During reduced energy intake, skeletal muscle maintains homeostasis by rapidly suppressing insulin-stimulated glucose utilization. Loss of this adaptation is observed with deficiency of the fatty acid transporter CD36. A similar loss is also characteristic of the insulin-resistant state where CD36 is dysfunctional. To elucidate what links CD36 to muscle glucose utilization, we examined whether CD36 signaling might influence insulin action. First, we show that CD36 deletion specific to skeletal muscle reduces expression of insulin signaling and glucose metabolism genes. It decreases muscle ceramides but impairs glucose disposal during a meal. Second, depletion of CD36 suppresses insulin signaling in primary-derived human myotubes, and the mechanism is shown to involve functional CD36 interaction with the insulin receptor (IR). CD36 promotes tyrosine phosphorylation of IR by the Fyn kinase and enhances IR recruitment of P85 and downstream signaling. Third, pretreatment for 15 min with saturated fatty acids suppresses CD36-Fyn enhancement of IR phosphorylation, whereas unsaturated fatty acids are neutral or stimulatory. These findings define mechanisms important for muscle glucose metabolism and optimal insulin responsiveness. Potential human relevance is suggested by genome-wide analysis and RNA sequencing data that associate genetically determined low muscle CD36 expression to incidence of type 2 diabetes.
CD36 (SR-B2) has high affinity for long-chain fatty acids (FA) and facilitates tissue FA uptake in rodents (1,2) and humans (3,4). The protein also transduces signaling initially documented to mediate its role in immunity and atherosclerosis (5–7). However, accumulating evidence supports the importance of CD36 signaling in regulating metabolic pathways such as FA oxidation (8), fatty taste perception (9,10), eicosanoid formation (11), and chylomicron production (12), among others.
CD36 is important for muscle metabolic adaptation (7). In wild-type mice, fasting causes muscle to reduce glucose utilization, whereas in CD36−/− mice, muscle glucose utilization persists despite high circulating FA causing hypoglycemia and increasing risk of sudden death (13). CD36−/− mice have accelerated depletion of glycogen stores during exercise and fail to increase muscle FA oxidation (14). Also, beneficial changes in substrate utilization and muscle performance induced by exercise training are not observed in CD36−/− mice (15).
The mechanisms associating CD36 to regulation of muscle glucose metabolism are unknown. CD36 signaling was shown to regulate FA oxidation by directly modulating AMPK activation (8,14). We examined whether it might influence insulin action on muscle glucose utilization. Using a mouse with conditional deletion of CD36 in muscle and primary-derived human myotubes, we show that CD36 regulates insulin stimulation of glucose metabolism. CD36 interacts with the insulin receptor (IR) and augments its insulin-induced phosphorylation by the kinase Fyn. Saturated FA rapidly dissociate Fyn, transducing pathway inhibition. The findings provide insight into mechanisms important for optimal muscle insulin responsiveness.
Research Design and Methods
All protocols for animal experiments in this study were approved by the Washington University in St. Louis Animal Studies Committee.
Generation of Skeletal Muscle-Specific CD36−/− Mice
C57BL/6 Cd36-floxed (Cd36fl/fl) mice (17) were crossed with mice expressing human skeletal actin–reverse tetracycline-controlled transactivator (HSA-rtTA) Cre (18) for more than five generations. Skeletal muscle Cd36 deletion (smCd36−/−) was induced by giving 8-week-old Cre-positive and Cre-negative (Cd36fl/fl) mice one intraperitoneal doxycycline injection (100 mg/kg) then doxycycline (2 g/L) in drinking water for 7 days, followed by a washout period of >7 days. The mice were used for studies between 10 and 18 weeks, a range that includes mice fed a high-fat diet (HFD) for 5 weeks (Surwit, D12331; Research Diets). Male mice were used for most studies unless indicated, but key findings on muscle insulin signaling and glucose disposal were reproduced in females. Combined male/female data are presented as indicated.
Intragastric Palm Oil and Glucose Tolerance Test
Palm kernel oil, blended at 57°C 1:1 with fat-free milk, or milk (vehicle), was administered to 10- to 15-week-old mice intragastrically (200 μL/mouse) after a 16-h fast. Glucose (2 g/kg) was given intraperitoneally 2 h later, and its clearance monitored in tail blood.
Cells and Treatments
Human skeletal muscle myotubes (HSMMs; Lonza), cultured and differentiated per Lonza’s instructions, were treated with lipofectamine RNAiMAX and 20 nmol/L CD36 (small interfering [si]RNA s2647 [siCD36_1], siRNA s2646 [siCD36_2]; Ambion) or nonspecific siRNAs and used 72 h after transfection. CHO cells with stable expression of human IR and human CD36 (CHO/IR/+CD36) and CHO/IR vector controls (11) were transiently transfected where indicated (lipofectamine LTX; Life Technologies) and used 48 h later. C2C12 myoblasts were cultured and differentiated as previously described (8). Unless indicated, all cells were serum-starved (16 h) before studies in low-glucose DMEM with 2 mmol/L l-glutamine, 100 μmol/L minimum essential medium nonessential amino acids, 100 units/mL penicillin, and 100 μg/mL streptomycin. All FA were added with BSA (2:1) for 15 min before insulin.
Quantitative Real-time PCR
RNA (TRIzol; Invitrogen) was subjected to cDNA reverse transcription and quantitative real-time PCR (ABI Prism 7000; Applied Biosystems) using Power SYBR Green PCR Master Mix and a 7500 Fast Real-Time PCR System (Applied Biosystems). Values (ΔΔCt) were normalized to 36B4 expression. Primers are listed in Supplementary Table 2.
Glycolysis in HSMMs and CHO cells was measured using Seahorse XF96 (Agilent) following the manufacturer’s protocols. Cells were seeded (3.4 × 104 cells/well) in XF96 microplates and switched 24 h later to serum-free medium (16 h) before experiments. For muscle explants, mice were euthanized by CO2, and dissected diaphragms were placed in warm (37°C) Seahorse XF medium with 2 mmol/L l-glutamine, 1 mmol/L sodium pyruvate, and 10 mmol/L glucose. Sections (∼2 × 2 × 1 mm) were transferred into XF24 islet capture microplates, seeded using islet capture screens, and processed (19).
For glucose uptake (20), cells were incubated with 1 μCi/mL 2-deoxy-d-[3H]glucose (2DG) in PBS and 0.1 mmol/L 2DG, washed with cold PBS containing 20 mmol/L 2DG, solubilized in 1% SDS, and counted (TriCarb 1600 TR; Packard). For glycogen (21), cells were incubated with d-[3H]glucose (1 μCi/mL) and 100 nmol/L insulin in PBS plus 0.2% BSA and 10 mmol/L glucose. Carrier glycogen in NaOH was added to lysates (35% weight [w]/volume [v]) before boiling (30 min, 95°C) and ethanol (95%) precipitation (−20°C). After centrifugation (12,000g for 10 min), ethanol-washed water-resuspended pellets were counted. Wheat germ agglutinin (WGA) staining adjusted for cell number (22).
Ceramides and diacylglycerols were measured by the Biomolecular Analysis Core at Washington University in St. Louis by liquid chromatography-tandem mass spectroscopy, as previously described (23). Muscles were homogenized in PBS (w/v: 1:4), and 50 μL homogenates plus internal standards (ceramides 17:0 and diacylglycerols 15:0–15:0) lipids were extracted. Peak area ratios of analytes to internal standards were used for analysis.
Plasma and muscle triglycerides (TG) were measured using the L-Type Triglyceride M kit and HR Series NEFA-HR (2) kit from Wako Diagnostics.
Positron Emission Tomography
Positron emission tomography imaging of fed mice used [18F]-2-fluoro-2-deoxy-d-glucose (FDG) to assess glucose metabolism (24). Uptake/transport of tracer was visualized and quantified by summing uptake kinetics up to 60 min after the tracer injection.
Tissue sections were processed as previously described (25). Slides were scanned (NanoZoomer 2.0 HT; Hamamatsu, Bridgewater, NJ), and images were analyzed to yield muscle cross-sectional area (CSA) using Visiomorph (Broomfield, CO).
Proximity Ligation Assay
For proximity ligation assay (PLA; Duolink In Situ Hybridization kit; Sigma-Aldrich), cells on coverslips were incubated with or without insulin, washed with PBS, fixed with ice-cold methanol (−20°C) for 20 min, and blocked for 1 h in PBS with 0.05% Tween 20, 1% BSA, and 5% goat serum. After incubation for 16 h with goat anti-CD36 (R&D Systems) and rabbit anti-IRβ (Cell Signaling) antibodies, processing followed the manufacturers’ instructions. For tissues, formalin-fixed paraffin-sections were deparaffinized and processed as above. Imaging used Nikon Eclipse TE2000-U, a Photometrics CoolSNAP cf camera and MetaMorph 6.2r6 (Molecular Devices).
Coimmunoprecipitation and Immunofluorescence
Tissues and cells were lysed (30–60 min) in ice-cold buffer (20 mmol/L Tris-HCL [pH 7.5], 150 mmol/L NaCL, 1% Triton X-100, 60 mmol/L octyl β-d-glucopyranoside, 200 μmol/L sodium orthovanadate, 50 mmol/L NaF, 1 mmol/L phenylmethylsulfonyl fluoride, and 1.0 µg/mL protease inhibitor mix) and cleared lysates (10,000g for 10 min) assayed for protein content (BCA 23225; Pierce Biotech). For immunoprecipitation (IP), cells or tissues were lysed in IP buffer containing 1% Anapoe-C12E8 (Anatrace), 0.1% Triton X-100, 150 mmol/L NaCl, 5 mmol/L MgCl2, and 25 mmol/L HEPES (pH 7.5) with phosphatase and protease inhibitors. Primary antibodies or isotype-matched IgG, prebound (30 min) to Dynabeads protein G magnetic beads (Thermo Fisher Scientific) were incubated overnight (4°C) with equal lysate proteins, washed with cold IP buffer, and boiled (5 min) in SDS sample buffer. Immunofluorescent microscopy was performed as previously described (8).
In-Cell Western Assay
Cells in black-walled 96-well microplates (Corning) at confluence were serum-starved, treated as indicated, fixed with 3% paraformaldehyde, and blocked (1 h, PBS containing 0.05% Tween 20, 1% BSA, and 5% goat serum). Incubation (16 h) with monoclonal anti–phosphorylated (p)AKT(S) antibody (Cell Signaling) was followed by washing (PBS, Tween 0.05%), incubation (60 min) with horseradish peroxidase–anti-rabbit antibody, washing, then incubation (30 min, room temperature) with 3,3′,5,5′-tetramethylbenzidine (TMB) liquid substrate (Sigma-Aldrich) before adding 1 N NaOH and reading absorbance at 450 nmol/L. To normalize signals, cells were incubated (60 min) with WGA Alexa Fluor 680 (Invitrogen), rinsed (PBS), and fluorescence was measured at 700 nmol/L (LI-COR). Backgrounds for TMB (primary antibody omitted) and WGA Alexa Fluor 680 staining (WGA omitted) were subtracted from the data.
Analyses were performed with GraphPad Prism 7.2 software (GraphPad Software). Unpaired two-tailed Student t tests and two-way ANOVA, with the Bonferroni correction for multiple comparisons, were used as appropriate. Data are presented as the means ± SE unless otherwise indicated. A P value <0.05 was considered significant.
CD36 Deletion in Skeletal Muscle
We generated the smCd36−/− mouse using has-rtTA Cre (18). Adult (8-week-old) Cre-positive and floxed controls were administered doxycycline (detailed under 2research design and methods) and used after a minimum 7-day washout period. Muscle CD36 expression was assayed for all cohorts. Typically, CD36 mRNA (Fig. 1A) and protein (Fig. 1B) were reduced by 50–60% in the oxidative (slow-twitch) diaphragm and the metabolically mixed (slow and fast twitch fibers) gastrocnemius, and no reduction was observed in the heart, as expected (Fig. 1A). No reduction was observed in mixed quadriceps and abdominal muscles (rectus abdominus), where CD36 expression is normally low (Fig. 1A and B). This is consistent with minor mosaicism of the expression cassette (18). Residual CD36 expression in the diaphragm and gastrocnemius is likely accounted for by blood vessel CD36, because immunostained myotubes appeared CD36 depleted (Fig. 1C). All studies were conducted with diaphragm or gastrocnemius, with quadriceps or rectus abdominus used as negative controls.
CD36 Deletion Reduces Muscle Ceramides
Chow-fed smCd36−/− mice and Cd36fl/fl littermates weighed the same (data not shown) and had comparable fat and lean body mass (Fig. 1D). Muscle morphology and fiber CSA was unaltered (Fig. 1E). Fasting plasma levels of glucose, TG, and unesterified FA (Fig. 1F–H) were similar. Diaphragm and quadriceps TG content was equivalent for both groups (Fig. 1I). However, most ceramide species measured were reduced (Fig. 1J), and several diacylglycerol species (Fig. 1K) trended lower in smCd36−/− diaphragms; however, no changes were measured in quadriceps, where CD36 knockdown is not observed (Supplementary Fig. 1A and B).
Reduced Glucose Uptake by Muscle
CD36 deletion reduced muscle expression of glucose metabolism and insulin signaling genes (Fig. 2A). Expression of FA metabolism genes (Fig. 2A) and mitochondrial genes and proteins (Supplementary Fig. 1C and D) was unchanged.
Glucose tolerance was similar between genotypes (Fig. 2B). Although females from both genotypes had slightly better glucose disposal than males, there were no sex-driven genotype-related differences (Supplementary Fig. 2A–C). In vivo uptake of FDG measured in hind-limb muscles was reduced in smCd36−/− mice, whereas heart glucose uptake increased (Fig. 2C), suggesting that less glucose uptake by skeletal muscle makes more available for the heart. Glucose uptake by abdominal muscle, where CD36 expression was not decreased (Fig. 1A), was unchanged (Fig. 2C).
Glucose metabolism assayed using muscle explants ex vivo showed insulin stimulation of glycolysis (extracellular acidification rate [ECAR]) and glucose oxidation (oxygen consumption rate) was diminished in explants from smCd36−/− mice compared with explants from Cd36fl/fl controls (Fig. 2D).
Impaired Disposal of Postprandial Glucose
Postprandial glucose disposal was tested during absorption of a fat-rich meal to engage mixed and oxidative muscles where CD36 expression is deleted. The mice were given an intragastric bolus of palm oil with skim milk (1:1), and 2 h later, during peak absorption (26) (Supplementary Fig. 1E), intraperitoneal (i.p.) glucose and its blood clearance were monitored. Glucose disposal was impaired by 40% (Fig. 3A, top panel), and the area under the curve (AUC) (Fig. 3B) was reduced (P = 0.002) in smCd36−/− compared with Cd36fl/fl controls. Clearance was similar when intragastric skim milk was given as a vehicle control (Fig. 3A, bottom panel). Plasma TG (Fig. 3C) and free FA (Fig. 3D) levels determined after glucose injection did not differ between groups.
To examine whether the impaired glucose disposal in smCd36−/− mice reflects diminished muscle insulin signaling, mice were given the palm oil, then 2 h later insulin i.p., and diaphragms were harvested after 15 min. Diaphragm and gastrocnemius, but not quadriceps, of smCd36−/− mice had attenuated insulin-stimulated AKT phosphorylation (Fig. 3E and F). Phosphorylation of AKT targets glycogen synthase kinase 3 (GSK3α/βS21/9) and acyl citrate lyase (ACLyS455) (Fig. 3E and G) was also diminished. These data suggest CD36 deletion reduced insulin signaling in oxidative muscle during the high-fat meal.
SmCd36−/− Mice Are Not Protected From High-Fat Feeding
The mice were challenged with an HFD for 5 weeks, and effects on glucose metabolism were examined. Weight gain by smCd36−/− and Cd36fl/fl mice was similar (Fig. 3H). TG content was reduced in diaphragm but not quadriceps of smCd36−/− mice compared with controls (Fig. 3I), but both groups had similarly impaired insulin-induced glucose clearance (Fig. 3J), irrespective of sex (Supplementary Fig. 2D and E). Muscle glucose uptake in vivo was similar (Fig. 3K). Overall, CD36 deletion did not protect against HFD-induced muscle insulin resistance.
Cell-Autonomous Regulation by CD36 of Insulin Signaling and Glucose Metabolism
The findings with smCd36−/− mice suggested that Cd36 deletion suppresses insulin signaling. To determine whether this effect is cell autonomous, we examined whether it can be reproduced in primary-derived HSMMs. CD36 is highly expressed in these cells and relocates to the plasma membrane after insulin (Fig. 4A), as reported for muscle and cardiomyocytes (22,27). Treatment with siRNA did not alter myotube morphology or myosin heavy-chain content (Fig. 4B) but reduced CD36 level by 60% (Fig. 4C and D). Insulin addition to HSMMs induced robust phosphorylation of AKTT308, S473 and its targets GSK3S21/9 and ACLyS45. However, the insulin response was suppressed in CD36-depleted myotubes (Fig. 4C and E). A second siRNA yielded similar data (Supplementary Fig. 3A and B). In contrast, expression of CD36 in HEK293 cells enhanced insulin signaling compared with controls (Supplementary Fig. 3C and D), suggesting CD36 regulation is not limited to myotubes.
We next examined whether acute CD36 inhibition was sufficient to blunt insulin signaling. The FA analog sulfosuccinimidyl-oleate (SSO) is a widely used irreversible and specific inhibitor of CD36 (28,29). Pretreating myotubes for 15 min with SSO (20 μmol/L) effectively reduced insulin-induced pAKT and pGSK3 (Fig. 4F and G).
CD36 depletion in HSMMs did not alter basal glycolysis but diminished insulin-stimulated glycolytic activity, capacity, and reserve (Fig. 4H and I). Insulin-stimulated glucose uptake (Fig. 4J) and incorporation into glycogen (Fig. 4K) were suppressed. Glucose uptake was also reduced by 15-min SSO pretreatment (Fig. 4L) in line with the reduced insulin signaling (Fig. 4F and G). CD36 expression also enhanced insulin-stimulated glycolysis in CHO cells expressing the IR (CHO/IR/+CD36) (11), as shown in Supplementary Fig. 3E.
Saturated FA Suppress Insulin Signaling via CD36
CD36 is a high-affinity receptor for saturated and unsaturated FA (30), so we examined whether it transduces FA effects on insulin signaling. HSMMs were pretreated (15 min) with the saturated FA palmitic acid (PA, 16:0) and myristic acid (MA, 14:0) or with monounsaturated oleic acid (OA, 18:1), complexed to BSA (2:1 molar ratio). Cells were then tested for insulin-stimulated (5 min) AKT phosphorylation. Interestingly, PA or MA suppressed insulin-induced pAKT but no suppression occurred with OA (Fig. 5A).
FA effects on insulin signaling were further validated using an in-cell Western (ICW) assay (Fig. 5B) sensitive for phosphorylated proteins (31). In control myotubes, insulin increased pAKT, and this was suppressed by pretreatment with PA but not OA. However, insulin stimulation and PA suppression were absent in CD36-depleted HSMMs, consistent with CD36 dependence of these effects (Fig. 5B). PA also reduced insulin-induced pAKT in CHO/IR/+CD36 cells but not in vector controls (Fig. 5C).
Additional FA types tested (Fig. 5D) included polyunsaturated FA (PUFA), eicosapentaenoic acid (EPA, 20:5), and linoleic acid (LA, 18:2), monounsaturated palmitoleic acid (PO, 16:1), a product of lipogenesis with positive effects on insulin responsiveness (32), and the nonmetabolized C16 FA analog β-β′-tetramethyl-hexadecanedioic acid (M16), an insulin sensitizer in rodents (33). In CHO/IR/+CD36 cells, 15-min pretreatment with EPA and LA did not alter insulin-stimulated phosphorylation of AKT or ACLy, whereas PO, OA, and M16 enhanced phosphorylation, and PA, as before, was inhibitory (Fig. 5D). These data show CD36 mediates differential effects of FA on insulin signaling, negative for PA and MA, and positive for PO, OA, and M16.
CD36 Enhances Tyrosine Phosphorylation of IR
Insulin signaling is initiated by tyrosine phosphorylation of IR. CD36 signal transduction involves interaction with specific signaling protein clusters (5,8–10), so we explored whether CD36 might functionally interact with IR. Immunoprecipitates (IPs) of IRβ, the regulatory IR subunit, from CHO/IR/+CD36 cells pulled down CD36 (Fig. 6A), and conversely, CD36 IPs pulled down IRβ (Fig. 6B). Proximity ligation assays for in situ visualization of protein interaction (34) showed the typical amplification patterns suggestive of CD36-IRβ proximity (Fig. 6C).
CD36 expression enhanced insulin-induced tyrosine phosphorylation of IRβ and recruitment of the phosphatidylinositide 3-kinase (PI3K) catalytic subunit P85 (Fig. 6D), supporting functional relevance of the CD36-IR interaction. We also examined whether CD36 interacts with IRβ in vivo. IPs of IRβ from mice gastrocnemius and quadriceps muscles pulled down CD36, consistent with interaction (Fig. 6E), and further validation was obtained using PLA, which showed the amplification pattern expected for interacting proteins (Fig. 6F).
Fyn Recruited by CD36 Phosphorylates IR
The mechanism for CD36 effect on IR phosphorylation was investigated next. Src tyrosine kinases mediate most CD36 signaling, and Fyn, in particular, was implicated in its metabolic effects (8,35). Insulin treatment of CHO/IR cells transiently transfected with Fyn resulted in recovery of more pIRβ (Fig. 7A). In contrast, IRβ phosphorylation was enhanced even without insulin when CHO/IR stably expressed CD36 (Fig. 7A), suggesting that some of the overexpressed Fyn was IRβ associated. In addition, more pIRβ was measured in these cells when insulin was added (Fig. 7A and B). These data suggested that Fyn phosphorylates IRβ and that CD36 recruits Fyn to the IR. Consistent with this, insulin treatment or CD36 expression each recruited Fyn to the plasma membrane (Fig. 7C), and cells expressing CD36 and Fyn showed extensive insulin-stimulated colocalization of Fyn and IRβ (Fig. 7C, bottom).
Saturated FA Dissociate Fyn From IR
We showed in Fig. 5 that saturated FA inhibit CD36 action to enhance insulin signaling, so we examined how PA affects Fyn phosphorylation of IRβ with or without insulin. CD36 IPs from CHO/IR/+CD36 cells treated with insulin contained pFynY416 (Fig. 7D), but pFynY416 was absent in IPs of cells pretreated (15 min) with PA. These data suggested insulin promotes interaction of activated Fyn with CD36, and this is disrupted by PA (Fig. 7D and E). Similarly, PA pretreatment abolished Fyn recovery in IPs of IRβ from C2C12 myotubes, which express high endogenous IRβ (Fig. 7F and G).
To further test Fyn’s mediation of CD36’s effect on insulin signaling, we used CHO/IR cells stably expressing the CD36 mutant CD36K/A (11,25). The COOH-terminal cytosolic segment of CD36 is required for Fyn-mediated effects, and this mutant has two COOH-terminal lysines substitution with alanine that impair CD36-induced Fyn signaling (11,25). Cells expressing CD36K/A had diminished insulin signaling compared with native CD36 (Fig. 8A and B) and reduced insulin’s ability to activate Fyn and phosphorylate IRβ (Fig. 8C and D). CD36K/A cells also did not show insulin stimulation of glycolysis (Fig. 8E). Together, the data in Figs. 7 and 8 suggest that insulin induces CD36-Fyn phosphorylation of IRβ, further stimulating glucose utilization. In contrast, PA dissociates the Fyn-CD36-IRβ complex, reducing IRβ phosphorylation and insulin-stimulated glucose metabolism (Fig. 8F).
Low Muscle CD36 Expression Associates With Insulin Resistance and Type 2 Diabetes in Humans
To explore potential human relevance of current findings, we examined whether genetically determined CD36 expression associates with the incidence of insulin resistance or type 2 diabetes (T2D). We applied PrediXcan analysis to a “discovery” cohort of 4,702 patients from Vanderbilt University’s BioVU genomic resource. We found significant associations (P = 10−3 to 10−7) between the genetic component of muscle CD36 expression and disease status. Notably, decreased CD36 expression in muscle associated with increased risk of T2D (Supplementary Table 3).
A similar analysis used the genome-wide association study summary statistics data from the Meta-Analyses of Glucose and Insulin-related traits Consortium. Decreased CD36 expression was found to associate with higher HOMA-insulin resistance (HOMA-IR) or insulin resistance (P = 0.03) (Supplementary Fig. 4A). Furthermore, the single nucleotide polymorphism within the cis region of CD36 (±1 mega base pairs of transcription start site) showed a distribution of P values significantly different from that expected by chance (Supplementary Fig. 4A). This gene region, from which gene expression estimation was generated, contains the rs17236824 variant that was highly significant for association with HOMA-IR (Supplementary Fig. 4B). These data linked genetically reduced CD36 expression with incidence of insulin resistance or T2D.
Major findings of this study are that the FA transporter CD36, which facilitates muscle FA uptake and oxidation, also enhances insulin action to stimulate muscle glucose utilization. CD36 functionally interacts with the IR to enhance its phosphorylation, P85 recruitment, and downstream signaling to promote glucose metabolism. These findings show that metabolic actions of CD36 are more complex than would be expected from its FA uptake function. We previously reported that CD36 signaling regulates oxidation of exogenous FA during fasting by modulating AMPK activation (8). Here we show that CD36 influences postprandial glucose metabolism by modulating insulin action. This dual role of CD36 signaling would influence energy adaptation and homeostasis.
The conditional deletion of CD36 in skeletal muscle showed that CD36 is required for optimal insulin stimulation of glucose metabolism. The systemic phenotype of the smCd36−/− was subtle, likely a limitation of the mosaic effect of the deletion. Still, this mouse provided opportunity to compare glucose metabolism in muscles with and without CD36 depletion from the same animal. We could document specific effect of Cd36 deletion to reduce ceramides and expression of genes of insulin and glucose metabolism. We also showed diminished glucose uptake by muscle in vivo and reduced insulin-stimulated glycolysis and glucose oxidation in muscle explants ex vivo. Together with the findings in myotubes and CHO/IR cells, these data provide strong support for physiological regulation of the IR pathway by CD36 signaling.
Sensitivity of the CD36-IR interaction to inhibition by saturated FA, especially PA, might be relevant to muscle glucose sparing, because PA, the main product of lipogenesis, is preferentially stored as TG by adipose tissue and released during fasting or exercise (36). These data suggest that PA might feedback in adipose tissue to limit lipogenesis by suppressing glucose uptake and that the data provide mechanistic insight into the glucose-FA cycle in muscle. This cycle initially proposed that FA metabolites inhibit muscle glucose utilization (37). Our data implicate membrane signaling as a major FA inhibition site, consistent with a primary role of membrane glucose transport (38,39). Saturated and unsaturated FA transduce markedly different CD36-mediated signaling (8,35), but the mechanisms for the distinctive effects are unclear. They might involve differential FA effects on CD36 localization in plasma membrane subdomains or restrictive properties of the lipid transport tunnel identified inside CD36 (40). CD36 regulation of IR (Fig. 8) and of AMPK (8) might explain CD36 influence on muscle fuel choice between FA and glucose. However, the mechanistic details of how this is accomplished, including downstream effects on transporter translocation, nutrient intracellular trafficking, and targeting to mitochondria (41–44), remain to be elucidated.
Our findings support a beneficial homeostatic role of muscle CD36. The associations identified between genetically determined low CD36 and insulin resistance (HOMA-IR) and T2D (Supplementary Table 3 and Supplementary Fig. 4) are consistent with this, although they are likely to reflect complex causality. Genetic CD36 variants are relatively common and often affect CD36 expression (45,46). These variants have been associated with risk of metabolic syndrome (47) and diabetes or insulin resistance (48,49). Dysfunctional interaction of CD36 with IR, Fyn, and P85 might contribute to its role in the etiology of metabolic disease.
Acknowledgments. The authors thank Dr. Jacob Bar-Tana (Hebrew University, Jerusalem) for the M16 and Meghan Lam (Washington University in St. Louis) for technical support.
Funding. This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases grants R01-DK-33301 and R01-DK-111175 and by pilot and feasibility awards to D.S. from P30-DK056341 (Nutrition Obesity Research Center), P30-DK020579 (Diabetes Research Center), and Longer Life Foundation (2017-007). This work was also supported by the Hope Center Alafi Neuroimaging Laboratory and National Institutes of Health Shared Instrumentation Grant (S10-RR-027552).
Duality of Interest. No potential conflicts of interest relevant to this article were reported.
Author Contributions. D.S. designed experiments, obtained and analyzed data, and wrote the manuscript. P.D. conducted experiments and data analysis. T.P., M.J.-S., N.-H.S., C.R.F., and K.I.S. helped conduct the experiments and analyze data. E.P. performed animal breeding and genotyping. K.L.H. conducted histological analysis. I.J.G. reviewed the manuscript. E.R.G. analyzed human genetic data. N.A.A. designed experiments, analyzed data, and wrote the manuscript. D.S. and N.A.A. are the guarantors of this work and, as such, had full access to all the data in the study and take responsibility for the integrity of the data and the accuracy of the data analysis.