Obesity and related inflammation are critical for the pathogenesis of insulin resistance, but the underlying mechanisms are not fully understood. Formyl peptide receptor 2 (FPR2) plays important roles in host immune responses and inflammation-related diseases. We found that Fpr2 expression was elevated in the white adipose tissue of high-fat diet (HFD)–induced obese mice and db/db mice. The systemic deletion of Fpr2 alleviated HFD-induced obesity, insulin resistance, hyperglycemia, hyperlipidemia, and hepatic steatosis. Furthermore, Fpr2 deletion in HFD-fed mice elevated body temperature, reduced fat mass, and inhibited inflammation by reducing macrophage infiltration and M1 polarization in metabolic tissues. Bone marrow transplantations between wild-type and Fpr2−/− mice and myeloid-specific Fpr2 deletion demonstrated that Fpr2-expressing myeloid cells exacerbated HFD-induced obesity, insulin resistance, glucose/lipid metabolic disturbances, and inflammation. Mechanistic studies revealed that Fpr2 deletion in HFD-fed mice enhanced energy expenditure probably through increasing thermogenesis in skeletal muscle; serum amyloid A3 and other factors secreted by adipocytes induced macrophage chemotaxis via Fpr2; and Fpr2 deletion suppressed macrophage chemotaxis and lipopolysaccharide-, palmitate-, and interferon-γ–induced macrophage M1 polarization through blocking their signals. Altogether, our studies demonstrate that myeloid Fpr2 plays critical roles in obesity and related metabolic disorders via regulating muscle energy expenditure, macrophage chemotaxis, and M1 polarization.

Obesity is a major risk factor for the development of insulin resistance and type 2 diabetes (1,2). Chronic low-grade inflammation in obesity is one of the most important causes of obesity-related complications (3). Macrophages play crucial roles in obesity-related inflammation and insulin resistance (4). In obese individuals, macrophages accumulate in adipose tissue and form crown-like structures (CLSs) by surrounding the dead adipocytes (5). Adipose tissue macrophages undergo a phenotypic switch from the alternatively activated M2 phenotype to the classically activated M1 phenotype during obesity (4). M1 macrophages secrete proinflammatory cytokines that aggravate tissue inflammation and cause local and systemic insulin resistance through interfering with insulin-signaling pathways (6). In contrast, M2 macrophages are constitutively present in the lean adipose tissue and express high levels of anti-inflammatory cytokines that are positively associated with insulin sensitivity (7). Studies have shown that the reduced macrophage infiltration into tissues and inhibited macrophage M1 polarization could attenuate obesity-related insulin resistance (8,9). Therefore, the identification of key molecules contributing to macrophage infiltration and M1 polarization may provide therapeutic targets against insulin resistance and type 2 diabetes.

Formyl peptide receptor 2 (FPR2) is a chemoattractant receptor that belongs to the FPR family. Human FPR2 and its mouse homolog Fpr2 are highly expressed in macrophages/monocytes and neutrophils, interact with peptide and lipid ligands, and transduce pro- or anti-inflammatory actions (10). The different effects of FPR2 on inflammation are dependent on the context and ligands that activate different signaling pathways (11,12). FPR2 is involved in multiple diseases, including bacterial infection, inflammation, asthma, Alzheimer disease, and cancers (13,14). Studies with Fpr2 knockout or Fpr2/Fpr3 double-knockout mice showed that Fpr2 played protective or detrimental roles in different disease models (1518). Annexin A1 (ANXA1) and lipoxin A4 (LXA4) are Fpr2 ligands with anti-inflammatory properties. ANXA1 reduces body weight gain in obese mice and inhibits hepatic inflammation during nonalcoholic steatohepatitis progression (19,20). LXA4 attenuates obesity-induced inflammation in adipose tissue and related diseases (21). Because ANXA1 and LXA4 can activate other receptors in addition to Fpr2 (11), it is of interest to determine the contribution of Fpr2 in obesity-related chronic inflammation and metabolic disorders by using Fpr2 knockout mice.

In the current study, our in vivo data from Fpr2-deficient mouse models and in vitro results from bone marrow-derived macrophages (BMDMs) demonstrated that Fpr2 plays vital roles in diet-induced obesity (DIO), inflammation, and metabolic disorders.

Animal Experiments

Fpr2−/− mice and wild-type (WT) littermates were generated by intercrossing Fpr2+/− mice (16). Myeloid-specific Fpr2 knockout (Fpr2MKO) mice were developed by crossing Fpr2flox/flox mice with LysM-Cre mice. C57BL/6 and db/db mice were obtained from Shanghai Laboratory Animal Company (Shanghai, China). Mice of different genotypes were housed in different cages. Eight-week-old male mice were fed a chow diet (10% fat calories) or high-fat diet (HFD; 60% fat calories) (Research Diets, New Brunswick, NJ) for 9–12 weeks.

Body composition was analyzed using EchoMRI (EchoMRI, Houston, TX). Glucose tolerance tests (GTTs) and insulin tolerance tests (ITTs) were conducted as described previously (22). Rectal temperatures were measured using a rectal probe attached to a digital thermometer (Physitemp Instruments, Clifton, NJ). To examine the tissue response to insulin, mice fasted for 4 h were injected with insulin (2 units/kg body wt) or PBS in the inferior vena cava, followed by collection of liver tissues at 3 min, epididymal white adipose tissue (WAT) at 5 min, and gastrocnemius muscle at 7 min after the injection. Akt phosphorylation was examined by Western blotting. All animal experiments were performed in accordance with the guidelines of the Institutional Animal Care and Use Committee at Shanghai Institute of Nutrition and Health, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences.

Tissue Uptake of 18F-Fluorodeoxyglucose

Mice fasted for 16 h were intravenously injected with 18F-fluorodeoxyglucose (5.8 ± 0.7 MBq/mouse). After 1 h, the tissue samples were collected and measured for radioactivity. The differential uptake ratio (DUR) was used as an index of radiotracer uptake in tissues. DUR = (tissue counts [cpm] per g of tissue)/(injected dose counts per g of body weight) (23).

Energy Expenditure Measurement

VO2, VCO2, and locomotive activities of mice were determined in the Comprehensive Lab Animal Monitoring System (Columbus Instruments, Columbus, OH). Data were collected for 48 h after the mice were acclimated to the system for 24 h with free access to food and water. The rate of energy expenditure (calories/min) was calculated as VO2 × (3.815 + [1.232 × {VCO2/VO2}]).

Biochemical Parameter Analysis

Serum and liver triglycerides (TGs), serum aspartate aminotransferase (AST), and alanine aminotransferase (ALT) were determined using the respective kits (Shensuoyoufu, Shanghai, China). Serum levels of nonesterified fatty acids (NEFA), insulin, and proinflammatory cytokines were measured with a NEFA assay kit (Wako Pure Chemicals, Osaka, Japan) and ELISA kits (Millipore, Billerica, MA; R&D Systems, Minneapolis, MN), respectively.

Histology and Immunohistochemistry

The tissues sections were stained with hematoxylin and eosin (H&E) or Oil Red O or immunostained with antibodies against CD68 and F4/80. Adipocyte diameter and CLSs in the WAT sections were analyzed using Image-Pro Plus software (Media Cybernetics, Rockville, MD). Macrophage infiltration in the liver sections was quantified by average optical density with Image-Pro Plus software.

mRNA and Protein Analysis

Quantitative real-time PCR was performed as described previously (24). The primer sequences are presented in the Supplementary Data. Western blot analysis was performed with primary antibodies against phosphorylated and total forms of Akt, nuclear factor-κB (NF-κB) p65, p38, extracellular signal–regulated kinase (ERK), c-Jun N-terminal kinase (JNK), transforming growth factor β-activated kinase 1 (TAK1) and STAT1, HSP90 (Cell Signaling, Danvers, MA), and β-actin (Sigma-Aldrich, St. Louis, MO). Signaling was visualized with ECL Plus Western Blotting Detection System (GE Healthcare, Salem, CT).

Cell Isolation, Culture, and Treatment

BMDMs prepared from bone marrow cells (25) were stimulated with lipopolysaccharide (LPS; Sigma-Aldrich), palmitate, IFN-γ (Peprotech, Rocky Hill, NY), or serum amyloid A3 (Saa3; CUSABIO, Wuhan, China), and examined for the phosphorylation of mitogen-activated protein (MAP) kinases, NF-κB p65, STAT1, and TAK1 by Western blotting, for expression of M1 macrophage-related proinflammatory cytokines by quantitative real-time PCR and ELISAs, respectively. Sodium palmitate (Sigma-Aldrich) was dissolved in fatty acid–free BSA (Sigma-Aldrich) solution with a 2.5:1 mol/L ratio.

Fpr2 was introduced into BMDMs by infecting the cells with Fpr2-lentiviruses. 3T3-L1 fibroblasts were differentiated into mature adipocytes as described previously (26) and transfected with Saa3 siRNA or negative control using Lipofectamine 3000 (Invitrogen, Thermo Fisher Scientific, Waltham, MA). The sequence of Saa3 siRNA is 5′-GCUGGUCAAGGGUCUAGAG-3′.

FACS Analysis

Stromal vascular cells (SVCs) were isolated from epididymal WAT by type I collagenase digestion (27,28), stained with an antibody cocktail containing anti–CD45-FITC, F4/80-phosphatidylethanolamine (PE), CD11b-eFluor 450, CD206-allophycocyanin (APC), and CD11c-PE-Cyanine7 (eBioscience, Thermo Fisher Scientific). M1 and M2 macrophages were identified with FACS analysis using the gate strategy as previously published (28).

Chemotaxis Assay

Chemotaxis of BMDMs in response to adipose tissue lysate, culture medium from adipocytes, or Saa3 was analyzed using a polycarbonate membrane with an 8-μm pore size in 48-well chemotaxis chambers (NeuroProbe, Gaithersburg, MD) (29). The migrated cells were analyzed with Image-Pro Plus software.

Bone Marrow Transplantations

Bone marrow transplantations (BMTs) between mice were performed as previously described (30). Fpr2 expression in neutrophils and monocytes isolated from bone marrows was detected to evaluate the efficiency of BMTs.

Statistics

All experiments were repeated at least three times. The results are presented as the mean ± SD or SEM. The statistical analysis was performed by using unpaired two-tailed Student t tests for two-group comparison and using two-way ANOVA or two-way repeated-measures ANOVA for multiple group comparison. The residual method (31) was used to control body weight in analyzing data of GTTs and ITTs and semiquantitative data of immunostainings. Body weight–adjusted blood glucose in GTTs/ITTs and semiquantitative data of immunostainings were computed as the residuals from the regression model with blood glucose or semiquantitative data of immunostainings as the independent variable and body weight as the dependent variable. Significance was accepted at P < 0.05.

Fpr2 Is Upregulated in Adipose Tissues of Diabetic Mice

We examined Fpr2 expression in major metabolic tissues of diabetic mice and found that Fpr2 expression was upregulated in WAT and gastrocnemius muscle of DIO mice compared with that of chow-fed mice (Fig. 1A). Fpr2 expression was also upregulated in WAT of db/db mice (Fig. 1A). We next examined the cellular source of Fpr2 in WAT and found that DIO mice had a higher level of Fpr2 mRNA in SVCs (Fig. 1A). These results show that Fpr2 expression is significantly increased in WAT of obese mice, especially in SVCs.

Figure 1

Deletion of Fpr2 attenuates DIO. A: Fpr2 mRNA expression in liver, epididymal WAT (eWAT), and gastrocnemius muscle of HFD mice (n = 6–10 per group), db/db mice, and their corresponding control mice (n = 6–7 per group), and in adipocytes (Ads) and SVCs isolated from adipose tissue of mice fed the HFD or chow (n = 3–4 per group). BI: Fpr2−/− and WT mice were fed the HFD or control chow. B: Body weight was measured weekly. Food intake over 24 h (C), body composition (D), and rectal temperature (RT) (F) were determined at 8 weeks on the HFD (n = 8–13 per group). Other parameters were examined at 10 weeks on HFD as follows: adipocyte diameter in H&E-stained eWAT sections (n = 5–9 per group) (E); average VO2, VCO2, the rate of energy expenditure, and locomotor activities of mice over 24 h (n = 6–8 per group) (G and H); and expression of thermogenic genes in gastrocnemius muscle, BAT, and sWAT (n = 7–12 per group) (F). Data are shown as the mean ± SEM. *P < 0.05, **P < 0.01 for comparison between WT and db/db mice (A) or between Fpr2−/− and WT mice fed the HFD (B–I). Two-way repeated-measures ANOVA was used in B, and the Student t test was used in A and CI.

Figure 1

Deletion of Fpr2 attenuates DIO. A: Fpr2 mRNA expression in liver, epididymal WAT (eWAT), and gastrocnemius muscle of HFD mice (n = 6–10 per group), db/db mice, and their corresponding control mice (n = 6–7 per group), and in adipocytes (Ads) and SVCs isolated from adipose tissue of mice fed the HFD or chow (n = 3–4 per group). BI: Fpr2−/− and WT mice were fed the HFD or control chow. B: Body weight was measured weekly. Food intake over 24 h (C), body composition (D), and rectal temperature (RT) (F) were determined at 8 weeks on the HFD (n = 8–13 per group). Other parameters were examined at 10 weeks on HFD as follows: adipocyte diameter in H&E-stained eWAT sections (n = 5–9 per group) (E); average VO2, VCO2, the rate of energy expenditure, and locomotor activities of mice over 24 h (n = 6–8 per group) (G and H); and expression of thermogenic genes in gastrocnemius muscle, BAT, and sWAT (n = 7–12 per group) (F). Data are shown as the mean ± SEM. *P < 0.05, **P < 0.01 for comparison between WT and db/db mice (A) or between Fpr2−/− and WT mice fed the HFD (B–I). Two-way repeated-measures ANOVA was used in B, and the Student t test was used in A and CI.

Deletion of Fpr2 Alleviates DIO, Insulin Resistance, and Impairment of Glucose and Lipid Metabolism

To study the contribution of Fpr2 to metabolic regulation, we fed WT and Fpr2−/−mice a chow diet or HFD. We found that systemic Fpr2 deletion had no significant effect on body weight and composition, food intake, energy expenditure, glucose, and lipid metabolism in chow-fed mice (Fig. 1B and Supplementary Fig. 1). When fed the HFD, Fpr2−/− mice presented similar food intake to WT mice but had lower body weight than WT mice after 3 weeks (Fig. 1B and C). Compared with WT mice, Fpr2−/− mice had lower fat mass percentage, higher lean mass percentage (Fig. 1D), and smaller adipocytes in WAT (Fig. 1E). Fpr2 deletion increased O2 consumption (O2), CO2 production (CO2), energy expenditure rate, and rectal temperature, but had no significant effect on physical activities (Fig. 1F–H). We further evaluated the expression of thermogenic genes in subcutaneous WAT (sWAT), brown adipose tissue (BAT), and the gastrocnemius muscle. Compared with WT mice, HFD-fed Fpr2−/− mice expressed higher levels of Pgc1α, Pparα, Ucp2, and Cd36 in the muscle, but expressed comparable thermogenic genes in sWAT and BAT (Fig. 1I). These results indicate that Fpr2 deficiency reduces body weight gain in HFD-fed mice through enhancing energy expenditure, especially in skeletal muscle.

Further studies showed that HFD-fed Fpr2−/− mice displayed lower blood glucose and serum insulin levels (Fig. 2A and B) and improved glucose tolerance and insulin sensitivity (Fig. 2C and E) compared with WT mice. Consistently, higher Akt phosphorylation in response to insulin was observed in WAT and gastrocnemius muscle of Fpr2−/− mice (Fig. 2G). After adjusting the data of GTTs and ITTs with body weight, the differences between WT and Fpr2−/− mice were decreased but still statistically significant (Fig. 2D and F), indicating that Fpr2 deficiency attenuates insulin resistance through reducing body weight gain and other mechanisms. In addition, Fpr2−/− deficiency reduced serum TG and NEFA levels (Fig. 2H), hepatic lipid accumulation (Fig. 2J and K), and serum AST and ALT levels (Fig. 2I). Collectively, these data demonstrate that Fpr2 deficiency improves insulin sensitivity and alleviates lipid and glucose dysregulation in DIO mice.

Figure 2

Fpr2−/− mice are protected from HFD-induced insulin resistance and hepatic steatosis. Fpr2−/− and WT mice were fed the HFD for 10 weeks. Metabolic phenotypes were examined as follows: blood glucose levels under fed and fasted conditions (A), serum insulin levels (n = 7–12 per group) (B); GTT and average area under the curve (AUC) adjusted with or without body weight (C and D); ITT and AUC adjusted with or without body weight (n = 10–14 per group) (E and F); Akt phosphorylation (p-) in the liver, epididymal WAT, and gastrocnemius muscle after insulin administration in the inferior vena cava (n = 4–6 per group) (G); serum levels of TG and NEFA (H); serum levels of AST and ALT (I); liver TG content (J); and H&E (HE) and Oil Red O staining of liver sections (n = 6–11 per group) (K). The results represent the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 for the comparison between Fpr2−/− and WT mice fed the HFD. Images in K are representatives of H&E and Oil Red O staining of liver sections. Scale bar = 100 μm. The Student t test was used in A and B and HJ. Two-way repeated-measures ANOVA was used in CF, and two-way ANOVA was used in G. Body weight–adjusted blood glucose levels (D) and blood glucose change (%) (F) were computed as the residuals from the regression model, with blood glucose level or change as the independent variable and body weight as the dependent variable.

Figure 2

Fpr2−/− mice are protected from HFD-induced insulin resistance and hepatic steatosis. Fpr2−/− and WT mice were fed the HFD for 10 weeks. Metabolic phenotypes were examined as follows: blood glucose levels under fed and fasted conditions (A), serum insulin levels (n = 7–12 per group) (B); GTT and average area under the curve (AUC) adjusted with or without body weight (C and D); ITT and AUC adjusted with or without body weight (n = 10–14 per group) (E and F); Akt phosphorylation (p-) in the liver, epididymal WAT, and gastrocnemius muscle after insulin administration in the inferior vena cava (n = 4–6 per group) (G); serum levels of TG and NEFA (H); serum levels of AST and ALT (I); liver TG content (J); and H&E (HE) and Oil Red O staining of liver sections (n = 6–11 per group) (K). The results represent the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 for the comparison between Fpr2−/− and WT mice fed the HFD. Images in K are representatives of H&E and Oil Red O staining of liver sections. Scale bar = 100 μm. The Student t test was used in A and B and HJ. Two-way repeated-measures ANOVA was used in CF, and two-way ANOVA was used in G. Body weight–adjusted blood glucose levels (D) and blood glucose change (%) (F) were computed as the residuals from the regression model, with blood glucose level or change as the independent variable and body weight as the dependent variable.

Deletion of Fpr2 Reduces HFD-Induced Systemic Inflammation, Tissue Macrophage Infiltration, and M1 Polarization

We next evaluated the contribution of Fpr2 to obesity-related inflammation. When fed the HFD, Fpr2−/− mice showed lower serum levels of interleukin 6 (IL-6), tumor necrosis factor-α (TNF-α), and chemokine (C-C motif) ligand 2 (CCL2) than WT mice (Fig. 3A). CLSs and CD68-positive macrophages in WAT and F4/80-positive macrophages in liver (Fig. 3B and C) were markedly reduced in Fpr2−/− mice. After adjustment with body weight, the differences of CLSs in WAT and infiltrated macrophages in liver were still statistically significant between WT and Fpr2−/− mice (Fig. 3D). Consistently, obese Fpr2−/−mice showed reduced expression of macrophage and M1 macrophage markers in WAT (Fig. 3E). However, the expression of M2 macrophage markers (Il-10, Ym1 except Retnla) in WAT was comparable between Fpr2−/− and WT mice (Fig. 3E). The expression of macrophage and M1 macrophage markers was slightly downregulated in the liver and muscle of Fpr2−/− mice (Fig. 3E). Peritoneal macrophages from HFD-fed Fpr2−/−mice also expressed lower mRNA levels of M1 macrophage-related genes than WT mice (Fig. 3F). These data demonstrate that Fpr2 deletion reduces tissue and systemic inflammation in DIO mice by inhibiting macrophage infiltration and M1 polarization.

Figure 3

Decreased tissue macrophage infiltration and M1 polarization in HFD-fed Fpr2−/− mice. Fpr2−/− and WT mice fed the HFD for 10 weeks were examined for serum levels of proinflammatory cytokines (n = 7–12 per group) (A); CD68-positive cells and CLSs in WAT adjusted with or without body weight (B and D); F4/80-positive cells in hepatic tissues adjusted with or without body weight (n = 6 per group) (C and D); mRNA expression of M1 and M2 macrophage markers in the epididymal WAT, liver, and gastrocnemius muscle (n = 9–12 per group) (E); and expression of M1 macrophage markers in peritoneal macrophages (pMac) (n = 5–10 per group) (F). Data are the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001. Images in B and C are representative immunostainings of WAT and liver sections with antibodies against CD68 and F4/80, respectively; the scale bar = 100 μm in B and 50 μm in C. CLSs and F4/80-positive cells per field were quantified using Image-Pro Plus software. The Student t test was used. Body weight–adjusted CLSs and F4/80-positive cells in D were computed as the residuals from the regression model, with CLSs or F4/80-positive cells as the independent variable and body weight as the dependent variable.

Figure 3

Decreased tissue macrophage infiltration and M1 polarization in HFD-fed Fpr2−/− mice. Fpr2−/− and WT mice fed the HFD for 10 weeks were examined for serum levels of proinflammatory cytokines (n = 7–12 per group) (A); CD68-positive cells and CLSs in WAT adjusted with or without body weight (B and D); F4/80-positive cells in hepatic tissues adjusted with or without body weight (n = 6 per group) (C and D); mRNA expression of M1 and M2 macrophage markers in the epididymal WAT, liver, and gastrocnemius muscle (n = 9–12 per group) (E); and expression of M1 macrophage markers in peritoneal macrophages (pMac) (n = 5–10 per group) (F). Data are the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001. Images in B and C are representative immunostainings of WAT and liver sections with antibodies against CD68 and F4/80, respectively; the scale bar = 100 μm in B and 50 μm in C. CLSs and F4/80-positive cells per field were quantified using Image-Pro Plus software. The Student t test was used. Body weight–adjusted CLSs and F4/80-positive cells in D were computed as the residuals from the regression model, with CLSs or F4/80-positive cells as the independent variable and body weight as the dependent variable.

Restoring Fpr2 Expression in Immune Cells in Fpr2-Deficient Mice Exacerbates DIO, Insulin Resistance, and Inflammation

Because deletion of Fpr2 improved inflammation in HFD-fed mice, we further investigated the contribution of Fpr2-expressing immune cells to metabolic dysregulation and inflammation in DIO mice through BMT. Bone marrows from Fpr2−/− mice were transplanted into irradiated WT recipients (Fpr2−/−-WT), or vice versa (WT-Fpr2−/−), and BMTs between WT mice (WT-WT) were performed as a control. The expression of Fpr2 mRNA in monocytes and neutrophils isolated from bone marrow was significantly decreased in Fpr2−/−-WT mice and was restored in WT-Fpr2−/− mice (Supplementary Fig. 2A). Food intake of WT-WT, Fpr2−/−-WT, and WT-Fpr2−/− mice was similar (Supplementary Fig. 2B). Compared with WT-WT mice, Fpr2−/−-WT mice were resistant to DIO (Fig. 4A and B), hyperglycemia, insulin resistance, dyslipidemia (Fig. 4C–F and Supplementary Fig. 2C and D), and systemic/tissue inflammation (Fig. 4G–L). The metabolic and inflammatory phenotypes of HFD-fed Fpr2−/−-WT mice were similar to those of HFD-fed Fpr2−/−mice. In contrast, WT-Fpr2−/− mice were prone to DIO (Fig. 4A and B), metabolic disturbances (Fig. 4C–F and Supplementary Fig. 2C and D), and tissue/systemic inflammation (Fig. 4G–L). Collectively, BMT studies demonstrate that Fpr2 in immune cells is deeply involved in DIO, metabolic disturbances, and inflammation.

Figure 4

Effect of deletion or restoration of Fpr2 in hematopoietic cells by BMT on HFD-induced insulin resistance and inflammation. WT mice received bone marrows from WT mice or Fpr2−/− mice (WT-WT, Fpr2−/−-WT), and Fpr2−/− mice received bone marrows from WT mice (WT-Fpr2−/−) fed the HFD for 9 weeks. Metabolic phenotypes were examined as follows: body weight (A), body composition (B), fed and fasted blood glucose levels (C), serum insulin levels (D), serum levels of TG and NEFA (E), and liver TG content (n = 7–12 per group) (F). Inflammatory parameters were measured as follows: serum levels of IL-6 and TNFα (n = 5–10 per group) (G), CLSs in epididymal WAT (H and J), F4/80-positive cells in hepatic tissues (n = 7–11 per group) (I and K), and mRNA expression of macrophage markers, and M1 and M2 macrophage markers in WAT, liver, and gastrocnemius muscle (n = 7–9 per group) (L). Data are the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 for the comparison between HFD-fed Fpr2−/−-WT and WT-WT mice; †P < 0.05 for the comparison between HFD-fed Fpr2−/−-WT and WT-Fpr2−/− mice. Scale bar = 100 μm in J and 50 μm in K. CLSs and F4/80-positive cells per field were quantified using Image-Pro Plus software. Two-way repeated-measures ANOVA was used in A and two-way ANOVA was used in B–I and L.

Figure 4

Effect of deletion or restoration of Fpr2 in hematopoietic cells by BMT on HFD-induced insulin resistance and inflammation. WT mice received bone marrows from WT mice or Fpr2−/− mice (WT-WT, Fpr2−/−-WT), and Fpr2−/− mice received bone marrows from WT mice (WT-Fpr2−/−) fed the HFD for 9 weeks. Metabolic phenotypes were examined as follows: body weight (A), body composition (B), fed and fasted blood glucose levels (C), serum insulin levels (D), serum levels of TG and NEFA (E), and liver TG content (n = 7–12 per group) (F). Inflammatory parameters were measured as follows: serum levels of IL-6 and TNFα (n = 5–10 per group) (G), CLSs in epididymal WAT (H and J), F4/80-positive cells in hepatic tissues (n = 7–11 per group) (I and K), and mRNA expression of macrophage markers, and M1 and M2 macrophage markers in WAT, liver, and gastrocnemius muscle (n = 7–9 per group) (L). Data are the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 for the comparison between HFD-fed Fpr2−/−-WT and WT-WT mice; †P < 0.05 for the comparison between HFD-fed Fpr2−/−-WT and WT-Fpr2−/− mice. Scale bar = 100 μm in J and 50 μm in K. CLSs and F4/80-positive cells per field were quantified using Image-Pro Plus software. Two-way repeated-measures ANOVA was used in A and two-way ANOVA was used in B–I and L.

Myeloid-Specific Deletion of Fpr2 Alleviates DIO, Insulin Resistance, and Inflammation

We generated Fpr2MKO mice to investigate which type of Fpr2-expressing immune cells participated in DIO, insulin resistance, and inflammation (Supplementary Fig. 3A). Consistent with the results from Fpr2−/− mice, myeloid-specific deletion of Fpr2 had no significant effect on metabolic phenotypes in mice fed the chow diet (Fig. 5A and Supplementary Fig. 3B–G) but alleviated obesity, insulin resistance, and glucose/lipid dysmetabolism and elevated body temperature, energy expenditure, and muscle thermogenesis in mice fed the HFD (Fig. 5 and Supplementary Fig. 3H–K). After adjustment with body weight, the improvement of glucose tolerance and insulin sensitivity in HFD-fed Fpr2MKO mice was slightly reduced but still statistically significant (Fig. 5G and I), indicating that myeloid Fpr2 deletion alleviates insulin resistance through mechanisms dependent and independent of body weight.

Figure 5

Myeloid-specific deletion of Fpr2 improves HFD-induced obesity and insulin resistance. Fpr2flox/flox and Fpr2MKO mice were fed control chow or the HFD for 10 weeks and examined for body weight (A), body composition (B), fed and fasted blood glucose levels (C), and serum insulin levels (n = 6–13 per group) (D); the DUR of 18F-fluorodeoxyglucose in the liver, WAT, and gastrocnemius muscle (E) (n = 5–7 per group); GTT and average area under the curve (AUC) adjusted with or without body weight (F and G); ITT and AUC adjusted with or without body weight (H and I); insulin stimulated Akt phosphorylation (p-) in liver, WAT, and gastrocnemius muscle (J); serum levels of TG and NEFA (K); serum levels of AST and ALT (L); and liver TG content (n = 6–13 per group) (M). The results represent the mean ± SEM, *P < 0.05, **P < 0.01, ***P < 0.001 for the comparison between Fpr2MKO and Fpr2flox/flox mice fed the HFD with or without insulin stimulation. Two-way repeated-measures ANOVA was used in A and F–I, and two-way ANOVA was used in J. The Student t test was used in BE and KM.

Figure 5

Myeloid-specific deletion of Fpr2 improves HFD-induced obesity and insulin resistance. Fpr2flox/flox and Fpr2MKO mice were fed control chow or the HFD for 10 weeks and examined for body weight (A), body composition (B), fed and fasted blood glucose levels (C), and serum insulin levels (n = 6–13 per group) (D); the DUR of 18F-fluorodeoxyglucose in the liver, WAT, and gastrocnemius muscle (E) (n = 5–7 per group); GTT and average area under the curve (AUC) adjusted with or without body weight (F and G); ITT and AUC adjusted with or without body weight (H and I); insulin stimulated Akt phosphorylation (p-) in liver, WAT, and gastrocnemius muscle (J); serum levels of TG and NEFA (K); serum levels of AST and ALT (L); and liver TG content (n = 6–13 per group) (M). The results represent the mean ± SEM, *P < 0.05, **P < 0.01, ***P < 0.001 for the comparison between Fpr2MKO and Fpr2flox/flox mice fed the HFD with or without insulin stimulation. Two-way repeated-measures ANOVA was used in A and F–I, and two-way ANOVA was used in J. The Student t test was used in BE and KM.

Myeloid-specific deletion of Fpr2 in HFD-fed mice also relieved systemic and tissue inflammation (Fig. 6A–C) and reduced the expression of M1 macrophage markers in WAT, liver, and muscle (Fig. 6E). The FACS analysis of SVCs from WAT of these mice consistently revealed a lower percentage of M1 macrophages and a comparable percentage of M2 macrophages compared with those of HFD-fed WT mice (Fig. 6F and G). The reduction of CLSs in adipose tissue and macrophage infiltration in hepatic tissues by myeloid Fpr2 deletion remained after adjustment with body weight (Fig. 6D). Collectively, these data indicate that myeloid Fpr2 plays an important role in DIO, insulin resistance, glucose/lipid dysmetabolism, and inflammation.

Figure 6

Myeloid Fpr2 deficiency reduces tissue macrophage infiltration and M1 polarization. Fpr2flox/flox and Fpr2MKO mice fed the HFD for 10 weeks were examined for serum levels of IL-6 and TNF-α (A); CD68-positive cells and CLSs in epididymal WAT adjusted with or without body weight (B and D); F4/80-positive cells in hepatic tissues adjusted with or without body weight (C and D); mRNA expression of macrophage markers, M1 and M2 macrophage markers in WAT, liver, and gastrocnemius muscle (E) (n = 7–12 per group); and flow cytometric analysis of M1 (CD45+F4/80+CD11b+CD11c+CD206) and M2 (CD45+F4/80+CD11b+CD11cCD206+) macrophages among adipose tissue macrophages (CD45+F4/80+CD11b+) (n = 5–7 per group) (F and G). Data are the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001. Images in B and C are representative immunostainings of WAT and liver sections with antibodies against CD68 and F4/80, respectively. The scale bar = 100 μm in B and 50 μm in C. CLSs and F4/80-positive cells per field were quantified using Image-Pro Plus software. The Student t test was used. Body weight–adjusted CLSs and F4/80-positive cells in D were computed as the residuals from the regression model, with CLSs or F4/80-positive cells as the independent variable and body weight as the dependent variable.

Figure 6

Myeloid Fpr2 deficiency reduces tissue macrophage infiltration and M1 polarization. Fpr2flox/flox and Fpr2MKO mice fed the HFD for 10 weeks were examined for serum levels of IL-6 and TNF-α (A); CD68-positive cells and CLSs in epididymal WAT adjusted with or without body weight (B and D); F4/80-positive cells in hepatic tissues adjusted with or without body weight (C and D); mRNA expression of macrophage markers, M1 and M2 macrophage markers in WAT, liver, and gastrocnemius muscle (E) (n = 7–12 per group); and flow cytometric analysis of M1 (CD45+F4/80+CD11b+CD11c+CD206) and M2 (CD45+F4/80+CD11b+CD11cCD206+) macrophages among adipose tissue macrophages (CD45+F4/80+CD11b+) (n = 5–7 per group) (F and G). Data are the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001. Images in B and C are representative immunostainings of WAT and liver sections with antibodies against CD68 and F4/80, respectively. The scale bar = 100 μm in B and 50 μm in C. CLSs and F4/80-positive cells per field were quantified using Image-Pro Plus software. The Student t test was used. Body weight–adjusted CLSs and F4/80-positive cells in D were computed as the residuals from the regression model, with CLSs or F4/80-positive cells as the independent variable and body weight as the dependent variable.

Adipocyte-Secreted Saa3 Induces Macrophage Chemotaxis Through Fpr2

We examined whether obese adipose tissue produced Fpr2 ligands to induce macrophage infiltration. We found that the chemotactic activity of the obese adipose tissue lysate (OATL) to BMDMs is higher than that of lean adipose tissue lysate. The chemotactic response of Fpr2−/− BMDMs to OATL is lower than that of WT BMDMs (Fig. 7A). These results indicate that obese adipose tissue contains a higher level of Fpr2 ligands than lean adipose tissue.

Figure 7

Saa3 induces macrophage migration via Fpr2. WT and Fpr2−/− BMDM migration in response to adipose tissue lysate from chow-fed mice (lean adipose tissue lysate [LATL]) and HFD-fed mice (OATL) (n = 4–6 per group) (A), or 10 µg/mL Saa3 (n = 8–10 per group) (B). Adipocytes differentiated from 3T3-L1 cells were transfected with siSaa3 or control siRNA (Scr) and then examined for gene expression (C) and the chemotactic activity of the culture medium to WT and Fpr2−/− BMDMs (n = 4–6 per group) (D). E: Proinflammatory cytokine expression in BMDMs induced by 100 µg/mL Saa3 for 6 h (n = 3 per group). F: Saa3 mRNA expression in the epididymal WAT of HFD-fed Fpr2−/−, Fpr2MKO and their corresponding control mice, as well as mice with BMT (n = 7–12 per group). Data are the mean ± SD (AE) or the mean ± SEM (F). *P < 0.05, **P < 0.01, ***P < 0.001, †P < 0.05, †††P < 0.001. Two-way ANOVA was used in A, B, D, and the right panel of F. The Student t test was used in C and the left and middle panels of F.

Figure 7

Saa3 induces macrophage migration via Fpr2. WT and Fpr2−/− BMDM migration in response to adipose tissue lysate from chow-fed mice (lean adipose tissue lysate [LATL]) and HFD-fed mice (OATL) (n = 4–6 per group) (A), or 10 µg/mL Saa3 (n = 8–10 per group) (B). Adipocytes differentiated from 3T3-L1 cells were transfected with siSaa3 or control siRNA (Scr) and then examined for gene expression (C) and the chemotactic activity of the culture medium to WT and Fpr2−/− BMDMs (n = 4–6 per group) (D). E: Proinflammatory cytokine expression in BMDMs induced by 100 µg/mL Saa3 for 6 h (n = 3 per group). F: Saa3 mRNA expression in the epididymal WAT of HFD-fed Fpr2−/−, Fpr2MKO and their corresponding control mice, as well as mice with BMT (n = 7–12 per group). Data are the mean ± SD (AE) or the mean ± SEM (F). *P < 0.05, **P < 0.01, ***P < 0.001, †P < 0.05, †††P < 0.001. Two-way ANOVA was used in A, B, D, and the right panel of F. The Student t test was used in C and the left and middle panels of F.

Saa3 upregulation has been reported in adipose tissue of obese mice (32). We found that Saa3 significantly induced WT BMDMs chemotaxis but had less effect on Fpr2−/− BMDMs (Fig. 7B). The conditioned medium (CM) from Saa3-knockdown adipocytes had lower chemotactic activity to WT BMDMs than CM from control adipocytes (Fig. 7C and D). The chemotactic response of Fpr2−/− BMDMs to the CM from Saa3-knockdown adipocytes is lower than that of WT BMDMs (Fig. 7D). These results indicate that adipocyte-secreted Saa3 and other factors induce BMDM migration through Fpr2. We found that Saa3 significantly induced Tnfα, Il-1β, Il-6, and Ccl2 expression in an Fpr2-independent manner (Fig. 7E). The expression of Saa3 in WAT was significantly decreased in HFD-fed Fpr2−/−, Fpr2−/−-WT, and Fpr2MKOmice but was recovered in Fpr2−/− mice who received BMTs from WT mice (Fig. 7F).

Fpr2 Promotes Macrophage M1 Polarization and Proinflammatory Cytokine Expression In Vitro

We further investigated the role of Fpr2 in macrophage M1 polarization and proinflammatory cytokine expression by stimulating BMDMs with LPS. Data showed that deletion of Fpr2 in BMDMs significantly reduced the expression of M1 macrophage-related genes (Il-1β, Il-6, Ccl2, and Tnfα) and proinflammatory cytokines (IL-6, CCL2, and TNF-α) (Fig. 8A and B). NF-κB and MAP kinases are important signaling molecules mediating proinflammatory cytokine expression and macrophage polarization. We found that Fpr2 deletion markedly inhibited NF-κB p65 and p38 phosphorylation and slightly inhibited ERK and JNK phosphorylation induced by LPS (Fig. 8C). The Fpr2 antagonist WRW4 consistently suppressed LPS-induced MAP kinases and NF-κB activation as well as M1 macrophage-related gene expression in BMDMs (Supplementary Fig. 4B and C). In contrast, Fpr2 overexpression enhanced LPS-stimulated signal transduction and proinflammatory cytokine expression (Fig. 8D–F). Fpr2 deficiency in BMDMs also reduced LPS-induced phosphorylation of TAK1 (Fig. 8C), an upstream molecule of NF-κB and MAP kinases in the LPS-stimulated signaling pathway (33). We also examined the involvement of Fpr2 in macrophage M1 polarization induced by palmitate and IFN-γ. Fpr2 deletion in BMDMs reduced M1 macrophage–related gene expression and phosphorylation of NF-κB p65 and MAP kinases in response to palmitate (Fig. 8G–I) and reduced STAT1 phosphorylation by IFN-γ (Supplementary Fig. 4D). These results indicate that Fpr2 deletion suppresses macrophage polarization toward an M1 phenotype via inhibiting TAK1-MAP kinase/NF-κB and STAT1-related signaling pathways.

Figure 8

Fpr2 mediates macrophage M1 polarization. WT and Fpr2−/− BMDMs were treated with 100 ng/mL LPS and examined for expression of M1 macrophage markers (treated for 6 h) (A), production of proinflammatory cytokines in the culture supernatant (treated for 9 h) (B), and phosphorylation of NF-κB p65, MAP kinases (ET, exposure time), and TAK1 (treated for 30 min) (C) (n = 3 per group). DF: BMDMs infected with Fpr2 expressing lentiviruses (Lenti-Fpr2) or control viruses (Lenti-NC) and examined for the expression of M1 macrophage markers (iNos, inducible nitric oxide synthase) (D) and phosphorylation (p-) of NF-κB p65 and MAP kinases after stimulation with 100 ng/mL LPS for 30 min (E and F). GI: BMDMs were stimulated with 400 μmol/L palmitate (Pal) (dissolved in BSA buffer) and examined for expression of M1-related proinflammatory cytokines (stimulated for 3 h) (G), and phosphorylation of NF-κB p65 and MAP kinases (treated for 1 h) (H and I) (n = 3 per group). Values shown represent the mean ± SD. *P < 0.05, **P < 0.01, ***P < 0.001. Two-way ANOVA was used in A, B, D, F, G, and I, and two-way repeated-measures ANOVA was used in C. Veh, vehicle.

Figure 8

Fpr2 mediates macrophage M1 polarization. WT and Fpr2−/− BMDMs were treated with 100 ng/mL LPS and examined for expression of M1 macrophage markers (treated for 6 h) (A), production of proinflammatory cytokines in the culture supernatant (treated for 9 h) (B), and phosphorylation of NF-κB p65, MAP kinases (ET, exposure time), and TAK1 (treated for 30 min) (C) (n = 3 per group). DF: BMDMs infected with Fpr2 expressing lentiviruses (Lenti-Fpr2) or control viruses (Lenti-NC) and examined for the expression of M1 macrophage markers (iNos, inducible nitric oxide synthase) (D) and phosphorylation (p-) of NF-κB p65 and MAP kinases after stimulation with 100 ng/mL LPS for 30 min (E and F). GI: BMDMs were stimulated with 400 μmol/L palmitate (Pal) (dissolved in BSA buffer) and examined for expression of M1-related proinflammatory cytokines (stimulated for 3 h) (G), and phosphorylation of NF-κB p65 and MAP kinases (treated for 1 h) (H and I) (n = 3 per group). Values shown represent the mean ± SD. *P < 0.05, **P < 0.01, ***P < 0.001. Two-way ANOVA was used in A, B, D, F, G, and I, and two-way repeated-measures ANOVA was used in C. Veh, vehicle.

In the current study, we found that Fpr2 was highly expressed in WAT of obese mice models, especially in the SVCs of WAT. Further studies of systemic Fpr2 deletion, BMTs, and myeloid-specific Fpr2 deletion in mice demonstrated that myeloid Fpr2 plays critical roles in DIO, metabolic disturbances, and inflammation.

The body weight change is associated with an imbalance between food intake and energy expenditure. Deletion of Fpr2 systemically or in myeloid cells in mice fed the HFD did not affect food intake but increased energy expenditure and body temperature. That BAT and skeletal muscle are important players in regulating thermogenesis is well known. Our results showed that Fpr2 deletion in obese mice increased fatty acid oxidation–related and thermogenic gene expression in muscle. Skeletal muscle is the largest organ in the body and is a major determinant of basal metabolic rate. An increase of energy expenditure in muscle through nonshivering thermogenesis can substantially affect whole-body metabolism and weight gain (34,35). Therefore, the increase of lean mass percentage and thermogenic gene expression in skeletal muscle may contribute to higher energy expenditure and lower body weight gain in HFD-fed Fpr2−/− and Fpr2MKO mice.

The mechanisms underlying the regulation of thermogenesis by Fpr2 in the muscle of obese mice is not clear. Studies by systemic knockout of TNF-α receptor or hypothalamic immunoneutralization of TNF-α in rodents indicate that obesity-related inflammation impairs whole-body energy expenditure, mitochondrial biogenesis, ATP production, and thermogenesis in BAT and muscle (36,37). Whether Fpr2 deficiency in DIO mice promotes muscle thermogenesis through alleviating inflammation remains to be further investigated.

In addition to energy expenditure, lipid accumulation in the liver is also associated with the changes of body fat mass (38). Therefore, the alleviation of hepatic steatosis in HFD-fed Fpr2−/− and Fpr2MKO mice may also contribute to lower fat mass. Body weight changes and inflammation have been reported to influence insulin sensitivity (39,40). Our studies showed that adjustment with body weight only slightly reduced the protective effect of Fpr2 deletion on HFD-induced insulin resistance and tissue inflammation, indicating that Fpr2 deficiency improves insulin resistance partly through reducing body weight gain and mainly through inhibiting inflammation.

Macrophage infiltration and polarization toward an M1 phenotype are the main drivers of insulin resistance in the context of obesity (3). We found that myeloid Fpr2 played critical roles in macrophage accumulation in the metabolic tissues of DIO mice. This conclusion is supported by the following evidence: First, Fpr2 was highly expressed in adipose tissue SVCs of DIO mice (Fig. 1A). Second, studies of Fpr2 deletion in three independent experiments demonstrated that myeloid Fpr2 contributed to macrophage infiltration in the metabolic tissues (Figs. 3B–D, 4H–K, and 6B–D). Third, obese adipose tissues contained higher levels of chemotactic ligands, which induced macrophage migration.

Our study for the first time demonstrated that adipocyte-released Saa3 induced macrophage chemotaxis in an Fpr2-dependent manner. Because Saa3 could stimulate macrophage migration in a short time (2 h) (Fig. 7B), we propose that Saa3 may be a chemotactic ligand for Fpr2. The direct interaction between Saa3 and Fpr2 awaits further verification. In addition, we found that systemic, bone marrow-, and myeloid-specific Fpr2 deletion in HFD-fed mice significantly reduced the expression of Saa3 in WAT and that the reduction of Saa3 was reversed after recovering Fpr2 expression in bone marrow cells (Fig. 7F). In obese mice, Saa3 has been reported to be highly expressed in the adipose tissue (41) and contributes to macrophage infiltration (32). Saa3 is upregulated by palmitate acid, glucose, LPS, TNF-α, and IL-1β in adipocytes (4245). Our studies showed that Saa3 induced proinflammatory cytokines expression in macrophages independent of Fpr2 (Fig. 7E). We thus hypothesize that Fpr2 deficiency alleviates DIO and inflammation, which results in the decrease of Saa3 in WAT and in turn blocks its contribution to WAT inflammation. Mitochondrial peptides released from ruptured cells have been reported to induce phagocyte migration and activation through Fpr2 (46). Therefore, Fpr2 ligands released by dead adipocytes and other cells of obese mice may also contribute to macrophage infiltration in metabolic tissues.

In addition to revealing the contribution of Fpr2 to obesity and macrophage infiltration in metabolic tissues of DIO mice, another important finding of our study is that Fpr2 could promote macrophage M1 polarization in DIO mice, which was supported by in vivo studies with Fpr2−/− and Fpr2MKO mice as well as mice with BMTs. This conclusion was further supported by in vitro studies with BMDMs stimulated by LPS, palmitate acid, and IFN-γ. First, Fpr2 expression in macrophages was upregulated by LPS (Supplementary Fig. 4A) and IFN-γ (47). Second, inhibition of Fpr2 by antagonist reduced LPS-stimulated proinflammatory cytokine expression and NF-κB/MAP kinase phosphorylation (Supplementary Fig. 4B and C). Third, gain- and off-Fpr2 studies showed that Fpr2 modulated the expression of M1 macrophage marker genes and proinflammatory cytokines and the activation of proinflammatory signaling pathways by LPS, palmitate, and IFN-γ (Fig. 8 and Supplementary Fig. 4D).

Jablonski et al. (48) recently reported that Fpr2 expression was upregulated during macrophage M1 differentiation but downregulated during M2 differentiation. They proposed Fpr2 as a new marker for M1 macrophages (48). These data support our finding that Fpr2 is an important regulator in macrophage M1 polarization. In addition, our studies showed that Fpr2 deletion had neither effect on hemogram profile and M2 macrophages in metabolic tissues of obese mice (Supplementary Fig. 5A and B and Figs. 3E, 4L, and 6E) nor effect on macrophage differentiation in vitro (Supplementary Fig. 5C), indicating that the effect of Fpr2 deficiency on macrophage M1 polarization is not mediated by altering macrophage differentiation and M2 polarization. Additional studies are needed to investigate the cross talk between Fpr2 and signaling pathways involved in macrophage M1 polarization induced by LPS, palmitate, and IFN-γ.

Finally, we checked whether other mechanisms are involved in the improvement of insulin resistance by Fpr2 deletion in obese mice. In vitro studies with 3T3L1 adipocytes showed that Fpr2 agonist and antagonist had no significant effect on insulin-induced phosphorylation of InsR, Akt, and GSK3β (Supplementary Fig. 6A). Insulin stimulation of primary adipocytes isolated from WT and Fpr2 KO mice consistently induced comparable phosphorylation of these proteins (Supplementary Fig. 6B). These results indicate that there is no cross talk between signalings of Fpr2 and insulin in adipose tissue. In addition, we found that depletion of gut microbiota with antibiotics had no effect on the improvement of obesity and insulin resistance by Fpr2 deficiency in HFD-fed mice (Supplementary Fig. 7), indicating that Fpr2 deletion regulates glucose and lipid metabolism in obese mice independent of gut microbiota.

Taken together, our study demonstrates that myeloid Fpr2 plays critical roles in DIO and its related complications by modulating energy expenditure as well as inflammation mediated by macrophage accumulation and M1 polarization in metabolic tissues. Our findings indicate that myeloid Fpr2 is a potential therapeutic target against obesity and related metabolic disorders. LXA4 and ANXA1, two anti-inflammatory ligands of Fpr2, have been reported to attenuate obesity-related inflammation and metabolic diseases (1921). Clarifying whether these two molecules inhibit obesity-related inflammation via activating the bias signaling of Fpr2 will be helpful for developing strategies against obesity-related metabolic disorders.

Acknowledgments. The authors thank Shengzhong Duan (Shanghai Jiao Tong University, Shanghai, China) for providing LysM-Cre mice and the animal facility staff at the Shanghai Institute of Nutrition and Health, Shanghai Institute for Biological Sciences, Chinese Academy of Science, for their support.

Funding. This study was supported by grants from the National Key Research and Development Program of China (2017YFC1601702) and the National Natural Science Foundation of China (31671232).

The funders had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. X.C. researched the data and wrote and edited the manuscript. S.Z. contributed to the experimental design. T.Z., P.Y., M.Y., H.M., N.L., F.M., and S.C. researched the data. J.M.W. provided the Fpr2flox/flox and Fpr2−/− mice and contributed to discussion. R.D.Y. and Y.Li contributed to discussion. Y.Le directed the project, contributed to discussions, and wrote, reviewed, and edited the manuscript. Y.Le is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

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