Inorganic Nitrate Promotes Glucose Uptake and Oxidative Catabolism in White Adipose Tissue Through the XOR-Catalyzed Nitric Oxide Pathway

According to the World Health Organization, there will be >650 million people worldwide with type 2 diabetes (T2D) by 2040. A lack of bioavailable nitric oxide (NO) observed in patients with T2D is considered a significant contributing risk factor for cardiovascular ill health (1). Polymorphisms in the endothelial nitric oxide synthase (eNOS) gene, responsible for in vivo NO production, are associated with cardiovascular disease, T2D, and insulin resistance in humans (2), supporting the view that perturbation of NO homeostasis contributes to the metabolic syndrome. Furthermore, eNOS-deficient mice are hypertensive, dyslipidemic, insulin resistant, and glucose intolerant (3). Inorganic nitrate, found in high concentrations in green leafy vegetables, can be reduced in vivo to form NO. Complementary studies of dietary nitrate administration to the eNOS-deficient mouse and Sprague-Dawley rats report reduced adiposity and improved glucose and insulin homeostasis after nitrate treatment (4,5). Inversely, long-term dietary nitrate deficiency induced the metabolic syndrome in mice (6). Nitrate may therefore provide a therapeutic avenue for the treatment of aspects of the metabolic syndrome.

Recently, the mechanisms behind the beneficial metabolic effects of nitrate have been explored. In skeletal muscle, nitrate induces effects associated with endurance exercise training, including fatty acid β-oxidation and fiber-type switching (7,8), which may contribute to observations that nitrate improves exercise tolerance (9). In the liver, nitrate may prevent steatohepatitis through effects on β-oxidation and lipogenesis (10).

In white adipose tissue (WAT), nitrate induces the browning response and β-oxidation (11). Whether nitrate directly regulates glucose metabolism in WAT remains unknown. We use metabolic profiling to identify a distinct glucose metabolic phenotype associated with WAT of nitrate-treated rats. We characterize the effect of nitrate on glucose uptake and metabolism in primary adipocytes using stable-isotope substrate labeling and define the mechanisms through which nitrate regulates glucose metabolism in WAT. This study provides evidence for a novel mechanism in WAT through which nitrate may mediate glucose metabolism.

Male Wistar rats (6 weeks old; 269 ± 2 g; n = 12) (Charles River Laboratories) were weight matched and received distilled water containing 0.7 mmol/L NaCl or water containing 0.7 mmol/L sodium nitrate (NaNO₃) (ultrapure; Sigma-Aldrich) ad libitum for 18 days, with food and water intake monitored (n = 6/group). Animals were housed in conventional cages at room temperature with a 12-h/12-h light/dark photoperiod.

All procedures involving live animals were performed by a license holder in accordance with U.K. Home Office regulations and underwent review by the University of Cambridge Animal Welfare and Ethical Review Committee. The rats used in this work were also used in separate but parallel studies (8,11) with a view to reducing the total numbers of animals used in accordance with U.K. Home Office best practice. Where relevant data have been reported previously (e.g., nitrate intakes), we refer to the previous paper (Supplementary Table 1).

Rats were fasted overnight and then euthanized with sodium pentobarbital (200 mg/mL) (Vétoquinol UK Ltd.). Blood was taken by cardiac puncture and processed for plasma, and WAT was removed and frozen in liquid nitrogen.

Primary WAT stromal vascular cells were fractionated from 6- to 10-week old C57BL6 mice or peroxisome proliferator–activated receptor-α (PPARα) null mice (Pparα null mice were a gift of Frank Gonzalez, National Institutes of Health, Bethesda, MD) as previously described (11,12). Cells were counted using a Scepter Cell Counter (Millipore) according to the manufacturer’s instructions. Cells were seeded at 10,000/cm² and then cultured and differentiated into adipocytes according to published methods (11,12). During the 6-day differentiation, cells were cultured with saline (control) or 500 μmol/L NaNO₃ (ultrapure; Sigma-Aldrich), 50 μmol/L 2-phenyl-4,4,5,5-tetramethylimidazoline-1-oxyl 3-oxide (PTIO), 1 μmol/L KT5823 (Santa Cruz Biotechnology), or 100 nmol/L insulin. Cells were treated with PTIO, KT5823, and insulin, with and without 500 μmol/L NaNO₃. NaNO₃ was added at day 1 and PTIO or KT5823 at day 5 of differentiation. For both Glut4 membrane biotinylation and immunoprecipitation assays and insulin-stimulated glucose uptake assays, after differentiation, primary mouse adipocytes were serum starved for 4 h before being treated with 100 nmol/L insulin or 500 μmol/L nitrate, or both 100 nmol/L insulin and 500 μmol/L nitrate together for 10 min.

Cell viability was determined using a resazurin assay. Briefly, cells were washed with PBS and incubated with 44 μmol/L resazurin in culture media for 2 h. Resazurin fluorescence was then measured (excitation 530 nm, emission 590 nm, cutoff 550 nm) with a fluorescence microplate reader.

FlexiTube siRNA against xanthine oxidoreductase (XOR), AllStars negative control siRNA, and HiPerFect Transfection Reagent were purchased from Qiagen. Adipocyte transfection was performed according to the manufacturer’s instructions (75 ng siRNA, 3 μL transfection reagent per well, 10 nmol/L final siRNA concentration) on days 2 and 4 of differentiation.

After the 6-day 500 μmol/L NaNO₃ treatment during differentiation, cells were cultured in low-glucose serum-free media supplemented with [U-¹³C]glucose (3,100 mg/L) for 24 h. Cellular metabolites were then extracted and analyzed by gas chromatography-mass spectrometry (GC-MS), as described below.

Metabolites were extracted from WAT, blood plasma, and primary adipocytes using a modified Bligh and Dyer method (13). Frozen WAT (20 mg) was pulverized using a Tissue Lyser II (Qiagen). Methanol:chloroform (2:1, 600 μL) was added to the WAT, plasma (50 μL), or primary adipocytes, and the samples were sonicated for 15 min. Chloroform:water (1:1) was then added (400 μL). Samples were centrifuged at 16,100g for 20 min, and the aqueous phase was separated, dried under nitrogen, and stored at −80°C until analysis.

Dried aqueous-phase samples were derivatized using methoxyamine hydrochloride solution (20 mg/mL in pyridine) (Sigma-Aldrich) and 30 μL N-methyl-N-trimethylsilyltrifluoroacetamide (Macherey-Nagel, Dueren, Germany) using the method described previously (14). GC-MS and data analysis were performed according to previously published methods (14). All GC-MS analyses were made using a Trace GC Ultra coupled to a Trace DSQ II single-quadrupole mass spectrometer (Thermo Scientific, Cheshire, U.K.). Derivatized aqueous samples were injected with a split ratio of 10 onto a 30-m × 0.25-mm 5% phenylpolysilphenylene-siloxane column with a 0.25-μm ZB-5 ms stationary phase (Phenomenex). The injector temperature was 230°C, and the helium carrier gas was used at a flow rate of 1.2 mL/min. The initial column temperature of 70°C was increased by 10°C/min to 130°C and then increased at a rate of 5°C/min to 230°C, followed by an increase of 20°C/min to 310°C, and held for 5 min (transfer line temperature = 250°C; ion source = 250°C; electron ionization = 70 eV). The detector was turned on after 240 s, and full-scan spectra were collected using 3 scans/s over a range of 50–650 m/z.

GC-MS chromatograms were processed using Xcaliber (version 2.0; Thermo Fisher Scientific). Each individual peak was integrated and then normalized. Overlapping peaks were separated using traces of single ions. Peak assignment was based on mass fragmentation patterns matched to the National Institute of Standards and Technology library and to previously reported literature.

Total RNA extraction from WAT and adipocytes, cDNA conversion, and quantitative real-time-PCR were performed according to published protocols (12). All data were normalized to 18S rRNA (mouse primary adipocytes) or RLPL1 (rat WAT) and quantitative measures obtained using the ΔΔcycle threshold method.

Surface Glut4 biotinylation, immunoprecipitation, and Glut4 Western blot were performed according to previously published protocols (15). Briefly, cells were incubated for 1 h at 4°C with 0.5 mg/mL biotin sulfo-NHS (Sigma-Aldrich). Cell lysates were precleared by incubation for 30 min with 0.5% (w/v) protein A–sepharose. The protein A–sepharose was pelleted by centrifugation for 1 min at 13,000g, and the supernatant was removed and incubated overnight with 0.5% (v/v) anti-Glut4 antibody (1F8; Cell Signaling Technology). Protein A–sepharose was added to 0.5% (w/v) to the samples, and incubation continued at 37°C for 1 h. Immunocomplexes were pelleted at 13,000g for 1 min, and the pellet was washed three times with 50 mmol/L Tris-HCl (pH 8.0), 150 mmol/L NaCl, 0.5% (w/v) sodium deoxycholate, 0.1% (w/v) SDS, and 1% (v/v) Nonidet P-40.

Immunoprecipitated biotinylated complexes were mixed with dissociation buffer (125 mmol/L Tris-HCl [pH 6.8], 2% (w/v) SDS, 20% (v/v) glycerol, 100 mmol/L dithiothreitol, bromophenol blue) and boiled for 5 min. Glut4 was resolved by electrophoresis through 10% polyacrylamide gels and then transferred to Hybond-P polyvinylidene difluoride membrane. The membrane was blocked for 1 h in PBS (1.5 mmol/L KH₂PO₄, 2.7 mmol/L Na₂HPO₄, 150 mmol/L NaCl [pH 7.4]) containing 5% (w/v) dried milk powder and 0.1% (v/v) Tween-20, followed by incubation with peroxidase-conjugated streptavidin (1:1,000 dilution in PBS containing 0.1% [v/v] Tween-20) for 1 h. Bound peroxidase conjugates were visualized using an enhanced chemiluminescence detection system (Amersham Biosciences). Quantitation of immunoblots was performed using ImageJ software.

Cells were grown and differentiated in 96-well plates. Cells were washed with Dulbecco’s PBS (DPBS) and placed in low-glucose (1 g/L) serum-free DMEM for 24 h. Media was replaced with low-glucose serum-free DMEM for 1 h. After media aspiration, DPBS containing 6-deoxy-6-[(7-nitro-2,1,3-benzoxadiazol-4-yl)amino]-d-glucose (6-NBDG) (200 μmol/L) was added for 1 h, and cells were kept at 37°C in 5% CO₂. Cells were washed three times with DPBS, and fluorescence was measured using a microplate reader (excitation 485 nm, emission 528 nm).

Metabolomics data analysis was performed using MetaboAnalyst, version 4.0 (16). Data sets were auto scaled and analyzed using partial least squares–discriminant analysis (PLS-DA). Metabolite changes responsible for clustering or regression trends within the pattern recognition models were identified by interrogating the corresponding loadings plot. Metabolites identified in the variable importance in projections/coefficients plots were deemed to have changed globally if they contributed to separation in the models with a confidence limit of 95%. Plasma metabolomics data were analyzed using univariate volcano plots with a fold change cutoff of 1.2 and P value cutoff of 0.05 to identify significantly different metabolites.

All data, including metabolomics data sets for the GC-MS analysis of nitrate-treated rat adipose tissue and nitrate-treated primary adipocytes, are available from the corresponding author on reasonable request.

Male Wistar rats were treated with 0.7 mmol/L NaCl or 0.7 mmol/L NaNO₃ via the drinking water for 18 days. Nitrate intake for the control group was 1 mg/kg/day compared with 8 mg/kg/day in the nitrate-treated group, whereas water and food intake was not significantly different between the groups (11) (Supplementary Table 1). Metabolomic profiling and analysis using PLS-DA multivariate statistics identified a distinct metabolic signature differentiating subcutaneous inguinal WAT of nitrate-treated animals from controls (Fig. 1A). Interrogation of the corresponding loadings plots identified a distinct reprogramming of glucose and fatty acid metabolism in the WAT of nitrate-treated rats (Fig. 1B). The decrease in the WAT concentration of the fatty acids propanoic, heptanoic, nonanoic, 3-hydroxyoctanoic, oleic, linoleic, and arachidonic acid in nitrate-treated animals was consistent with our previous findings that nitrate drives WAT browning and β-oxidation (11). In a novel observation, we identified a distinct glucose metabolism phenotype in the WAT of nitrate-treated rats. The key glycolytic intermediates glucose 6-phosphate (Fig. 1C) and 3-phosphoglycerate (Fig. 1D), alongside the pentose phosphate pathway–glycolysis metabolite glycerate (Fig. 1E) and the pentose phosphate pathway metabolite d-altro-heptulose (Fig. 1F), were decreased in nitrate-treated rat WAT. Metabolic profiling of the plasma from nitrate-treated animals indicated a corresponding decrease in glucose concentration (Fig. 1G and H).

To investigate whether nitrate functions to increase glucose uptake and metabolism in adipose tissue and possibly contributes to systemic glucose clearance, the expression of genes encoding the insulin-regulated GLUT type 4 (Glut4), the insulin-independent GLUT (Glut1), and a key rate-regulating enzyme in glycolysis, hexokinase 2 (Hk2), which converts glucose to glucose 6-phosphate, was interrogated using quantitative real-time-PCR (Fig. 1I). Nitrate significantly increased the expression of these glucose import and metabolism genes. These data suggest that nitrate increases glucose uptake and disposal in adipose tissue.

To determine whether nitrate functions directly on WAT to increase glucose uptake and metabolism, stromal vascular fraction–derived primary adipocytes isolated from inguinal WAT of mice were treated with nitrate. An NaNO₃ concentration of 500 μmol/L was chosen (11). Consistent with the results in vivo, metabolomic profiling identified that nitrate decreased the intracellular concentration of the glycolytic intermediates glucose 6-phosphate (Fig. 2A) and 3-phosphoglycerate (Fig. 2B) in adipocytes. Nitrate treatment also significantly decreased the glucose concentration of the media (Fig. 2C). The expression of genes key to glucose import and metabolism, Glut4, Glut1, and Hk2, was increased in the nitrate-treated adipocytes (Fig. 2D).

To functionally assess the effect on glucose metabolism observed in the WAT of nitrate-treated rats, the stable isotope substrate [U-¹³C]glucose was used to evaluate enrichment of glucose-derived carbon through glucose import, the glycolytic pathway, and into the tricarboxylic acid (TCA) cycle. Primary adipocytes were incubated in serum-free media containing [U-¹³C]glucose and treated with 500 μmol/L nitrate. GC-MS analysis was used to define the relative enrichment of metabolites. The labeled glucose enters the glycolytic pathway and is catabolized to pyruvate, which is converted to lactate or labeled acetyl-CoA, which enters the TCA cycle (Fig. 2E). An unlabeled metabolite is detected as the molecular ion (M) in the mass spectrum. Additional [¹³C]carbon atoms introduced to the specific molecule give rise to an increase in mass of 1 (M1, M2, M3, and so forth). Nitrate treatment increased the labeling of lactate (Fig. 2F) and the M2 isotopologues of TCA cycle intermediates, citrate (Fig. 2G), malate (Fig. 2H), and the amino acid glutamate (Fig. 2I), which is in fast exchange with 2-oxoglutarate from the TCA cycle. We noted a higher fractional enrichment of the M1 isotopologues of citrate (Fig. 4H) in nitrate-treated adipocytes, which may be indicative of a moderate increase in malic enzyme (ME) activity. Relative enrichment of the M3 isotopologues of TCA cycle intermediates was elevated in nitrate-treated cells (Fig. 2H and I). Increased labeling of M3 occurs through the action of pyruvate carboxylase (PC).

In summary, nitrate confers a functional effect on adipocytes, increasing not only glucose uptake and flux through glycolysis but also oxidative catabolism.

We then investigated whether nitrate increased Glut4 presentation at the plasma membrane of primary adipocytes at baseline and after insulin stimulation. Using a cell-surface protein biotinylation and immunoprecipitation approach, we identified that nitrate increased the adipocyte plasma membrane Glut4 concentration (Fig. 3A and B and Supplementary Fig. 1). We also observed that nitrate had an additive effect on plasma membrane Glut4 expression when combined with insulin (Fig. 3A and B). Next, the functional uptake of glucose into adipocytes treated with nitrate was measured with the fluorescent glucose analog 6-NDBG in combination with insulin (100 nmol/L) (Fig. 3C). Nitrate and insulin treatments did not affect cell viability (Supplementary Fig. 2). Nitrate increased adipocyte glucose uptake and had an additive effect on insulin-stimulated glucose uptake into primary adipocytes.

We next probed the mechanisms through which nitrate mediates its effects on glucose metabolism in adipocytes. Several of the metabolic effects of nitrate, including increased β-oxidation and lactate dehydrogenase activity in muscle, occur through downstream PPARα signaling (8,17). We examined the effect of nitrate on glucose import and catabolic gene expression in primary adipocytes differentiated from the stromal vascular fraction of inguinal WAT from PPARα null mice. The lack of PPARα had no effect on nitrate-induced expression of glucose metabolism genes within the adipocytes (Fig. 4A). Nitrate treatment significantly increased the expression of Glut4, Hk2, and Glut2. Thus, nitrate regulates glucose metabolic gene expression in adipocytes independently of PPARα.

NO can be generated in vivo from nitrate via XOR-catalyzed serial reduction of nitrate to nitrite and then to NO via the nitrate-nitrite-NO pathway (18,19). We found nitrate induced the browning process in WAT through this pathway (11). Therefore, we speculated that nitrate might be functioning via NO to increase glucose import and metabolism in WAT. Primary adipocytes were differentiated in the presence of nitrate and the NO scavenger PTIO. Nitrate and PTIO treatments did not affect cell viability (Supplementary Fig. 3). PTIO abrogated the nitrate-induced expression of glucose metabolism genes, Glut4, Glut1, and Hk2 (Fig. 4B). Next, the functional uptake of glucose into primary adipocytes treated with nitrate was measured with 6-NDBG in combination with PTIO (Fig. 4C). PTIO inhibited nitrate-mediated glucose uptake into adipocytes. NO increases glucose uptake through the activity of cyclic guanosine monophosphate (cGMP)–dependent protein kinase G (PKG) (20). We previously showed that nitrate increases intracellular cyclic guanosine monophosphate concentrations in adipocytes (11). We speculated that nitrate may signal via PKG to regulate adipocyte glucose metabolism. The pharmacological PKG inhibitor KT5823 blocked the nitrate-induced expression of Glut4, Glut1, and Hk2 in the adipocytes (Fig. 4D). These data demonstrate that nitrate signals via NO to enhance glucose uptake.

As mentioned above, the reduction of nitrate to NO in mammals can proceed via an enzymatic mechanism catalyzed by XOR (18,19). XOR is expressed in WAT and has a role in adipocyte homeostasis and the nitrate-induced browning mechanism (11,21). To determine the role of this enzyme in the nitrate-mediated enhancement of glucose catabolism, XOR in primary adipocytes was knocked down using siRNA (11) (Supplementary Fig. 4). Nitrate and XOR knockdown did not affect cell viability (Supplementary Fig. 5). Knockdown of XOR abrogated the increased expression of glucose import and the catabolism genes Glut4, Glut1, and Hk2 in adipocytes treated with nitrate (Fig. 4E). Functionally, the effect of XOR knockdown on glucose uptake into nitrate-treated primary adipocytes was measured using 6-NDBG (Fig. 4F). XOR knockdown inhibited nitrate-induced glucose uptake into primary adipocytes.

These data indicate that nitrate induces glucose uptake and oxidative catabolism in WAT via the XOR catalyzed reduction of nitrate and downstream NO signaling.

Inorganic nitrate, found in high concentration in green leafy vegetables, may restore impaired NO signaling to treat aspects of cardiometabolic diseases, including T2D (22). Nitrate regulates glucose homeostasis in rodent models of T2D (4–6,23); however, the mechanisms for nitrate-mediated improvements in metabolism remain to be fully elucidated. In WAT, nitrate promotes the browning response and enhanced fatty acid oxidation (11). The effects of nitrate on WAT glucose metabolism remained undefined. We are the first to determine that inorganic nitrate directly enhances both glucose uptake and oxidative catabolism through the activity of XOR, NO, and PKG signaling. These findings are consistent with the effects of NO on glucose uptake and Glut4 expression in WAT (24–26). Interestingly, NO exhibits both insulin-dependent and insulin-independent effects on glucose uptake (24). We observe that nitrate enhances insulin-stimulated glucose uptake and plasma membrane expression of the insulin-sensitive GLUT Glut4 in adipocytes. Nitrate also stimulates the expression of the insulin-independent transporter Glut1. This is consistent with our previous observation that nitrate induces WAT browning, since the browning phenomenon enhances Glut1 expression and may partly uncouple WAT glucose uptake from insulin action (27).

Several of the metabolic effects of nitrate, including enhanced lactate dehydrogenase activity, in skeletal muscle and heart are mediated through the nuclear receptor PPARα (8,17). Our data show that nitrate-mediated expression of glucose uptake and metabolism genes was independent of PPARα signaling. The effects of nitrate on Glut4, Glut1, and Hk2 were preserved in PPARα null adipocytes. This may partly reflect the low levels of expression of this nuclear receptor in WAT.

Although both WAT browning and enhanced mitochondrial biogenesis and β-oxidation in muscle are likely to substantially contribute to the antiobesity and antidiabetic effects of nitrate, increased glucose uptake and oxidative catabolism will be a distinct factor mediating the effect of the bioactive anion on glucose homeostasis.

Funding. Funding was received from the Medical Research Council (grants MC_UP_A090_1006, MC_PC_13030, MR/P011705/1, and MR/P01836X/1 to J.L.G. and MR/R014086/1 to L.D.R.), the Biotechnology and Biological Sciences Research Council (grants BB/H013539/2 to J.L.G. and BB/R013500/1 to L.D.R.), and the Diabetes UK RD Lawrence Fellowship (16/0005382 to L.D.R.)

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

Author Contributions. B.D.M., A.M., N.T.W., T.A., A.W., and L.D.R. performed most of the experiments. B.D.M., S.A.M., J.L.G., and L.D.R. assisted with metabolomic screens and [¹³C] isotope studies. T.A. and A.J.M. designed and led the animal studies. S.A.M. assisted with experiments throughout. M.T.K. and R.M.C. provided intellectual input. L.D.R. designed and led the studies, interpreted the results, and wrote the manuscript with input from all coauthors. L.D.R. 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.