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NK cells link obesity-induced adipose stress to inflammation and insulin resistance

Nature Immunology volume 16, pages 376–385 (2015)Cite this article

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Abstract

An important cause of obesity-induced insulin resistance is chronic systemic inflammation originating in visceral adipose tissue (VAT). VAT inflammation is associated with the accumulation of proinflammatory macrophages in adipose tissue, but the immunological signals that trigger their accumulation remain unknown. We found that a phenotypically distinct population of tissue-resident natural killer (NK) cells represented a crucial link between obesity-induced adipose stress and VAT inflammation. Obesity drove the upregulation of ligands of the NK cell–activating receptor NCR1 on adipocytes; this stimulated NK cell proliferation and interferon-γ (IFN-γ) production, which in turn triggered the differentiation of proinflammatory macrophages and promoted insulin resistance. Deficiency of NK cells, NCR1 or IFN-γ prevented the accumulation of proinflammatory macrophages in VAT and greatly ameliorated insulin sensitivity. Thus NK cells are key regulators of macrophage polarization and insulin resistance in response to obesity-induced adipocyte stress.

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Figure 1: A high-fat diet increases IFN-γ production by NK cells in VAT.
Figure 2: VAT-resident NK cells are activated by HFD feeding and promote insulin resistance.
Figure 3: VAT NK cells are required and sufficient for the induction of insulin resistance.
Figure 4: VAT-resident NK cells expand locally in response to an HFD.
Figure 5: Obese VAT induces expression of NKp46-l and stimulates IFN-γ production by NK cells.
Figure 6: NCR1 deficiency delays the development of insulin resistance after DIO.
Figure 7: NK cell–derived IFN-γ induces M1 macrophage formation in response to NCR1 activation.

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Acknowledgements

We thank S. Slavić-Stupac, M. Samsa and K. Miklić for technical assistance and D. Jurišić-Eržen and M. Zelić for help with study setup. Supported by the European Foundation for the Study of Diabetes (New Horizons Program), the Unity through Knowledge Fund (15/13 to B.P.), the University of Rijeka (13.06.1.1.03 to B.P.), the Netherlands Organization for Scientific Research (91614029 to F.M.W.) and the European Commission (PCIG14-GA-2013-630827 to F.M.W.).

Author information

Author notes

  1. Vedrana Jelenčić, Sonja Valentić and Marko Šestan: These authors contributed equally to this work.

Authors and Affiliations

  1. Department of Histology and Embryology, Faculty of Medicine, University of Rijeka, Rijeka, Croatia

    Felix M Wensveen, Vedrana Jelenčić, Sonja Valentić, Marko Šestan & Bojan Polić

  2. Department of Internal Medicine, University Hospital Rijeka, Rijeka, Croatia

    Tamara Turk Wensveen & Davor Štimac

  3. Max Planck Institute for Metabolism Research Cologne, Cologne, Germany

    Sebastian Theurich, F Thomas Wunderlich & Jens C Brüning

  4. The Lautenberg Center for General and Tumor Immunology, The Hebrew University Hadassah Medical School, Jerusalem, Israel

    Ariella Glasner & Ofer Mandelboim

  5. Department of Surgery, University Hospital Rijeka, Rijeka, Croatia

    Davor Mendrila

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  1. Felix M Wensveen
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Contributions

F.M.W. designed and carried out most of the experiments and analyzed data. V.J., S.V., M.Š., T.T.W. and V.S. performed and analyzed experiments. A.G. generated key research reagents. D.M. obtained informed consent from patients and obtained human samples. D.Š. supervised the setup of the clinical study. F.T.W. and J.C.B. coordinated the metabolic studies. O.M. helped design studies with NCRGFP mice. B.P. directed the research and wrote the paper with F.M.W., with input from all coauthors.

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Correspondence to Bojan Polić.

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Integrated supplementary information

Supplementary Figure 1 Phenotype and function of VAT-resident NK cells.

(a) VAT leukocytes were stimulated in vitro with the indicated stimuli, and IFN-γ production was assessed in NK cells and macrophages (n = 3). (b) VAT leukocytes were stimulated in vitro with PMA and ionomycin, and IFN-γ production was assessed (n = 3). (c) Splenic and VAT-resident NK cells were isolated, and the relative number of cells expressing the indicated markers was determined by flow cytometry. Gated for CD3–NK1.1+NCR1+ cells, except for the markers NK1.1 (CD3–NCR1+) and NCR1 (CD3–NK1.1+). Representative plots are shown of one of seven independent experiments, with three to five mice used per experiment. (d) Total splenocytes or VAT-derived leukocytes were stimulated with PMA and ionomycin, and the expression of CD107a on NK cells was determined by flow cytometry (n = 5). (e) The relative number of NK cells in total splenocytes or VAT-derived NK cells was determined and cells were mixed in the indicated ratio with CFSE-labeled target cells. After 5 h, effector cell–induced cell death (Topro3+) of target cells was determined by flow cytometry (n = 5). (f) Splenic and VAT-resident NK cells were stimulated in vitro with IL-15 or agonistic NCR1 antibodies, and expression of Ki67 was analyzed after 48 h. Gated for CD3–NK1.1+ cells (n = 3). (g) Kinetics of the absolute number of γδ T cells in VAT after 4 weeks of NCD or HFD feeding (n = 5). (h) Kinetics of the absolute number of the indicated immune cell subsets in the spleens of mice after 12 weeks of NCD or HFD feeding (n = 5). (i) Relative number of regulatory T cells (CD3+CD4+FoxP3+) in VAT after 12 weeks of NCD or HFD feeding (n = 5). Shown are data from one of two to seven experiments with similar results (mean ± s.e.m., n = 4–5, *P < 0.05, **P < 0.01, ***P < 0.001). FMO, fluorescence minus one.

Supplementary Figure 2 VAT-resident NK cells are activated upon HFD feeding and promote M1 macrophage formation.

(a) Mice were fed an NCD or an HFD for 4 weeks, after which VAT leukocytes were stimulated in vitro with PMA and ionomycin and cytokine production was assessed in NK cells and CD4+ TH cells (n = 5). (b) Mice were fed an NCD or an HFD and received PBS or α-NK1.1 antibodies once every 5 d. After 12 weeks, numbers of NK (CD3–NK1.1+NCR1+), NKT (CD3+NK1.1+NCR1–) and CD8+ T cells (CD8+CD3+) were quantified (n = 4–5). (c) Gating strategy for adipose tissue macrophages and NK cells. M1 macrophages (M1) are defined as CD19–CD11b+F4/80+GR1–/DimCD11c+CD86Dim. M2 macrophages (M2) are defined as CD19–CD11b+F4/80+GR1–/DimCD11c–CD86+. NK cells are defined as CD3–NK1.1+NCR1+. (d) Macrophage subsets (as defined in c) were analyzed for the expression of M1 and M2 markers under homeostatic conditions (arginase, IL-4Rα) and after 2 d of in vitro stimulation with LPS (iNOS, n = 3). (e) Mice were fed an NCD or an HFD and received PBS or α-NK1.1 antibodies once every 5 d. After 6 weeks, the ratio between M1 and M2 macrophages in VAT was determined (n = 4–5). (f,g) Mice were HFD-fed for 2 weeks and injected with PBS or neutralizing α-IFN-γ antibodies once every other day (n = 5). (f) Ratio between M1 and M2 macrophages in VAT. (g) Phenotype of NK cells in VAT. (h–k) Mice were fed an NCD or an HFD for 12 weeks and received depleting α-NK1.1 or isotype control antibodies every 5 d. (h) Mice were fasted overnight and injected i.p. with either PBS (–) or 1.0 U/kg insulin (+). Immunoblot was used to quantify phosphorylated p70 (Thr389) in tissue samples of liver and VAT. β-Actin was used as a loading control. (i) Quantification shows pooled data from two independent experiments (n = 6). (j) Body weight and (k) visceral fat pad weight of indicated groups after 12 weeks of feeding (n = 5). Shown are data from one of two to five experiments with similar results (mean + s.e.m., n = 5, *P < 0.05, **P < 0.01, ***P < 0.001).

Supplementary Figure 3 NK cells do not require the adaptive immune system to promote HFD-induced M1 macrophage formation in VAT.

(a,b) Mice underwent either sham operation or removal of abdominal fat (VATectomy; n = 5). (a) 12 weeks after surgery, mice were subjected to GTT. (b) Weight of subcutaneous fat pads. (c) Wild-type and Prkdcscid/scid (SCID) mice were fed an NCD or an HFD for 12 weeks and subjected to GTT. Shown are plasma glucose levels and AUC (n = 3–5). (d,e) NCD- or HFD-fed SCID mice were injected with α-NK1.1 antibodies or PBS once every 5 d. After 14 weeks, (d) the phenotype of NK cells and (n = 4–5) (e) the number of total and M1 macrophages in VAT were determined (n = 3–5). (f,g) SCID mice were fed an NCD or an HFD for 12 weeks. Mice received depleting α-NK1.1 or isotype control antibodies every 5 d. After 12 weeks the (f) body weight and (g) visceral fat pad weight of mice were determined (n = 4–5). Shown are data from one of two experiments with similar results (mean ± s.e.m., n = 3–5, *P < 0.05, **P < 0.01). AUC, area under the curve.

Supplementary Figure 4 HFD promotes proliferation of NK cells in VAT.

(a) Fraction of leukocytes as a percentage of the stromal vascular fraction (SVF) in spleen, brown adipose tissue (BAT), VAT and subcutaneous (SC) fat (n = 14). (b) Fraction of NK cells as a percentage of leukocytes in indicated organs under homeostatic conditions (n = 14). (c) Mice were HFD-fed for 4 weeks and subsequently injected with 2 mg BrdU. BrdU incorporation in VAT NK cells was determined after 4 h (n = 5). Shown are pooled data from three independent experiments (a,b) or one experiment (c).

Supplementary Figure 5 VAT-resident NK cells are activated by stress ligands in adipose tissue upon HFD feeding.

(a) Wild-type mice were fed an NCD or an HFD for 4 weeks. VAT sections were stained with mNCR1-immunoglobulin or mNKG2D-immunoglobulin fusion proteins and visualized with DAB. Irrelevant fusion proteins were used as negative controls. As a positive control for NCR1-L staining, 3T3-L1 cells were included. Arrow shows fat capsule. Scale bar, 100 μm. (b) Concentration-dependent IFN-γ production in NK cells after in vitro NCR1 stimulation (n = 3). (c) Splenic NK cells from Ncr1gfp/gfp mice were stimulated in vitro with plate-bound NK1.1 or NCR1 antibodies, and IFN-γ production was assessed (n = 3–4). (d) The SVF of VAT was stimulated for 48 h with agonistic NCR1 antibodies in the presence or absence of neutralizing IFN-γ antibodies. After 48 h, the fraction of iNOS+ and CD11c+ macrophages was determined (n = 3). (e) NK cells were activated in vitro for 48 h with agonistic NCR1 antibodies in the presence or absence of neutralizing IFN-γ antibodies. Subsequently, supernatants of these cells were taken and used to stimulate macrophages. After 48 h, iNOS production in macrophages was assessed (n = 3). Shown are data from one of two to four experiments with similar results (mean ± s.e.m., n = 5, *P < 0.05, **P < 0.01, ***P < 0.001). n.d., not determined.

Supplementary Figure 6 NCR1 deficiency or NK cell depletion reduces HFD-induced insulin resistance but does not affect weight gain.

(a–c) Mice were fed an NCD or an HFD and received PBS or α-NK1.1 antibodies once every 5 d (n = 4–5). (a) After 5 weeks, the number of M1 macrophages was determined in VAT. (b,c) After 12 weeks, (b) total body weight and (c) visceral adipose fat pad weight were determined. (d) Mice were fasted overnight and injected i.p. with either PBS (–) or 1.0 U/kg insulin (+). Immunoblot was used to quantify p-Akt (Ser473), p-GSK3α (Ser21/9), p-p70 (Thr389), pERK1/2 (Thr202/Tyr204) and total ERK1/2 in tissue samples of liver and VAT. Calnexin was used as a loading control (n = 2–3). Shown are data from one of two experiments with similar results (mean ± s.e.m., n = 4–5, *P < 0.05, **P < 0.01, ***P < 0.001).

Supplementary Figure 7 Therapeutic blocking of NCR1 prevents M1 macrophage formation.

(a) HFD-fed Ifng–/– mice (CD45.2) received PBS, wild-type (CD45.1+) or NCR1GFP NK cells (4 × 105) every 4 d. Shown are representative plots of the experiment shown in Figure 7a,b. Donor cells were identified based on their expression of CD45.1 or GFP. (b) Indicated groups were fed an NCD or an HFD for 12 weeks and subjected to GTT (n = 3–5). (c,d) HFD-fed Ifng–/– mice received PBS or wild-type GFP+ NK cells (5 × 105) once every 5 d and were compared with PBS-injected, NCD-fed animals. After 2 weeks, the phenotypes of (c) donor and recipient NK cells and (d) macrophages in VAT were analyzed (n = 3–4). (e) NK cells were stimulated in vitro with plate-bound α-NCR1, in the presence or absence of the same clone of α-NCR1 antibodies soluble in the medium. (f) Wild-type NK cells, NCR1 NK cells, BW cells and BW cells transgenically expressing NCR1ζ were stained with the antibodies mNCR15 and mNCR16 to demonstrate the specificity of these reagents for NCR1. (g) Six cell lines that are known to express NCR1 ligands were stained with Ncr1-immunoglobulin alone or in combination with NCR1-specific antibodies. NCR1-specific antibodies blocked binding of NCR1-immunoglobulin to target cell lines, indicating that this reagent shows target binding that is identical to that of NCR1. (h) NK cells were stimulated with PMA and ionomycin or agonistic Ncr1 antibodies in the presence or absence (control) of Ncr1-immunoglobulin. NCR1-immunoglobulin is able to reduce NK cell activation upon NCR1 stimulation, again demonstrating the specificity of NCR1-immunoglobulin fusion protein for NCR1 ligands. (i–k) Mice were fed an NCD or an HFD for 2 weeks. Animals received PBS or NCR1 ligand–blocking NCR1-immunoglobulin fusion proteins twice per week. Ncr1GFP/GFP mice are genetically deficient for NCR1. M1 macrophage and NK cell numbers were analyzed after 2 weeks (n = 5). (i) NK cell numbers and (j) M1 macrophage numbers in VAT. (k) M1/M2 ratio in VAT showed that NCR1 deficiency or its blocking prevented the absolute (Fig. 7e) and relative increase of M1 cells in VAT. (l) Fat pad weight of mice after 12 weeks of feeding (n = 5). Shown are data from one of one to two experiments with similar results (mean ± s.e.m., n = 5, *P < 0.05, **P < 0.01, **P < 0.001).

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Wensveen, F., Jelenčić, V., Valentić, S. et al. NK cells link obesity-induced adipose stress to inflammation and insulin resistance. Nat Immunol 16, 376–385 (2015). https://doi.org/10.1038/ni.3120

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