Glucose Metabolism Is Required for Platelet Hyperactivation in a Murine Model of Type 1 Diabetes

Patients with type 1 diabetes mellitus (T1DM) display increased platelet activation and thrombosis (1–3). The mechanisms underlying this dysfunction are incompletely understood. T1DM is accompanied by multiple systemic abnormalities, including hyperglycemia, dyslipidemia, increased inflammation, and an altered hormonal milieu, any of which could contribute to platelet dysfunction. Platelets from diabetic rats display an enrichment of glycolytic intermediates (4); however, whether this is due to increased glycolytic flux or an impaired ability to metabolize glucose is unclear. Furthermore, platelet activation correlates with glucose metabolism. Platelets incubated in hyperglycemic conditions demonstrate increased platelet activation (5), and platelets lacking the ability to take up glucose display decreased platelet activation and in vivo thrombosis (6). In T1DM, the specific contribution of glucose uptake to platelet dysfunction is unknown.

To determine the direct consequence of altered glucose utilization on platelet function, we induced insulin-deficient diabetes (T1DM) in mice lacking the ability to metabolize glucose in platelets. These mice with platelet-specific double-knockout (DKO) of GLUT1 and GLUT3 are unable to take glucose up from the extracellular compartment (6). Using these mice, we evaluated the hypothesis that increased glucose metabolism in platelets from T1DM mice promoted platelet hyperactivation and thrombosis. Here we show that platelets from mice given streptozotocin (STZ) to model T1DM exhibited increased glucose uptake and glycolysis that correlated with increased GLUT3 protein expression. Furthermore, platelets from STZ-induced diabetic mice exhibited increased platelet activation and thrombosis, which was prevented when platelet glucose metabolism was abrogated.

All animals were generated on a C57Bl6 background and housed under normal light and temperature conditions. DKO mice were generated as previously described (6) by crossing mice expressing a platelet factor 4 driven Cre recombinase to mice harboring homozygous GLUT1 and GLUT3 floxed alleles.

For STZ administration, 8- to 12-week-old mice were injected i.p. with 50 mg/kg STZ or citrate buffer daily for 5 days (7). All studies were conducted on mice between 7 and 12 weeks after STZ administration. The institutional animal care and use committee at the University of Iowa and the University of Utah approved all animal studies.

Blood was isolated into 1:20 acid citrate dextrose through carotid artery cannulation as previously described (8). For GLUT1 protein analysis and Seahorse analysis, platelets were negatively depleted of leukocytes and red blood cells by incubation with CD45 and Terr119 microbeads (Milteny Biotec, Cambridge, MA). Platelet counts were determined by Cellometer Auto M10 (Nexcelom Bioscience, Lawrence, MA).

Platelet counts were obtained using the Advia 120 whole-blood analyzer (Siemens, Berlin, Germany). For circulating half-life and depletion assays, platelet concentrations were determined as CD41-APC⁺ events normalized to fluoSpheres (BD Biosciences, San Jose, CA). For circulating half-life, mice were injected i.v. with anti-GPIbβ-FITC antibody (Emfret Analytics, Eibelstadt, Germany), and blood was obtained via cheek bleed. For depletion studies, DKO and littermate control mice were injected i.v. with 2 μg/g anti-GPIbα antibody (Emfret Analytics), and platelet counts were obtained every 24 h.

Plasma glucose concentrations were determined with a Contour clinical glucometer (Bayer, Leverkusen, Germany). Serum free fatty acids (FFA) were measured using an FFA assay kit (MAK044; MilliporeSigma, St. Louis, MO).

Proplatelet formation assays were determined as previously described with some modifications (6,9). Bone marrow from DKO or control mice was cultured in DMEM with 5 mmol/L glucose, glutamate, and recombinant thrombopoietin for 5 days. At day 6, megakaryocytes were enriched using a gravity BSA gradient and plated into fibrinogen-coated chamber slides. Megakaryocytes were then treated with 42 μmol/L BSA (vehicle), 250 μmol/L palmitate complexed with BSA (Cat #102720-100; Agilent, Santa Clara, CA), 250 μmol/L oleic acid, or 250 μmol/L myristic acid (Nu-Chek Prep, Elysian, MN) for 24 h. Oleic and myristic acid were conjugated to BSA in the presence of NaOH, then neutralized to pH 7.2. The fraction of proplatelet-forming megakaryocytes to total megakaryocytes was then quantified in a blinded manner examining >100 megakaryocytes/biological replicates.

To determine glucose uptake, washed platelets in 1 mmol/L glucose DMEM were incubated with 10 mmol/L [³H]2-deoxy-d-glucose for 10 min with or without 0.5 units/mL thrombin. Platelets were washed then lysed in 1 mol/L NaOH. Glucose uptake was normalized to protein content using bicinchoninic acid analysis.

Seahorse analysis was conducted as previously described (6). Briefly, platelets were negatively depleted of leukocytes and red blood cells. Platelets bioenergetics were evaluated by Seahorse XF96 Analyzer (Agilent Technologies). Data were normalized to platelet counts.

Platelets were lysed in radioimmunoprecipitation assay buffer and analyzed via Western blot. For GLUT1 protein analysis, platelets were depleted of leukocytes and red blood cells. Primary antibodies to GLUT1 (07-1401) and GLUT3 (AB1344) were purchased from MilliporeSigma. ImageJ software (National Institutes of Health) was used for densitometry quantification.

Washed platelets were incubated in HEPES Tyrode’s buffer for 1 h, then incubated in the presence of the indicated agonist with JonA-PE, CD62p-FITC (Emfret Analytics), and CD41-APC (eBioscience) for 10 min at 37°C. Reactions were stopped by the addition of FACS lysis buffer (Becton Dickinson, Franklin Lakes, NJ) and analyzed using flow cytometry LSR II (Becton Dickinson).

Collagen/epinephrine-induced pulmonary embolism was conducted as previously described (6,10). Briefly, mice were injected i.v. with 430 μg/kg collagen (Chrono-Log, Columbia, MD) and 20 μg/kg epinephrine (MilliporeSigma) in PBS. Survival was determined when spontaneous respiratory chest expansions ceased for 1 min.

Statistical analyses were performed using GraphPad 7 or Microsoft Excel 2011 software. Data are presented as mean ± SD. The statistical significance threshold of P < 0.05 was determined.

To evaluate platelet glucose metabolism in a model of T1DM, we gave mice lacking both GLUT1 and GLUT3 specifically in platelets (DKO) and littermate controls STZ or citrate buffer. Mice subjected to STZ (control-STZ and DKO-STZ) displayed an approximately twofold increase in circulating glucose concentrations at 4 weeks, which was sustained for up to 12 weeks (Fig. 1A). STZ administration did not alter circulating platelet counts in control mice (Fig. 1B). We previously reported decreased circulating platelets in DKO mice (6). Interestingly, 6 and 8 weeks after STZ administration, circulating platelet counts were increased in DKO-STZ mice to levels similar to controls (Supplementary Table 1 and Fig. 1B). To determine whether increased platelet counts were due to decreased clearance, we monitored the platelet circulating half-life. DKO mice exhibited decreased circulating half-life; however, STZ administration did not increase survival (Fig. 1C). We therefore investigated platelet biogenesis. To do this we depleted platelets by injecting mice with anti-GPIbα antibodies and monitored the time of platelet regeneration. No change in platelet regeneration was observed in control-STZ mice. DKO mice demonstrated a significantly longer time to platelet regeneration (Fig. 1D). Platelet regeneration was increased in DKO-STZ mice to levels similar to controls. These data indicate that increased platelet counts in DKO-STZ mice are due to increased platelet production relative to nondiabetic DKO mice.

We sought to determine mechanisms responsible for normalizing platelet production in diabetic DKO mice. Because we have previously demonstrated that DKO platelets and megakaryocytes have very little glucose uptake and metabolism (6), we thought it unlikely that hyperglycemia was directly influencing megakaryocyte function. We therefore considered the possibility that increased availability of FFA in the diabetic state enhanced megakaryocyte FFA metabolism, which compensated for the absence of glucose uptake. Circulating FFA were increased in diabetic mice (Fig. 2A). Incubation of bone marrow–derived megakaryocyte cultures with BSA-conjugated palmitic acid, oleic acid, and myristic acid increased proplatelet formation in DKO megakaryocytes (Fig. 2B and C); however, no FFA-mediated increase was observed in controls. These data suggest that in the absence of glucose metabolism, increased FFA availability and presumably increased utilization can promote proplatelet formation.

Platelets from diabetic wild-type mice exhibited significantly increased basal glucose uptake relative to nondiabetic controls (Fig. 3A). After thrombin administration, glucose uptake increased approximately threefold and twofold, respectively, in nondiabetic and STZ-administered wild-type mice, such that absolute levels of glucose uptake were equivalent (Fig. 3A). Similar to our previous findings, DKO platelets displayed negligible glucose uptake (6), which was not altered by diabetes (Fig. 3A). We next evaluated glycolysis rates using Seahorse flux analysis. DKO platelets demonstrated significantly impaired glycolysis (Fig. 3B). Under basal conditions, hyperglycemia did not alter glycolysis in platelets isolated from wild-type mice (Fig. 3B); however, after administration of mitochondrial inhibitors, platelets from diabetic wild-type mice displayed increased glycolysis relative to nondiabetic controls (Fig. 3B). These data support the conclusion that platelet glucose utilization is increased in a murine model of T1DM.

GLUT1 and GLUT3 are the physiologically relevant GLUTs in platelets (6). Because glucose uptake was increased, we examined GLUT expression. GLUT1 is expressed in red blood cells and leukocytes; therefore, to examine GLUT1 protein expression, platelet preparations were depleted of cells positive for Ter119 and CD45. GLUT1 protein content was unchanged in platelets isolated from diabetic wild-type mice relative to nondiabetic controls (Fig. 3C and D). However, GLUT3 platelet protein content was significantly increased by diabetes (Fig. 3C and E). Diabetes increases GLUT3 protein content in the placenta (11) and neuronal tissue (12), suggesting glucose-dependent regulation of GLUT3. Together these findings reveal that platelets from mice given STZ increase GLUT3 protein expression that may contribute to increased glucose uptake and glycolysis.

Platelets from patients with T1DM and platelets incubated under hyperglycemic conditions display increased platelet activation (1–3,5). Therefore, we hypothesized that increased platelet glucose metabolism results in increased activation. Under basal conditions, no differences in platelet activation marked by GPIIbIIIa activation (relative JonA geometric mean fluorescence intensity) or α-granule release (CD62p geometric mean fluorescence intensity) were observed (Fig. 4A and B). In response to submaximal concentrations of protease-activated receptor 4 (PAR4) peptide and convulxin, nondiabetic and diabetic wild-type platelets both revealed increased activation. However, platelet activation was significantly higher in diabetic mice relative to similarly treated controls (Fig. 4A and B). Consistent with our previous reports, DKO platelets demonstrated blunted activation (6). Importantly, diabetic DKO platelets did not display an additional increase in agonist-mediated activation relative to DKO platelets (Fig. 4A and B). This insensitivity was not due to STZ-induced toxicity or suppression of platelet activation, because in response to maximal doses of PAR4 peptide, DKO and DKO-STZ platelets exhibit equivalent degrees of activation relative to submaximal agonist concentrations (Fig. 4A and B). Together, these data indicate that glucose metabolism is a critical mediator of the hyperglycemia-associated increase in agonist-mediated activation.

To determine whether diabetes increases in vivo thrombosis, we subjected mice to collagen/epinephrine-induced pulmonary embolism. This in vivo assay is believed to be largely driven by platelet activation versus endothelial changes that could also develop in diabetes. Control-STZ mice demonstrated significantly reduced length of survival relative to controls (Fig. 4C). In contrast, DKO and DKO-STZ mice both exhibited prolonged survival relative to control groups; however, no significant change in survival was observed between DKO and DKO-STZ mice (Fig. 4C). Thus, reducing platelet glucose utilization protects against diabetes-associated platelet hyperactivation.

This study identifies an important role for glucose utilization in the increased platelet activation that characterizes diabetes and also identifies a potential role for FFA in maintaining proplatelet biogenesis when platelet glucose utilization is impaired. Diabetes reversed thrombocytopenia in mice lacking the ability to import glucose into platelets and megakaryocytes, likely through increased FFA utilization. Thus, hyperglycemia and increased FFA availability exhibit distinct effects on platelet biogenesis versus activation, with glucose or FFA being interchangeable for proplatelet biogenesis from megakaryocytes, whereas glucose is required for platelet activation.

Here we demonstrate that in a model of T1DM, platelets increase glucose uptake and glycolysis in concert with increased GLUT3 expression. Importantly, when glucose utilization is blocked, the hyperglycemia-associated increase in platelet function and thrombosis is abolished, indicating that glucose metabolism in the T1DM model drives platelet hyperactivation. These findings indicate that inhibition of glucose metabolism in platelets in individuals with diabetes could ameliorate platelet hyperfunction, providing a potential therapeutic opportunity to reduce the associated increased risk of thrombosis. Metabolic inhibitors have become a promising target in cancer therapeutics, that some of these glycolytic inhibitors could be repurposed to decrease glucose metabolism in diabetic platelets may be possible. Future work to elucidate the mechanisms responsible for GLUT3 protein induction in T1DM may also reveal additional therapeutic targets.

Acknowledgments. The data presented herein were obtained at the Flow Cytometry Facility, which is a Carver College of Medicine core research facility at the University of Iowa.

Funding. This work was supported by National Institutes of Health grants TL1-TR-001875 to T.P.F., R01-AG-048022 to M.T.R., R01-HL-126547-01 to A.S.W., and U54-HL-112311 to A.S.W. and E.D.A., who are both established investigators of the American Heart Association.

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

Author Contributions. T.P.F. contributed to writing the original draft. T.P.F., A.M., K.G., E.A.M., and R.A.C. contributed to the investigation. T.P.F., R.A.C., M.T.R., A.S.W., and E.D.A. conceptualized the study and contributed to study methodology. T.P.F., A.S.W., and E.D.A. contributed to writing, reviewing, and editing the manuscript. R.A.C., M.T.R., A.S.W., and E.D.A. supervised the study. A.S.W. and E.D.A. acquired funding and contributed to resources. T.P.F. and E.D.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.

Prior Presentation. Parts of this study were presented in abstract form at the American Heart Association’s Vascular Discovery: From Genes to Medicine Scientific Sessions 2018, San Francisco, CA, 10–12 May 2018.