Abstract
The adipokine leptin is primarily produced by white adipose tissue (AT) and is a potent monocyte/macrophage chemoattractant in vitro. The long form of the leptin receptor (LepR) is required for monocyte/macrophage chemotaxis towards leptin. In this study, we examined the effects of haematopoietic LepR as well as LepR with C–C chemokine receptor 2 (CCR2) deficiency (double knockout (DKO)) on macrophage recruitment to AT after two different periods of high fat diet (HFD) feeding. Briefly, 8-week-old C57BL/6 mice were transplanted with bone marrow (BM) from Lepr+/+, Lepr−/− or DKO donors (groups named BM-Lepr+/+, BM-Lepr−/− and BM-DKO respectively), and were placed on an HFD for 6 or 12 weeks. At the end of the study, macrophage infiltration and the inflammatory state of AT were evaluated by real-time RT-PCR, histology and flow cytometry. In addition, glucose and insulin tolerance were assessed at both time points. Our results showed no differences in macrophage accumulation or AT inflammatory state between the BM-Lepr+/+ and BM-Lepr−/− mice after 6 or 12 weeks of HFD feeding; any effects observed in the BM-DKO were attributed to the haematopoietic deficiency of CCR2. In addition, no changes in glucose or insulin tolerance were observed between groups after either period of HFD feeding. Our findings suggest that although leptin is a potent chemoattractant in vitro, haematopoietic LepR deficiency does not affect macrophage accumulation in AT in early to moderate stages of diet-induced obesity.
Introduction
Accumulation of inflammatory macrophages in adipose tissue (AT) during obesity has been shown to correlate with AT inflammation and subsequent insulin resistance (IR; Weisberg et al. 2003, Xu et al. 2003). In addition, the polarisation of the infiltrating macrophages is known to be an important factor in the inflammatory state of AT (Lumeng et al. 2007, 2008). Thus, understanding the mechanism(s) of macrophage recruitment to AT could lead to the development of therapies against IR and type 2 diabetes. Adipokines, which are hormones secreted from AT, could act as chemoattractants and contribute to macrophage recruitment to AT. The adipokine leptin has emerged as a strong candidate, given its known role in immune regulation (La Cava & Matarese 2004). Leptin is a 16 kDa soluble protein encoded by the Ob gene and is expressed primarily by white AT, such that circulating levels of leptin are positively correlated with levels of adiposity and AT macrophage accumulation (Halaas et al. 1995, Coenen et al. 2007).
Leptin acts primarily through the long form of the leptin receptor (LepR) and plays an important role in weight regulation through its action in the hypothalamus (Leshan et al. 2006). Mice that lack either leptin or the LepR are characterised by severe hyperphagia and morbid obesity, as well as other hormonal abnormalities (Robertson et al. 2008). Previous studies have shown that leptin promotes neutrophil and monocyte chemotaxis as well as the activation of monocytes/macrophages leading to secretion of pro-inflammatory cytokines and the up-regulation of inducible nitric oxide synthase (iNOS; Zarkesh-Esfahani et al. 2001, Dixit et al. 2003, Gruen et al. 2007). We have shown that leptin is a potent monocyte chemoattractant in vitro and that this chemotaxis is LepR-dependent (Gruen et al. 2007). In addition, we have shown that expression of Emr1 (the gene for F4/80) in AT, an indicator of macrophage infiltration, is positively correlated with circulating levels of leptin during obesity (Supplementary Figure 1, see section on supplementary data given at the end of this article; r2=0.6973; P<0.0001; Coenen et al. 2007). Given that leptin is primarily secreted from AT and that its circulating levels are positively correlated with increases in macrophage infiltration, we hypothesised that leptin could act as a monocyte chemoattractant that regulates macrophage recruitment to AT during obesity. We designed a bone marrow transplant (BMT) study to determine if haematopoietic deficiency of the functional long form of the LepR, which we have shown to be required for monocyte chemotaxis towards leptin in vitro (Gruen et al. 2007), would lead to decreases in macrophage recruitment to AT during high fat diet (HFD) feeding. We transplanted bone marrow from Lepr−/− and double knockout (DKO) mice into wild type (WT) recipients and placed these mice on an HFD for 6 or 12 weeks. These time points were chosen because of previously shown kinetics of macrophage recruitment to AT (Nishimura et al. 2009). We examined macrophage infiltration into AT by flow cytometry, histology and real-time RT-PCR. Our data provides evidence that haematopoietic deficiency of the LepR does not affect macrophage recruitment to AT or insulin sensitivity during HFD-induced obesity.
Materials and Methods
Mice and diets
All animal care and experimental procedures were performed with approval from the Institutional Animal Care and Use Committee of Vanderbilt University. The Lepr+/+, Lepr−/− and Ccr2−/− mice were purchased from Jackson Laboratories (Bar Harbor, ME, USA) and were on a C57BL/6 background. DKO donor mice were generated by crossing Ccr2−/− with Lepr+/− mice. All studies were performed with 8-week-old male donor and recipient mice. Mice were fed ad libitum and given free access to water. Mice were kept on antibiotic water for 1 week prior to BMT until 4 weeks post-BMT. Mice were then placed on a 60% fat diet from Research Diets, Inc. New Brunswick, NJ, USA, (D12492) with a calorie density of 5.24 kcal/g; starting at 4 weeks post-BMT for a total of 6 or 12 weeks (Fig. 1). Because we were only interested in changes between the two genotypes and not the diet effects we did not use chow-fed mice in the original study; however, we performed an additional study in which we showed that HFD-fed Lepr+/+ mice have increased macrophage infiltration and become insulin resistant in comparison to Lepr+/+ mice fed a chow diet (Supplementary Figure 2, see section on supplementary data given at the end of this article). Thus, we had the ability to identify any changes in these parameters in mice fed HFD for only 6 weeks.
Study design. C57BL/6 mice were transplanted with either Lepr+/+, Lepr−/− or DKO (Lepr−/−; Ccr2−/−) bone marrow at 8 weeks of age. After 4 weeks mice were placed on a 60% diet (HFD) for 6 or 12 weeks (n=10 per group). GTTs and ITTs were performed after 6 and 12 weeks. Mice were sacrificed after the 6 or 12 week period for analysis of AT macrophage infiltration and inflammation.
Citation: Journal of Endocrinology 212, 3; 10.1530/JOE-11-0338
Bone marrow transplantation
Bone marrow cells were collected from Lepr+/+, Lepr−/− and DKO donor mice and were injected into the retro-orbital venous plexus of lethally irradiated WT recipient mice. Complete reconstitution of the desired bone marrow was confirmed by PCR of splenic cells using primers specific for the db allele and the Neo cassette insertion in the Ccr2−/− deficient mice (data not shown). The BM-Lepr+/+ controls in this study were also used as negative controls for a different study performed simultaneously (Gutierrez et al. 2011). In addition, recipients of Ccr2−/− marrow (BM-Ccr2−/−) were included in the previous paper. There were no significant differences in any parameter studied between the Ccr2−/− and DKO recipients. Thus, only the DKO mice are shown in the current report.
Body weight, body composition and food intake
Body weight and food intake were measured weekly for the duration of the study. Total lean fat and free fluid mass were measured by nuclear magnetic resonance (NMR) using a Bruker Minispec (Woodlands, TX, USA) in the Vanderbilt Mouse Metabolic Phenotyping Centre (MMPC) at baseline, 6 and 12 weeks after placement on an HFD.
Insulin and glucose tolerance tests
Mice were fasted for 5 h and baseline glucose levels were measured using a LifeScan One Touch Ultra Glucometer (Johnson & Johnson, Northridge, CA, USA) via the tail vein. Mice were then injected i.p. with either 0.4 U (6 weeks) and 0.5 U (12 weeks) insulin per kilogram of lean mass; or 2 g (6 weeks) and 1 g (12 weeks) dextrose per kilogram of lean mass. Glucose levels were then assessed at 15, 30, 45, 60, 90 and 150 min post-injection.
Plasma collection and measurements
Mice were fasted for 5 h and bled from the retro-orbital venous plexus using heparinised collection tubes. Plasma was separated by centrifugation and stored at −80 °C. Plasma insulin levels were measured using an ELISA kit from Millipore, Inc. (Billerica, MA, USA). Leptin levels were assessed at the Vanderbilt University Hormone Core.
Real-time RT-PCR
RNA extraction was performed with an RNeasy kit (Qiagen) according to the manufacturer's instructions. The iScript cDNA synthesis kit (Bio-Rad) was used for reverse transcriptase reactions. Real-time RT-PCR experiments were performed using an iQ5 thermal cycler. The reactions were carried out using IQ Supermix (Bio-Rad) and FAM-conjugated Assay-on-Demand (Applied Biosystems, Foster City, CA, USA) primer/probe sets normalized to 18S. The following genes were assessed: 18S (4352930E), Emr1 (Mm00802530_m1), Arg1 (Mm01190441_g1), Cd68 (Mm00839636_g1), Tnf (Mm00443258_m1), Nos2 (Mm00440502_m1), Cd163 (Mm0118511), Mgl1 (Mm00546125_g1), Mgl2 (Mm00460844_m1), Il6 (Mm00446190_m1) and Il1b (Mm9999061_mH). The data was analysed using the Pfaffl method (Pfaffl 2001) and presented as expression relative to the BM-Lepr+/+ controls at each respective time point.
Histology
Epididymal AT was harvested from mice, weighed and a portion was fixed overnight in 10% (v/v) formalin, transferred to 70% (v/v) ethanol and paraffin embedded. Tissue was cut into 7 μm sections and stained with 0.1% (w/v) toluidine blue O (TBO) solution (Newcomer Supply, Middleton, WI, USA) as previously described (Coenen et al. 2007).
Stromal vascular fraction separation
Epididymal AT pads were excised and minced in 3 ml of 0.5% (w/v) BSA in PBS and placed on ice. Subsequently, 3 ml of 2 mg/ml collagenase was added to the minced fat and incubated at 37 °C for 20 min with shaking. The cell suspension was filtered through a 100 μm filter and then spun at 300 g for 10 min to separate floating adipocytes from the stromal vascular fraction (SVF) pellet. The SVF pellet was then resuspended in 3 ml ACK lysis buffer and incubated at room temperature (RT) for 5 min for red blood cell lysis, followed by dilution in 10 ml PBS. The cells were then washed twice with PBS.
Flow cytometry
Stromal vascular and white blood cells were first incubated with Fc block for 5 min at RT, followed by incubation for 30 min at 4 °C with fluorophore-conjugated antibodies: F4/80-APC (eBioscience, San Diego, CA, USA), Ly6C-FITC (BD Biosciences, Franklin Lakes, NJ, USA), Ly6G-PE (BD Biosciences) and CD11b-APC-Cy7 (BD Bioscience). DAPI (0.2 μg/ml) was added to each sample as a viability stain 10 min before flow cytometry analysis. Stromal vascular cells (SVCs) were processed on an MACS Quant analyzer (Miltenyi Biotech, Bergisch Gladbach, Germany). Blood cells were processed using a 5 Laser LSRII machine in the Vanderbilt Flow Cytometry Core. All data was analysed using FlowJo Software version 7.6.1 (Ashland, OR, USA).
Statistical analysis
GraphPad Prism 4.0 software, La Jolla, CA, USA was used for all statistical analyses. Data was analysed using one-way ANOVA to test differences between the three experimental groups and two-way ANOVA to compare measurements with two different variables (i.e. glucose and insulin tolerance tests (GTTs and ITTs)). Outliers were excluded from the data for each individual parameter if outside the range of the mean±2 s.d. and P≤0.05 was considered significant.
Results
Metabolic profile
Male C57BL/6 mice were transplanted with Lepr+/+, Lepr−/− or DKO marrow followed by HFD feeding for 6 or 12 weeks as described in the Materials and Methods section (Fig. 1). There were no differences in body weight (Fig. 2A) or food intake (data not shown) between the groups at any point during the study. Likewise, there were no differences in adiposity among genotypes at any time although all groups increased in adiposity after 6 and 12 weeks of HFD feeding (Fig. 2B). Epididymal fat pad weight (Fig. 2C) and circulating leptin levels (Fig. 2D) were not different among genotypes at any time point. In accordance with the NMR adiposity data, epididymal AT weight and leptin levels were generally elevated between the 6 and 12 week time points for each group (Fig. 2C and D). Glucose and insulin tolerance tests were performed after 6 weeks (Fig. 3A and C) and 12 weeks (Fig. 3B and D) of HFD feeding. All groups responded similarly to the intraperitoneal glucose and insulin challenges after both periods of HFD feeding. In general, fasting insulin levels were increased for all groups between the baseline measurements and 12 weeks of HFD feeding (Fig. 3E). Fasting glucose levels were significantly increased (P<0.001) in all groups between the pre-HFD and both the 6 and 12 week time points (Fig. 3F).
Weight gain was similar among groups upon HFD feeding. (A) Average weekly body weight of BM-Lepr+/+, BM-Lepr−/− and BM-DKO mice pre- and post-HFD. (B) Percent adiposity (total fat tissue divided by total body weight) determined by NMR. (C) Epididymal AT weight. (D) Circulating plasma leptin levels assessed by ELISA. Data are presented as mean±s.e.m. For pre-HFD up to 6 weeks post-HFD, n=20 mice per group. For 12 weeks post-HFD, n=9–10 mice per group.
Citation: Journal of Endocrinology 212, 3; 10.1530/JOE-11-0338
All groups have similar glucose and insulin tolerance. Glucose tolerance tests were performed by injecting mice i.p. with dextrose at a concentration of 2 g (6 weeks) or 1 g (12 weeks) per kilogram of lean mass after a 5 h morning fast. Glucose tolerance curves for all groups after (A) 6 weeks and (B) 12 weeks of HFD feeding. Insulin tolerance tests were performed by injecting mice i.p. with 0.4 U (6 weeks) or 0.5 U (12 weeks) human insulin per kilogram of lean mass after a 5 h morning fast. Insulin tolerance curves for all groups after (C) 6 weeks and (D) 12 weeks of HFD feeding. (E) Fasting insulin levels were measured by ELISA at the pre-HFD, 6 and 12 weeks time points. (F) Fasting glucose levels were assessed at the pre-HFD, 6 and 12 weeks time points. Data are presented as mean±s.e.m. For GTT and ITT data, n=9–10 mice per group for 6 week data and n=5 mice per group for 12 week data. For fasting insulin and glucose data, n=8–10 per group. ***P<0.001 for all groups between Pre-HFD and 6 or 12 weeks.
Citation: Journal of Endocrinology 212, 3; 10.1530/JOE-11-0338
Blood monocytes
Two phenotypic and functional subsets of mature blood monocytes have been described: Ly6Clo and Ly6Chi (Geissmann et al. 2010). The Ly6Chi circulating monocyte subset is thought to be the source of recruited inflammatory macrophages to inflamed tissues. The circulating levels of blood monocytes and neutrophils were assessed by flow cytometry after 12 weeks of HFD feeding by first gating on all live CD11b+ cells. The BM-Lepr−/− mice had a ∼40% (P<0.05) reduction in circulating Ly6Chi monocytes, and BM-DKO mice had a ∼90% (P<0.001) reduction, when compared with the BM-Lepr+/+ controls (Fig. 4A, B, C and D). No differences were observed in the Ly6Clo monocyte subset or the percentage of circulating neutrophils between groups (Fig. 4E and F).
BM-Lepr−/− mice have decreased circulating ‘classical’ monocytes. Blood cells were isolated from 12 week HFD-fed recipient mice and analysed by flow cytometry. Representative plots showing expression of Ly6C after gating on all CD11b+ live cells. (A) BM-Lepr+/+, (B) BM-Lepr−/− and (C) BM-DKO. Quantification of the percentage of (D) CD11b+Ly6Chi, (E) CD11b+Ly6Clo and (F) CD11b+Ly6CmidLy6Ghi cells in all groups (mean±s.e.m.; n=4–5 mice per group). *P<0.05 between BM-Lepr+/+ and BM-Lepr−/− mice. ***P<0.001 between BM-Lepr+/+ and BM-DKO mice. ^P<0.05 between BM-Lepr−/− and BM-DKO mice.
Citation: Journal of Endocrinology 212, 3; 10.1530/JOE-11-0338
Macrophage accumulation in AT
The accumulation of macrophages in AT after 6 and 12 weeks of HFD feeding was evaluated by histology, real-time RT-PCR and flow cytometry. TBO staining showed that there were no overt differences in overall immune infiltration between groups after either period of HFD feeding (Fig. 5A, B, C, D, E and F). Real-time RT-PCR gene expression analysis in total AT for the macrophage markers Emr1 and Cd68 showed no differences between groups after 6 weeks (Fig. 5G) or 12 weeks (Fig. 5H) of HFD feeding.
Histology and real-time RT-PCR analysis of macrophages in AT. Representative AT TBO images of (A) BM-Lepr+/+, (B) BM-Lepr−/− and (C) BM-DKO mice after 6 weeks of HFD feeding. Representative AT TBO images of (D) BM-Lepr+/+, (E) BM-Lepr−/− and (F) BM-DKO mice after 12 weeks of HFD feeding. Real-time RT-PCR gene expression analysis of Emr1 (F4/80) and Cd68 in total AT after (G) 6 and (H) 12 weeks of HFD feeding (mean±s.e.m.; n=10 mice per group). Full colour version of this figure available via http://dx.doi.org/10.1530/JOE-11-0338.
Citation: Journal of Endocrinology 212, 3; 10.1530/JOE-11-0338
The SVF was separated from the adipocytes of the epididymal AT by collagenase digestion. Macrophages from the SVF were quantified by flow cytometry using an antibody against F4/80. The representative histograms illustrate the percentage of live F4/80hi cells in the SVF after 6 weeks (Fig. 6A, B and C) and 12 weeks (Fig. 6D, E and F) of HFD feeding. Quantification of the percentage of F4/80hi cells in the SVF showed a significant decrease (P<0.05) in the BM-DKO group when compared with the BM-Lepr+/+ and BM-Lepr−/− groups after 6 weeks of HFD feeding (Fig. 6G). There were no differences in the percentage of F4/80hi cells in the AT of any group after 12 weeks of HFD feeding (Fig. 6H). The BM-DKO mice had an increase in a discrete F4/80lo myeloid population (Fig. 6C, arrow) which is due to the deficiency in haematopoietic C–C chemokine receptor 2 (CCR2), as we have previously reported (Gutierrez et al. 2011).
Flow cytometry analysis of stromal vascular cells (SVCs). SVCs were isolated and the percentage of live F4/80+ cells was quantified. Representative histograms of F4/80hi cells for (A) BM-Lepr+/+, (B) BM-Lepr−/− and (C) BM-DKO mice after 6 weeks of HFD feeding; (D) BM-Lepr+/+, (E) BM-Lepr−/− and (F) BM-DKO mice after 12 weeks of HFD feeding. (G and H) Quantification of the percentage of F4/80hi live cells in the SVF of all groups after 6 or 12 weeks of HFD feeding. Data represent mean±s.e.m. of four to five mice per group. Arrow indicates F4/80lo cells found in BM-DKO mice. *P<0.05 between BM-Lepr+/+ and BM-DKO mice. ^P<0.05 between BM-Lepr−/− and BM-DKO mice.
Citation: Journal of Endocrinology 212, 3; 10.1530/JOE-11-0338
AT inflammatory state
The inflammatory state of total AT was determined by assessing the gene expression of several pro- and anti-inflammatory markers by real-time RT-PCR. Analysis of the expression of the inflammatory markers Tnf and Nos2 showed no differences in these genes among groups after 6 weeks on the HFD (Fig. 7A). There was a significant decrease (P<0.001) in Nos2 expression in the BM-DKO group when compared with the BM-Lepr+/+ and BM-Lepr−/− groups at the 12 week time point (Fig. 7B). Additionally, assessment of the anti-inflammatory ‘M2’ markers Arg1, Mgl1 and Mgl2 showed that there was a significant increase (P<0.05) in Arg1 expression in the BM-DKO mice in comparison to the BM-Lepr+/+ mice after 6 weeks of HFD feeding (Fig. 7A). Arg1, Mgl1 and Mgl2 were all significantly increased (P<0.05) in the BM-DKO group in comparison to the BM-Lepr+/+ and/or BM-Lepr−/− groups after 12 weeks of HFD (Fig. 7B). Bone marrow LepR deficiency alone did not impact anti-inflammatory gene expression in AT.
Pro- and anti-inflammatory gene expression. Real-time RT-PCR gene expression analysis of pro- and anti-inflammatory markers was performed on RNA isolated from total AT. Expression of Tnf, Nos2, Arg1, Mgl1 and Mgl2 in all groups after (A) 6 weeks and (B) 12 weeks of HFD feeding (mean±s.e.m.; n=10 mice per group). *P<0.05 between BM-Lepr+/+ and BM-DKO mice. **P<0.01 between BM-Lepr+/+ and BM-DKO mice. ***P<0.01 between BM-Lepr+/+ and BM-DKO mice. ^P<0.05 between BM-Lepr−/− and BM-DKO mice.
Citation: Journal of Endocrinology 212, 3; 10.1530/JOE-11-0338
Discussion
Inflammatory macrophage infiltration into AT during obesity has been shown to be temporally correlated with local and systemic inflammation and IR (Weisberg et al. 2003, Xu et al. 2003, Hotamisligil 2006, Lumeng et al. 2007). Several genetic models that either disrupt macrophage recruitment to AT (Kanda et al. 2006, Weisberg et al. 2006, Odegaard et al. 2007, Surmi et al. 2010) or promote an AT macrophage switch towards an ‘M2’ or anti-inflammatory phenotype (Bouhlel et al. 2007, Odegaard et al. 2007, Westcott et al. 2009) have been explored. Despite the great progress made in this field in the past decade, methods to actively decrease macrophage recruitment to AT or diminish their inflammatory impact on this tissue are poorly understood.
We and others have shown that the adipokine leptin plays an important role in macrophage chemotaxis in vitro directly via its ability to act as a macrophage chemoattractant (Gruen et al. 2007) or indirectly through the up-regulation of adhesion molecules on endothelial cells (Curat et al. 2004). Additionally, leptin has been shown to play a role in eosinophil (Kato et al. 2011) and neutrophil (Caldefie-Chezet et al. 2001, 2003) chemotaxis. These findings have led to the speculation that leptin, which has potent monocyte chemotactic activity, could be an important mediator of monocyte/macrophage chemotaxis to AT, where this adipokine is mainly produced (Surmi & Hasty 2008, Bourlier & Bouloumie 2009, Gutierrez et al. 2009, Maeda et al. 2009, Vona-Davis & Rose 2009, Wood et al. 2009, Conde et al. 2010, Fernandez-Riejos et al. 2010). This is of particular interest during the initial stages of inflammatory macrophage accumulation in AT when recruitment is positively correlated with adiposity and circulating leptin levels (Supplementary Figure 1; Coenen et al. 2007). However, this hypothesis had not been tested in vivo. In this study we assessed whether haematopoietic LepR deficiency led to a disruption in macrophage recruitment to AT during obesity with the hypothesis that BM-Lepr−/− chimeras would have reduced macrophage accumulation and improvements in glucose and insulin tolerance. Given that compensation by other cytokines was possible, we also generated BM-DKO chimeras that have a haematopoietic deficiency in both Lepr and Ccr2.
Our results showed that haematopoietic LepR deficiency led to a significant decrease in the percentage of circulating Ly6Chi monocytes (Fig. 4B and D). A similar but more dramatic phenotype has been observed in mice with CCR2 deficiency (Tsou et al. 2007, Gutierrez et al. 2011), in which Ly6Chi monocytes fail to egress from the bone marrow. In fact, the BM-DKO mice showed an identical phenotype to that of the BM-Ccr2−/− mice (Fig. 4C and D), indicating that having both CCR2 and LepR deficiency does not confer an additive or synergistic effect on the decreases in circulating inflammatory monocyte levels. The mechanism responsible for the reduction of Ly6Chi monocytes in BM-Lepr−/− mice is unknown; however, because the levels of circulating monocyte in the BM-DKO mice are identical to those of the BM-Ccr2−/− mice, it is likely that the BM-Lepr−/− mutation is not affecting inflammatory monocyte egress but instead their proliferation and/or differentiation in the bone marrow. This hypothesis remains to be tested.
The main end point of this study was to determine if the deficiency of LepR led to a reduction in macrophage recruitment to AT. Previous studies have shown that the initial process of inflammatory macrophage accumulation in AT occurs between 6 and 12 weeks of HFD feeding (Nishimura et al. 2009, Shaul et al. 2010); thus we used these time points to assess macrophage accumulation using multiple approaches. Our histology (Fig. 5A, B, C, D, E and F), gene expression (Fig. 5G and H) and flow cytometry (Fig. 6) data showed no protective effect of LepR deficiency on macrophage accumulation in AT after 6 or 12 weeks of HFD feeding. There was a significant decrease (P<0.05) in the percentage of F4/80hi macrophages in the BM-DKO group after 6 weeks of HFD feeding (Fig. 6G); however, this can be accounted for by the haematopoietic CCR2 deficiency, which leads to a decrease in F4/80hi cells and the accumulation of F4/80lo cells (Gutierrez et al. 2011). In addition, no changes in glucose or insulin tolerance were observed between the three groups (Fig. 3A, B, C and D) despite the fact that all mice became hyperglycaemic and hyperinsulinemic in response to the HFD feeding (Fig. 3E and F). Thus, we can conclude that haematopoietic LepR deficiency alone or in combination with haematopoietic CCR2 deficiency does not affect inflammatory macrophage recruitment to AT or insulin sensitivity after 6 or 12 weeks of HFD feeding.
The negative results are not a consequence of the inability of these mice to have increased macrophage infiltration, become insulin resistant or glucose intolerant during HFD feeding. We performed an additional study in which we show that Lepr+/+ mice fed an HFD have increased macrophage infiltration, glucose intolerance and IR when compared with Lepr+/+ mice fed a chow diet (Supplementary Figure 2). Thus, within the limits of the sensitivity of our assays, haematopoietic LepR deficiency does not affect macrophage recruitment or insulin sensitivity during obesity.
Similar negative results have been obtained by our group and others when assessing inflammatory macrophage recruitment to AT after deletion of a single chemokine or chemokine receptor (Chen et al. 2005, Inouye et al. 2007, Kirk et al. 2008, Surmi et al. 2010). Accordingly, the current study provides evidence for the redundancy of chemokines and chemokine receptors in the innate immune system. Despite the elimination of two receptors that mediate the function of four different chemokines (leptin, CCL2, CCL7 and CCL8) that are up-regulated in the AT during obesity, macrophage infiltration into AT is not disrupted. This redundancy in the chemokine system contributes to the requirement for leukocyte chemotaxis in normal immune system function, making it ‘robust’ to naturally occurring mutations (Mantovani 1999).
Previously published studies suggest that in vitro leptin treatment leads to the activation of monocyte/macrophages and have shown a dose-dependent induction of inflammatory mediators such as interleukin 6 and tumor necrosis factor α (Zarkesh-Esfahani et al. 2001). Macrophages are primarily responsible for AT inflammation during obesity (Hotamisligil et al. 1993, 1995, Xu et al. 2003); thus, we tested whether haematopoietic LepR deficiency led to changes in the inflammatory state of total AT by real-time RT-PCR. Our results showed no differences in pro- or anti-inflammatory marker gene expression between BM-Lepr+/+ and BM-Lepr−/− mice after any period of HFD feeding (Fig. 7A and B). Significant differences were observed in the inflammatory gene expression of BM-DKO mice; however, these are similar to the changes in gene expression found in the BM-Ccr2−/− mice (Gutierrez et al. 2011). As with macrophage chemotaxis, the effects of leptin on inflammatory gene expression that have been found in vitro are likely compensated in vivo by other cytokines found in the AT environment during obesity.
In conclusion, our study provides evidence that despite the potency of leptin as a monocyte/macrophage chemoattractant (Gruen et al. 2007) in vitro, haematopoietic deficiency of its functional receptor does not affect macrophage accumulation in AT. Our current finding that haematopoietic LepR deficiency decreases the number of circulating Ly6Chi monocyte levels should be further studied as it could provide a novel role for leptin in haematopoiesis and/or myeloid cell migration. Our results do not rule out the possibility that leptin can affect macrophage infiltration into AT through the up-regulation of adhesion molecules in the endothelial cells of the SVF, as has been previously suggested by Curat et al. (2004).
Supplementary data
This is linked to the online version of the paper at http://dx.doi.org/10.1530/JOE-11-0338.
Declaration of interest
The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Funding
This work was supported by NIH (grant number HL089466), a supplement to this grant to D A G, and by grant F31DK091128 to D A G. A H H is also supported by an American Diabetes Association Career Development Award (1-07-CD-10); and D A G was also supported by the Molecular Endocrinology Training Grant (NIH grant number DK07563-23).
Author contribution statement
D A G designed, performed and analysed all experiments and wrote the manuscript. A H H aided in designing the experiments and edited the manuscript.
Acknowledgements
Flow Cytometry experiments were performed in the VUMC Flow Cytometry Shared Resource supported by the Vanderbilt Digestive Disease Research Centre (DK058404). Leptin ELISAs were performed at the Vanderbilt Hormone Assay Core which is part of the Vanderbilt Mouse Metabolic Phenotyping Centre supported by grant U24 DK59637. We would like to thank the Major laboratory at Vanderbilt University for their assistance with flow cytometry.
References
Bouhlel MA, Derudas B, Rigamonti E, Divart R, Brozek J, Haulon S, Zawadzki C, Jude B, Torpier G & Marx N 2007 PPARgamma activation primes human monocytes into alternative M2 macrophages with anti-inflammatory properties. Cell Metabolism 6 137–143. doi:10.1016/j.cmet.2007.06.010.
Bourlier V & Bouloumie A 2009 Role of macrophage tissue infiltration in obesity and insulin resistance. Diabetes and Metabolism 35 251–260. doi:10.1016/j.diabet.2009.05.001.
Caldefie-Chezet F, Poulin A, Tridon A, Sion B & Vasson MP 2001 Leptin: a potential regulator of polymorphonuclear neutrophil bactericidal action? Journal of Leukocyte Biology 69 414–418.
Caldefie-Chezet F, Poulin A & Vasson MP 2003 Leptin regulates functional capacities of polymorphonuclear neutrophils. Free Radical Research 37 809–814. doi:10.1080/1071576031000097526.
Chen A, Mumick S, Zhang C, Lamb J, Dai H, Weingarth D, Mudgett J, Chen H, MacNeil DJ & Reitman ML et al. 2005 Diet induction of monocyte chemoattractant protein-1 and its impact on obesity. Obesity Research 13 1311–1320. doi:10.1038/oby.2005.159.
Coenen KR, Gruen ML, Chait A & Hasty AH 2007 Diet-induced increases in adiposity, but not plasma lipids, promote macrophage infiltration into white adipose tissue. Diabetes 56 564–573. doi:10.2337/db06-1375.
Conde J, Scotece M, Gómez R, Gómez-Reino JJ, Lago F & Gualillo O 2010 At the crossroad between immunity and metabolism: focus on leptin. Expert Review of Clinical Immunology 6 801–808. doi:10.1586/eci.10.48.
Curat CA, Miranville A, Sengenès C, Diehl M, Tonus C, Busse R & Bouloumié A 2004 From blood monocytes to adipose tissue-resident macrophages: induction of diapedesis by human mature adipocytes. Diabetes 53 1285–1292. doi:10.2337/diabetes.53.5.1285.
Dixit VD, Mielenz M, Taub DD & Parvizi N 2003 Leptin induces growth hormone secretion from peripheral blood mononuclear cells via a protein kinase C- and nitric oxide-dependent mechanism. Endocrinology 144 5595–5603. doi:10.1210/en.2003-0600.
Fernandez-Riejos P, Najib S, Santos-Alvarez J, Martín-Romero C, Pérez-Pérez A, González-Yanes C & Sánchez-Margalet V 2010 Role of leptin in the activation of immune cells. Mediators of Inflammation 2010 568343 doi:10.1155/2010/568343.
Geissmann F, Manz MG, Jung S, Sieweke MH, Merad M & Ley K 2010 Development of monocytes, macrophages, and dendritic cells. Science 327 656–661. doi:10.1126/science.1178331.
Gruen ML, Hao M, Piston DW & Hasty AH 2007 Leptin requires canonical migratory signaling pathways for induction of monocyte and macrophage chemotaxis. American Journal of Physiology. Cell Physiology 293 C1481–C1488. doi:10.1152/ajpcell.00062.2007.
Gutierrez DA, Puglisi MJ & Hasty AH 2009 Impact of increased adipose tissue mass on inflammation, insulin resistance, and dyslipidemia. Current Diabetes Reports 9 26–32. doi:10.1007/s11892-009-0006-9.
Gutierrez DA, Kennedy A, Orr JS, Anderson EK, Webb CD, Gerrald WK & Hasty AH 2011 Aberrant accumulation of undifferentiated myeloid cells in the adipose tissue of CCR2-deficient mice delays improvements in insulin sensitivity. Diabetes 60 2820–2829. doi:10.2337/db11-0314.
Halaas JL, Gajiwala KS, Maffei M, Cohen SL, Chait BT, Rabinowitz D, Lallone RL, Burley SK & Friedman JM 1995 Weight-reducing effects of the plasma protein encoded by the obese gene. Science 269 543–546. doi:10.1126/science.7624777.
Hotamisligil GS 2006 Inflammation and metabolic disorders. Nature 444 860–867. doi:10.1038/nature05485.
Hotamisligil GS, Shargill NS & Spiegelman BM 1993 Adipose expression of tumor necrosis factor-alpha: direct role in obesity-linked insulin resistance. Science 259 87–91. doi:10.1126/science.7678183.
Hotamisligil GS, Arner P, Caro JF, Atkinson RL & Spiegelman BM 1995 Increased adipose tissue expression of tumor necrosis factor-alpha in human obesity and insulin resistance. Journal of Clinical Investigation 95 2409–2415. doi:10.1172/JCI117936.
Inouye KE, Shi H, Howard JK, Daly CH, Lord GM, Rollins BJ & Flier JS 2007 Absence of CC chemokine ligand 2 does not limit obesity-associated infiltration of macrophages into adipose tissue. Diabetes 56 2242–2250. doi:10.2337/db07-0425.
Kanda H, Tateya S, Tamori Y, Kotani K, Hiasa K, Kitazawa R, Kitazawa S, Miyachi H, Maeda S & Egashira K et al. 2006 MCP-1 contributes to macrophage infiltration into adipose tissue, insulin resistance, and hepatic steatosis in obesity. Journal of Clinical Investigation 116 1494–1505. doi:10.1172/JCI26498.
Kato H, Ueki S, Kamada R, Kihara J, Yamauchi Y, Suzuki T, Takeda M, Itoga M, Chihara M & Ito W et al. 2011 Leptin has a priming effect on eotaxin-induced human eosinophil chemotaxis. International Archives of Allergy and Immunology 155 335–344. doi:10.1159/000321195.
Kirk EA, Sagawa ZK, McDonald TO, O'Brien KD & Heinecke JW 2008 Monocyte chemoattractant protein deficiency fails to restrain macrophage infiltration into adipose tissue (corrected). Diabetes 57 1254–1261. doi:10.2337/db07-1061.
La Cava A & Matarese G 2004 The weight of leptin in immunity. Nature Reviews. Immunology 4 371–379. doi:10.1038/nri1350.
Leshan RL, Bjornholm M, Münzberg H & Myers MG Jr 2006 Leptin receptor signaling and action in the central nervous system. Obesity 14 (Suppl 5) 208S–212S. doi:10.1038/oby.2006.310.
Lumeng CN, Bodzin JL & Saltiel AR 2007 Obesity induces a phenotypic switch in adipose tissue macrophage polarization. Journal of Clinical Investigation 117 175–184. doi:10.1172/JCI29881.
Lumeng CN, DelProposto JB, Westcott DJ & Saltiel AR 2008 Phenotypic switching of adipose tissue macrophages with obesity is generated by spatiotemporal differences in macrophage subtypes. Diabetes 57 3239–3246. doi:10.2337/db08-0872.
Maeda T, Kiguchi N, Kobayashi Y, Ikuta T, Ozaki M & Kishioka S 2009 Leptin derived from adipocytes in injured peripheral nerves facilitates development of neuropathic pain via macrophage stimulation. PNAS 106 13076–13081. doi:10.1073/pnas.0903524106.
Mantovani A 1999 The chemokine system: redundancy for robust outputs. Immunology Today 20 254–257. doi:10.1016/S0167-5699(99)01469-3.
Nishimura S, Manabe I, Nagasaki M, Eto K, Yamashita H, Ohsugi M, Otsu M, Hara K, Ueki K & Sugiura S et al. 2009 CD8+ effector T cells contribute to macrophage recruitment and adipose tissue inflammation in obesity. Nature Medicine 15 914–920. doi:10.1038/nm.1964.
Odegaard JI, Ricardo-Gonzalez RR, Goforth MH, Morel CR, Subramanian V, Mukundan L, edEagle A, Vats D, Brombacher F & Ferrante AW et al. 2007 Macrophage-specific PPARgamma controls alternative activation and improves insulin resistance. Nature 447 1116–1120. doi:10.1038/nature05894.
Pfaffl MW 2001 A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Research 29 e45 doi:10.1093/nar/29.9.e45.
Robertson SA, Leinninger GM & myers MJ Jr 2008 Molecular and neural mediators of leptin action. Physiology and Behavior 94 637–642. doi:10.1016/j.physbeh.2008.04.005.
Shaul ME, Bennett G, Strissel KJ, Greenberg AS & Obin MS 2010 Dynamic, M2-like remodeling phenotypes of CD11c+ adipose tissue macrophages during high-fat diet–induced obesity in mice. Diabetes 59 1171–1181. doi:10.2337/db09-1402.
Surmi BK & Hasty AH 2008 Macrophage infiltration into adipose tissue: initiation, propagation and remodeling. Future Lipidology 3 545–556. doi:10.2217/17460875.3.5.545.
Surmi BK, Webb CD, Ristau AC & Hasty AH 2010 Absence of macrophage inflammatory protein-1{alpha} does not impact macrophage accumulation in adipose tissue of diet-induced obese mice. American Journal of Physiology. Endocrinology and Metabolism 299 E437–E445. doi:10.1152/ajpendo.00050.2010.
Tsou CL, Peters W, Si Y, Slaymaker S, Aslanian AM, Weisberg SP, Mack M & Charo IF 2007 Critical roles for CCR2 and MCP-3 in monocyte mobilization from bone marrow and recruitment to inflammatory sites. Journal of Clinical Investigation 117 902–909. doi:10.1172/JCI29919.
Vona-Davis L & Rose DP 2009 Angiogenesis, adipokines and breast cancer. Cytokine and Growth Factor Reviews 20 193–201. doi:10.1016/j.cytogfr.2009.05.007.
Weisberg SP, McCann D, Desai M, Rosenbaum M, Leibel RL & Ferrante AW Jr 2003 Obesity is associated with macrophage accumulation in adipose tissue. Journal of Clinical Investigation 112 1796–1808. doi:10.1172/JCI19246.
Weisberg SP, Hunter D, Huber R, Lemieux J, Slaymaker S, Vaddi K, Charo I, Leibel RL & Ferrante AW Jr 2006 CCR2 modulates inflammatory and metabolic effects of high-fat feeding. Journal of Clinical Investigation 116 115–124. doi:10.1172/JCI24335.
Westcott DJ, Delproposto JB, Geletka LM, Wang T, Singer K, Saltiel AR & Lumeng CN 2009 MGL1 promotes adipose tissue inflammation and insulin resistance by regulating 7/4hi monocytes in obesity. Journal of Experimental Medicine 206 3143–3156. doi:10.1084/jem.20091333.
Wood IS, de Heredia FP, Wang B & Trayhurn P 2009 Cellular hypoxia and adipose tissue dysfunction in obesity. Proceedings of the Nutrition Society 68 370–377. doi:10.1017/S0029665109990206.
Xu H, Barnes GT, Yang Q, Tan G, Yang D, Chou CJ, Sole J, Nichols A, Ross JS & Tartaglia LA et al. 2003 Chronic inflammation in fat plays a crucial role in the development of obesity-related insulin resistance. Journal of Clinical Investigation 112 1821–1830. doi:10.1172/JCI19451.
Zarkesh-Esfahani H, Pockley G, Metcalfe RA, Bidlingmaier M, Wu Z, Ajami A, Weetman AP, Strasburger CJ & Ross RJM 2001 High-dose leptin activates human leukocytes via receptor expression on monocytes. Journal of Immunology 167 4593–4599.
