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. 2017 Nov 21;595(24):7427–7439. doi: 10.1113/JP275133

Nitric oxide is required for the insulin sensitizing effects of contraction in mouse skeletal muscle

Xinmei Zhang 1,2,✉, Danielle Hiam 1, Yet‐Hoi Hong 1, Anthony Zulli 3, Alan Hayes 1,3, Stephen Rattigan 4, Glenn K McConell 1,3
PMCID: PMC5730845  PMID: 29071734

Abstract

Key points

  • People with insulin resistance or type 2 diabetes can substantially increase their skeletal muscle glucose uptake during exercise and insulin sensitivity after exercise.

  • Skeletal muscle nitric oxide (NO) is important for glucose uptake during exercise, although how prior exercise increases insulin sensitivity is unclear.

  • In the present study, we examined whether NO is necessary for normal increases in skeletal muscle insulin sensitivity after contraction ex vivo in mouse muscle.

  • The present study uncovers, for the first time, a novel role for NO in the insulin sensitizing effects of ex vivo contraction, which is independent of blood flow.

Abstract

The factors regulating the increase in skeletal muscle insulin sensitivity after exercise are unclear. We examined whether nitric oxide (NO) is required for the increase in insulin sensitivity after ex vivo contractions. Isolated C57BL/6J mouse EDL muscles were contracted for 10 min or remained at rest (basal) with or without the NO synthase (NOS) inhibition (N G‐monomethyl‐l‐arginine; l‐NMMA; 100 μm). Then, 3.5 h post contraction/basal, muscles were exposed to saline or insulin (120 μU ml−1) with or without l‐NMMA during the last 30 min. l‐NMMA had no effect on basal skeletal muscle glucose uptake. The increase in muscle glucose uptake with insulin (57%) was significantly (P < 0.05) greater after prior contraction (140% increase). NOS inhibition during the contractions had no effect on this insulin‐sensitizing effect of contraction, whereas NOS inhibition during insulin prevented the increase in skeletal muscle insulin sensitivity post‐contraction. Soluble guanylate cyclase inhibition, protein kinase G (PKG) inhibition or cyclic nucleotide phosphodiesterase inhibition each had no effect on the insulin‐sensitizing effect of prior contraction. In conclusion, NO is required for increases in insulin sensitivity several hours after contraction of mouse skeletal muscle via a cGMP/PKG independent pathway.

Keywords: insulin sensitivity, muscle contraction, L‐NMMA

Key points

  • People with insulin resistance or type 2 diabetes can substantially increase their skeletal muscle glucose uptake during exercise and insulin sensitivity after exercise.

  • Skeletal muscle nitric oxide (NO) is important for glucose uptake during exercise, although how prior exercise increases insulin sensitivity is unclear.

  • In the present study, we examined whether NO is necessary for normal increases in skeletal muscle insulin sensitivity after contraction ex vivo in mouse muscle.

  • The present study uncovers, for the first time, a novel role for NO in the insulin sensitizing effects of ex vivo contraction, which is independent of blood flow.

Introduction

Increased physical activity is important for both the prevention and management of type 2 diabetes (T2D) (Wojtaszewski & Richter, 2006). After the initial insulin independent increases in glucose uptake post‐contraction have worn off in 2–3 h (Gao et al. 1994; Funai et al. 2010), skeletal muscle remains more sensitive to insulin for 24–48 h in both rodents (Cartee et al. 1989) and humans (Mikines et al. 1988). Some 3–4 h after a 60 min bout of single leg exercise in humans, glucose uptake during a hyperinsulinaemic euglycaemic clamp (‘insulin clamp’) increases substantially more in the exercised leg than in the rested leg (Richter et al. 1989; Wojtaszewski et al. 2000). Importantly, acute exercise increases skeletal muscle insulin sensitivity in both individuals with T2D and matched controls (Devlin et al. 1987). Although the insulin sensitizing effect of acute contraction/exercise has been known for many years, the mechanisms involved are unclear.

Insulin activates insulin signalling pathways in skeletal muscle that results in glucose transporter type 4 (GLUT‐4) translocation to the plasma membrane and increased glucose transport. Even though there are increases in insulin‐stimulated glucose uptake after acute contraction or exercise, there is little evidence of greater proximal insulin signalling (Wojtaszewski et al. 2000; Wojtaszewski & Richter, 2006). However, there are indications that more distal insulin signalling may be increased by acute exercise (e.g. phosphorylation of Akt substrate of 160 kDa; AS160, also referred to as TBC1D4) (Arias et al. 2007; Funai et al. 2009, 2010; Treebak et al. 2009; Castorena et al. 2014; Kjobsted et al. 2015; Sjoberg et al. 2017). Some 6–24 h after an acute exercise bout, increases in protein expression of some of key proteins such as GLUT‐4 are sometimes observed (Hood, 2001). Because this introduces a confounding variable, studies attempting to uncover the mechanism(s) that acute exercise increases skeletal muscle insulin sensitivity are generally conducted 3–4 h after exercise (Richter et al. 1989; Wojtaszewski & Richter, 2006).

Although never specifically examined, there are some findings in the literature which suggest that increases in nitric oxide (NO) during contraction/exercise could be involved in the increase in insulin sensitivity after contraction/exercise. Both neuronal NOS (nNOS) and endothelial NOS (eNOS) deficient mice are insulin resistant (Shankar et al. 2000) and eNOS deficient mice supplemented with nitrate (NO3), an inorganic anion abundant in vegetables that can be converted in vivo to NO, show improved glucose tolerance (Carlstrom et al. 2010). In addition, the content of nNOS in skeletal muscle tends to change in parallel with skeletal muscle insulin sensitivity (Shankar et al. 2000; Bradley et al. 2007). Supporting this notion, we have found that endurance trained humans, who are known to be insulin sensitive, have increased skeletal muscle nNOS protein (McConell et al. 2007), whereas individuals with insulin resistance/T2D have reduced nNOS protein levels (Bradley et al. 2007). Acute and long‐term administration of l‐arginine, the substrate for NO formation from NOS, improves insulin secretion and insulin sensitivity in healthy individuals and in individuals with diabetes (Piatti et al. 2001). NO also increases insulin transport in endothelial cells in skeletal muscle (Wang et al. 2013), and therefore presumably skeletal muscle insulin exposure. Finally, we have shown that NO synthase (NOS) inhibition attenuates increases in skeletal muscle glucose uptake during contraction in mice and rats (Stephens et al. 2004; Ross et al. 2007; Merry et al. 2010b) and also during exercise in healthy controls and in people with T2D (Bradley et al. 1999; Kingwell et al. 2002). Therefore, we hypothesized that NOS inhibition during contraction would attenuate the increase in insulin sensitivity 3.5 h after ex vivo contraction. Ex vivo contractions were chosen because this eliminates any potential confounding effects of blood flow.

Methods

Ethical approval

Animal care and experimental protocols and collection of human serum for the present study were approved by both the Animal Experimentation Ethics Committee and the Human Research Ethics Committee of Victoria University and conformed with the Australian National Code of Practice for the Care and Use of Animals for Scientific Purposes, as described by the National Health and Medical Research Council (NHMRC) of Australia.

Animals and experimental design

Male C57BL/6J mice aged 12–14 weeks were purchased from Animal Resources Centre (Perth, WA, Australia). The mice were individually housed in groups of two to four and maintained in an environmentally controlled animal room under a 12:12 h light/dark cycle at 21°C with ad libitum access to standard rodent chow (Specialty Feeds, Western Australia) and water. Food was removed from 08.30 h to 12.30 h on the day of an experiment. After the mice were deeply anaesthetized with pentobarbital sodium (26 G needle, 60 mg kg−1 i.p.; Rhone Merieux, Pinkenba, Queensland, Australia), they were constantly monitored for depth of anaesthesia by monitoring their plantar flexion and response to tail and paw pinch. When slight reflex/response was detected, supplemented doses (one‐tenth of the original dose) of anaesthesia were administered before tissue removal. Under deep anaesthesia, the skin of the hind limbs was removed exposing the limb muscles. Extensor digitorum longus (EDL) muscles were carefully excised from the mice. Following the removal of muscles and when they were deeply anaesthetized, mice were humanely killed by decapitation.

Materials and antibodies

All chemicals were purchased from Sigma‐Aldrich Chemicals (St Louis, MO, USA) unless otherwise stated. 2‐Deoxy‐d‐[1,2‐3H]‐glucose and d‐[1‐14C]‐mannitol were purchased from Perkin Elmer (Waltham, MA, USA). Reagents and apparatus for SDS‐PAGE and immunoblotting were purchased from Bio‐Rad (Hercules, CA, USA). The RED 660 Protein Assay Reagent Kit and Neutralizer were purchased from GBiosciences (St Louis, MO, USA). SuperSignal West Femto Chemiluminescent Substrate was provided by Thermo Scientific (Waltham, MA, USA). Primary antibodies for p‐Akt (Ser473 and Thr308), Akt, p‐TBC1D1 (Thr590, Thr596 and Ser660), TBC1D1, p‐TBC1D4 (Thr642), TBC1D4 and actin used in the western blotting were purchased from Cell Signaling Technology (Danvers, MA, USA). HRP conjugated goat anti‐rabbit IgG (H + L) secondary antibody was obtained from Thermo Scientific (Waltham, MA).

Collection and treatment of serum

As reported previously (Gao et al. 1994), a serum factor is required for an increase in insulin sensitivity after ex vivo rat skeletal muscle contraction and we also found that serum alone has no effect on mouse skeletal muscle glucose uptake at rest (Levinger et al. 2016). Whether serum is required during ex vivo contraction of mouse skeletal muscle for increases in insulin‐stimulated glucose uptake has not previously been examined. After an overnight fast, blood was collected from four healthy men via venepuncture. The blood was allowed to clot at room temperature then centrifuged at 3000 g for 30 min. The serum was collected and frozen at −80 °C until use. All serum used was from the same individuals. Repeat freeze‐thawing of serum was avoided.

Muscle dissection, incubation and contraction

Under deep anaesthesia, both EDL muscles were rapidly dissected. The proximal and distal tendons were tied using a 5/0 silk suture with two small aluminum hooks tied to each tendon. For all incubation steps, buffer was continuously maintained at 30°C (Merry et al. 2010b) and gassed with carbogen (Carbogen; BOC Gases, North Ryde, NSW, Australia). Muscles were pre‐incubated with or without 50% human serum in Buffer 1 (KHB in mm: 119 NaCl, 4.7 KCl, 2.5 CaCl2, 1.2 MgSO4, 1.2 KH2PO4 and 25 NaHCO3, pH 7.4 + 0.01% bovine serum albumin + 2 mm glucose + 8 mm mannitol) for 30 min. For muscle contraction, muscles were mounted in incubation chambers containing buffer 1 with or without serum and stimulating platinum electrodes (Zultek Engineering, Melbourse, VIC, Australia) and stimulated for 10 min with the parameters (12 V, train durations: 350 ms at a frequency of 60 Hz, 12 contractions min‐1) (Merry et al. 2010b). Non‐contracted muscles were treated the same as contracted muscles, except that they were not stimulated to contract. Muscles were incubated in the presence or absence of the NOS inhibitor N G‐monomethyl‐l‐arginine (l‐NMMA) (100μM) (Merry et al. 2010a) during the pre‐incubation and contraction periods.

Muscle treatment post‐electrical stimulation and glucose uptake measurements

Immediately after electrical stimulation, all muscles (regardless of whether the previous incubation was with or without l‐NMMA) were transferred to a vial containing buffer 1 for a 1 min wash. Muscles were then transferred to other baths containing buffer 1 for 3 h with the buffer changed every 30 min.

After 3 h, all muscles were incubated with buffer 2 containing 2 mm pyruvate + 8 mm mannitol with or without insulin for 30 min. For glucose uptake analysis, all muscles were incubated for 10 min with buffer 3 containing 2 mm 2‐deoxy‐d‐[1,2‐3H]‐glucose (2‐DG, 0.256 μCi ml−1) and 16 mm D‐[1‐14C]‐mannitol (0.166 μCi ml−1), as well as insulin if it was present during the previous incubation with buffer 2. For some muscle pairs, l‐NMMA (100 μm) was also present during this incubation.

To determine whether NO during insulin exposure was acting via the NO/cGMP/protein kinase G (PKG) pathway, the GC inhibitor 1H‐[1,2,4]oxadiazolo‐[4,3‐a]quinoxalin‐1‐one (ODQ, which blocks the NO‐mediated increase in cGMP; 10 μm) (Merry et al. 2010a), or the phosphodiesterase type 5 inhibitor (T‐1032, which inhibits cGMP breakdown and therefore raises cGMP levels; 27 μm) (Mahajan et al. 2003) or the cGMP‐dependent PKG inhibitor (Rp‐8‐Br‐PET‐cGMPS; 5 μm) (Merry et al. 2010a) were used to block of the NO/cGMP/PKG pathway. The concentrations of ODQ and Rp‐8‐Br‐PET‐cGMPS used in the present study were based on our previous studies using isolated ex vivo muscles (Merry et al. 2010a, b). In addition, the cyclic nucleotide phosphodiesterase (PDE5) inhibitor T1032 was used in the present study rather than another PDE5 inhibitor, zaprinast, because zaprinast has been shown in our previous study to have no inhibitory effect on insulin‐mediated glucose uptake by muscles in vivo, whereas T‐1032 showed inhibitory effects (Mahajan et al. 2003). The muscle pairs were incubated in the presence or absence of the inhibitor ODQ, or T‐1032 or Rp‐8‐Br‐PET‐cGMPS during the 30 min of insulin incubation and 10 min of 2‐DG incubation.

Given that 120 μU ml−1 of insulin results in maximum insulin‐stimulated glucose uptake (Hamada et al. 2006), it was anticipated that both ODQ and Rp‐8‐Br‐PET‐cGMPS would attenuate the increase in insulin‐stimulated glucose uptake after contraction; thus, 120 μU ml−1 of insulin was used for ODQ and Rp‐8‐Br‐PET‐cGMPS treatments. On the other hand, given that we anticipated that T‐1032 would increase insulin‐stimulated glucose uptake after contraction, we used a submaximal dose of insulin (60 μU ml−1) (Hamada et al. 2006) with T‐1032 treatment to provide a greater opportunity to observe any increase in glucose uptake.

After the 10 min incubation with radioisotopic tracers, muscles were rapidly rinsed, trimmed and cut in halves and frozen in liquid nitrogen. One half was kept for immunoblotting and the other half was used for glucose uptake determination. The muscle for glucose uptake was homogenized in 1 m NaOH at 95°C for 10 min and then neutralized by 1 m HCl followed by centrifugation. The supernatant (200 μl) was added to 4 ml of liquid scintillation cocktail (Perkin Elmer). The radioactivity of both tracers was measured using a β scintillation counter (Tri‐Carb 2910TR; Perkin Elmer) and glucose uptake was calculated as described previously (Merry et al. 2010a; Zhang et al. 2011).

NOS activity assay

NOS activity was determined in separate EDL muscles based on the catalytic reaction of NOS converting radiolabelled l‐[14C]‐arginine to radiolabelled l‐[14C]‐citrulline, as described previously (Merry et al. 2010a). NOS activity was determined from the difference between samples incubated with and without l‐NAME and was expressed as picomol of l‐[14C] citrulline formed min–1 milligram muscle protein–1.

Sample preparation and immunoblotting

Sample preparation for immunoblotting was initially described by Murphy (2011). Briefly, 10 muscle sections of 20 μm thickness were homogenized with 100 μl of solubilizing buffer [0.125 m Tris–Cl (pH 6.8), 4% w/v SDS, 10% glycerol, 10 mm EGTA, 0.1 m DTT and protease inhibitor cocktail]. The protein concentration was determined using a Red 660 assay kit (G‐Biosciences, St Louis, MO, USA). Proteins (5 μg loaded per well) were separated with 10% SDS/PAGE gels and then transferred to polyvinylidene fluoride for 120 min at 100 V. Following transfer, the membrane was blocked with 5% (w/v) skim milk powder dissolved in Tris‐buffered saline, 0.1% Tween‐20 (TBST) at room temperature for 1 h. The primary antibodies were diluted in 5% (w/v) bovine serum albumin in TBST and applied and incubated overnight at 4°C. After 1 h of incubation with secondary antibody at room temperature, images were exposed to SuperSignal West Femto Chemiluminescent Substrate (Thermo Scientific) and the VersaDoc Imaging System (Bio‐Rad)and densitometry was performed using Quantity One (Bio‐Rad). All of the phosphorylation data are presented relative to the total protein of the protein of interest.

Statistical analysis

All data are expressed as the mean ± SEM. Statistical testing was performed using SPSS, version 22 (IBM Corp., Armonk, NY, USA) or Prism, version 6 (GraphPad Software Inc., San Diego, CA, USA). For multiple comparisons, one‐way ANOVA and two‐way ANOVA with or without repeat measurement (between factor: insulin and treatment condition – for glucose uptake and protein expression) were used. Tukey's post hoc test or Fisher's least significant difference test was performed when ANOVA revealed statistical significance. P ≤ 0.05 was considered statistically significant.

Results

The effect of serum exposure during ex vivo contraction on mouse skeletal muscle insulin sensitivity 3.5 h post‐contraction

First, we examined the effect of serum on mouse skeletal muscle insulin‐stimulated glucose uptake post ex vivo contraction with slight modifications in comparison with a previously described method (Funai et al. 2010) (Fig. 1 A). It was shown previously that, during an insulin dose response (0, 60, 120 and 20,000 μU ml−1), glucose uptake in isolated mouse skeletal EDL from sedentary mice is maximal at 120 μU ml−1 and tends (P = 0.08) to be increased at the submaximal dose of 60 μU ml−1 (Hamada et al. 2006). In addition, Kjobsted et al. (2017) recently reported that submaximal insulin (100 μU ml−1) and, to a greater extent, maximal insulin (10,000 μU ml−1) enhanced glucose uptake ex vivo in isolated EDL muscle from wild‐type mice 3 h after in situ contraction.

Figure 1. Effect of ex vivo muscle contraction with and without serum on insulin sensitivity of glucose uptake.

Figure 1

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Insulin (120 μU ml−1). A, experimental design. B, 2‐DG uptake. Data are the mean ± SEM, n = 3 or 4 per group. * P < 0.05 vs. no insulin, #P < 0.05 vs. no serum. White bars: no insulin; black bars: insulin.

We anticipated that l‐NMMA would attenuate the insulin‐stimulated glucose uptake after prior contraction. Therefore, 120 μU ml−1 of insulin was used in the present study except where indicated. Our data showed that electrical stimulated contraction in serum‐free buffer did not increase basal (no insulin) or 120 μU ml−1 insulin‐stimulated skeletal muscle glucose uptake in mouse EDL measured 3.5 h post electrical stimulation (Fig. 1 B). By contrast, stimulation of glucose uptake by insulin was markedly (P < 0.05) enhanced 3.5 h post ex vivo contractile activity in muscles stimulated to contract when immersed in 50% human serum in buffer 1 (Fig. 1 B). Therefore, 50% human serum in buffer 1 was used for all experiments, which differs from the 100% serum used previously in rats (Gao et al. 1994; Funai et al. 2010).

NOS inhibition during insulin exposure blocks the increase in the insulin‐stimulated glucose uptake after contraction

We have shown previously that NOS inhibition attenuates the increase in skeletal muscle glucose uptake during contraction in mice and rats (Stephens et al. 2004; Ross et al. 2007; Merry et al. 2010b), as well as during exercise in healthy controls and in people with T2D (Bradley et al. 1999; Kingwell et al. 2002). To examine whether NO is required for the increase in insulin sensitivity post ex vivo contraction (Fig. 2 A), muscles were treated with the NOS inhibitor l‐NMMA (100 μm) either (i) during the period of pre‐incubation (30 min) and muscle contraction (10 min) (NOS inhibition during contraction) or (ii) during vehicle or the 120 μU ml−1 insulin incubation (30 min) and 2‐DG tracer incubation (10 min) (NOS inhibition during contraction). In the absence of insulin skeletal muscle, glucose uptake was similar (P > 0.05) 3.5 h after no contraction, contraction, NOS inhibition during contraction and NOS inhibition during insulin (Fig. 2 B). This indicates that the effect of prior contraction had worn off. Contraction significantly (P < 0.01) increased insulin‐stimulated glucose uptake 3.5 h post‐contraction and this increase was not affected by NOS inhibition during the pre‐incubation and contraction periods (Fig. 2 B). Surprisingly, NOS inhibition during insulin (and 2‐DG tracer) incubation prevented the increase in insulin‐stimulated glucose uptake in response to prior contraction (Fig. 2 B). The incremental (delta) increase in insulin‐stimulated glucose uptake (insulin‐stimulated glucose uptake minus basal glucose uptake) was significantly higher in the contraction and the contraction plus NOS inhibition during contraction groups than in the non‐contraction and contraction plus NOS inhibition during insulin groups (Fig. 2 B).

Figure 2. NOS inhibition during insulin exposure prevents the increase in insulin‐stimulated glucose uptake and NOS activity 3.5 h post‐contraction in mouse skeletal EDL muscles.

Figure 2

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A, experimental design. B, the effect of NOS inhibition (L‐NMMA; 100 μm) during contraction and during insulin (120 μU ml−1) exposure on glucose uptake 3.5 h after ex vivo contraction. Data are the mean ± SEM, n = 6–12. * P < 0.05 vs. no insulin treatment; #P < 0.05 vs. rest plus insulin group and vs. contraction and then NOS inhibition during insulin group. C, NOS activity of EDL muscles in the presence of insulin. Data are the mean ± SEM, n = 6 per group. #P < 0.05 vs. rest and vs. contraction and then NOS inhibition during contraction group.

NOS activity

NOS activity was significantly reduced by NOS inhibition during insulin treatment to a level significantly below the basal state (Fig. 2 C). NOS activity has a tendency to increase in the NOS inhibition during contraction group, although this was not significant (P = 0.08) (Fig. 2 C).

The NO‐mediated insulin‐sensitizing effect of prior contraction does not involve cGMP/PKG downstream signalling

Because NO signalling involves activation of the soluble form of guanylate cyclase to produce cGMP, the NO/cGMP/PKG signalling pathway is generally considered to be the major downstream target of NO (Stamler & Meissner, 2001) (Fig. 3 A). To explore the mechanism(s) by which NO acts to increase insulin‐stimulated skeletal muscle glucose uptake post contraction, and specifically whether this NO signalling occurs via cGMP/PKG, the soluble guanylate cyclase (sGC) inhibitor ODQ (which blocks the NO‐mediated increase in cGMP), the PDE5 inhibitor T1032 (which inhibits cGMP breakdown and therefore raises cGMP levels) and the cGMP‐dependent PKG inhibitor Rp‐8‐Br‐PET‐cGMPS were applied to block this pathway in acordance with our previous studies (Mahajan et al. 2003; Merry et al. 2010a, b). We found that the insulin sensitizing effects of prior contraction were not affected by the presence of these inhibitors during insulin incubation 3.5 h post‐contraction (Fig. 3 B).

Figure 3. Agents modifying the cGMP/PKG pathway have no effect on insulin‐stimulated glucose uptake 3.5 h after contraction.

Figure 3

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sGC inhibition by ODQ (10 μm), PDE5 inhibition by T‐1032 (27 μm) and PKG inhibition by Rp‐8‐Br‐PET‐cGMPS (5 μm). In all experiments, 120 μU ml−1 of insulin was used, except in the T‐1032 treatment where 60 μU ml−1 was used. A, relationship of the inhibitors used with the cGMP/PKG pathway. B, 2‐DG glucose uptake. Data are the mean ± SEM, n = 4–6 per group. #P < 0.05 vs. rest. White bars: vehicle; black bars: inhibitor. C, experimental design to examine any possible physical interaction between insulin and the inhibitors used. The inhibitors (l‐NMMA, ODQ and T1032) were incubated with insulin for 30 min. D, no physical interaction between insulin and the examined inhibitors. Data are the mean ± SEM, n = 4–6. * P < 0.05 vs. no insulin.

To exclude the possibility that there was a physical interaction between insulin and the inhibitors possibly preventing them from having an effect on insulin‐stimulated glucose uptake, the resting muscles were co‐incubated with or without l‐NMMA, ODQ or T1032 with insulin for 30 min, and then they were incubated with [3H]‐2‐deoxyglucose and [14C]‐mannitol for 10 min to measure glucose uptake (Fig. 3 C). As shown in Fig. 3 D, there was no difference between insulin and insulin plus any of these inhibitors, indicating that no physical interaction could explain the effect of l‐NMMA and the lack of effect of these other agents.

Insulin signalling

There was little Akt Thr308 and Akt Ser473 phosphorylation in the absence of insulin and no significant differences between treatments (Fig. 4). Insulin significantly (P < 0.001) increased the phosphorylation of Akt at both Thr308 and Ser473, with no differences observed between the four treatments (Fig. 4 B and C). Insulin significantly increased the phosphorylation of TBC1D1 at Thr590 (P < 0.01) and Thr596 (P < 0.001) but not at Ser660, with no greater insulin‐stimulated phosphorylation at these sites 3.5 h following prior contraction (Fig. 5 A–D). Although TBC1D4 Thr642 phosphorylation per se did significantly (P < 0.05) increase with insulin (data not shown), given the variability of the total TBC1D4 data this increase was not significant when TBC1D4 Thr642 phosphorylation was presented relative to the total TBC1D4 (Fig. 5 E and F). NOS inhibition either during contraction or during insulin had no significant effect on TBC1D1 or TBC1D4 phosphorylation at the sites examined (Fig. 5).

Figure 4. Akt phosphorylation 3.5 h after ex vivo contraction in mouse skeletal muscle.

Figure 4

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Insulin (120 μU ml−1), n = 6 per group. Data are the mean ± SEM. A, representative blots of pAkt and Akt in the EDL. B–C, statistical results for pAkt Thr308 (B) and pAkt Ser473 (C). * P < 0.05 or ** P < 0.01 or *** P < 0.001 vs. no insulin.

Figure 5. TBC1D1 and TBC1D4 phosphorylation in response to insulin 3.5 h after ex vivo contraction in mouse skeletal muscle.

Figure 5

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A, representative blots of pTBC1D1 and TBC1D1 in the EDL. B–D, statistical results for pTBC1D1 Thr590 (B), Thr596 (C) and Ser660 (D). E, representative blots of pTBC1D4 and TBC1D4 in the EDL. F, statistical results for pTBC1D4 Thr642. Insulin (120 μU ml−1), n = 6 in each group. Data are the mean ± SEM. * P < 0.05 or ** P < 0.01 or *** P < 0.001 vs. no insulin.

Discussion

We report that, in mouse muscle, as has been shown in rat muscle, ex vivo contraction increases insulin sensitivity several hours after contraction. By contrast to our hypothesis, NOS inhibition during contraction had no effect on insulin‐stimulated glucose uptake 3.5 h later. However, remarkably, NOS inhibition during the insulin treatment 3.5 h after contraction prevented the insulin sensitizing effect of the prior contraction. The results of the present study also suggest that the effects of NO on insulin sensitivity after contraction may not act via the classic NO/cGMP/PKG signalling pathway. Given that the measurements were conducted in isolated muscles, these observed effects of NOS inhibition cannot be a result of alterations in other confounders such as blood flow and so must relate to muscle effects per se.

Several previous studies in rats (Gao et al. 1994; Funai et al. 2010) have reported that ex vivo muscle contraction increases skeletal muscle insulin‐stimulated glucose uptake ∼3 h later, which is consistent with human exercise studies (Richter et al. 1989; Wojtaszewski et al. 2000). The results of the present study extend these findings to mice, which is important because this means that studies with genetically modified mice are now possible. As shown in rats (Gao et al. 1994; Funai et al. 2010), we found that it was necessary to include serum during the ex vivo muscle contractions in mice to observe the insulin sensitizing effects of contraction. Furthermore, we found that a mixture of 50% serum with 50% KHB buffer, rather than the 100% serum used in rats, was sufficient to induce greater insulin‐stimulated glucose uptake ∼3 h after ex vivo contraction in mouse skeletal muscle (Fig. 1).

NOS inhibition during contraction in mice and during exercise in humans attenuates the increase in glucose uptake during contraction/exercise (Bradley et al. 1999; Kingwell et al. 2002; Ross et al. 2007; Merry et al. 2010a, b). As such, we hypothesized that NOS inhibition during contraction would attenuate the increase in insulin sensitivity 3.5 h after contraction. However, our hypothesis was not confirmed because NOS inhibition during contraction had no effect on later insulin sensitivity. We previously reported that the addition of l‐arginine can overcome the inhibitory effects of NOS inhibition during contraction (Hong et al. 2015). Therefore, it is possible that the effects of the NOS inhibitor were somewhat nullified by the presence of serum during contraction because l‐arginine is present in healthy human serum at a concentration of ∼100 μm.

Importantly, NOS inhibition during insulin incubation blocked the increase in insulin sensitivity in response to earlier contraction (Fig. 2 B). The mechanism(s) involved are unclear at this stage. The relationships between skeletal muscle, NO production, NOS activity, diabetes, exercise and insulin sensitivity are complex. Insulin has been shown to increase nNOS phosphorylation in C2C12 muscle cells and in mouse skeletal muscle (Hinchee‐Rodriguez et al. 2013) and skeletal muscle NOS activity is reported to increase during a euglycaemic hyperinsulinaemic clamp in healthy humans (Kashyap et al. 2005). Therefore, it is possible that insulin activates increases in skeletal muscle NO production to increase glucose uptake and also that the NOS inhibitor subsequently prevented this effect. Indeed, in line with the prevention of the contraction‐stimulated increase in insulin sensitivity, NOS activity was significantly reduced in the presence of NOS inhibition during insulin treatment (Fig. 2 C).

Most studies in rodents and humans find little effect of prior exercise or contraction on proximal insulin signalling (Wojtaszewski et al. 2000; Hamada et al. 2006; Funai et al. 2010; Castorena et al. 2014). In line with this, we found that there was no difference in insulin‐stimulated Akt phosphorylation with or without prior ex vivo contraction (Fig. 4). Despite unaltered proximal signalling, some studies have reported greater downstream insulin signalling at the level of TBC1D4 3 h after exercise in rats and humans (Funai et al. 2009; Treebak et al. 2009; Castorena et al. 2014). Although previous studies reported increases in mouse EDL TBC1D4 Thr642 phosphorylation with insulin (Chen et al. 2011; Kjobsted et al. 2015; Kjobsted et al. 2017), we found no significant increase in TBC1D4 Thr642 phosphorylation with insulin in the present study when TBC1D4 Thr642 phosphorylation was presented relative to the total TBC1D4. However, TBC1D4 Thr642 phosphorylation per se did increase with insulin but, given the variability with total TBC1D4, this effect was lost when TBC1D4 Thr642 phosphorylation was divided by total TBC1D4 (Fig. 5 F).

In our human study, we have now shown that skeletal muscle pTCB1D4 Thr704 (pTCB1D4 Thr711in mice) is increased 4 h after exercise (Sjoberg et al. 2017). In addition, the increase in pTCB1D4 Thr704 during an euglycaemic hyperinsulinaemic clamp is greater in previously exercised muscle than in non‐exercised muscle in humans (Sjoberg et al. 2017). It is not known whether similar responses of pTCB1D4 Thr711 are observed in mice because, unfortunately, an antibody for TBC1D4 704/711 phosphorylation was not commercially available when we conducted our study. Future mouse studies should aim to examine this site.

It is important to note that Funai et al. (2010) reported additive effects of prior in vivo exercise and ex vivo contraction on insulin stimulated glucose uptake, suggesting that in vivo exercise and ex vivo contraction may enhance insulin sensitivity by different mechanisms. Along these lines, we recently found (Sjoberg et al. 2017) that NOS inhibition in humans overcomes the greater insulin sensitivity in a leg that exercise 4 h earlier compared to a rested leg. In that previous study (Sjoberg et al. 2017), and similar to the present study, NOS inhibition had no effect on insulin signalling in either the contracted on non‐contracted muscle. However, it appeared that the reduction in blood flow with NOS inhibition (especially in microvascular blood flow) in the previous study (Sjoberg et al. 2017) was the major reason for NOS inhibition, as in the present study, overcoming/preventing the increased insulin sensitivity because of earlier exercise. However, in the present study, there is no blood flow component. These results support the suggestion that in vivo exercise and ex vivo contraction may enhance insulin sensitivity by different mechanisms, with both involving NO. Further research is required to clarify this.

Akt, TBC1D1 and TBC1D4 phosphorylation were not affected by NOS inhibition during insulin treatment and therefore do not appear to account for the observed effects of NOS inhibition preventing the increase in insulin sensitivity after contraction. The mechanisms responsible for this remarkable effect of NOS inhibition on insulin‐stimulated glucose uptake after contraction are not clear. Recent evidence indicates that the cytoskeleton is important for skeletal muscle glucose uptake in response to both contraction and insulin (Su et al. 2005; Wang et al. 2011; Sylow et al. 2013a) and, given that skeletal muscle nNOS is associated with the cytoskeleton (Percival et al. 2010), it is possible that this could be playing a role. Depolymerization of the actin cytoskeleton decreases glucose uptake (Sylow et al. 2013b) and rearrangement of the actin cytoskeleton by Rac1 (Ras‐related C3 botulinum toxin substrate 1), a small Rho family GTPase, is necessary for insulin‐stimulated GLUT4 translocation in L6 myotubes (Ueda et al. 2008). In addition, Rac1 and its downstream target, PAK1, are activated by contraction/exercise in human and mouse skeletal muscle and insulin‐stimulated GLUT4 translocation is impaired in skeletal muscle from Rac1 knockout (KO) mice (Sylow et al. 2013a, b). Inhibition of Rac1 or Rac1 KO reduces both contraction‐stimulated and insulin‐stimulated glucose uptake in mouse muscle (Sylow et al. 2013a, b). There is also some evidence of interactions between Rac1 and NO, including in C2C12 muscle cells (Su et al. 2005; Cheng et al. 2006; Godfrey & Schwarte, 2010). Follow‐up studies should examine whether NOS inhibition during insulin exposure attenuates the observed increases in pPAK1 after prior ex vivo skeletal muscle contraction.

The cGMP/PKG pathway, which is present in skeletal muscle, is generally considered to be the major downstream signalling pathway of NO (Stamler & Meissner, 2001). However, modification of cGMP/PKG signalling with the sGC inhibitor ODQ (guanylate cyclase produces cGMP in response to NO), the PDE5 inhibitor T1032 (PDE5 breaks down cGMP) and the cGMP‐dependent PKG inhibitor Rp‐8‐Br‐PET‐cGMPS showed no significant effect on the insulin‐sensitizing effects of prior contraction in mouse muscle ex vivo (Fig. 3 A and B). These results suggest that NO increases skeletal muscle insulin sensitivity post‐contraction via cGMP/PKG independent mechanism(s). This is similar to that previously during ex vivo contractions where l‐NMMA attenuates the increase in skeletal muscle glucose uptake during ex vivo contractions, whereas there is no effect of inhibition of sGC or PKG (Merry et al. 2010a). Moreover, in endothelial cells (Wang et al. 2013) and in adipocytes (Kaddai et al. 2008), the stimulatory effect of NO donors on insulin transport occurred not via cGMP/PKG but instead via S‐nitrosylation.

The alternatively‐spliced isoform of nNOS, nNOSμ, is the primary source of skeletal muscle NO during contraction in mouse muscle (Silvagno et al. 1996) and in contracting muscle cells (Hirschfield et al. 2000). Indeed, it has been shown that contraction increases cGMP during ex vivo skeletal muscle contraction in normal mice and eNOS KO mice but not in nNOSμ KO mice (Lau et al. 2000). Therefore, it is possible that, in the present study, skeletal muscle NO production was from nNOSμ. Follow‐up studies should aim to examine whether the increase in insulin sensitivity after ex vivo contraction is attenuated in nNOSμ mouse muscle. In addition, studies with NOS inhibition in humans could be conducted to determine whether NO production plays a role in the insulin sensitizing effects of exercise in humans. We have infused local NOS inhibitors into the femoral artery of humans during exercise in studies examining the role of NO in glucose uptake during exercise (Bradley et al. 1999). Similar methods could be used with the infusion of a NOS inhibitor during insulin several hours after acute exercise. It has been shown that, 4 h after single leg exercise, there is 50% or greater increases in insulin‐stimulated glucose uptake into the exercised leg compared to the rested leg (Richter et al. 1989).

As a result of technical difficulties and the small muscle mass in the present study, we were unable to measure sGC activity to confirm the efficacy of ODQ or measure PKG activity to confirm the efficacy of Rp‐8‐Br‐PET‐cGMP. However, the same concentration of ODQ used in the present study should be considered to prevent NO donor stimulated increases in glucose uptake in EDL muscle (Merry et al. 2010a).

In conclusion, we have shown that NO is required for normal increases in insulin sensitivity several hours after the ex vivo contraction of mouse muscle. NOS inhibition during contraction had no effect on insulin sensitivity 3.5 h later but, remarkably, NOS inhibition during insulin exposure post‐contraction prevented the increases in insulin sensitivity following ex vivo contraction. Although we found that NOS inhibition during insulin treatment post‐contraction had no effect on Akt, TBC1D1 or TBC1D4 phosphorylation at the sites examined, future mouse studies should investigate other sites of TBC1D4 phosphorylation, and especially whether the increase in pTCB1D4 Thr704 in response to insulin in humans (pTCB1D4 Thr711 in mice) is greater ∼5 h after exercise. Finally, given that blocking sGC and PKG during insulin exposure had no effect on the increase in insulin sensitivity after contraction, this suggests that NO acts independently of the cGMP/PKG pathway to increase insulin sensitivity after contraction.

Additional information

Competing interests

The authors declare that they have no competing financial interests.

Author contributions

XZ and GKM were responsible for the conception and design of the study. XZ, DH and YHH conducted the experiments. XZ, DH, SR and GKM contributed to the analysis of data. AZ and AH contributed to set up ex vivo contraction apparatus. XZ and GKM wrote the first version of the manuscript. All authors contributed to the review and editing of the manuscript. All authors have approved the final version of the manuscript and agree to be accountable for all aspects of the work. All persons designated as authors qualify for authorship, and all those who qualify for authorship are listed.

Funding

The present study was funded by the National Health and Medical Research Council (NHMRC, Project grant number 1012181 to GKM) and Biomedical & Lifestyle Diseases (BioLED) to XZ in Australia.

Acknowledgements

The authors thank Associate Professor Itamar Levinger for fruitful discussion and helpful suggestions.

Edited by: Michael Hogan & Paul Greenhaff

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