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Enhanced Glucose Transport, but not Phosphorylation Capacity, Ameliorates Lipopolysaccharide-Induced Impairments in Insulin-Stimulated Muscle Glucose Uptake

Otero, Yolanda F.; Mulligan, Kimberly X.; Barnes, Tammy M.; Ford, Eric A.; Malabanan, Carlo M.; Zong, Haihong; Pessin, Jeffrey E.; Wasserman, David H.; McGuinness, Owen P.

doi: 10.1097/SHK.0000000000000550
Basic Science Aspects
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ABSTRACT Lipopolysaccharide (LPS) is known to impair insulin-stimulated muscle glucose uptake (MGU). We determined if increased glucose transport (GLUT4) or phosphorylation capacity (hexokinase II; HKII) could overcome the impairment in MGU. We used mice that overexpressed GLUT4 (GLUT4Tg) or HKII (HKTg) in skeletal muscle. Studies were performed in conscious, chronically catheterized (carotid artery and jugular vein) mice. Mice received an intravenous bolus of either LPS (10 μg/g body weight) or vehicle (VEH). After 5 h, a hyperinsulinemic-euglycemic clamp was performed. As MGU is also dependent on cardiovascular function that is negatively affected by LPS, cardiac function was assessed using echocardiography. LPS decreased whole body glucose disposal and MGU in wild-type (WT) and HKTg mice. In contrast, the decrease was attenuated in GLUT4Tg mice. Although membrane-associated GLUT4 was increased in VEH-treated GLUT4Tg mice, LPS impaired membrane-associated GLUT4 in GLUT4Tg mice to the same level as LPS-treated WT mice. This suggested that overexpression of GLUT4 had further benefits beyond preserving transport activity. In fact, GLUT4 overexpression attenuated the LPS-induced decrease in cardiac function. The maintenance of MGU in GLUT4Tg mice following LPS was accompanied by sustained anaerobic glycolytic flux as suggested by increased muscle Pdk4 expression, and elevated lactate availability. Thus, enhanced glucose transport, but not phosphorylation capacity, ameliorates LPS-induced impairments in MGU. This benefit is mediated by long-term adaptations to the overexpression of GLUT4 that sustain muscle anaerobic glycolytic flux and cardiac function in response to LPS.

*Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Tennessee

College of Medicine, University of Tennessee Health Science Center, Memphis, Tennessee

Department of Internal Medicine, University of Michigan, Ann Arbor, Michigan

§Department of Medicine and Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, New York

Address reprint requests to Owen P. McGuinness, PhD, Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, 702 Light Hall, Nashville, TN 37232-0615. E-mail: owen.mcguinness@vanderbilt.edu

Received 25 September, 2015

Revised 14 October, 2015

Accepted 9 December, 2015

This work was supported by National Institutes of Health Grants DK-078188, DK-043748, and DK-033823. OPM also received support from DK020593 and DK059637. KXM and TMB were supported by the Molecular Endocrinology Training Grant T32-DK-07563.

YFO and KXM contributed equally to this manuscript.

The authors report no conflicts of interest.

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INTRODUCTION

Hospitalized individuals especially in those with an accompanying infection are prone to hyperglycemia. Although there was initial enthusiasm that severe glucose control in the hospitalized setting would improve outcomes in the critically ill patient, subsequent data have observed negative outcomes (1, 2). The main approach to normalize glycemia is to administer exogenous insulin. Unfortunately, this increases the risk of hypoglycemia (3, 4). Thus, a better understanding of the factors that contribute to infection-induced insulin resistance is needed to better control hyperglycemia in the hospitalized setting.

Infection and lipopolysaccharide (LPS), a potent proinflammatory factor of gram-negative bacteria, impair muscle glucose uptake (MGU) (5–7). Skeletal muscle represents the bulk of insulin-sensitive tissue in the body, and is a major site of glucose disposal. The regulation of glucose uptake in the skeletal muscle is distributed over three steps: vascular delivery of glucose to the tissue and its subsequent diffusion into the interstitium; facilitated transport into the intracellular space by glucose transporters (GLUT); and irreversible phosphorylation of glucose to glucose-6-phosphate by hexokinase (HK) (8). Each step is regulated by a number of factors that can be modulated by insulin as and impaired by LPS (9, 10).

It is unknown whether targeting one or more of those steps would improve MGU in response to LPS. Several studies in other settings where insulin action in muscle is altered (obesity and exercise) have increased or decreased the amounts of GLUT4 and HK to assess the importance of transport and glucose phosphorylation in determining insulin action (11–13). GLUT4 is the main GLUT responsible for insulin-stimulated glucose uptake, and it is mainly expressed in skeletal muscle, heart, and adipose tissue, and translocates to the plasma membrane upon insulin stimulation. In the current study, we used transgenic mouse models that overexpress HKII or GLUT4 in skeletal muscle to determine if lowering these barriers to glucose uptake will improve insulin-stimulated MGU in the presence of LPS. We observed that improving glucose transport (GLUT4 overexpression), but not phosphorylation capacity (HKII overexpression), improved basal and insulin-stimulated MGU. Surprisingly, GLUT4 overexpression did not attenuate the LPS-induced decrease in membrane-associated GLUT4. It did, however, attenuate the LPS-induced decrease in cardiac function, and upregulate anaerobic glycolysis, which may have contributed to the improvement in MGU.

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METHODS

Animal care and husbandry

Mice on a C57Bl/6J background selectively overexpressing HKII (HKIITg) in skeletal muscle using a transgene containing the human HKII cDNA driven by the rat muscle creatine kinase promoter, and mice transgenic for the hGLUT4-11.5 construct (GLUT4Tg) were studied (14, 15). At 3 weeks of age, mice were separated by sex and maintained in microisolator cages on a 12-h light/dark cycle with free access to food and water. All experiments were performed on male mice that were HKIITg, GLUT4Tg, or wild-type (WT) littermates at approximately 3 months of age. All procedures performed were approved by the Vanderbilt University Institutional Animal Care and Use Committee.

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Surgical procedures

Chronic catheters were surgically implanted as previously described (16). Briefly, mice were anesthetized with isoflurane. The left carotid artery and right jugular vein were catheterized for sampling and infusions, respectively. The free ends of the catheters were tunneled under the skin to the back of the neck where the loose ends of the catheters were attached via stainless steel connectors to tubing made of Micro-Renathane, which were exteriorized and sealed with stainless steel plugs. Animals were individually housed after surgery, and body weight was recorded daily.

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In vivo metabolic experiments

All metabolic experiments were performed following an approximately 5-day postoperative recovery period as previously described (5). For metabolic studies, conscious, unrestrained catheterized mice were placed in approximately 1-L plastic container lined with bedding and fasted at 7:00 AM (t = −300 min). The animals were immediately connected to a Dual Channel Stainless Steel Swivel (Instech Laboratories, Plymouth Meeting, Pa) to allow simultaneous jugular vein infusion and sampling of arterial blood. Mice were not handled on the day of study and allowed to move freely to eliminate stress. At t = −240 min, the mice received a bolus of vehicle (VEH; saline) or Escherichia coli endotoxin (E. coli 011:B4; Lot#129K4025, Sigma-Aldrich, St. Louis, MO) at 10 μg/g BW into the jugular vein catheter. A 5 μCi bolus of 3-[3H]-D-glucose was given into the jugular vein at t = −120 min followed by a constant infusion at a rate of 0.05 μCi/min. Following a 2-h equilibrium period (t = 0 min), a baseline arterial blood sample was drawn for measurement of blood glucose, 3-[3H]-D-glucose, hematocrit, plasma insulin, and plasma cytokine levels. Red blood cells from a donor mouse on a C57Bl/6j background washed with 0.9% heparinized saline were infused at a rate of 4 μL/min for the duration of the study to minimize falls in the hematocrit.

Plasma glucose levels and tissue glucose uptake were assessed in six groups [WT VEH (n = 7), WT LPS (n = 8), HKIITg VEH (n = 8), HKIITg LPS (n = 8), GLUT4Tg VEH (n = 8), GLUT4Tg LPS (n = 7)] 6 h after administration of LPS or VEH before initiation of the clamp (i.e., preclamp). A hyperinsulinemic-euglycemic (120 mg/dL) clamp was used to assess insulin action in six groups (WT VEH [n = 8], WT LPS [n = 8], HKIITg VEH [n = 9], HKIITg LPS [n = 9], GLUT4Tg VEH [n = 13], GLUT4Tg LPS [n = 10]). As we previously observed that LPS impairs insulin clearance, a lower dose of insulin (2 vs. 4 mU/kg/min) was used in LPS-treated animals to match arterial insulin concentration between groups (5). Arterial blood samples were taken every 10 min to determine blood glucose levels. At t = 70, 80, 90, and 100 min, blood samples were taken to determine 3-[3H]-D-glucose. At t = 100 min, a 13 μCi bolus of 2-deoxy [14C] glucose ([2-14C]DG) was administered into the jugular vein catheter to assess tissue glucose uptake. At t = 102, 105, 110, 115, and 125 min, arterial blood was sampled to determine blood glucose, plasma 3-[3H]-D-glucose, and [2-14C]DG. At t = 125 min, a final arterial blood sample was taken to assess all hormones and plasma [2-14C]DG. The mice were then anesthetized. The soleus, gastrocnemius, superficial white vastus lateralis (SVL), gonadal adipose tissue (AT), liver, heart, and brain were excised, immediately frozen in liquid nitrogen, and stored at −70°C until future tissue analysis.

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Plasma and muscle sample analysis

Immunoreactive insulin was assayed using a Linco Rat Radioimmunoassay kit (Linco Research, Inc, St. Charles, Mo) and a double antibody method. Plasma free fatty acid levels were measured enzymatically using a commercially available kit (NEFA C; Wako Chemicals, Richmond, Va). Plasma lactate levels were determined in a 96-well plate using a modification of the method of Lloyd et al. (17). To measure 3-[3H]-D-glucose and [2-14C]DG in the plasma, samples were deproteinized with barium hydroxide and zinc sulfate, dried, and radioactivity was determined using liquid scintillation counting (TriCarb Liquid Scintillation Analyzer; PerkinElmer Life and Analytical Sciences, Downers Grove, Ill). Excised soleus, gastrocnemius, SVL, gonadal AT, heart, and brain were deproteinized with perchloric acid and subsequently neutralized to a pH of approximately 7.5. A portion of the sample was counted ([2-14C]DG and [2-14C]DG-G-phosphate ([2-14C]DGP), whereas a portion was treated with Ba(OH)2 and ZNSO4, and the supernatant was counted ([2-14C]DG). Both [2-14C]DG and [2-14C]DG-G-phosphate ([2-14C]DGP) radioactivity levels were determined using liquid scintillation counting.

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Calculations

Glucose flux rates were assessed using non–steady-state equations assuming a volume of distribution (130 mL/kg). Tissue-specific clearance (Kg) of [2-14C]DG and glucose uptake (Rg) were calculated as previously described (5):

where [2-14C]DGPtissue is the [2-14C]DGP radioactivity (dpm/g) in the tissue, AUC [2-14C]DGplasma is the area under the plasma [2-14C]DG disappearance curve (dpm/mL/min), and [glucose]plasma is the average blood glucose (μg/μL) during the experimental period.

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Echocardiography

Transthoracic echocardiography was performed using a system (Sonos 5500; Agilent, Andover, Mass) with a 15-MHz high frequency linear transducer at a frame rate of 100 frames/s. All images were acquired at a depth setting of 20 mm. Images were acquired at two time points: t = 0 min, which represents the time immediately before an injection of VEH or LPS and at t = 360 min. The mouse was picked up at the nape of its neck, and an ultrasound-coupling gel was applied to the precordium with the ultrasound probe. Two-dimensional targeted M-mode echocardiographic images were obtained at the level of the papillary muscles from the parasternal short-axis views and recorded at a speed of 150 cm/s (maximal temporal resolution) for measurements of heart rate. All other measurements were made on-screen using the digitally recorded signals. Stroke volume and cardiac output were calculated using the equations shown below:

where LVIDd and LVIDs are left ventricular internal dimensions during diastole and systole, respectively.

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Real-time quantitative PCR

Heart tissues were homogenized in TRIzol (Ambion RNA, Carlsbad, Calif), and mRNA was extracted. cDNA was obtained using a cDNA Reverse Transcription kit (Applied Biosystems, Lot# 66105, Foster City, Calif) and stored at −20°C. PCR amplification via Applied Biosystems system and TaqMan probes (for Pdk4 and Nos2) allowed mRNA quantification using the ΔΔCt method. Gene expression was normalized using Gapdh as a housekeeping gene.

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Glucose-6-phosphate assay

Heart and muscle were homogenized in perchloric acid, and supernatant was neutralized with KOH (1 M). Samples and standards were incubated in a buffer containing ATP (adenosine triphosphate), NADP (nicotinamide adenine dinucleotide phosphate), and glucose-6-phosphate dehydrogenase. NADPH (dihydronicotinamide adenine dinucleotide phosphate) fluorescence was measured.

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Immunofluorescence microscopy

Tissues isolated from mice that underwent a hyperinsulinemic-euglycemic clamp were embedded in optimal cutting temperature compound. The frozen tissue cross-sections (10 μM) were blocked with 3% bovine serum albumin in phosphate buffered saline (pbs) for 60 min at room temperature. Primary antibodies were used at the following dilutions: GLUT4 polyclonal antibody (1:1,000) was obtained from East Acres Biologicals Inc, Southbridge, MA, and α-dystroglycan (α-DG) monoclonal antibody (1:50) was a kind gift from Dr. Kevin Campbell (The University of Iowa, Iowa City, Iowa), and used as previously described (18). Fluorescently conjugated secondary antibodies (1:100, Jackson ImmunoResearch Laboratories, West Grove, Pa) were added to the sections for 30 min at room temperature. After extensive washing with PBS, the slides were mounted with Vectashield Mounting Medium (Vector Laboratories, Burlingame, Calif). The slides were visualized by confocal fluorescent microscopy (model LSM510; Carl Zeiss MicroImaging, Inc, Thornwood, NY). Quantification of cell surface immunofluorescence was performed using NIH Image software. Briefly, individual muscle fibers were delineated by α-DG labeling and were circumscribed to obtain the total GLUT4 fluorescent intensity. The intracellular GLUT4 fluorescent intensity of the same fiber was determined by circumscribing approximately 1 μm below the α-DG-labeled surface. The difference between the total and intracellular signals was defined as the cell surface GLUT4. The cell surface GLUT4 intensity normalized for the intracellular GLUT4 intensity was plotted as the relative cell surface GLUT4.

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Data analysis

Data are presented as the mean ± SEM. Differences between groups were compared by two-way ANOVA and by Student t test when appropriate. The significance level was set at P < 0.05.

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RESULTS

GLUT4 but not HKII overexpression inhibits the LPS-induced decrease in baseline MGU

It has previously been shown that the GLUT4 transgene increases GLUT4 protein content 2- to 4-fold in skeletal muscle of C57BL/6 mice (14). In the presence of a saline infusion, Glut4Tg mice had the expected lower baseline (i.e., without exogenous insulin stimulation) insulin and glucose levels 6 h after VEH treatment than their WT counterparts. Although LPS led to a significant decrease in insulin levels in WT mice, the insulin levels observed in Glut4Tg mice treated with LPS during saline infusion failed to decrease further relative to VEH controls (Fig. 1). This was also reflected in the fasting glucose concentrations. WT mice that received VEH had higher baseline glucose concentrations than WT mice that received LPS and Glut4Tg mice that received either VEH or LPS. Tracer-determined glucose turnover rate was increased in Glut4Tg compared with WT mice treated with VEH (13.2 ± 2.0 vs. 16.4 ± 1.3 mg/kg/min; WT vs. Glut4Tg), but it decreased to similar levels in the presence of LPS (9.8 ± 3.1 vs. 11.6 ± 1.3 mg/kg/min). LPS decreased baseline MGU in WT mice. However, Glut4Tg mice were protected from the LPS-induced decrease in baseline MGU (Fig. 2, A and B).

Fig. 1

Fig. 1

Fig. 2

Fig. 2

In contrast to GLUT4 overexpression, HKII overexpression did not prevent the LPS-induced decrease in basal MGU. The HKII transgene is overexpressed 3-, 5-, and 7-fold in the soleus, gastrocnemius, and SVL, respectively (15). The basal glucose concentrations were similar between WT and HKII with VEH and LPS treatment (Fig. 1), and there was neither significant improvement in baseline tissue glucose uptake nor LPS-induced tissue glucose uptake (Fig. 2, C and D). There were no significant differences in insulin levels between HKIITg and WT mice with VEH treatment. LPS decreased baseline insulin to a similar extent as WT (Fig. 1).

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GLUT4 but not HKII overexpression inhibits LPS-induced decreases in insulin-stimulated MGU

A hyperinsulinemic-euglycemic clamp was used to assess insulin action in GLUT4Tg, HKIITg, and WT mice. Insulin levels increased during the clamp and were matched between both groups with VEH and LPS treatment, and glucose concentrations were maintained at approximately 120 mg/dL (Fig. 3 and Fig. 4A). Both GLUT4Tg and WT mice had similar glucose infusion rate (GIR; mg/kg/min) with VEH treatment (59.5 ± 0.2 vs. 54.2 ± 0.5, respectively; Fig. 4B). However, in GLUT4Tg mice, LPS did not impair the GIR to the same extent as WT mice (decreased by 23% vs. 35%; 45.6 ± 0.2 vs. 35.4 ± 0.6; Fig. 4B). LPS decreased glucose uptake in the gastrocnemius and SVL of WT mice compared with VEH treatment. In contrast, in GLUT4Tg mice, LPS failed to suppress MGU (Fig. 4, C and D).

Fig. 3

Fig. 3

Fig. 4

Fig. 4

On the contrary, under hyperinsulinemic-euglycemic clamp conditions, HKIITg and WT mice had a similar GIR with VEH treatment, and GIR decreased similarly with LPS treatment (54.6 ± 0.5 vs. 54.2 ± 0.5; 35.6 ± 0.4 vs. 35.4 ± 0.6, respectively; Fig. 5B). This translated into a significant decrease in glucose uptake in the gastrocnemius, SVL, and heart in both HKIITg and WT mice with LPS treatment (Fig. 5, C and D).

Fig. 5

Fig. 5

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GLUT4 overexpression increases arterial lactate levels

GLUT4Tg mice had significantly higher arterial lactate levels following VEH treatment than WT mice (Fig. 6). LPS modestly increased lactate concentration in WT mice. LPS did not further increase the already elevated lactate in GLUT4Tg mice. However, during the clamp, lactate concentration markedly increased in GLUT4Tg mice. In HKTg lactate was similar to that seen in WT mice. As overexpression of HKII had no phenotype in terms of alterations in MGU, GIR, or plasma lactate, we focused on further characterizing the phenotype of the GLUT4 overexpressing mice.

Fig. 6

Fig. 6

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GLUT4 overexpression increases plasma nonesterified free fatty acids (NEFAs)

Increased NEFAs can impair MGU. We determined if the improvement in insulin-stimulated MGU in response to LPS observed with overexpression of GLUT4 could be due to improved suppression of plasma NEFAs during the hyperinsulinemic-euglycemic clamp. In fact, this was not the case. During the basal period, GLUT4Tg tended to have higher plasma NEFA levels with VEH treatment (Table 1). However, during the clamp, plasma NEFAs did not decrease in GLUT4Tg to the same extent as observed in WT mice. With LPS treatment, plasma NEFA levels were increased approximately 2-fold during the preclamp period (i.e., before initiation of the clamp but 6 h after treatment with LPS) and decreased during the clamp to a similar extent in GLUT4Tg compared with WT mice.

Table 1

Table 1

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GLUT4 overexpression does not prevent LPS-induced decrease in plasma membrane-associated GLUT4

GLUT4 association with the plasma membrane in gastrocnemius muscle of mice that underwent a hyperinsulinemic-euglycemic clamp was examined by immunohistochemistry. The muscle fiber cell surface distribution of GLUT4 was co-localized with the sarcolemma marker protein α-DG. With VEH treatment, there was a relative increase in cell surface-associated GLUT4 in transgenic compared with WT mice. LPS decreased cell surface GLUT4 to similar levels in both WT and transgenic mice (Fig. 7). As an increase in transport activity can be offset by a corresponding defect in downstream glycolysis, we examined whether overexpression of GLUT4 would lead to a corresponding increase in glucose-6-phosphate in the presence of inflammatory stress. In fact, glucose-6-phosphate content (μmol/mg) in the SVL (3.2 ± 0.1 vs 3.7 ± 0.1; WT vs. GLUT4Tg) and heart (5.9 ± 0.8 WT vs. 6.6 ± 1.0 GLUT4Tg) in LPS-treated animals were unaltered by overexpression of GLUT4. This suggests that events occurring outside the myocyte contributed to the protection afforded by overexpression of GLUT4.

Fig. 7

Fig. 7

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GLUT4 overexpression improves cardiac function and increases Pdk4 expression

The effect of LPS treatment on cardiovascular parameters in GLUT4Tg compared with WT mice was examined. Stroke volume, heart rate, and cardiac output decreased in both groups treated with LPS compared with their VEH-treated counterparts (Table 2). However, although LPS treatment led to an approximately 50% decrease in stroke volume and cardiac output in WT animals, the fall was attenuated in GLUT4Tg mice.

Table 2

Table 2

To determine if the cardiac protection for LPS with GLUT4 overexpression was due to altered cardiac glucose oxidation or attenuated cardiac inflammation, we assessed cardiac gene expression of Pdk4 and Nos2 (i.e., iNOS). Cardiac muscle Pdk4 expression was increased in GLUT4Tg following VEH treatment, which is indicative of a decrease in glucose oxidation (Fig. 8). LPS treatment markedly increased Pdk4 expression in WT, but did not further elevate expression in GLUT4Tg. Cardiac muscle Nos2 expression was decreased in GLUT4Tg following VEH (Fig. 8). LPS increased Nos2 expression in WT and GLUT4Tg to a similar level, suggesting cardiac inflammation was augmented to a similar extent.

Fig. 8

Fig. 8

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DISCUSSION

LPS induces marked insulin resistance. We enhanced the glucose transport or phosphorylation capacity of the muscle to determine if either could overcome the LPS-induced impairment in insulin action in vivo. We demonstrate that increasing the ability to transport glucose, but not phosphorylate glucose, prevents the LPS-induced impairment in MGU in vivo. Surprisingly, GLUT4 overexpression did not preserve membrane-associated GLUT4 at the sarcolemmal membrane in the presence of LPS. It did, however, attenuate the LPS-induced impairment in cardiac output and induced adaptations that sustained anaerobic glycolysis. Our previous findings suggested that alterations in muscle glucose delivery play a critical role in insulin resistance resulting from LPS (5, 19). These results suggest that the protective effect of GLUT4 overexpression is not due to maintenance of transport capacity but, in part, due to the maintenance of cardiac function in response to LPS.

Overexpression of GLUT4, but not HKII, attenuated the LPS-induced decrease in baseline MGU. In GLUT4Tg mice, MGU was not altered in the basal state of VEH-treated animals. However, in contrast to WT and HKIITg, GLUT4Tg mice were protected from the LPS-induced decrease in baseline MGU. Some studies suggested GLUT4 overexpression augmented baseline MGU, implying that glucose transport is the primary site of resistance in the basal state (20, 21). Consistent with our prior work we did not see an increase in MGU in response to saline infusion (i.e., in the absence of exogenous insulin stimulation) with GLUT4 overexpression (21). It is possible that the lower glucose concentrations in the current study may have limited our ability to detect an increase in basal MGU. One possible explanation as to how overexpression of GLUT4 sustained MGU in the presence of LPS is that chronic overexpression of GLUT4 augmented muscle glycolytic capacity. As has been previously reported, plasma lactate was in fact increased in GLUT4Tg mice with VEH, and this was not observed in HKIITg mice (22). Thus, as MGU was normal, a greater fraction of the basal MGU was not oxidized but diverted to anaerobic glycolysis (i.e., lactate release); this has been observed ex vivo in muscle from GLUT4Tg mice (23). The corresponding elevation of NEFA may have helped to further limit glucose oxidation and favor lactate release. The increase in plasma lactate may also serve to maintain the increase in basal endogenous glucose production in GLUT4Tg mice with VEH treatment (24). However, as will be discussed later, accompanying improvements in the cardiovascular response to LPS in GLUT4Tg would also contribute to the maintenance of MGU in response to LPS.

Overexpression of GLUT4, but not HKII, also protected animals from the LPS-induced decrease in insulin action. During the hyperinsulinemic-euglycemic clamp, WT, HKIITg, and GLUT4Tg mice had similar GIRs and MGU following VEH treatment. This is consistent with our prior studies, demonstrating that increased GLUT4 does not augment insulin-stimulated MGU in unstressed WT mice in vivo(21). However, GLUT4Tg mice, but not HKIITg mice, were resistant to the LPS-induced decrease in whole body GIR and MGU. These results indicate that strategies that augment glucose transport capacity can overcome barriers to insulin-stimulated glucose uptake induced by LPS. On the contrary, overexpression of GLUT4 could not prevent the LPS-induced decrease in membrane-associated GLUT4. Thus, in GLUT4Tg mice, the maintenance of MGU in response to LPS may be secondary to adaptations to the overexpression of GLUT4 that can serve a protective role from other effects limiting MGU during inflammation. As HKII overexpression did not modify the response to LPS, we focused on the mechanism for the improvement in glucose disposal in the GLUT4Tg mice.

Greater suppression of NEFA availability could not explain the improvement in MGU with GLUT4 overexpression in response to LPS. The transgenic mouse model used for these studies increased GLUT4 protein not only in muscle but also in AT (14). Plasma NEFA levels in the basal state were increased in GLUT4Tg mice. In response to an increase in insulin, NEFA decreased in all groups. However, in GLUT4Tg mice, NEFA remained elevated relative to WT animals. Increased basal NEFA levels in GLUT4Tg mice could be due to an increase in lipolysis and/or a decrease in re-esterification (25, 26). We found no significant difference in either basal or insulin-stimulated glucose uptake by AT in the GLUT4Tg mice compared with WT mice. The lower insulin and glucose concentration in GLUT4Tg mice may, in part, explain the higher NEFA levels in the basal state. The higher NEFA in GLUT4Tg mice may reflect long-term adaptions of the AT to the overexpression of GLUT4, or possibly to alterations in autonomic tone to the AT. However, as NEFAs were higher both in the basal and clamp states in GLUT4Tg mice, greater suppression of NEFA cannot explain the improvement in MGU in response to LPS.

Improved insulin-stimulated glucose disposal in GLUT4Tg mice receiving LPS was not associated with a maintenance of membrane-associated glucose transport. Similar to other observations, using immunohistochemistry we observed that GLUT4Tg mice had increased membrane-associated GLUT4 protein during the clamp (27, 28). However, we observed that cell surface GLUT4 was decreased by LPS in both WT and GLUT4Tg mice. Inflammation is a potent inhibitor of insulin-stimulated GLUT4 translocation (29). Thus, although overexpression of GLUT4 increased total cell GLUT4 content, it could not prevent the decrease in membrane-associated GLUT4 in the setting of inflammation. It is possible that a small increase of membrane-associated transporter in GLUT4Tg mice treated with LPS could have gone undetected with this method. These results were interesting in light of our previous data, which showed that insulin signaling through the PI3-K pathway was not impaired with LPS treatment (5). This pathway is believed to be the major stimulator of GLUT4 translocation to the plasma membrane under insulin-stimulated conditions. Our data are consistent with a recent report that tumor necrosis factor alpha (TNF-α) impairs GLUT4 translocation independent of the PI3-K pathway (30). This would suggest that other factors, besides preserving insulin-stimulated membrane-associated transport activity, contributed to the improvement in MGU in GLUT4Tg mice treated with LPS. As glucose-6-phosphate is a negative regulator of HK, it is possible that the chronic overexpression of GLUT4 leads to a compensatory upregulation of muscle glycolytic capacity that was preserved in response to LPS. We measured the content of glucose-6-phosphate in heart and skeletal muscle to determine if the content of the substrate in muscle was lower, which would reflect augmentation of downstream pathways. Both LPS-treated WT mice and GLUT4Tg mice showed a similar content of glucose-6-phosphate. Therefore, the amelioration of the deleterious effect of LPS in MGU occurring in GLUT4Tg mice cannot be solely explained by upregulation of glucose transport and downstream metabolic pathways in skeletal muscle. As already mentioned, glucose conversion to lactate is increased in GLUT4Tg mice. Insulin can augment muscle glycogen synthesis, glucose oxidation, and lactate release. It is possible that this adaptation of enhanced diversion of glucose to lactate was maintained in response to LPS, thus limiting the decrease in insulin-stimulated MGU. As basal MGU was also improved, it would suggest that the protection was not directly linked to improved insulin signaling. This is consistent with the lack of an alteration in insulin signaling pathway. Thus, it is likely that events outside skeletal muscle (i.e., glucose delivery) were altered to sustain MGU in response to LPS.

The GLUT4 transgene was also overexpressed in the heart, which might be cardioprotective. Improved cardiac function may have contributed to the maintenance of MGU in GLUT4Tg mice treated with LPS. The dose of LPS we used impairs cardiac function and tissue perfusion as is seen in critically ill patients. These are important determinates of MGU (5). Overexpression of GLUT4 attenuated the impairment in cardiac output following LPS and improvements in insulin signaling can be cardioprotective (31). This is consistent with prior reports showing that loss of cardiac GLUT4 transport activity impairs cardiac function (32, 33), whereas the overexpression of GLUT protects both cardiac and mitochondrial function in response to stressors (34). It is unclear why overexpression of GLUT4 was protective of cardiac function. We observed that GLUT4 overexpression lowered basal cardiac Nos2; however, LPS augmented Nos2 to the same extent in WT and GLUT4Tg mice, suggesting that cardiac inflammation, which can impair cardiac function, was comparable. GLUT4Tg had markedly upregulated Pdk4 expression (which would impair cardiac pyruvate oxidation), and other reports suggest overexpression of GLUT preserves cardiac mitochondrial function (34). As LPS impairs, cardiac fat oxidation maintenance of cardiac glucose uptake may be cardioprotective (35). The sustained GLUT4 expression did not increase glucose-6-phosphate in the heart, but it may have served to protect mitochondrial function and cardiac function even though cardiac glucose oxidation was not augmented and only diversion to lactate was sustained. As MGU is very sensitive to tissue perfusion, the secondary benefit of the improved cardiac function was maintenance of MGU during inflammation. Capillary density is not increased in GLUT4Tg(23). Future studies will need to be done to determine if muscle perfusion is preserved in GLUT4Tg and whether this is linked to the cardiac protection in response to LPS.

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CONCLUSIONS

Overexpression of muscle-specific HKII does not benefit insulin-stimulated glucose uptake in the presence of LPS, whereas overexpression of GLUT4 exerts a protective role. Surprisingly, this protection is not mediated solely by a preservation of membrane-associated GLUT4. Rather, preservation of cardiac function and long-term metabolic adaptations in muscle limit the impairment in MGU. Thus, the endotoxin-induced impairment in both basal and insulin-stimulated MGU is driven by a complex interaction of cardiovascular and metabolic events.

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Acknowledgments

The authors thank the members of the Mouse Metabolic Phenotyping Center (DK059637; Carlo Malabanan, Deanna Bracy, and Freyja James) who implanted the catheters as well as Lin Zhong who performed the echocardiography studies, and also thank the Vanderbilt University Hormone Assay and Analytical Core for performing insulin assays (DK059637 and DK020593).

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Keywords:

Cardiac output; endotoxin; inflammation; insulin resistance

© 2016 by the Shock Society