Cell-intrinsic lysosomal lipolysis is essential for macrophage alternative activation

M2 macrophage activation is dependent on fatty acid oxidation

Using extracellular flux analysis, we compared oxygen consumption by M0 (unactivated), M1 and M2 bone marrow-derived macrophages. We found that M2 macrophages had enhanced mitochondrial oxygen consumption rates (OCR), and markedly increased spare respiratory capacity (SRC^10,18 the quantitative difference between maximal uncontrolled OCR, and the initial basal OCR), indicative of increased commitment to OXPHOS (Fig. 1a). In contrast, we found no evidence of mitochondrial oxygen consumption in M1 macrophages (Fig. 1a), which rather rely on aerobic glycolysis, measured as the extracellular acidification rate, ECAR (Fig. 1b), to meet their bioenergetic needs¹⁹. The extent of the metabolic difference between M1 and M2 cells was apparent in the overall ratio of OXPHOS to aerobic glycolysis, which was 10-fold higher in M2 than M1 macrophages, reflecting polar opposite core metabolic programs (Fig. 1c). Consistent with previous reports, we found that commitment of M2 macrophages to FAO and OXPHOS was evident in the increased expression of genes in these pathways (Supplementary Fig. 1a,c)^8,20. This pathway was also sensitive to the suppression of mitochondrial oxygen consumption by etomoxir, an inhibitor of Cpt1, a key FAO enzyme (Fig. 1d)⁸. We found that etomoxir also eradicated SRC in M2 macrophages (Fig. 1e), validating the link between FAO and heightened OXPHOS. Enhanced SRC was previously linked to longevity in memory CD8⁺ T cells¹⁰. We found this increased longevity to also be true in macrophages, where M2 cells exhibited a significant survival advantage compared to M0 and M1 cells in culture (Supplementary Fig. 2).

M2 and M1 macrophage activation are marked by the differential expression of a large panel of genes, including Mrc1 (CD206), Clec10a (CD301), Retlna (RELMα), Pdcd1lg2

(PD-L2) and Nos2 (iNOS) (Supplementary Fig. 1b,c). Using flow cytometry, we found that etomoxir remarkedly inhibited the expression of CD301, CD206 and RELMα in macrophages cultured in M2 conditions (Fig. 1f,g), but had comparatively little effect on the expression of iNOS in cells cultured in M1 conditions (Fig. 1h). Thus, FAO, measureable as increased baseline OCR and increased SRC, is essential for M2 activation.

Lipolysis contributes to M2 activation

Cellular requirements for fatty acids can be met by enzymatically regulated lipolysis of triacylglycerols into diacylglycerols and monoacylglycerols, accompanied by fatty acid release¹¹; an endpoint of this process is the release of glycerol from cells in which lipolysis is occurring¹¹. We performed global metabolite profiling using mass spectrometry and found significantly increased amounts of 3 out of 4 monoacylglycerols measured in M2 cells, indicating that lipolysis is enhanced in these cells (Fig. 2a). Consistent with this conclusion, we found increased concentrations of extracellular glycerol were present in cultures of M2 compared with M0 macrophages (Fig. 2b). To test whether lipolysis is linked to FAO and OXPHOS, we stimulated cells with IL-4 in the presence or absence of tetrahydrolipistatin (Orlistat), a clinically utilized active site-directed lipase inhibitor²¹. We found that Orlistat inhibited IL-4-induced increases in lipolysis, as measured by glycerol production (Fig. 2b), and suppressed associated changes in OXPHOS (Fig. 2c), although it had no effect on the metabolic changes induced by M1 conditions (Fig. 2d). These reductions in OXPHOS and SRC in Orlistat-treated M2 cells were linked to significantly reduced survival, evident as in decreased viability by day 2 of culture (Fig. 2e). Orlistat had less marked effects on the survival of M0 cells (Fig. 2e). Within the short timeframe that M1 cells remained alive, we could detect no significant effect of inhibiting lipolysis on their survival (data not shown). Given the close link between FAO and M2 activation, we next tested the effects of inhibiting lipolysis on macrophages stimulated with IL-4. We found that Orlistat suppressed the IL-4 induced expression of CD206, CD301, PD-L2 and RELMα(Fig. 2f), but had no effect on iNOS expression in M1 macrophages (Fig. 2g). Together, these data indicate that lipolysis supports FAO and in this way is critical for M2 activation.

Lysosomal acid lipase is responsible for lipolysis upstream of FAO

Our data implicated lipolysis as a critical mechanism for the generation of fatty acids for FAO and M2 activation in macrophages stimulated with IL-4. In other cells fatty acids for FAO are released from triacylglycerols stored with cholesterol esters in LDs by a process of lipolysis initiated by ATGL and continued by hormone-sensitive lipase (HSL), encoded by Pnpla2 and Lipe, respectively^11,12,22. However, amongst genes encoding lipases, we found that Pnpla2 and Lipe were expressed more strongly in M1 cells than in M2 cells (Fig. 3a), suggesting that these genes might not play a role in M2 activation. To directly test this, we measured RELMα expression in peritoneal macrophages (PM) from Pnpla2^–/– mice following the i.p. injection of IL-4–anti-IL-4 complexes (IL-4c), a procedure shown previously to induce peritoneal macrophage M2 activation²³. We found that >90% of macrophages in Pnpla2^–/– mice or in wild-type mice expressed RELMα following IL-4c injection, indicating that the loss of ATGL has no effect on M2 activation (Supplementary Fig. 3a). A substantial body of literature has associated M1 activation with the development of lipid droplets²⁴, but whether or not lipid droplets are prevalent in M2 macrophages is unclear. Using transmission electron microscopy (TEM) we found that M1 macrophages contained easily identifiable lipid droplets, whereas M2 cells did not (Fig. 3b). Together with the lipase expression data and data from experiments using Pnpla2^–/– macrophages, these findings suggest that M2 macrophages are not utilizing canonical lipid droplet-triacylglycerol hydrolysis pathways to generate fatty acids for FAO. Nevertheless, we did observe that neutral lipids as measured by flow cytometric detection of BODIPY stainin, the appearance of lipid droplets as visualized by electron microscopy, and the quantitation of triacylglycerols and cholesterol esters by mass spectrometry, accumulated in macrophages stimulated with IL-4 in the presence of Orlistat (Supplementary Fig. 3b-d). Mass spectrometry additionally confirmed that neutral lipid stores are greater in M1 macrophages than in M2 macrophages (Supplementary Fig. 3d). These findings indicate that M2 macrophages accumulate neutral lipid stores when lipolysis is inhibited.

We noted that, unlike genes encoding many other cellular lipases (including Pnpla2, Lipe, Lipc, Lipg), expression of Lipa was upregulated in M2 compared to M1 macrophages (Fig. 3a, Supplementary Fig. 1c), suggesting that it could be important for M2 activation. Lipa encodes LAL, a triacylglycerol lipase that is able to breakdown triacylglycerols delivered into the lysosomes by endocytosis of low density lipoproteins (LDLs and VLDLs)²⁵. Since LAL is a known target of Orlistat²⁶, we reasoned that the observed effects of Orlistat on macrophage FAO and M2 activation (Fig. 2) could reflect the inhibition of LAL function in this system. To investigate the role of LAL, which unlike the other cellular lipases has optimum activity at acidic rather than neutral pH, in M2 activation, we first neutralized lysosomal pH using chloroquine or Bafilomycin A1. We found that these conditions completely inhibited the IL-4–induced increases in OXPHOS, as measured by OCR, OCR/ECAR ratio and SRC (Fig. 3c). These effects were accompanied by the inhibition of IL-4–induced increases in CD206, CD301, PD-L2 and RELMα expression (Fig. 3d) and a decrease in longevity (Fig. 3e). These data support the view that an acidic compartment is essential for M2 activation and are consistent with a role for LAL in this process. To formally address this possibility, we targeted LAL using shRNAs, and found that increased OXPHOS, expression of CD206, CD301, RELMα and PD-L2, and increased longevity in response to IL-4 were significantly inhibited by the suppression of LAL expression (Fig. 3f-h). Suppression of LAL expression also resulted in the accumulation of lipid droplets in macrophages stimulated with IL-4 (Supplementary Fig. 3b). Additionally, we examined the ability of Lipa^–/– macrophages to become M2 activated and found that increases in expression of CD206, CD301 and RELMα were significantly muted in Lipa^–/– macrophages versus wild-type cells (Fig. 3i). Finally, we made bone marrow chimeras in which irradiated CD45.2 mice were reconstituted with CD45.1 bone marrow that had been bone transduced with retrovirus encoding shRNAs targeting Lipa or a control gene (luciferase). Progeny from transduced cells could be distinguished via expression of human CD8 encoded as a reporter by the retrovirus. We examined M2 activation of peritoneal macrophages derived from CD45.1⁺ transduced versus untransduced bone marrow cells following i.p. injection of IL-4c. We found that peritoneal macrophage RELMα expression in response to IL-4 was inhibited by the suppression of LAL expression (Fig. 3j). Together, these data indicate that cell-intrinsic lipolysis mediated by LAL plays a critical role in optimal macrophage M2 activation in response to IL-4.

Elevated OXPHOS and optimal M2 activation are dependent on CD36

The important role for LAL in M2 macrophages suggested that triacylglycerol delivery to lysosomes is a key process in the generation of fatty acids to support FAO and M2 activation. We examined the expression of receptors that mediate the uptake of triacylglycerol-carrying serum lipoproteins in M2 macrophages and observed that CD36 was upregulated in M2 cells (Fig. 4a)¹⁵. The same was true for the related scavenger receptor Scarb1, but not for Ldlr1, Vldlr, or Lrp8 (Fig. 4a, Supplementary Fig. 1c). Based on these observations, and the fact that CD36 has been previously implicated in M2 activation¹⁶, we hypothesized that CD36 may play a role in increased FAO in M2 macrophages by facilitating the uptake of triacylglycerols. To examine this, we assessed responses to IL-4 of macrophages made from the bone marrow of Cd36^–/– mice. We found that compared to M0 macrophages, wild-type M2 macrophages, but not Cd36^–/– M2 macrophages, took up more LDL (Fig. 4b) and VLDL (Fig. 4c). In addition to being a receptor for LDL and VLDL, CD36 is a fatty acid translocase^13,14. While we found M2 macrophages to be more capable than M0 cells of taking up palmitate, we observed no difference between wild-type and Cd36^–/– cells in this regard (Supplementary Fig. 4a), suggesting that the fatty acid translocase activity of CD36 is not accounting for increased free fatty acid uptake by M2 cells. Compared to wild-type macrophages, Cd36^–/– macrophages exhibited significantly lower IL-4 induced increases in OXPHOS as measured by OCR, the OCR/ECAR ratio and SRC (Fig. 4d), but were substantially similar in terms of the metabolic markers of M1 activation (Fig. 4e). Consistent with the critical role for FAO in M2 activation, Cd36^–/– macrophages exhibited substantially reduced levels of expression of PD-L2 and RELMαin response to IL-4 (Fig. 4f). In comparison, iNOS expression was similar in wild-type and Cd36^–/– macrophages stimulated in M1 conditions (Fig. 4g). We further addressed the role of CD36 by examining the effect on wild-type macrophage responses to IL-4 of inhibiting CD36 with sulfo-N-succinimidyl oleate (SSO)²⁷. The findings from these experiments confirmed those from using Cd36^–/– macrophages. We found that LDL uptake was substantially inhibited by SSO (Supplementary Fig. 4b). Moreover, SSO completely blocked IL-4-induced increases in OXPHOS as measured by OCR, the OCR/ECAR ratio and SRC (Supplementary Fig. 4c) but had no effect on metabolic markers of M1 activation (Supplementary Fig. 4d). SSO substantially inhibited expression of CD206, CD301, PD-L2 and RELMαin response to IL-4 (Supplementary Fig. 4e), but had no effect on iNOS expression in M1 macrophages (Supplementary Fig. 4f). Together, these data support the view that elevated CD36 expression in macrophages in response to IL-4 plays an important role in increased uptake of LDL and VLDL, and that this pathway supports FAO and M2 activation.

Next we tested the ability of monocyte-derived macrophages from a CD36-deficient patient to become alternatively activated in response to IL-4. We found that SRC was substantially diminished in human CD36-deficient M2 macrophages compared to CD36-sufficient human M2 macrophages (Fig. 4h). Expression of CD206, the mannose receptor C type 1 (MRC1), is a marker of human M2 activation, as is expression of CD36²⁸. Thus we used flow cytometry to examine CD36 and CD206 expression in M0, M1 and M2 macrophages from CD36-sufficient and -deficient subjects. We found that CD36 and CD206 expression were elevated in M2 versus M0 and M1 macrophages, and that expression of CD206 was substantially lower in CD36-deficient macrophages stimulated with IL-4 compared to CD36-sufficient macrophages (Fig. 4i). Together with our findings from mouse Cd36^–/– macrophages and the SSO inhibitor studies, these data from human CD36-deficient macrophages strongly support a significant role for CD36 in M2 activation.

Our data suggested that the lysosomal lipolysis of exogenous triacylglycerols acquired through a CD36-dependent mechanism is important for optimal M2 activation. In this case, we reasoned that depriving macrophages of exogenous triacylglycerols should impact M2 activation. We cultured macrophages in complete medium containing or lacking serum for 24 h and then measured the expression of key M2 marker genes following an additional 24 h of stimulation with IL-4. In these experiments, M2 activation was substantially inhibited, but not prevented, by the absence of serum (Supplementary Fig. 5a). Consistent with our data implicating CD36-mediated uptake of LDL and VLDL in M2 macrophages, we found that under serum-free conditions, M2 activation could be promoted by the addition of LDL and VLDL (Supplementary Fig. 5a). The finding that M2 activation could proceed to some extent in the absence of serum, suggested that macrophages have an alternative pathway to fuel FAO. A candidate for such an alternative pathway is fatty acid synthesis. To explore this possibility, we examined how M2 activation was affected by (5-(Tetradecyloxy)-2-furoic Acid (TOFA), an inhibitor of acetyl-CoA carboxylase, which catalyzes an early step in the synthesis of fatty acid from acetyl-CoA. TOFA completely blocked M2 activation in serum-free conditions and, in the presence of serum, reduced activation to levels observed in inhibitor-free, serum-free conditions (Supplementary Fig. 5b). These data suggest that macrophages possess an underlying pathway through which, in the presence of IL-4, they synthesize fatty acids to support M2 activation, and that optimal M2 activation requires contributions of exogenous triacylglycerols and de novo synthesized fatty acids.

These findings raised the question of whether M2 macrophages first incorporate endogenously synthesized fatty acids into triacylglycerols prior to liberating them by lipolysis for FAO, as has been suggested to be the dominant pathway in other cell types¹¹. We examined this by first testing the effect of Orlistat on M2 activation in serum-free conditions. We found that the inhibition of lipolysis under these conditions did inhibit M2 activation (Supplementary Fig. 5b), indicating that endogenously synthesized fatty acids are incorporated into triacylglycerols in M2 macrophages. Triacylglycerols synthesized in this way are stored as lipid droplets, but our ultrastructural analysis failed to identify lipid droplets in M2 cells (Fig. 3b). However, we did observe lipid droplets in macrophages stimulated with IL-4 in the presence of Orlistat and in macrophages transduced with shRNA targeting Lipa (Supplementary Fig. 3b). Since the inhibition of lipolysis precludes the synthesis of these neutral lipid stores from fatty acids liberated from acquired triacylglycerols, we assume that they are produced from de novo synthesized fatty acids. Taken together, our data raise the possibility that in addition to a primary pathway in which exogenous triacylglycerols are broken down in the lysosomal compartment to support FAO, M2 macrophages are also able to synthesize triacylglycerols from de novo synthesized fatty acids and hydrolyze these via the same pathway.

M2 activation during helminth infection is LAL-dependent

Our data indicated that lipolysis is essential for the metabolic switch and associated changes in the expression of key genes that occur as macrophages become alternatively activated in response to IL-4. To assess whether lipolysis is critical for M2 activation in vivo, we used the Heligmosomoides polygyrus bakeri (H. polygyrus) mouse model of human intestinal helminth infection, in which M2 macrophages play an essential role in immunity²⁹. We found that peritoneal macrophage numbers increased significantly over time during this infection (Fig. 5a), and that peritoneal macrophages from infected mice exhibited the metabolic profile of IL-4-activated bone marrow macrophages, with heightened baseline OCR and marked SRC (Fig. 5b). Consistent with previous observations, a substantial percentage of peritoneal macrophages from H. polygyrus-infected mice expressed the M2 marker RELMα (Fig. 5c)³⁰. We found that maintenance of the peritoneal M2 macrophage activation phenotype is LAL-dependent in this setting since increases in OCR and RELMα expression were significantly inhibited when peritoneal macrophages were treated ex-vivo with siRNAs targeting Lipa (Fig. 5d,e). The dependence of M2 activation on LAL was also apparent when peritoneal cells were maintained in a type 2 environment through the addition of IL-4 in vitro (Fig. 5f,g). These data indicate that macrophages in a type 2 cytokine environment in vivo exhibit lipolysis-dependent changes in metabolism and M2 marker gene expression that are equivalent to those induced by IL-4 in bone marrow derived macrophages in vitro.

Inhibition of lipolysis suppresses parasite elimination

Primary infection with H. polygyrus in B6 mice is chronic despite the development of a type 2 cytokine response. However, injection of IL-4c into infected mice results in reductions in adult parasite numbers, and M2 macrophages are implicated in the protective mechanism induced by this cytokine^3,29,31. Given our findings, we reasoned that lipolysis should be crucial for the development of M2 macrophages in these settings, and therefore that inhibition of this pathway would limit the expression of M2 macrophage-dependent protective immune responses. We found that IL-4c injected after primary infection markedly enhanced peritoneal macrophage OXPHOS (Fig. 6a) and increased RELMα expression (Fig. 6b), while simultaneously significantly reducing parasite burden, as measured by egg and adult worm counts (Fig. 6c,d). Consistent with a major role for lipolysis in M2 polarization, we found that enhanced M2 activation and concomitant protective effects associated with IL-4c injection were significantly inhibited by parenteral Orlistat treatment (Fig. 6a-e).

Mice in which primary H. polygyrus is cleared by chemotherapy exhibit a degree of immunity to secondary infection²⁹. We compared the peritoneal macrophage response during primary H. polygyrus infection to that of a secondary re-infection following parasite clearance by administration of pyrantel pamoate and resting the animals for five weeks. We found that peritoneal macrophage numbers (Fig. 7a) and the magnitude of the increase in OXPHOS and RELMαexpression (Fig. 7b,c) were all significantly greater in mice responding to secondary infection. This correlated with resistance to secondary infection, apparent as a >50% drop in adult worm burdens arising from secondary vs. primary infections (Fig. 7d). We then asked whether inhibition of lipolysis following secondary infection affected enhanced M2 activation, or resistance to secondary infection. We found that Orlistat treatment prevented the increase in peritoneal macrophage numbers that occurred during secondary infection (Fig. 7a), inhibited all measured parameters of enhanced M2 activation (Fig. 7b,c) and suppressed resistance to reinfection in immune mice (Fig. 7d).

Given the experimental approach used here, there was a possibility that Orlistat was affecting not only macrophage function, but also adaptive immunity. To address this, we used 4get/KN2 IL-4 reporter mice to enumerate CD4⁺ type 2 helper T cells (T_H2) cells in mesenteric lymph nodes following primary or secondary infections with or without Orlistat treatment. We found that inhibition of lipolysis during the development of the effector or secondary effector response had no measurable effect on the adaptive response, as measured by the overall cellularity of the MLN (Supplementary Fig. 6a), or the numbers of IL-4–secreting T_H2 cells in the MLN (Supplementary Fig. 6b). Furthermore, examining IL-4 reporter expression in all live cells from mice treated with Orlistat failed to reveal any effects on non-T_H2 sources of IL-4 (data not shown). In summary, our data show that the inhibition of lipolysis is able to block macrophage M2 activation associated with resistance to the helminth parasite H. polygyrus, and in so doing suppress immunity to this pathogen

Animals and in vivo experiments

C57BL/6J mice, 4get/KN2 mice, B6 Cd36^–/– mice and B6.129P2-Pnpla2tm^1Rze/J (Pnpla2^–/–) mice were bred and maintained in specific pathogen free conditions under protocols approved by the institutional animal care committee at Washington University School of Medicine, and were used at 8-12 weeks of age. Lipa^+/+ and Lipa^–/– mice were maintained at Indiana State University under SPF conditions. In some experiments, mice were injected i.p. with tetrahydrolipistatin (Orlistat; 200 mg/kg, Cayman Chemical) or with 300 μl of 3% thioglycollate (Sigma) immediately prior to IL-4 complexed to mAb anti-IL-4 (IL-4c; containing 5 μg of IL-4, PeproTech, and 25 μg of anti-IL-4 clone 11B11, BioXcell)⁴⁹. For infection with H. polygyrus bakeri, we followed protocols described in detail previously⁵⁰. Mice were infected orally by gavage with 200 infective L3 stage larvae. For secondary infection experiments, adult H. polygyrus were eliminated from infected mice by oral administration of pyrantel pamoate (1 mg/mouse; Columbia Laboratory) at day 14 post-primary infection, and > 5 weeks later the mice were challenge-infected with 200 L3 stage larvae by gavage. At day 9 post challenge infection, mice were sacrificed and parasite burdens were measured. To count worm, intestines were removed, opened longitudinally, and placed into a metal strainer on top of a 50 ml tube filled with phosphate buffered saline (PBS; Corning Inc.) for 4 h at 37 °C. Parasites dropped through the filter into the tube and were recovered for counting on a dissecting microscope. Parasite eggs were enumerated by floating eggs in feces collected from individual mice on saturated sodium chloride prior to collection, and counting under a microscope.

Isolation of cells from mice

Peritoneal exudate cells (PECs) were harvested from sacrificed naïve or infected mice by peritoneal lavage with 10 ml of sterile PBS containing 5% FBS (HyClone) per mouse. Total numbers of PECs and peritoneal macrophages were determined by cell counting in combination with multi-color flow cytometry. For analysis of lymphocyte activation by flow cytometry, single cell suspensions were prepared from lymphoid organs by forcing them through 70 μm strainers, and washing with RPMI containing 5% FBS (Corning Inc). Red blood cells were lysed with ACK lysis buffer (0.15 M NH₄Cl, 1 mM KHCO₃, 0.1 mM EDTA). Bone marrow was flushed from the femur and tibia using RPMI containing 5% FBS and dissociated into a single cell suspension by repeated pipetting. Cells were maintained on ice until use or analysis.

Preparation of macrophages from bone marrow and macrophage activation

Bone marrow cells were differentiated in the presence of recombinant mouse M-CSF (20 ng/ml; R&D Systems) in complete medium (RPMI 1640 containing 10 mM glucose, 2 mM L-glutamine, 100 U/mL of penicillin/streptomycin and 10% FBS) for 7 days. Day 7 macrophages were washed and variously stimulated with IL-4 (20 ng/ml; PeproTech) or lipopolysaccharide (LPS, 20 ng/ml; Sigma) plus IFN-γ (50 ng/ml; R&D Systems) in the presence or absence of 200 μM etomoxir (ETO; Sigma), 20 μM sulfo-n-succinimidyl oleate (SSO), 30 μM chloroquine (CLQ; Sigma), 0.1 μM bafilomycin A1 (Baf; Tocris Bioscience), 100 μM Orlistat (Cayman), 20 μM 5-(Tetradecyloxy)-2-furoic Acid (TOFA, Sigma), 5 or 50 μg/ml of LDL (Kalen Biomedical, LLC) and VLDL (Kalen Biomedical, LLC) for 24 h. Macrophages were then harvested and analyzed by flow cytometry for expression of markers of M1 or M2 activation. In some experiments the survival of macrophages was monitored following M2 or M1 activation. For these experiments, bone marrow-derived macrophages (0.5 × 10⁶ cells per well of a 48-well plate) were cultured in complete medium with mouse M-CSF (20 ng/ml) and stimulated with IL-4 or LPS plus IFN-γ. Cell viability was monitored by flow cytometric analysis of 7-amino-actinomycin D (7-AAD; BioLegend) staining of F4/80⁺ cells over time. Culture medium and stimulation signals were replenished every 1 or 2 days.

Identification of human CD36-sufficient and -deficient subjects

Peripheral blood mononuclear cells (PBMC) were isolated from venous blood samples obtained from a CD36-deficient adult subject and an adult subject with normal CD36 expression using Ficoll-Paque (GE Health Care) and density gradient centrifugation. The CD36-deficient subject is homozygous for the G allele (allele frequency ~10%) of coding SNP rs3111938 (T/G), which introduces a stop codon and results in a truncated protein that is degraded. Homozygosity at the G allele results in complete CD36 deficiency⁵¹. The study was approved by the Washington University School of Medicine Institutional Review Board with written informed consent obtained from each subject.

Isolation of mononuclear cells from human peripheral blood and human macrophage activation

Monocytes were purified from PBMCs using human CD14 MicroBeads (MACS) and then differentiated into macrophages in complete RPMI medium containing 20 ng/ml recombinant human M-CSF (R&D Systems) for 8 days. Human macrophages were harvested on day 8, and stimulated with 20 ng/ml human IL-4 (R&D Systems) or 50 ng/ml human IFN-γ (R&D Systems) plus 20 ng/ml LPS for 24 h.

Retroviral transduction

Retroviral transduction of macrophages was accomplished using a protocol that we have used previously to transduce bone marrow derived dendritic cells⁵². Sequences for luciferase and for Lipa short hairpin RNAs (shRNAs) were obtained from Open Biosystems and cloned into the MSCV-LTRmir30-PI8 retroviral vector, encoding huCD8 as a reporter. Two independent sequences were used to target Lipa: TCAAGTCCAGCATTCACTG, and TGTGCTTCAGAGACCAGGT. Recombinant retroviruses were used for spin infection (800 x g, 2 h) of day 3 bone marrow macrophage cultures. After 7 days in culture with M-CSF, macrophages were harvested and transduced macrophages were gated based on huCD8 expression. Both shRNAs were used for experiments, with similar results, but only data from experiments that utilized TGTGCTTCAGAGACCAGGT are shown. For the bone marrow chimera, bone marrow cells from donor mice were collected and cultured in complete RPMI medium containing 10 ng/ml IL-3, 10 ng/ml IL-6 and 50 ng/ml murine c-kit ligand for 24 h. Cells were then transduced with retroviral shRNAs targeting Lipa or luciferase, and 5 × 10⁵ transduced cells were intravenously injected into irradiated recipients. Recipients were used 6 – 10 weeks later.

Treatment of peritoneal macrophages with small interfering RNA (siRNA)

An siRNA targeting mouse LAL (Lipa; AAAGCUAUGAAACCAUGGTg) was purchased from Applied Biosystems, and Silencer Negative control siRNA#1, which is not matched to any sequence in the mouse genome, was also provided by the manufacturer and used as a control. siRNAs (50 nM) were complexed with Lipofectamine 2000 and delivered into peritoneal macrophages as per the manufacturer's instructions. siRNA-treated peritoneal macrophages were then stimulated with 20 ng/ml of IL-4 for 48 h prior to analysis.

Metabolism assays

For real-time analysis of extracellular acidification rates (ECAR) and oxygen consumption rates (OCR), macrophages were analyzed using an XF-96 Extracellular Flux Analyzer (EFA, Seahorse Bioscience) as described in detail for bone marrow-derived dendritic cells previously⁵². Three or more consecutive measurements were taken under basal conditions and following the sequential addition of 1 μM oligomycin to inhibit the mitochondrial ATP synthase, 1.5 μM fluoro-carbonyl cyanide phenylhydrazone (FCCP), a protonophore, which uncouples ATP synthesis from oxygen consumption by the electron transport chain, and 100 nM rotenone plus 1 μM antimycin A (Rot/Ant; all drugs for this assay were purchased from Sigma), which inhibit the electron transport chain. In this assay, basal oxygen consumption can be established by measuring OCR in the absence of drugs. Declines in OCR following the addition of oligomycin and rotenone and antimycin are expected and indicate that cells are consuming oxygen for mitochondrial OXPHOS. Spare respiratory capacity (SRC) is calculated as the difference between basal OCR and maximal OCR following the addition of FCCP. Maximal OCR occurs following the addition of FCCP since cells attempt to maintain a proton gradient across the inner mitochondrial membrane by increasing the consumption of oxygen by the electron transport chain^10,53. Global metabolite profiling of stimulated macrophages was performed using mass spectrometry by Metabolon, Durham, NC. Glycerol in macrophage supernatants was measured using absorbance at 570 nm after incubation with Free Glycerol Reagent (Sigma) and by comparison to a glycerol standard (Sigma). For the lipidomic analysis, washed cell pellets were frozen and thawed and homogenized in 500 μl of PBS using an Omni Bead Ruptor 24 (Omni International, Inc.). 50 μl of each homogenate was reserved for protein measurement. 450 μl of each homogenate was used for lipidomic analysis. A modified Bligh-Dyer extraction method was used to extract lipids in the presence of an internal standard mixture. Mass spectrometric measurement was performed with a Shimadzu 10A HPLC system coupled to a TSQ Quantum Ultra triple quadrupole mass spectrometer operated in SRM mode under ESI(+). Different analytical HPLC columns and mobile phases were employed for different analytes to achieve the optimal sensitivity and separation. A pooled lipid extract from study samples was used as a quality control (QC), and was injected between every sample to verify the instrumental consistency, which is described as %CV of QC. Only the species with (%CV of QC) < 15% were reported. Data processing was conducted with Xcalibur software (Thermo). The concentration of an analyte was calculated as the concentration of its corresponding internal standard multiplied by the peak area ratio of the analyte to the internal standard, based on the assumption that the MS responses of the analyte and internal standard are the same.

Flow Cytometry

Cells were kept at 4 °C and blocked with 5 μg/ml of anti-CD16/32 (clone 93, eBiosciences) before surface staining with antibodies to CD11b (clone M1/70, eBiosciences), F4/80 (clone BM8, eBiosciences), CD206 (clone C068C2, Biolegend), CD301 (clone ER-MP23, AbD Serotec), PD-L2 (clone TY25, eBiosciences), CD62L (clone MEL-14, BD Biosciences), CD44 (clone IM7, BD Biosciences), CD4 (clone RM4-5, BD Biosciences), and/or human CD2 (clone RPA-2.10, BD Biosciences). For intracellular staining of RELMα and iNOS, cells were fixed in 4% ultrapure paraformaldehyde, and stained with rabbit anti-RELMα (PeproTech) and mouse anti-Nos2 (clone C-11; Santa Cruz Biotechnology) for 1 h at 22°C in 0.2% saponin buffer, followed by incubation with appropriate fluorochrome-conjugated anti-rabbit or anti-mouse IgG (both Jackson Immunoresearch) in the same buffer. BODIPY labeled LDL (10 μg/ml, Invitrogen), BODIPY labeled VLDL (10 μg/ml, Kalen Biomedical, LLC) and BODIPY labeled palmitate (1 μM, Invitrogen) were used in conjunction with flow cytometry for uptake experiments. Intracellular neutral lipids were stained with 500 ng/ml BODIPY 493/503 (Invitrogen) and measured by flow cytometry. All macrophage staining data shown represent cells that were gated as CD11b⁺F4/80⁺, and living based on staining with LIVE/DEAD (Invitrogen) or 7-amino-actinomycin D (eBiosciences). Human macrophages were stained with antibodies to human CD14 (clone MφPg, BD Biosciences), human CD36 (clone CB38, BD Biosciences) and human CD206 (clone 15-2, Biolegend). Staining results are shown for CD14⁺ cells. Data were acquired on a FACSCanto II flow cytometer (BD Biosciences), and analyzed with FlowJo v.9.5.2 (Tree Star Inc.).

RNA-sequencing

mRNA was extracted from lysates of cells that had been stimulated for 24 h, using oligodT beads (Invitrogen). cDNA synthesis, sequencing and sequence analysis were performed as described⁵³. Raw and processed sequencing data were deposited to Pubmed.

Transmission electron microscopy

For ultrastructural analysis, cells were fixed in 2% paraformaldehyde, 2.5% glutaraldehyde (Polysciences Inc.), 0.05% malachite green (Sigma) in 100 mM sodium cacodylate buffer, pH 7.2 for 1 h at 22°C. The malachite green was incorporated into the fixative to stabilize lipid constituents soluble in aqueous glutaraldehyde. Samples were washed in cacodylate buffer and post-fixed in 1% osmium tetroxide (Polysciences Inc.) for 1 h. Samples were then rinsed extensively in distilled H₂O prior to en bloc staining with 1% aqueous uranyl acetate (Ted Pella Inc.) for 1 h. Following several rinses in dH₂O, samples were dehydrated in a graded series of ethanol and embedded in Eponate 12 resin (Ted Pella Inc.). Sections of 95 nm were cut with a Leica Ultracut UC7 ultramicrotome (Leica Microsystems Inc.), stained with uranyl acetate and lead citrate, and viewed on a JEOL 1200 EX transmission electron microscope (JEOL USA Inc.) equipped with an AMT 8 megapixel digital camera (Advanced Microscopy Techniques).

Statistical analysis

Data were analyzed using Graphpad Prism (v5). Comparisons for two groups were calculated using One-way ANOVA and, where indicated, unpaired two-tailed Student's t-tests. Differences were considered significant when P-values were below 0.05. No randomization or exclusion of data points was used. Pilot in vivo studies were used for estimation of the sample size required to ensure adequate power.