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Regulation of injury-induced skeletal myofiber regeneration by glucose transporter 4 (GLUT4)

Skeletal Muscle volume 14, Article number: 33 (2024) Cite this article

Abstract

Background

Insulin resistance and type 2 diabetes impair cellular regeneration in multiple tissues including skeletal muscle. The molecular basis for this impairment is largely unknown. Glucose uptake via glucose transporter GLUT4 is impaired in insulin resistance. In healthy muscle, acute injury stimulates glucose uptake. Whether decreased glucose uptake via GLUT4 impairs muscle regeneration is presently unknown. The goal of this study was to determine whether GLUT4 regulates muscle glucose uptake and/or regeneration following acute injury.

Methods

Tibialis anterior and extensor digitorum longus muscles from wild-type, control, or muscle-specific GLUT4 knockout (mG4KO) mice were injected with the myotoxin barium chloride to induce muscle injury. After 3, 5, 7, 10, 14, or 21 days (in wild-type mice), or after 7 or 14 days (in control & mG4KO) mice, muscles were isolated to examine [3H]-2-deoxyglucose uptake, GLUT4 levels, extracellular fluid space, fibrosis, myofiber cross-sectional area, and myofiber centralized nuclei.

Results

In wild-type mice, muscle glucose uptake was increased 3, 5, 7, and 10 days post-injury. There was a rapid decrease in GLUT4 protein levels that were restored to baseline at 5–7 days post-injury, followed by a super-compensation at 10–21 days. In mG4KO mice, there were no differences in muscle glucose uptake, extracellular fluid space, muscle fibrosis, myofiber cross-sectional areas, or percentage of centrally nucleated myofibers at 7 days post-injury. In contrast, at 14 days injured muscles from mG4KO mice exhibited decreased glucose uptake, muscle weight, myofiber cross sectional areas, and centrally nucleated myofibers, with no change in extracellular fluid space or fibrosis.

Conclusions

Collectively, these findings demonstrate that glucose uptake via GLUT4 regulates skeletal myofiber regeneration following acute injury.

Background

Insulin resistance and type 2 diabetes impair cellular regeneration in multiple tissues including skeletal muscle [1,2,3]. In mice fed a high fat diet to induce insulin resistance, muscles injured by injection of the myotoxic agent, cardiotoxin, exhibited lower muscle weights [3], smaller myofiber cross-sectional areas [3], greater areas of muscle necrosis [2], and increased fibrosis [3] compared to injured muscles from control mice. Consistent with these findings, cardiotoxin-injured muscles from genetically modified type 2 diabetic mice (i.e., ob/ob and db/db mice) exhibited a reduction in the number and cross-sectional area of myofibers with centralized nuclei [1]. While collectively these findings demonstrate that factors common to insulin resistance and type 2 diabetes impair muscle regeneration following acute injury, the molecular mechanisms underlying this impairment are not well understood.

Skeletal muscle is the primary tissue responsible for glucose uptake in the body [4, 5]. Prior work has demonstrated that glucose uptake and/or glucose utilization are increased in rodent muscles 2–5 days following burn [6, 7], blunt trauma [8], or injection of the myotoxic agent, λ-carrageenan [9]. Since muscle glucose uptake is increased across multiple injury models, these findings suggest that glucose may play a key role in one or more of the cellular processes needed for muscle repair. However, to date, no studies have investigated whether enhanced muscle glucose uptake is critical for muscle regeneration following acute injury.

Glucose transporter 4 (GLUT4) is the most abundant glucose transporter in skeletal muscle [10], and the translocation of GLUT4 from intracellular locations to the cell surface is the primary mechanism by which insulin stimulates muscle glucose uptake (see recent reviews [11,12,13]). Insulin-stimulated GLUT4 translocation and muscle glucose uptake are well-known to be impaired in type 2 diabetes [11, 14]. Whether acute injury stimulates muscle glucose uptake via GLUT4 is currently unknown. Prior work has shown that rodent muscles injured with the myotoxic agents, cardiotoxin or bupivacaine, exhibit very low levels of GLUT4 protein at 3 days post-injury [15, 16]. However as muscle regeneration progresses, GLUT4 protein levels are rapidly restored to non-injured muscle levels by 6–7 days [15, 16], before peaking above non-injured control levels at 14 days post-injury [15]. Together these findings suggest that the rapid induction and restoration of GLUT4 may play a critical role in the complete and healthy regeneration of skeletal muscle. Whether impaired muscle regeneration in type 2 diabetes is due to an impairment in muscle glucose uptake via GLUT4 is presently unknown.

The purpose of this study was to determine if GLUT4 regulates skeletal muscle glucose uptake and/or regeneration following acute injury. We hypothesized that loss of glucose uptake via GLUT4 would impair muscle regeneration following acute muscle injury. To test the hypothesis, skeletal muscles from wild-type mice were first injected with the myotoxin barium chloride (BaCl2) and examined 3, 5, 7, 10, 14 and 21 days later to assess the timing and magnitude of changes in ex vivo muscle [3H]-2-deoxyglucose uptake and GLUT4 levels. Skeletal muscles from muscle-specific GLUT4 knockout mice were then injected with BaCl2 and muscles examined 7 and 14 days later to assess glucose uptake and muscle regeneration.

Methods

Mice

Wild-type C57BL/6J mice (male, 10–11 wks old; strain# 000664), and muscle creatine kinase promoter-driven Cre recombinase mice (MCK-Cre; Tg(Ckmm-cre)5Khn; C57BL/6 background; strain# 006475 [17]) were obtained from The Jackson Laboratory. GLUT4 LoxP mice (mixed FVB, 129 & C57BL/6J background [18]) were obtained from Dr. Barbara B. Kahn (Beth Israel Deaconess Medical Center). MCK-Cre mice were bred to GLUT4 LoxP mice to generate wild-type, MCK-Cre, GLUT4 LoxP and muscle-specific GLUT4 knockout (mG4KO) mice. Weaned mice were housed 2–5 mice/cage in ventilated cages containing environmental enrichment in a specific pathogen free facility. Housing rooms were maintained at 21–22 °C with a 12 h light/dark cycle, and mice assessed daily. A chow diet (Tekland Global 2018SX) and water were provided ad libitum. For this study, data from sex and age-matched MCK-Cre and/or GLUT4 LoxP littermate mice were pooled into a single control (CON) group. See Suppl Fig. 1, for the figures in which both MCK-Cre and GLUT4 LoxP mice were utilized within the pooled control group.

Body weight and body composition

Mice were weighed to ± 0.1 g. Lean and fat mass was assessed using an EchoMRI™-700 body composition analyzer.

Fasting blood glucose

Mice were fasted for 12–14 h overnight, and blood was sampled from the tail to measure glucose levels with a glucometer (OneTouch® Ultra®2, Milpitas, CA, USA).

Tibia length

Mice were anesthetized by either inhalation of isoflurane or intraperitoneal injection of pentobarbital sodium (80–90 mg/kg body weight; Covetrus North America, Dublin, OH), and euthanized by cervical dislocation. Tibias were dissected from both legs, and the length of each tibia measured to ± 0.01 mm with a digital micrometer by three individuals blinded to the experiment group. The average of all six tibia length measures was calculated and used as the single tibia length measure for that mouse.

Barium chloride-induced acute skeletal muscle injury

Mice were anesthetized by inhalation of 2.5-3% isoflurane. The fur covering the tibialis anterior muscles was shaved and the skin disinfected with 10% povidone-iodine solution. The tibialis anterior and extensor digitorum longus muscles of one leg were injected with 50 µl of a 0.2 μm filter sterilized, 1.2% (w/v) BaCl2 (Sigma-Aldrich, cat# B0750) saline solution. The contralateral leg was not injected. Mice were allowed to recover on an ~ 37 °C heated pad and returned to home cages. Moistened chow diet pellets were added to cages to assist in food availability during initial recovery. Mice were examined at least once per day for 3 days to monitor recovery.

Muscle [3H]-2-deoxyglucose uptake & extracellular fluid space

Ex vivo muscle [3H]-2-deoxy-D-glucose uptake and extracellular fluid space was assessed as described by our group [19, 20]. Mice were fasted overnight (12–14 hrs) and anesthetized with pentobarbital sodium (80–90 mg/kg body weight; intraperitoneal). After 40 min, mice were euthanized by cervical dislocation, and extensor digitorum longus muscles excised. Muscles were pre-incubated in 95% O2 5% CO2-gassed, 37˚C Krebs-Ringer-Bicarbonate (KRB) buffer [117 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl2, 1.2 mM KH2PO4, 1.2 mM MgSO4, 24.6 mM NaHCO3] supplemented with 2 mM sodium pyruvate for 90 min. Muscles were incubated at 30˚C in KRB buffer containing 1.5 µCi/ml [3H]-2-deoxyglucose, 1 mM 2-deoxyglucose, 0.45 µCi/ml [14C]-mannitol, and 7 mM mannitol for 10 min and frozen in liquid nitrogen. Muscles were solubilized in 1 N NaOH and neutralized with 1 N HCl. Aliquots of solubilized samples were centrifuged at 10,000 x g for 1 min, and muscle [3H] and [14C] levels measured using a liquid scintillation counter (Tri-Carb 4910TR, Perkin Elmer Inc., Hebron, KY, USA). Muscle [3H]-2-deoxyglucose uptake rates were calculated utilizing [14C]-mannitol levels to account for extracellular fluid space.

RT-qPCR analyses

Mice were anesthetized with isoflurane gas (2–3%) or pentobarbital sodium (80–90 mg/kg body weight) and euthanized by cervical dislocation. Tibialis anterior muscles were dissected and ≤ 30 mg segments placed in RNAprotect (Qiagen, cat#76104), kept at 4˚C for 20–30 h, and stored at -80˚C. Muscles were homogenized using a Bullet Blender tissue homogenizer (Next Advance, Troy, NY, USA) and RNA isolated using a RNeasy fibrous tissue kit including DNase I treatment (Qiagen, cat#74704). Total RNA was quantified using a NanoDrop One-C (Thermo Fisher Scientific) with sample yields between 350 and 600 ng RNA/mg non-injected muscles, 1100–1600 ng RNA/mg for 3 day post-BaCl2 injected muscles, 1500–2200 ng RNA/mg for 5 day post-BaCl2 injected muscles, 1600–2250 ng RNA/mg for 7 day post-BaCl2 injected muscles, 1500–2300 ng RNA/mg for 10 day post-BaCl2 injected muscles, 1000–1200 ng RNA/mg for 14 day post-BaCl2 injected muscles, and 600–800 ng RNA/mg for 21 day post-BaCl2 injected muscles and RNA purity as measured by A260/A280 was between 2.0 and 2.1. RNA integrity was visually assessed by running samples on an agarose gel. RNA was reverse transcribed to cDNA using an iScript cDNA synthesis kit and manufacturer standard cycling conditions (Bio-Rad, cat#1708891).

All RT-qPCR procedures were done according to the Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines [21]. Samples of 50 ng cDNA were manually pipetted in triplicate in 384 well plates with transcript-specific primers and iTaq™ Universal SYBR® Green Supermix (Bio-Rad cat #1725121). Reactions were run on a Bio-Rad CFX384 Touch Real-Time PCR Detection System (BioRad, Hercules, CA, USA) with the following cycle conditions: 95˚C for 30 s, 95˚C for 5 s and 62˚C for 30 s (repeated 40x); and analyzed with CFX Maestro software (Bio-Rad). PCR product specificity was confirmed by melt curves. Primer PCR efficiency was calculated from calibration curves [parameters (ranges): slope (-3.53 to -3.30), y-intercept (30.0 to 38.3), and r2 values (0.99)] and verified to be linear over a ≥ 100-fold range for all primers. Relative SLC2A/GLUT gene expression was determined based on a normalization factor that was calculated as the geometric mean of the Cq values of the selected reference genes [i.e., hydroxyacyl glutathione hydrolase (HAGH), ribosomal protein S17 (RPS17), and signal recognition particle 14 (SRP14)]. Primer sequences and efficiencies for reference genes were previously published [22]. Sequences and primer efficiency information for the SLC2A/GLUT primers can be found in Suppl. Table 1.

Immunoblot analyses

Mouse tissues were homogenized in lysis buffer [20 mM Tris-HCl pH 7.4, 5 mM EDTA, 10 mM Na4P2O7, 100 mM NaF, 2 mM Na3VO4, 1% IGEPAL® CA-630, 1.5 µM aprotinin, 3 mM benzamidine, 23.4 µM leupeptin and 1 mM phenylmethylsulfonyl fluoride] using a Bullet Blender tissue homogenizer (Next Advance). Samples were rotated end-over-end at 4˚C for 60 min, centrifuged at 13,000 x g for 30 min, and lysate protein concentrations determined using Bio-Rad protein assay dye reagent (cat#5000006).

GLUT4 immunoblot analyses were performed using standard methods described by our group [19, 20]. Lysates (10–20 µg total protein) were subjected to SDS-PAGE using 10% acrylamide gels and proteins transferred onto 0.2 μm nitrocellulose membranes. Equal protein loading and transfer was assessed using Ponceau S solution stain (Sigma-Aldrich, cat#P7170) as shown in Suppl Fig. 2. Membranes were blocked with 5% bovine serum albumin [dissolved in a 1x Tris-buffered-saline + 0.1% Tween-20 (1xTBST) solution] for 1 h at room temperature. Membranes were incubated in a 1:2000 (v/v) GLUT4 primary antibody (EMD Millipore, cat#07-1404, lot#2890837 RRID: AB_1587080) in 1xTBST solution overnight at 4˚C. Membranes were incubated in a 1:5000 (v/v) rabbit-horseradish peroxidase-conjugated secondary antibody (Cell Signaling Technology, cat#7074, lot#30) in 1xTBST solution for 1 h at room temperature. Bands were visualized using enhanced chemiluminescence substrate (Perkin Elmer Life Sciences, cat# NEL105001EA). Imaging and densitometric analyses were performed using a Bio-Rad Laboratories Chemidoc imager and Image Lab™ software. Raw unedited GLUT4 immunoblot images are provided in Suppl Fig. 2.

Immunohistological analyses

Tibialis anterior muscles were dissected from euthanized mice, embedded in Optimal Cutting Temperature (O.C.T.) compound, and rapidly frozen in liquid nitrogen-cooled 2-methylbutane. Transverse cross-sections 10 μm thick were taken from the mid-belly of the muscle and adhered to slides. Muscle sections were permeabilized with 0.1% Triton-X100 [diluted in 1x phosphate buffered saline (PBS)] for 2 min, washed for 5 min in 1x PBS, and blocked in either 10% donkey or goat serum for 20 min at room temperature. Sections were incubated in either laminin primary antibody (1:500, Sigma-Aldrich, cat#L9393, RRID: AB_477163) in a 2% goat serum, 0.5% BSA, 1x PBS solution or a type 1 collagen primary antibody (1:500, SouthernBiotech, cat#1310-01, RRID: AB_2753206) in a 2% donkey serum, 1x PBS solution overnight at 4 °C. Sections were incubated with either an anti-rabbit Alexa Fluor 647 secondary antibody (1:250, Invitrogen, cat#A21244, RRID: AB_2535812) or an anti-goat Alex Fluor 488 secondary antibody (1:250, Invitrogen, cat#A11055, RRID: AB_2534102) for 1 h at room temperature, and nuclei labeled with DAPI (2 μg/ml in 1x PBS). Sections were visualized on a Zeiss Axioobserver 7 microscope and images acquired using Zen Blue 3.0 software. Muscle collagen content was quantified from type 1 collagen-immunostained images using ImageJ software and expressed as the percent area of collagen relative to the area of the whole muscle. Myofiber cell perimeters were digitized from laminin-immunostained images using Cellpose v2.2.3 software and myofiber cross-sectional area quantified using ImageJ software. The average number of myofibers counted per muscle was > 800 for each group. Non-muscle cells with cross-sectional areas < 50 µm2 were excluded from quantification. The average myofiber cross-sectional area for each muscle was calculated. Myofibers were separated into groups of 450–500 µm2 and the percent frequency of distribution calculated. Centralized nuclei were identified from DAPI-stained images and quantified using QuantiMus software [23].

Statistical analyses

Data sets were screened once for outliers defined as data points whose value exceeded the mean ± 2x standard deviation for that group to avoid the possibility of a statistical type 2 error. Any identified outliers were removed prior to statistical analyses with GraphPad Prism v10 software. Statistical significance was defined as P < 0.05. During manuscript peer review, a re-analysis of the data sets was performed with the outliers included. This reanalysis demonstrated that the removal of any identified outliers did not alter the interpretation or conclusion of any of the data sets. The data are reported as mean ± standard deviation. The statistical test and sample size for each data set are indicated in the figure legends.

Results

BaCl2-induced injury stimulates a rapid and sustained increase in muscle glucose uptake

Intramuscular injection of the myotoxic agent BaCl2 induces acute injury and regeneration of rodent skeletal muscle [24,25,26]. While prior work had demonstrated that muscle injury induced by burn, blunt trauma, or λ-carrageenan injection increase muscle glucose uptake [6,7,8,9]; to date no studies have examined whether BaCl2-induced injury elicits similar effects. Thus, the first objective of this study was to investigate the timing and magnitude of changes in muscle glucose uptake stimulated by BaCl2-induced muscle injury. Muscles from one leg of wild-type mice were injected with BaCl2. The contralateral leg was not injected and served as the control. After 3, 5, 7, 10, 14 or 21 days, muscles were incubated in [3H]-2-deoxyglucose to assess glucose uptake. As shown in Fig. 1, BaCl2-induced muscle injury increased glucose uptake at 3, 5, 7, and 10 days post injection compared to the non-injured muscles, with trends towards increased glucose uptake still observed at 14 and 21 days.

Fig. 1
figure 1

BaCl2-induced muscle injury stimulates a rapid and sustained increase in muscle glucose uptake. In male, wild-type C57BL/6J mice, 50 µl of a 1.2% BaCl2 solution was injected into the extensor digitorum longus (EDL) muscles of one leg. The contralateral leg was not injected. After 3, 5, 7, 10, 14 or 21 days, mice were fasted overnight, anesthetized with pentobarbital sodium, and euthanized. EDL muscles were incubated in [3H]-2-deoxyglucose to assess muscle glucose uptake. Data are presented as individual data points plus the mean ± standard deviation, with N = 5–6 muscles/group. Statistical significance was defined as P < 0.05, determined by Two-Way ANOVA with Tukey’s posthoc analysis, and denoted by ‘*’ vs. non-injected control muscle (-), ‘a’ vs. 3 days, ‘b’ vs. 5 days, and ‘c’ vs. 7 days

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Multiple glucose transporters are altered by BaCl2-induced muscle injury

There are fourteen members of the facilitative glucose transporter (GLUT) family in mammalian cells, GLUTs 1–14 (see recent review [13]). While GLUT11 and GLUT14 are not present in the mouse genome [27, 28], studies have demonstrated that GLUT7 and GLUT13 (also known as the H+/myoinositol transporter) do not transport 2-deoxyglucose [29, 30]. To begin to assess the contribution of the remaining GLUTs to injury-induced muscle glucose uptake, muscles from wild-type mice were injected with BaCl2 and isolated 3, 5, 7, 10, 14 and 21 days later to examine GLUT mRNA levels by RT-qPCR analyses. As shown in Fig. 2A-I, all of the GLUTs exhibited an increase or decrease in expression at varying time points following BaCl2 injury. The GLUTs that exhibited an early (< 7 day) post-BaCl2 injury-induced increase in expression included GLUT1, GLUT5, GLUT6, GLUT9, and GLUT10, whereas those that exhibited a later (> 7 day) post-BaCl2 injury-induced increase in expression were GLUT4 and GLUT8 whose mRNA levels exceeded non-injured control levels at 21 days post-injury.

Fig. 2
figure 2

BaCl2-induced injury stimulates time-dependent alterations in the expression of multiple glucose transporters in skeletal muscle. In male, wild-type C57BL/6J mice, 50 µl of a 1.2% BaCl2 solution was injected into the tibialis anterior (TA) muscles of one leg. The contralateral leg was not injected (Day 0 muscles). After 3, 5, 7, 10, 14 or 21 days, TA muscles were collected and RNA isolated to assess glucose transporter (GLUT) levels by RT-qPCR. A: GLUT1; B: GLUT3; C: GLUT4; D: GLUT5; E: GLUT6; F: GLUT8; G: GLUT9; H: GLUT10; and I: GLUT12. Data are presented as individual data points plus the mean ± standard deviation, with N = 6 muscles/group. Statistical significance was defined as P < 0.05, determined by One-Way ANOVA with Tukey’s posthoc analysis, and denoted by ‘*’ vs. non-injected control muscle, ‘a’ vs. 3 days, ‘b’ vs. 5 days, ‘c’ vs. 7 days, ‘d’ vs. 10 days, and ‘e’ vs. 14 days

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Glucose transporter 4 (GLUT4) expression is altered by BaCl2-induced muscle injury

Prior work had demonstrated a rapid decline, restoration, and then supercompensation of GLUT4 protein levels by 14 days post-cardiotoxin-induced injury [15]. Thus, the next objective was to determine whether BaCl2-induced muscle injury induced similar alterations in muscle GLUT4 protein levels. Muscles from wild-type mice were injected with BaCl2 and isolated 3, 5, 7, 10, 14 and 21 days later to assess GLUT4 protein levels by immunoblot analyses. As shown in Fig. 3, there was a time-dependent change in GLUT4 protein levels with the results demonstrating a ~ 70% decrease at 3 days post-BaCl2 injection, a restoration to non-injured levels by 5–7 days post-injection, and last a 40–45% super-compensation at 10, 14, and 21 days post-injection.

Fig. 3
figure 3

BaCl2-induced muscle injury stimulates a time-dependent alteration in muscle GLUT4 protein levels. In male, wild-type C57BL/6J mice, 50 µl of a 1.2% BaCl2 solution was injected into the tibialis anterior (TA) muscles of one leg. The contralateral leg was not injected. After 3, 5, 7, 10, 14 or 21 days, TA muscles were collected and protein extracted to assess total muscle GLUT4 protein levels by immunoblot (IB) analyses. Data are presented as individual data points plus the mean ± standard deviation, with N = 6 muscles/group. Statistical significance was defined as P < 0.05, determined by Two-Way ANOVA with Tukey’s posthoc analysis, and denoted by ‘*’ vs. non-injected control muscle (-), ‘a’ vs. 3 days, and ‘b’ vs. 5 days

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Muscle-specific deletion of GLUT4 reduces lean mass

To assess the role of GLUT4 in skeletal muscle regeneration, muscle-specific GLUT4 knockout (mG4KO) mice were generated by breeding two established mouse models: a GLUT4 LoxP mouse [18, 31] with a muscle creatine kinase promoter-driven Cre recombinase (MCK-Cre) mouse [17, 19, 31, 32]. As previously described for the mG4KO model [31], while GLUT4 protein was observed in adipose tissue, it was absent from cardiac and skeletal muscle (Fig. 4A).

Fig. 4
figure 4

Body weight and body composition are impaired in muscle-specific GLUT4 knockout mice. All measures were made in 11–12 week old, male control (CON) and muscle-specific GLUT4 knockout (mG4KO) mice. (A) Tissues were collected to assess GLUT4 protein levels by immunoblot (IB) analysis. Images from N = 3 mice/group. (B) Body weight. (Panels C-F) Body composition was assessed by EchoMRI™. (C) Lean mass. (D) Fat mass. (E) Percent lean mass relative to body weight. (F) Percent fat mass relative to body weight. (G) Tibias were isolated and tibia length measured with a micrometer. Data are presented as individual data points plus the mean ± standard deviation, with N = 10–13 mice/group. Statistical significance was defined as P < 0.05, determined by t-tests, and denoted by ‘*’ vs. CON mice

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Prior work had demonstrated that mice with germline deletion of GLUT4 exhibit a growth-retarded phenotype with mice exhibiting lower body weights [33,34,35,36]. To assess the role of muscle GLUT4 expression in the regulation of body weight and lean mass, 11–12 week old control and mG4KO mice were weighed and body composition assessed. As shown in Fig. 4B-F, there was a reduction in body weight and lean mass in the mG4KO mice, but no change in fat mass. To assess whether the mG4KO mice had reduced body weight due to allometric scaling, tibia length was examined as an indicator of mouse length. Tibia length was not different in the mG4KO mice compared to littermate controls (Fig. 4G), indicating that the muscle-specific loss of GLUT4 reduces lean mass independent of overall mouse size.

Muscle-specific depletion of GLUT4 impairs BaCl2 injury-induced muscle glucose uptake

To assess the role of GLUT4 in injury-induced muscle glucose uptake, muscles from mG4KO mice were injected with BaCl2. The contralateral leg was not injected and served as the control. After 7 or 14 days, muscles were incubated in [3H]-2-deoxyglucose to assess glucose uptake. At 7 days post BaCl2-injection, glucose uptake was not different between the injured muscles from the control and mG4KO mice (Fig. 5A). In contrast, at 14 days post-injection, glucose uptake was reduced ~ 40% in the injured muscles from the mG4KO mice compared to the injured muscles from the control mice (Fig. 5B).

Fig. 5
figure 5

Muscle-specific deletion of GLUT4 impairs muscle glucose uptake at 14 days post BaCl2-induced injury. In 13–14 week old, male control (CON) and muscle-specific GLUT4 knockout (mG4KO) mice, BaCl2 solution was injected into the muscles of one leg. The contralateral leg was not injected. Muscles were examined at 7 days or 14 days post-injury. (Panels A + B) Extensor digitorum longus muscles were incubated in [3H]-2-deoxyglucose to assess glucose uptake. (Panels C + D) Tibialis anterior muscles were frozen to assess GLUT4 protein levels by immunoblot (IB) analyses. Representative images and quantification of band intensity. Data are presented as individual data points plus the mean ± standard deviation, with N = 7–9 muscles/group. Statistical significance was defined as P < 0.05, determined by Two-Way ANOVA with Tukey’s posthoc analysis, and denoted by ‘*’ vs. non-injected control muscle (-), and ‘#’ vs. CON mice

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To determine if the BaCl2-induced injury led to a transient expression of GLUT4 in the regenerating muscles from the mG4KO mice, immunoblots were performed. GLUT4 protein was not detected in the non-injured muscles from the mG4KO mice (Fig. 5C + D). In contrast, GLUT4 protein was detected in the BaCl2-injected muscles from the mG4KO mice. Compared to the BaCl2-injured muscles from the control mice, GLUT4 protein levels were reduced ~ 50% at 7 days post-injection, and ~ 90% at 14 days post-injection (Fig. 5C + D). This transient expression of GLUT4 protein in the injured muscles from the mG4KO mice was expected since Cre recombinase would not have been present in the myofibers until muscle creatine kinase expression was induced.

Muscle-specific deletion of GLUT4 impairs BaCl2 injury-induced muscle regeneration

To determine if GLUT4 regulates muscle regeneration, BaCl2-injured muscles from mG4KO mice were examined for changes in weight, extracellular fluid space, fibrosis, myofiber cross-sectional area, and percentage of myofibers with centralized nuclei. As shown in Fig. 6A + 6B, the weight of non-injured muscles was lower in the mG4KO mice compared to the controls. At both 7 and 14 days post-injury, BaCl2-injected muscles from the mG4KO mice weighed less than the BaCl2-injected muscles from the control mice.

Fig. 6
figure 6

Muscle-specific deletion of GLUT4 reduces muscle weight but not extracellular fluid space post BaCl2-induced injury. In 13–14 week old, male control (CON) and muscle-specific GLUT4 knockout (mG4KO) mice, BaCl2 solution was injected into the muscles of one leg. The contralateral leg was not injected. Extensor digitorum longus (EDL) muscles were examined at 7 days (panels A + C) or 14 days (panels B + D) post-injury. (A + B) EDL muscles were excised and immediately weighed (N = 11–14 muscles/group). (C + D) EDL muscles were incubated in [14C]-mannitol to assess extracellular fluid space (N = 7–8 muscles/group). Data are presented as individual data points plus the mean ± standard deviation. Statistical significance was defined as P < 0.05, determined by Two-Way ANOVA with Tukey’s posthoc analysis, and denoted by ‘*’ vs. non-injected control muscle (-), and ‘#’ vs. CON mice

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Muscle weight could be influenced by multiple factors including edema, fibrosis, and myofiber size. To determine if the loss of GLUT4 altered injury-induced edema, extracellular fluid space was measured. As shown in Fig. 6C, at 7 days post BaCl2 injection there was an ~ 80% increase in extracellular fluid space in the muscles from both the control and mG4KO mice. At 14 days post-injection, extracellular fluid space was not different between the non-injured and BaCl2-injected muscles from either genotype (Fig. 6D).

To determine if the loss of GLUT4 altered injury-induced fibrosis, muscle cross-sections were immunolabeled for type 1 collagen. Representative images are provided in Fig. 7A + B. As shown in Fig. 7C + D, there was no difference in the percent area of collagen detected in the non-injured muscles from the mG4KO mice compared to the control mice. At both 7 and 14 days post-BaCl2-induced injury, there was an increase in type 1 collagen content compared to the non-injured contralateral control muscles; however, there was no difference between the control and mG4KO muscles at either time point (Fig. 7C + D).

Fig. 7
figure 7

Muscle-specific deletion of GLUT4 does not alter muscle fibrosis post BaCl2-induced injury. In 13–14 week old, male control (CON) and muscle-specific GLUT4 knockout (mG4KO) mice, BaCl2 solution was injected into the muscles of one leg. The contralateral leg was not injected. (Panels A + B) Representative images of tibialis anterior muscle cross-sections immunolabeled for type 1 collagen (red) and stained with DAPI to label nuclei (green) for the (A) 7 day post-BaCl2 injured muscles and for the (B) 14 day post-BaCl2 injured muscles. Scale bar = 200 μm. (Panels C + D) Quantification of the percent area of type 1 collagen in the muscles from the (C) 7 day post-BaCl2 injected mice (N = 4–5 muscles/group), and (D) 14 day post-BaCl2 injected mice (N = 4–6 muscles/group). Data are presented as individual data points plus the mean ± standard deviation. Statistical significance was defined as P < 0.05, determined by Two-Way ANOVA with Tukey’s posthoc analysis, and denoted by ‘*’ vs. non-injected control muscle (-)

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To determine if the loss of GLUT4 altered myofiber size, muscle cross-sections were immunolabeled for laminin and myofiber cross-sectional areas measured (Fig. 8A-F). Representative images are provided in Fig. 8A + B. Non-injured muscles from the mG4KO mice had a ~ 15% reduction in average myofiber cross-sectional area; with a higher frequency of myofibers with cross-sectional areas between 50 and 1000 µm2 and a reduced frequency of myofibers between 2000 and 4000 µm2 (Fig. 8C-F). At 7 days post-BaCl2 injection, the average myofiber cross-sectional area was not different between the control and mG4KO mice, and there were no major differences in myofiber size distribution (Fig. 8G + H). In contrast, at 14 days post-BaCl2 injection there was a ~ 30% decrease in the average myofiber cross-sectional area in the mG4KO mice compared to the controls (Fig. 8I). This was observed with a higher frequency of myofibers with cross-sectional areas between 50 and 1000 µm2 and a reduced frequency of myofibers with cross-sectional areas between 2000 and 3500 µm2 (Fig. 8J).

Fig. 8
figure 8

Muscle-specific deletion of GLUT4 impairs myofiber size at 14 days post BaCl2-induced injury. In 13–14 week old, male control (CON) and muscle-specific GLUT4 knockout (mG4KO) mice, BaCl2 solution was injected into the muscles of one leg. The contralateral leg was not injected. (A + B) Representative images of muscle cross-sections with the myofiber cell perimeter immunolabeled for laminin (violet) and the nuclei stained with DAPI (green) for the (A) 7 day post-BaCl2 injured muscles, and for the (B) 14 day post-BaCl2 injured muscles. Scale bar = 200 μm. (Panels C-J) Quantification of the average myofiber cross-sectional areas (CSA) and percent frequency of myofibers with CSAs divided into groups of 450–500 um2. (C + D & G + H) Data from mice euthanized 7 days after BaCl2 injections (N = 4–6 muscles/group). (E + F & I + J) Data from mice euthanized 14 days after BaCl2 injections (N = 6–7 muscles/group). Data are presented as individual data points plus the mean ± standard deviation, or as mean ± standard deviation. Statistical significance was defined as P < 0.05, determined by t-tests, and denoted by ‘*’ vs. non-injected control muscle (-)

Full size image

The relocalization of nuclei from the center to the periphery of a myofiber is an indicator of myofiber maturation. To assess whether the smaller myofiber cross-sectional areas observed in the mG4KO mice were due to impaired myofiber maturation, the percentage of fibers with centralized nuclei were quantified. At 7 days post-BaCl2 injection, the percentage of myofibers with centralized nuclei was not different between the control and mG4KO mice (Fig. 9A). In contrast, at 14 days post-injection the BaCl2-injured muscles from the mG4KO mice had fewer centrally nucleated myofibers (Fig. 9B), suggesting that loss of GLUT4 expression led to a premature maturation of the myofibers post-injury.

Fig. 9
figure 9

Muscle-specific deletion of GLUT4 reduces the number of centrally nucleated myofibers post BaCl2-induced injury. In 13–14 week old, male control (CON) and muscle-specific GLUT4 knockout (mG4KO) mice, BaCl2 solution was injected into the muscles of one leg. The contralateral leg was not injected. Quantification of the percentage of myofibers with centralized nuclei in muscles from mice euthanized at (A) 7 days (N = 4–6 muscles/group), and (B) 14 days post-injection (N = 6–7 muscles/group). Data are presented as individual data points plus the mean ± standard deviation. Statistical significance was defined as P < 0.05, determined by Two-Way ANOVA with Tukey’s posthoc analysis, and denoted by ‘*’ vs. non-injected control muscle (-), and ‘#’ vs. CON mice

Full size image

Discussion

The major conclusion of the present study is that glucose uptake via GLUT4 plays a key role in controlling skeletal muscle regeneration following acute injury. This conclusion is based on findings from muscle-specific GLUT4 knockout mice which exhibited decreased muscle glucose uptake, reduced muscle weights, reduced myofiber cross sectional areas, and a reduced percentage of myofibers with centralized nuclei 14 days post BaCl2-induced muscle injury. This experimental evidence strongly supports the overall concept that increased glucose uptake into injured muscle is important for the healthy and complete regeneration of skeletal myofibers. These findings are also in agreement with studies that demonstrated impaired muscle regeneration in pathological conditions such as insulin resistance and type 2 diabetes in which muscle glucose uptake via GLUT4 is impaired.

Prior work in rat muscle had demonstrated an increase in glucose uptake 3–5 days post-injury [6,7,8,9]. While the findings from this study demonstrated a role for GLUT4 in the regulation of muscle glucose uptake at 14 days post-injury, the glucose transporter(s) responsible for the increased glucose uptake at earlier time points is less clear. Prior work and our findings found that cardiotoxin, glycerol, and/or BaCl2 injury induce the rapid expression of several alternative glucose transporters in muscle tissue, including: GLUT1, GLUT5, GLUT6, GLUT9, and GLUT10 [37]. Given the rapid and robust decrease in GLUT4 levels following muscle injury, and the lack of an impairment in BaCl2-injury induced glucose uptake at 7 days post-injection in the mG4KO mice, it is likely that one of more of these GLUTs is mediating the early induction of glucose uptake in the injured muscle tissue. Unfortunately, well-validated, commercially available antibodies for these glucose transporters are not currently available and thus whether their protein levels mirror their mRNA levels is currently not known. In addition, despite the similarity in name, GLUTs can exhibit highly divergent preferences for substrates. For example, GLUT5 prefers to transport D-fructose over D-glucose [38], GLUT9 prefers to transport urate over D-glucose [39], and GLUT10 prefers to transport D-galactose over D-glucose [40]. Thus, while expression levels provide important first information about the possible involvement of these alternative GLUTs in injury-induced muscle glucose uptake, future studies will need to selectively knockout each GLUT to fully assess whether it is responsible for mediating glucose uptake at any time point post-injury.

While prior work in the Akita mouse model of type 1 diabetes had suggested that hyperglycemia was associated with increased muscle fibrosis following cardiotoxin-induced injury [41], no difference in type 1 collagen content was observed in the muscles from the mG4KO mice. The level of hyperglycemia was much lower in the mG4KO mice (i.e., 5.9±1.1 mM blood glucose) (Suppl. Figure 3) than in the Akita mice (i.e., blood glucose ~ 32 mM) [41], suggesting that large elevations in blood glucose levels are needed to alter collagen deposition in muscle. However, other work detected increased type 1 collagen content in the muscle of obese, insulin resistant human subjects that were not hyperglycemic [42]. Since the mG4KO mice utilized in this study were not obese, collectively these findings suggest that systemic factors and not muscle insulin resistance per se underlie the increased fibrosis seen in the muscle of type 2 diabetic subjects. Future studies would need to feed the mG4KO mice a high fat diet to assess the contribution of obesity to the impaired muscle regeneration seen in insulin resistant, human skeletal muscles.

While the results from this study demonstrate that loss of GLUT4 expression reduces myofiber size, the molecular mechanisms underlying this impairment are not yet known. Prior work using mG4KO mice demonstrated a decrease in insulin-stimulated muscle glycolytic flux [32], which could reduce ATP generation. Since muscle protein synthesis is a high energy demanding cellular process [43], a decrease in ATP production would be expected to limit protein synthesis rates and subsequently myofiber area following an injury. In addition, recent work demonstrated that glucose-derived carbons can be incorporated into lipids, RNA, and protein in cultured mouse C2C12 myoblasts and myotubes [44]. Thus, a decrease in muscle glucose uptake could also reduce the generation of metabolites critical for biosynthetic reactions post-injury. For example, the glycolytic intermediate 3-phosphoglycerate can be dehydrogenated by the enzyme phosphoglycerate dehydrogenase (PHGDH) thereby diverting glucose flux towards serine/glycine biosynthesis. With recent work demonstrating that knockdown of PHGDH expression impairs insulin-like growth factor-stimulated C2C12 myotube growth [45], it is possible that the decrease in glucose uptake seen in the injured muscles from the mG4KO mice may reduce myofiber area via a decrease in glucose-derived serine/glycine production. Future work is currently focused on determining the metabolic consequences of impaired glucose uptake via GLUT4 during muscle regeneration.

The findings of this study advance the field of skeletal muscle regeneration by identifying glucose uptake and GLUT4 expression as novel factors that could be targeted to improve muscle recovery following an injury in individuals with insulin resistance and type 2 diabetes. However, there were several aspects of the experimental design of this study that limit this overall conclusion. A single method of injury (i.e., BaCl2 injection) was utilized to induce muscle damage and thus it is presently unknown whether glucose uptake via GLUT4 plays a role in muscle regeneration following injury by another approach (e.g., blunt trauma, burn, etc.). Prior work showed that muscle creatine kinase expression was decreased 45–80% in mouse muscles 3 days post-myotoxin-induced injury [37], which may explain why GLUT4 protein was detected in the injured muscles from the mG4KO mice at 7 days post-BaCl2 injection. Thus, while a strength of this study was the determination of a role for GLUT4 in mediating injury-induced myofiber maturation, the role of GLUT4 in regulating muscle satellite cell activation, myocyte proliferation, or myoblast differentiation is not entirely clear. Future studies would need to utilize a global GLUT4 knockout mouse model to fully assess how impaired glucose uptake via GLUT4 may regulate these early stages of muscle regeneration following an injury. However, male global GLUT4 knockout mice exhibit a more pronounced growth retarded phenotype than then mG4KO mice [33], which could confound the interpretation of those studies.

Conclusions

The major findings of the current study demonstrated that GLUT4 regulates muscle repair following acute injury. These findings are significant in that they advance the field of skeletal muscle regeneration by identifying glucose uptake via GLUT4 as a novel therapeutic target to improve muscle recovery, and specifically myofiber maturation, following an injury.

Data availability

The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.

Abbreviations

BaCl2 :

Barium Chloride

CON:

Control

GLUT4:

Glucose Transporter 4

IACUC:

Institutional Animal Care and Use Committee

KRB:

Krebs-Ringer-Bicarbonate

MCK-Cre:

Muscle creatine kinase promoter-driven Cre recombinase

mG4KO:

Muscle-specific GLUT4 knockout

OCT:

Optimal Cutting Temperature

PBS:

Phosphate Buffered Saline

SDS-PAGE:

Sodium Dodecyl Sulfate–Polyacrylamide Gel Electrophoresis

TBST:

Tris-buffered-saline + 0.1% Tween-20

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Acknowledgements

The authors thank Kaitie N. Phillips (Indiana University) for her help with mouse genotyping.

Funding

This research was funded by the National Institute of Diabetes and Digestive and Kidney Diseases R01DK106210 (B.B.K.), R01DK043051 (B.B.K.), R01DK103562 (C.A.W.); the National Heart, Lung, and Blood Institute R01HL158647 (S.S.W.); the Muscular Dystrophy Association MDA603201 (S.S.W.); and the JPB Foundation (B.B.K.). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Diabetes and Digestive and Kidney Diseases, the National Institutes of Health, the Muscular Dystrophy Association, or the JPB Foundation. Additional support was provided by the Indiana University School of Medicine and the Indiana Center for Musculoskeletal Health as new laboratory start-up funds to S.S.W. and C.A.W.

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Authors and Affiliations

  1. Department of Anatomy, Cell Biology & Physiology, Indiana University School of Medicine, Indianapolis, IN, USA

    Tyler J. Sermersheim, LeAnna J. Phillips, Parker L. Evans, Steven S. Welc & Carol A. Witczak

  2. Indiana Center for Musculoskeletal Health, Indianapolis, IN, USA

    Steven S. Welc

  3. Indiana Center for Diabetes & Metabolic Diseases, Indianapolis, IN, USA

    Tyler J. Sermersheim, LeAnna J. Phillips, Parker L. Evans & Carol A. Witczak

  4. Division of Endocrinology, Diabetes & Metabolism, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, 02215, USA

    Barbara B. Kahn

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Contributions

Study conceptualization and design, T.J.S., L.J.P., S.S.W., and C.A.W.; methodology, acquisition of data, and/or interpretation of data, T.J.S., L.J.P., P.L.E., B.B.K., S.S.W., and C.A.W.; formal analysis, T.J.S., L.J.P., P.L.E., and C.A.W.; writing—original draft preparation, C.A.W.; writing—review and editing, T.J.S., L.J.P., P.L.E., B.B.K., S.S.W., and C.A.W.; funding acquisition, B.B.K., S.S.W., and C.A.W. All authors have read and approved the final version of the manuscript. All authors have agreed to be personally accountable for their contributions to ensure that questions related to the accuracy or integrity of any part of the work are appropriately investigated, resolved, and the resolution documented in the literature.

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Correspondence to Carol A. Witczak.

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Procedures were performed in accordance with the Indiana University School of Medicine Institutional Animal Care and Use Committee (IACUC, protocol# 21053 approved on 17 May 2021 and protocol# 21118 approved on 31 August 2021), and the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals.

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Sermersheim, T.J., Phillips, L.J., Evans, P.L. et al. Regulation of injury-induced skeletal myofiber regeneration by glucose transporter 4 (GLUT4). Skeletal Muscle 14, 33 (2024). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s13395-024-00366-y

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Skeletal Muscle

ISSN: 2044-5040

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