were incubated for indicated times.

Zymosan A (Z4250; Sigma-Aldrich) was resuspended at 20 mg/ml in 500 µL PBS by vigorous vortexing. The zymosan suspension was centrifuged at 1000 × g for 15 min and resuspended again at 20 mg/ml in 500 µL PBS. An equal volume of opsonizing reagent (Z2850; Thermo Fisher) was added, vortexed, and incubated at 37°C for 60 min. The opsonized zymosan (opZym) was then washed 2× with PBS and resuspended at 10 mg/ml in PBS. Particles of opZym were counted with a hemocytometer and used at 10 particles/cell. To assay phagocytosis in rpMACs, we incubated opsonized zymosan-FITC (Z2841; Thermo Fisher) with cells for 1 h. Fluorescence of nonphagocytosed zymosan-FITC particles was quenched with trypan blue (1 mg/ml) for 1 min at room temperature, and cells were then imaged with a Zeiss Axiovert 200 fluorescent microscope (32).

Isolated rpMACs were adhered to Corning 12-well plates for 45 min and were then given new DMEM containing 8.75 µM 1-¹⁴C-AA (0.5 µCi/ml) (NEC661050UC; PerkinElmer) bound 3:1, mol:mol, with FA-free BSA and incubated at 37°C in 5% CO₂ for 18 h (pulse). Cells were then washed with PBS containing 2% BSA three times to remove any free AAs, and fresh DMEM was added to the cells containing 0.5% BSA and incubated for 6 h. Medium was then collected, cells were lysed using Qiagen RLT buffer, and radioactivity from both was measured with a liquid scintillation counter.

Multidimensional mass spectrometry–based shotgun lipidomics analysis of lipids for macrophages samples was performed as described previously (33, 34). In brief, premixed internal standard was added to 10⁶ isolated macrophages for quantitation of lipid species and then normalized with the protein content (per milligram protein), which was performed following the instructions of the Pierce BCA protein assay kit (Cat #23225; Thermo Scientific). The lipids were extracted using a modified Bligh and Dyer procedure (35), and each lipid extract was reconstituted in chloroform/methanol (1:1, v/v) at a volume of 50 µL.

For shotgun lipidomics, lipid extracts were diluted to a final concentration of ∼500 fmol total lipids per microliter. Mass spectrometric analysis was performed on a triple-quadrupole mass spectrometer (TSQ Altis; Thermo Fisher Scientific, San Jose, CA) and a Q Exactive mass spectrometer (Thermo Scientific, San Jose, CA), both of which were equipped with an automated nanospray device (TriVersa NanoMate; Advion Bioscience, Ithaca, NY) as described previously (36). Identification and quantification of lipid species were performed using an automated software program (37, 38). Data processing (ion peak selection, baseline correction, data transfer, peak intensity comparison, and quantitation) was performed as described previously (38).

The rpMACs were isolated as described earlier and plated in six-well tissue culture plates in phenol red–free DMEM for 45 min. After adhering to the plate, macrophages were washed with PBS 2×, and media were replaced with 2 ml PBS (with 0.9 mM CaCl₂ and 0.49 mM MgCl₂-6H₂O) with or without opsonized zymosan (10 particles/cell) for 2 h at 37°C. The cells and medium were then combined with 2 vol of ice-cold methanol containing commercially available deuterium-labeled internal standards (d₄-LTB₄, d₈-5-HETE, d₄-PGE₂, and d₅-LXA₄; Cayman Chemical) for calculation of extraction efficiency. After centrifugation (13,000 rpm), supernatants were carefully separated from the resulting pelleted material and subjected to solid-phase extraction (SPE) (39). The pellets were stored at −80°C and later used for protein determination via Pierce BCA protein assay (Thermo). SPE and LC-MS/MS analysis was conducted as described in part previously (39). In brief, the pH of the samples was reduced via the addition of acidified water (pH 3.5). Acidified samples were then added to conditioned C18 SPE columns (Biotage). After a column wash with n-hexanes, lipid mediators were liberated from the column and collected via the addition of methyl formate. Samples were concentrated by evaporating the solvent using N₂ gas and then resuspended in a solution of equal parts methanol and water (50:50). To ensure the elimination of any precipitate, samples were then centrifuged and transferred to clean glass inserts and amber autosampler vials for liquid chromatography tandem mass spectrometry (LC-MS/MS) analysis. Using a high-performance liquid chromatograph (Shimadzu) coupled to a QTrap5500 mass spectrometer (AB Sciex) operating in negative ionization mode, we identified lipid mediators using scheduled multiple reaction monitoring transitions and by matching their retention time with synthetic standards run in parallel. The multiple reaction monitoring transitions used included: m/z 351/189 (PGE₂, PGD₂), 353/193 (PGF_2α), 369/169 (TXB₂), 335/195 (LTB₄ and isomers), 351/115 (LXA₄, 15epi-LXA₄), 351/221 (LXB₄), 303/259 (AA), 335/115 (5, 15-diHETE), 319/219 (15-HETE), 319/179 (12-HETE), and 319/115 (5-HETE). Lipid mediators were quantified by accounting for the extraction recovery of internal deuterated standards and by calibration curves of external standards for each individual mediator.

ACSL activity was measured as previously described (26, 40); in brief, rpMACs from three mice were pooled, and the cells were sheared with 30 passes through a 26-gauge needle in sucrose buffer (10 mM Tris [pH 7.4], 1 mM EDTA, 0.25 M sucrose). Lysates were centrifuged at 100,000 × g, and the pellet containing the membrane fraction was resuspended in sucrose buffer. Protein was quantitated with Pierce BCA Protein Assay Kit (Thermo Fisher). Membrane protein fractions (5 µg) were added to 250 µL assay buffer (10 mM ATP, 250 µM CoA, 5 mM DTT, 8 mM MgCl₂*6H₂O, 175 mM Tris [pH 7.4], and 0.03% Triton X-100) containing 10 µM 1-¹⁴C-AA (PerkinElmer) and incubated at room temperature for 20 min. The reaction was stopped with the addition of 2.25 ml modified Dole’s reagent (80:20:1, isopropanol:heptane:1 M H₂SO₄), incubated at room temp for 10 min, followed by the addition of 2 ml Heptane and 0.5 ml H₂O. After vortexing, phases separated for 15 min, and the aqueous phase was measured with a liquid scintillation counter.

Mice were injected i.p. with 10 µg/kg LPS and euthanized 4 h later by cardiac puncture to collect blood while anesthetized with 2.5% isoflurane, followed by cervical dislocation. The mouse peritoneum was exposed and injected with 3 ml PBS containing 0.5% BSA and 2 mM EDTA. The peritoneal lavage fluid (PLF) was collected and centrifuged for 10 min at 400 × g to pellet cells. Cells were then stained with fluorochrome-conjugated REAfinity Abs (2 µl/10⁶ cells) against mouse CD11b (VioGreen), F4/80 (PE), and Ly6G (allophycocyanin) (Miltenyi Biotec) for 15 min at 4°C. Cells were washed and centrifuged before FACS analysis on a BD LSR II. The gating strategy was determined by using florescence-minus-one staining (Supplemental Figure 1A). Analysis of the FACS data was performed with FlowJo software.

PLF from mice after 4-h LPS-induced peritonitis was collected and centrifuged for 10 min at 400 × g to remove cells. The PLF was acidified by addition of an equal volume of 1 M acetate buffer (pH 4.0). Purification of the lipid mediators was accomplished using SPE (C18) columns (400020; Cayman Chemical) and eluted with ethyl acetate containing 1% methanol. The ethyl acetate was evaporated under nitrogen gas, and the concentrations of LTB₄ and PGE₂ were measured by ELISA according to the manufacturer’s protocols (LTB₄, 520111; PGE₂, 514010; Cayman Chemical). No purification of lipid mediators was used for measurement of LTB₄ and PGE₂ in culture medium from rpMACs treated with E. coli O55:B5-derived LPS (1 μg/ml) (L6529; Sigma Aldrich) for 6 h. The ELISAs were performed according to the manufacturer’s protocols as described earlier.

Isolated rpMACs were lysed in RLT buffer (Qiagen) containing 1% 2-ME. RNA was isolated using the Qiagen RNeasy Mini Kit according to the manufacturer’s protocol. The RNA concentration was quantified using a NanoDrop spectrophotometer (NanoDrop 1000; Wilmington, DE), and 500 ng of RNA was used for the generation of cDNA with the Thermo Fisher High-Capacity cDNA Reverse Transcription kit. cDNA was further diluted 10× in nuclease-free water. Quantitative real-time PCR was performed using SybrGreen (Thermo Fisher) with the Applied Biosystems 7300 Real-Time PCR System. All primers were used at a concentration of 1 µM (primer sequences are listed in Table I). Gene expression was calculated using the 2^−ΔΔCt method and normalizing to Gapdh.

We generated a mouse line with a myeloid-specific deficiency of Acsl4 (Acsl4^mKO) by transferring the LysM-Cre transgene into our line of Acsl4 floxed mice. We used the littermate floxed males without Cre as controls (Acsl4^Flox). Acsl4 is located on the X chromosome, and therefore males have only one copy of the gene. First, we analyzed Acsl mRNA expression in isolated rpMACs from male mice between the ages of 12 and 16 wk. RT-PCR analysis revealed that the rpMACs from Acsl4^mKO mice had an 80–90% reduction of Acsl4 mRNA as compared with the control Acsl4^Flox mice (Fig. 1A). Furthermore, we observed no significant differences in mRNA expression between groups for other Acsl isoforms (1, 3, 5, or 6), demonstrating that our approach to reduce Acsl4 expression in rpMACs was specific and did not result in any compensatory increases in other ACSL isoforms.

We next measured total membrane ACSL activity toward AA in rpMACs. We observed that Acsl4^mKO rpMACs exhibited a 56.5% decrease in ACSL activity that catalyzes the addition of a CoA moiety to AA, confirming a role for ACSL4 in regulating arachidonyl-CoA (AA-CoA) (Fig. 1B). To determine whether expression of Acsl4 regulated the rate of AA retention within the rpMACs, we performed pulse-chase experiments to quantitate the turnover of radiolabeled AA within cells. For these studies, cells were incubated with 1-¹⁴C-labeled AA bound to BSA for 18 h. The cells were then washed and chased with medium containing 0.5% BSA without labeled AA for 6 h. The rate of release of radiolabel from Acsl4^mKO rpMACs into the medium was 1.88-fold higher than that of Acsl4^Flox cells (Fig. 1C). There was also a concurrent 33% reduction in the radiolabel measured within the Acsl4^mKO rpMACs after the 6-h chase (Fig. 1D). The radiolabel released in the medium likely consists of both oxidized AA metabolites and free AA bound to albumin in the medium, but those individual molecular species are not distinguished with the pulse-chase assay. Consistent with our pulse-chase data, previous studies have also demonstrated that the generation of AA-CoA by ACSLs traps the FA within cells, resulting in increased uptake of AA into cells (41–43).

In separate studies, we harvested rpMACs from the two lines of mice and quantitated the level of free AAs by LC-MS/MS. We found that levels of free AAs were significantly reduced in the Acsl4^mKO rpMACs as compared with the floxed controls (Fig. 1E). To determine whether the reduced levels of AA within Acsl4^mKO rpMACs were due to a reduction in the expression of enzymes involved in the synthesis of AA from LA, we quantitated mRNA expression of the relevant enzymes by RT-PCR analysis. Surprisingly, expression of the FA Δ6-desaturase (Fads2), the initial and rate-limiting enzyme in the synthesis of AA from LA, was increased 77-fold in Acsl4^mKO rpMACs as compared with the floxed controls (Fig. 1F). Also, mRNA expression of the elongation of very long-chain FAs 5 (Elovl5) enzyme was upregulated 2.7-fold, although the difference was not significant (p = 0.056). Fads1, the Δ5-desaturase, and last enzyme involved in the synthesis of AA, was slightly, but significantly, upregulated in Acsl4^mKO rpMACs (Fig. 1F). In summary, our results demonstrate that Acsl4 deficiency increases the rate that radiolabeled-AA is released from cells, which is likely due to the reduced ability of these cells to generate AA-CoA that cannot diffuse out of the cell. Furthermore, these data demonstrate that the reduced levels of AA within cells are not secondary to decreased synthesis of AA from LA. In fact, the deficiency of Acsl4 led to increased expression of the enzymes involved in AA synthesis, which is likely an attempt to compensate for the reduction of AAs within the cells.

Our work and that of others have shown that deficiency of Acsl4 results in decreased incorporation of AAs within PL (23, 26). To evaluate the effects of Acsl4 deficiency on PL remodeling, we used rpMACs and performed shotgun lipidomic analysis with MS/MS to determine the profile of FA content of PL species. Our studies demonstrated that deficiency of Acsl4 in rpMACs potently reduced the levels of AA between 75–90% in all PL classes (Fig. 2, Supplemental Fig. 2), with a 98% reduction in phosphatidic acid (PA-18:0-20:4) (Fig. 2C). DPA (22:5n-3 or n-6) and the ω-3 PUFA, DHA (22:6n-3), were also significantly reduced in all PLs, although ω-3 PUFA was only a minor fraction of the total PUFAs in the cells. We did not detect EPA (20:5n-3) in any PLs from rpMACs of either line of mice. Conversely, LA (18:2) and oleic acid (OA; 18:1) were significantly increased in all PL classes, with the largest increases found in phosphatidylethanolamine (Fig. 2A, 2B, Supplemental Fig. 2). Deficiency of Acsl4 also resulted in decreased diacylpalmitoyl phosphatidylcholine (PC-16:0-16:0) and, to a lesser extent, 16:0-16:0-PA, without significant changes to 16:0-16:0 in other PL classes. These studies demonstrate that Acsl4 is the predominant Acsl isoform in macrophages that facilitates the incorporation of AA into PL.

In our studies investigating the effects of Acsl4 deficiency on the PL profile in rpMACs, our most robust observation was a 75–90% reduction in the incorporation of AAs into PLs. Given that AAs within PLs serve as the primary source for the generation of lipid mediators after an inflammatory stimulation, we assessed the production of eicosanoids. To determine how reductions in AA content in PL, caused by Acsl4 deficiency, altered the generation of lipid mediators, we incubated rpMACs in the absence and presence of opZym, as an inflammatory stimulus. Of relevance to our studies, others have demonstrated that zymosan increases production of lipid mediators generated through the cyclooxygenase-1/2, 5-LOX, and 12/15-LOX pathways to a much greater extent as compared with LPS in rpMACs (44). Indeed, after a 2-h incubation with opZym (10 particles/cell), several eicosanoids were increased, including the PGs (PGE₂, PGD₂, PGF_2α), leukotrienes (LTB₄ and trans isomers), LXs (LXA₄, 15epi-LXA₄, LXB₄), and HETEs (5-HETE, 12-HETE, 15-HETE, 5,15-diHETE) (Fig. 3). Remarkably, all eicosanoids in our targeted analysis were significantly reduced in Acsl4^mKO rpMACs as compared with Acsl4^Flox rpMACs (Fig. 3). Importantly, there was no difference in the uptake of opZym between Acsl4^Flox and Acsl4^mKO rpMACs that would explain the reductions in eicosanoids (Supplemental Fig. 3).

We next investigated whether mRNA expression of various enzymes involved in the biosynthesis of eicosanoids was altered. RT-PCR analysis revealed no differences in 5-lipoxygenase activating protein (Alox5ap), leukotriene A₄ hydrolase (Lta₄h), Alox5, or Ptgs2 mRNA (Fig. 4A). We did observe a significant 2.5-fold increase in Alox15 (Fig. 4A), although this did not overcome the suppression of eicosanoid biosynthesis in Acsl4^mKO rpMACs. Collectively, these results suggest that deficiency of Acsl4 in rpMACs impairs biosynthesis of eicosanoids by reducing the bioavailability of AA.

Given the decreased production of both inflammatory eicosanoids (leukotrienes, PGs, and HETEs) and some proresolving mediators (LXs) in Acsl4^mKO rpMACs after opZym treatment, we investigated whether deficiency of Acsl4 alters cytokine expression in vitro. Expressions of Il6, Nos2, Ccl5, and Ccl2 were significantly increased in rpMACs stimulated with zymosan at 6 h, while they all returned to baseline by 24 h poststimulation (Fig. 4B). We observed significant reductions in Il6, Nos2, Ccl5, and Ccl2 mRNA expression at 6 h in the Acsl4^mKO rpMACs as compared with the Acsl4^Flox controls. The increase in Nos2 mRNA levels was completely blunted in the Acsl4^mKO rpMACs. Also, Nos2 mRNA levels were significantly decreased at 24 h poststimulation in the Acsl4^mKO as compared with Acsl4^Flox rpMACs, but no differences were observed in expression of any of the other genes at 24 h (Fig. 4B). In summary, these data demonstrate that Acsl4 deficiency reduced inflammatory cytokine expression in response to acute incubation with opZym.

We next extended our studies to investigate the TLR4 signaling pathway through LPS activation to determine whether we could observe similar results as demonstrated with opZym and because rpMACs are the primary cell type that contributes to peritoneal neutrophil (PMN) recruitment during LPS-induced peritonitis (4). Ex vivo, Acsl4^mKO rpMACs treated with LPS generated significantly lower levels of LTB₄ and PGE₂ (Fig. 5A), although LPS did not stimulate LTB₄ production to the same extent as opZym. Given the reduction of the potent neutrophil chemotactic signaling molecule, LTB₄, in Acsl4^mKO rpMACs treated with LPS or opZym, we investigated how neutrophil recruitment into the peritoneum after i.p. injection of LPS would be affected by Acsl4 deficiency. For these studies, mice were injected i.p. with LPS (10 μg/kg); 4 h later, peritoneal fluid was harvested to measure levels of LTB₄ and PGE₂ by ELISA, and FACS analysis was performed to quantitate the number of neutrophils. Both LTB₄ and PGE₂ levels were significantly reduced in peritoneal fluid of Acsl4^mKO mice as compared with the Acsl4^Flox controls after i.p. LPS injections (Fig. 5B). Significantly, the total number of neutrophils (CD11b⁺ Ly6G⁺ F4/80⁻) in peritoneal fluid after LPS injection was reduced by 80% in Acsl4^mKO mice (Fig. 5C). The total number of peritoneal macrophages (CD11b⁺ F4/80⁺ Ly6G⁻) was 55% higher in the Acsl4^mKO mice after LPS stimulation, although it did not reach statistical significance. In the absence of LPS, we observed negligible numbers of neutrophils in the Acsl4^Flox mice (average of 4.6 neutrophils per 10⁵ cells). In summary, these studies demonstrated that, in response to LPS, Acsl4-deficient rpMACs produced reduced levels of LTB₄ and PGE₂ in vitro and in vivo and reduced recruitment of neutrophils in vivo.

Macrophages are immune cells that respond to infection and various other insults in part by generating proinflammatory lipid mediators and cytokines. Pathogen-associated molecular patterns, such as yeast and bacterial cell wall components, can bind TLRs on macrophages and activate phospholipase A₂, which then cleaves PUFAs, including AA, out of PL. The AA can be subsequently oxidized by cyclooxygenases or lipoxygenases into the PGs, leukotrienes, thromboxanes, and HETEs, as well as other proinflammatory lipid mediators. For AA and other FAs to be incorporated into PL, they must first be activated by ACSL enzymes that catalyze the thioesterification of a CoA to a FA, which generates a fatty-acyl-CoA that is trapped within the cell and can be incorporated into PL by members of the lysophosphatidylcholine acyltransferase (Lpcat) family (29). ACSL4 has been established as having a strong preference for AA and other PUFAs with three or more double bonds to generate PUFA-CoA, and in particular AA-CoA. Our current findings involving both in vitro and in vivo studies demonstrate that Acsl4 deficiency reduces the rate of AA-CoA formation and dramatically reduces AA incorporation into PL in rpMACs.

We previously generated floxed Acsl4 mice with adipocyte-specific deficiency of Acsl4 to elucidate the in vivo role of ACSL4. Those studies revealed that adipocytes from mice fed a high-fat diet had a 30–50% reduction in the levels of AA incorporated into PL (26). Acsl4 deficiency did not completely reduce the incorporation of AA into adipocyte PL, suggesting that other members of the ACSL family, in addition to ACSL4, may participate in the generation of AA-CoA necessary for incorporation into PL. In contrast to those observations in adipocytes, we now show that Acsl4 deficiency in rpMACs results in a robust 75–90% reduction in the levels of AA incorporated into all classes of PL. Furthermore, we observed a 98% reduction of AA within PA, which is the precursor for all other PLs (45), indicating that Acsl4 is necessary for the de novo generation of PL containing AA through the Kennedy pathway. Thus, our studies demonstrated that ACSL4 is the predominant ACSL isoform regulating the incorporation of AA into macrophage PL under homeostatic conditions. Consistent with this, we demonstrated with our pulse-chase experiment that the total retention of radiolabeled AA in cellular extracts was reduced, whereas the radiolabel present in the cells during the pulse was released into the chase medium from Acsl4-deficient rpMACs at nearly twice the rate of the floxed controls. We note, however, that the radiolabel measurements cannot solely be attributed to native AA and could include further oxidized products of AA.

In addition to the reduced levels of AA incorporated into PL, we also found reduced free AA in the basal state of the rpMACs. One possible explanation for a reduction of total AA in the cells could have been that there was decreased expression of the enzymes involved in the generation of AA from LA. However, interestingly, our studies revealed a robust increase in the mRNA expression of the enzymes involved in the conversion of LA into AA, perhaps indicating a physiological attempt by the cells to increase cellular AA levels. We therefore conclude that the most likely explanation for the reduced AA content in Acsl4-deficient rpMACs is that Acsl4 deficiency results in a decreased ability to form AA-CoA, and therefore free AA is not activated or trapped within the cell.

Along with the decrease of AA, we observed a reciprocal increase of OA and LA in the PL of Acsl4-deficient rpMACs. The increase in those unsaturated FA may be of relevance because previous studies have shown that reduced incorporation of both AA and LA in PL in mice with Lpcat3 deficiency resulted in decreased membrane fluidity and diminished diffusion of FA into enterocytes (46). It is likely that the increase in OA and LA in Acsl4-deficient rpMACs may act to maintain membrane fluidity, although the mechanism for how this occurs is not currently understood and is a focus of investigation.

In our studies, Acsl4 deficiency in isolated rpMACs resulted in dramatic reductions in nearly all AA-derived lipid mediators produced in response to inflammatory stimulation by opZym, which is consistent with the reductions of AA in PL. The decreased levels of eicosanoids included both proinflammatory (PGs, leukotrienes) and proresolving lipid mediators (LXs). Of significance, in studies from the laboratories of Peters-Golden (47) and Olefsky (48), LTB₄ has been previously demonstrated to regulate the expression of Il6, Ccl5, Nos2, and Ccl2 in response to various TLR ligands in part by maintaining MyD88 expression and NF-κB activation in macrophages. The reduction of LTB₄ that we observed in Acsl4-deficient rpMACs after opZym stimulation thus may be one factor that contributes to the observed reductions in mRNA expression of Il6, Ccl5, Nos2, and Ccl2. Nonetheless, it is also possible that the resulting alterations in membrane PL remodeling with Acsl4-deficient rpMACs may dampen intracellular signaling pathways in response to TLR ligands and/or eicosanoid G protein–coupled receptors. The reduction in 12-HETE may also contribute to the reduced mRNA levels of Ccl2 (49). In addition, LTB₄ is a potent neutrophil chemoattractant, and the reduction of LTB₄ in Acsl4^mKO mice after LPS stimulation likely underlies the reduced neutrophil recruitment in vivo.

The LXs are SPMs derived from AAs that can act to resolve inflammation and stimulate tissue regeneration (50). We found that the AA-derived LXs, including LXA₄, 15epi-LXA₄, and LXB₄, were significantly reduced in Acsl4^mKO rpMACs stimulated with zymosan. These mediators are potent regulators of inflammation resolution and counterregulate proinflammatory actions of LTB₄ and PGE₂ (17, 51). The results demonstrate that, by acting to regulate substrate availability in membrane PL, ACSL4 regulates downstream production of both proinflammatory and proresolving lipid mediators derived from AA.

Our research focus was to investigate the effects of ACSL4 on rpMACs freshly isolated from the animal and in vivo. In contrast, a previous study by Kuwata et al. (22) investigating the function of ACSL4 used bone marrow–derived macrophages (BMDMs) that were cultured and differentiated in vitro from mice with whole-body knockout of Acsl4. In those studies, the Acsl4 knockout BMDMs were cultured in the presence of serum that could contain AA and other FAs, which contrasts with our studies done in serum-free media. Lipidomic analysis of these Acsl4-deficient BMDMs demonstrated reductions in incorporation of AA and other PUFAs into PL that were similar to our results, but to a lesser extent. They also noted a small reciprocal increase in OA and LA that were not as robust as what we observed. In direct opposition to our studies, when the Acsl4 knockout BMDMs were stimulated with LPS, they had significant increases in PGs, thromboxane, and HETEs, which occurred in the presence of serum. Importantly, our in vivo studies demonstrated a reduction of PGE₂ and LTB₄ in the PLF from Acsl4^mKO mice after LPS injection that corroborated our ex vivo results that found similar decreases in PGE₂ and LTB₄ in the LPS-treated Acsl4^mKO rpMACs. We hypothesize that the observed increase in AA-derived lipid mediators measured in the study by Kuwata et al. in BMDMs could be a result of the utilization of unesterified AA in serum. Conversely, long-term knockdown of Acsl4 in smooth muscle cells, cultured in the presence of serum, was shown to reduce PGE₂ to a similar extent as we observed after acute stimulation of rpMACs with opZym in serum-free medium (30), indicating that the effects of Acsl4 deficiency are time and cell type dependent.

In the BMDM study by Kuwata et al. (22), the authors did not find differences in expression of Nos2 or Ccl2 after LPS treatment, whereas we saw significantly decreased expression of these proinflammatory genes in the Acsl4-deficient rpMACs after opZym treatment. BMDMs, as compared with rpMACs, are known to have diminished 5-LOX activity necessary to produce LTB₄, especially with LPS as an inflammatory stimulus (44). As discussed, LTB₄ can increase expression of Nos2 and Ccl2, and we measured high levels of LTB₄ after opZym stimulation in the control Acsl4^Flox rpMACs compared with the Acsl4^mKO rpMACs, which might explain why we saw differences in expression of those cytokines and the other researchers did not.

In summary, our studies have shown that ACSL4 is critical for maintaining normal PUFA content in PL within peritoneal macrophages through incorporation of AA into PA, a precursor for generation of diverse PL, as well as esterification of PUFA containing more than three double bonds during homeostatic PL remodeling and generation of AA metabolites in response to TLR activation. Future directions aimed at investigating membrane fluidity caused by the drastic alterations in PUFA content within PL of Acsl4-deficient macrophages will be important to explore, as well as the effects of these alterations on mobility, energy metabolism, and various signaling pathways. Lastly, the reductions in AA-derived lipid mediators and cytokines in Acsl4-deficient rpMACs after an inflammatory stimulus suggest that ACSL4 may be a good target for drug therapies aimed at treating diseases associated with chronic inflammation.

The authors have no financial conflicts of interest.