| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
SKELETAL MUSCLE AND EXERCISE |
1 Department of Physiology and Pharmacology, Karolinska Institutet, 171 77, Stockholm, Sweden
2 Department of Laboratory Medicine, Karolinska Institutet, Novum, Karolinska University Hospital, 141 86, Stockholm, Sweden
3 Department of Physiology, University of Kentucky Medical Center, Lexington, KY 40436-0298, USA
 |
Abstract |
|---|
 Top Abstract IntroductionMethodsResultsDiscussionReferences |
|---|
50% (P < 0.05 versus control values), but did not significantly affect basal 2-DG uptake or the uptake induced by insulin, hypoxia or 5-aminoimidazole-4-carboxamide-1-ß-D-ribofuranoside (AICAR, which mimics AMP-mediated activation of AMP-activated protein kinase, AMPK). Ebselen, a glutathione peroxidase mimetic, also inhibited contraction-mediated 2-DG uptake (by almost 60%, P < 0.001 versus control values). Muscles from mice overexpressing Mn2+-dependent superoxide dismutase, which catalyses H2O2 production from superoxide anions, exhibited a 25% higher rate of contraction-mediated 2-DG uptake versus muscles from wild-type control mice (P < 0.05). Exogenous H2O2 induced oxidative stress, as judged by an increase in the [GSSG]/[GSH + GSSG] (reduced glutathione + oxidized glutathione) ratio to 2.5 times control values, and this increase was substantially blocked by NAC. Similarly, NAC significantly attenuated contraction-mediated oxidative stress as judged by measurements of glutathione status and the intracellular ROS level with the fluorescent indicator 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein (P < 0.05). Finally, contraction increased AMPK activity and phosphorylation 10-fold, and NAC blocked 50% of these changes. These data indicate that endogenously produced ROS, possibly H2O2 or its derivatives, play an important role in contraction-mediated activation of glucose transport in fast-twitch muscle.
(Received 29 March 2006;
accepted after revision 13 June 2006;
first published online 15 June 2006)
Corresponding author A. Katz: Department of Physiology and Pharmacology, Karolinska Institutet, 171 77 Stockholm, Sweden. Email: abram.katz{at}ki.se
|
Introduction |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
|
Methods |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
2-Deoxy-D-[1,2-3H]glucose (2-DG), carboxy-[14C]inulin and [
-32P]ATP were from Amersham Biosciences. Ebselen (E3520) and N-acetylcysteine (NAC, A8199) were from Sigma. Human insulin (Actrapid) was from Novo Nordisk. 5-(and-6)-Chloromethyl-2',7'-dichloro-dihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) was from Invitrogen. Antibodies against pan-
-AMPK, phosphorylated -AMPK (T172), acetyl CoA carboxylase (ACC) and phosphorylated ACC (S79) were from Cell Signalling Technology. All other reagents were from either Sigma or Boehringer Mannheim.
Male NMRI mice weighing 25–30 g were housed at room temperature with a 12 h–12 h light–dark cycle. Food and water were provided ad libitum. The mice were purchased from B&K Universal (Sollentuna, Sweden). Mice overexpressing Mn2+-dependent superoxide dismutase (mitochondrial isoform; SOD2) and wild-type (WT) littermates were generated as described elsewhere (Silva et al. 2005). Superoxide dismutase catalyses the following reaction:
|
|
We used mice from the PAC662D1 line exhibiting an almost sixfold increased SOD2 activity (Silva et al. 2005). The mice weighed 25–30 g, and the SOD2 mice did not differ from the WT littermates with respect to body weight and whole body oxygen consumption at 4 or 24°C either before or after noradrenaline injection (Silva et al. 2005). Furthermore, there were no significant differences in twitch kinetics, force–frequency relationship, fatigue development or recovery from fatigue between WT and SOD2 extensor digitorum longus (EDL) muscles (data not shown). This demonstrates that SOD2 overexpression did not alter contractile function, which suggests that muscle fibre composition was also not altered. Animals were killed by rapid cervical dislocation, and the EDL muscles were isolated. All procedures were approved by the Stockholm North ethics committee.
Experimental design
Contraction experiments.
For contraction studies, stainless-steel hooks were tied with nylon thread to the tendons of the muscles. Muscles were then transferred to a stimulation chamber (volume 
10 ml) and mounted between a force transducer and an adjustable holder (World Precision Instruments). The chamber temperature was set at 25°C with a water-jacketed circulation bath. The muscle was bathed in a Tyrode solution with the following composition (mM): NaCl, 121; KCl, 5; CaCl2, 1.8; NaH2PO4, 0.4; MgCl2, 0.5; NaHCO3, 24; EDTA, 0.1; glucose, 5.5; and 0.1% fetal calf serum. The pH of the Tyrode solution was set to 7.4 by continuously and directly gassing the solution with 95% O2–5% CO2. Muscles were stimulated with current pulses (0.5 ms duration; 150% of the current required for maximum force response) via plate electrodes lying parallel to the fibres. Muscles were set to the length at which tetanic force was maximum and then bathed in a Tyrode solution that contained one of two antioxidants: either 20 mM
N-acetylcysteine (NAC; control being addition of 10 mM NaCl) or 30 µM ebselen in DMSO (control being an equivalent volume of DMSO). As a control for the osmotic effect of NAC, 10 mM NaCl was always added to the 121 mM NaCl Tyrode solution bathing the contralateral muscle. Muscles were then allowed to rest for 60 min. Thereafter, the muscles were stimulated at 50 Hz (tetanic duration 100 ms, 2 trains s–1) for 10 min. This protocol decreases muscle glycogen by 80% (Sandström et al. 2004). For measurements of AMPK and glutathione status (see below), muscles were quick-frozen in liquid N2 within 5 s after the last tetanus. For measurement of glucose uptake, muscles were transferred at the end of the 10 min stimulation to vials containing 1.5 ml Tyrode solution lacking glucose and containing 2 mM pyruvate with or without the test drug (or vehicle) and incubated in a shaking water-bath (110 oscillations min–1, 35°C, air phase in vial was continuously gassed with 95% O2–5% CO2) for 40 min and frozen in liquid nitrogen. Radiolabelled 2-DG (1 mM) and inulin were added 20 min before freezing as described elsewhere (Shashkin et al. 1995).
To assess intracellular ROS formation in contracting muscle, small bundles of 5–20 fibres were mechanically dissected from the EDL muscle. Bundles were loaded with the ROS-sensitive indicator by incubation for 90–120 min in 10 µM CM-H2DCFDA at room temperature. Thereafter, bundles were transferred to the muscle trough and stretched to the length at which tetanic force was maximal. Muscles were then washed for 30 min with 5.5 mM glucose Tyrode solution (± 20 mM NAC). A BIO-RAD MRC 1024 and a Nikon Diaphot 200 inverted microscope with a x20 objective lens (NA 0.75) were used. In the cell, esterases cleave CM-H2DCFDA to release CM-H2DCFH, which is converted to a fluorescent product (CM-H2DCF) when exposed to ROS (Xie et al. 1999). CM-H2DCF was excited with 488 nm light, and the emitted light collected through a 515 nm long-pass filter. Confocal images were taken of the muscle fibres at rest and then after 10 min of intermittent tetanic contractions (same stimulation protocol as for the whole EDL muscle). When NAC was added, it was after the loading period and remained in the medium until after the postcontraction scans were made. CM-H2DCF fluorescence was measured before and after contraction and after the bundle was exposed to 1 mM H2O2 for 5 min. Confocal images were stored and analysed offline with ImageJ (available at http://rsb.info.nih.gov/ij/). Changes in intracellular CM-H2DCF fluorescence intensity are expressed as a percentage of that before the series of contractions or H2O2 exposure.
Rested muscle experiments.
A series of glucose uptake experiment was also performed in resting muscle preparations. These muscles were incubated in 1.5 ml of the pyruvate-supplemented Tyrode solution at 35°C in the shaking water-bath as described above. To examine the effects of antioxidants on basal glucose uptake, muscles were incubated in the presence of NAC or ebselen (with appropriate controls) in the shaking bath for 80 min and then frozen. To examine the effects of hypoxia with or without NAC on glucose uptake, muscles were continuously gassed with 95% N2–5% CO2 for a total of 80 min (Cartee et al. 1991). N-Acetylcysteine was present throughout the 80 min incubation period in half the samples. Since 60 min are required to observe the maximal effect of hypoxia on glucose transport in isolated muscle preparations (Cartee et al. 1991), no preincubation with NAC in 95% O2–5% CO2 was performed. To examine the effects of 5-aminoimidazole-4-carboxamide-1-ß-D-ribofuranoside (AICAR; final concentration, 2 mM) on glucose uptake, muscles were incubated first for 30 min and then AICAR was added to the muscles and the incubation was continued for an additional 80 min (Kurth-Kraczek et al. 1999). N-Acetylcysteine was present throughout the 110 min incubation period in half the samples. AICAR is converted in the cell to AICAR-monophosphate, which is an AMP mimetic that activates AMPK and AMPK kinase (Young et al. 1996; Kurth-Kraczek et al. 1999). To examine the effects of insulin (final concentration, 20 mU ml–1) on glucose uptake, muscles were preincubated for 30 min; insulin was added and the incubation was continued for an additional 50 min. N-Acetylcysteine was present throughout the 80 min incubation period in half the samples. 2-Deoxyglucose and inulin were present during the last 20 min of incubation in these experiments. During the insulin experiments, bovine serum albumin (0.1% v/v) was present in the pyruvate-supplemented Tyrode solution to inhibit insulin binding to the glass vial.
To study the effects of exogenous H2O2 (final concentration = 3 mM) on glutathione status, muscles were incubated in 1.5 ml of the 5.5 mM glucose Tyrode solution in the shaking bath at 35°C for 90 min and then frozen. H2O2 was added after the initial 60 min of incubation. NAC was present throughout the 90 min incubation period in half the samples. To study the effects of NAC on glutathione status and AMPK activity in resting muscles, muscles were incubated in 1.5 ml of the 5.5 mM glucose Tyrode solution in the shaking bath at 25°C for 60 min and then frozen. NAC was present throughout the 60 min incubation period in half the samples. The latter experiments were performed at 25°C, because the contraction-mediated changes in glutathione status and AMPK activity were also studied at this temperature.
To study the effects of exogenous H2O2 (final concentration, 3 mM) on glucose uptake, muscles were incubated in pyruvate-supplemented Tyrode solution in the shaking bath for 80 min at 35°C (see above). Hydrogen peroxide was added to half the muscles after 30 min. Isotopes were present during the last 20 min of incubation. To examine the effects of exogenous H2O2 on AMPK activity, muscles were incubated in the shaking bath for 60 min at 25°C in 5.5 mM glucose Tyrode solution (since basal and contraction-mediated AMPK activities were studied under these conditions). Hydrogen peroxide was added to half the muscles after 30 min.
Analytical
2-Deoxyglucose uptake. For analysis of 2-DG uptake, frozen muscles were added to preweighed Eppendorf tubes containing 0.5 ml of 1 N NaOH. The muscle was weighed and then digested at 70°C for 15 min. The tubes were cooled on ice and centrifuged at 23 000g for 5 min. Aliquots of the supernatant were added to scintillation cocktail and counted for 14C and 3H as described earlier (Shashkin et al. 1995).
Glutathione. For analysis of glutathione, a kit was used (Biooxytech GSH/GSSG-412, Oxis Health Products, Portland, OR, USA). Muscles were freeze-dried, dissected free of non-muscle constituents, powdered and thoroughly mixed. The powders were divided into two aliquots, which were homogenized in ground glass homogenizers containing ice-cold 5% metaphosphoric acid (80 µl (mg dry weight)–1) with and without 1-methyl-2-vinyl-pyridinium trifluoromethane sulphonate (M2VP; 10% v/v), a scavenger of reduced glutathione (GSH). The homogenates were centrifuged at 23 000g for 15 min at 4°C. The pellets were digested with 1 N NaOH (60°C) and assayed for protein with the Bio-Rad assay (BIO-RAD). For measurement of reduced + oxidized glutathione (TGSH = GSH + oxidized gluthathione [GSSG]), 4 µl supernatant were mixed with 96 µl assay buffer, and for GSSG estimation, 5 µl of the supernatant with M2VP were mixed with 95 µl GSSG assay buffer. For both assays, the samples were mixed with 300 µl of chromagen, glutathione reductase and NADPH, and absorbance (reduction of dithiobis-2-nitrobenzoic acid at 412 nm) was measured after 4.5 min in a spectrophotometer. Preliminary experiments demonstrated that the assays were linear with respect to the extract volume used and reaction time, and GSH was not detectable in the presence of M2VP (data not shown).
AMPK.
AMPK activity was analysed by assessing the incorporation of radiolabelled phosphate from ATP into SAMS peptide (Winder & Hardie, 1996) with some modifications. Briefly, muscles were freeze-dried and treated as above. Muscles were homogenized in ice-cold buffer (100 µl (mg dry weight)–1) consisting of (mM): Tris, 10; sucrose, 250; NaF, 50; EDTA, 1; ß-mercaptoethanol, 10; and one tablet protease inhibitor cocktail (Roche) per 50 ml of buffer, pH 7.5. The homogenate was centrifuged at 23 000g for 30 min at 4°C. The supernatant was divided into aliquots. One aliquot was assayed for protein (see above). Another aliquot was diluted with seven volumes of homogenization buffer, and 10 µl of the diluted extract were mixed with 30 µl reaction buffer, resulting in the following final concentrations (mm): Hepes (pH 7.0), 40; SAMS peptide, 0.2; NaCl, 80; EDTA, 0.8; AMP, 0.2; dithiothreitol (DTT), 0.8; MgCl2, 5; ATP, 0.2; [-32P]ATP, 2 µCi; and glycerol, 8% (v/v);. The assay was performed at 37°C for 10 min. Thereafter, 30 µl of the mixture were spotted onto Whatman P81 discs, washed in 1% phosphoric acid, dried and counted. Blanks consisted of mixtures spotted without incubation. The assay was linear with respect to the used extract volume and reaction time, and no activity was detected in the absence of SAMS peptide (data not shown). In preliminary experiments, we also assayed the AMPK activity in ammonium-sulphate-precipitated extracts as described by others (Winder & Hardie, 1996). While this procedure also yielded linearity with respect to extract volume and reaction time, there was a large loss of enzyme activity when compared to results from untreated extracts (data not shown). These findings are in agreement with results recently reported by others (Derave et al. 2000). Because of the prevention of enzyme loss and the decrease in preparative steps, assays were performed on untreated supernatants.
Western blots were performed for phosphorylated and total AMPK and ACC. Briefly, 20 (total and phosphorylated ACC) or 25 µg (total and phosphorylated AMPK) of supernatant (see above) protein were separated by SDS-PAGE (4–12% Bis-Tris Gels; Invitrogen) and transferred onto polyvinylidine fluoride (PVDF) membranes (BIO-RAD). Membranes were blocked in 5% (w/v) non-fat milk in Tris-buffered saline containing 0.05% Tween 20 followed by incubation with primary antibodies (all at 1:1000 dilution) overnight. Membranes were then washed and incubated with secondary antibody (donkey antirabbit at 1:2000 dilution for all). Immunoreactive bands were visualized using enhanced chemiluminescence (Super Signal, Pierce, Rockford, IL, USA.). Band densities were analysed with ImageJ (NIH, USA; http://rsb.info.nih.gov/j/).
Values for glutathione and AMPK are expressed relative to dry weight. Differences between groups were maintained also if the results were adjusted for protein. Force was sampled on-line and stored on a desktop computer for subsequent analysis. Tetanic force was measured as the peak force during the 100 ms of stimulation.
Statistics
Signficant differences between means were determined with Student's t test for paired samples. P < 0.05 was regarded as significant. Values are presented as means ± S.E.M.
|
Results |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Contraction increased 2-DG uptake roughly threefold (Fig. 1A). In the presence of NAC, the rate of 2-DG uptake after contraction was significantly lower than during control conditions. Thus, NAC abolished 50% of contraction-mediated 2-DG uptake. N-Acetylcysteine did not affect basal 2-DG uptake, which is primarily ascribed to the availability of Glut-1 transport proteins in the cell membrane (Mueckler, 1994). However, contraction accelerates glucose transport via translocation of Glut-4 transport proteins to the cell surface. Therefore, we determined whether other interventions that increase glucose transport via translocation of Glut-4 proteins (Holloszy, 2003; Jessen & Goodyear, 2005; Rose & Richter, 2005) are affected by NAC. We found that insulin, hypoxia and AICAR increased 2-DG uptake to values similar to that seen with contraction, but NAC did not significantly affect these increases. It could be argued that NAC interfered with contraction-mediated 2-DG uptake owing to an inhibition of force production. However, force generation during the repeated contractions in the presence of NAC was virtually identical to control values (Fig. 1B).
|
20 min at 35°C (Zhang et al. 2006), and a 60 min preincubation period was required, the contraction-mediated 2-DG uptake was induced at 25°C. However, insulin-, hypoxia- and AICAR-mediated 2-DG uptake were induced at 35°C. All 2-DG uptake measurements were performed at 35°C. Therefore, caution should be exerted in directly comparing 2-DG uptake between different modes of glucose uptake induced at different temperatures. N-Acetylcysteine and ROS formation
The above results with NAC implicated endogenously produced ROS in contraction-mediated glucose uptake. Glutathione status is commonly used as a measure of intracellular oxidative stress and was therefore studied in the next series of experiments. Glutathione peroxidase catalyses the following reaction: H2O2
+ 2GSH 
GSSG + 2H2O; thus changes in GSH and GSSG will reflect changes in H2O2. First, we incubated muscles with 3 mM H2O2 in the absence and presence of NAC. Hydrogen peroxide resulted in a marked degree of oxidative stress as evidenced by a decrease in TGSH (Fig. 2). The decrease in TGSH was probably associated with an increase in mixed disulphides, since protein glutathionylation occurs with excessive oxidative stress (Ghezzi, 2005). Hydrogen peroxide also increased the GSSG/TGSH ratio to 250% of control values, and NAC counteracted the H2O2-dependent effect. We then investigated the effect of contraction in the absence and presence of NAC on glutathione status. N-Acetylcysteine did not affect glutathione status in the basal state, but resulted in significantly smaller changes in GSSG and the GSSG/TGSH ratio following contraction (Fig. 3). In summary, NAC decreased the oxidative stress associated with addition of exogenous H2O2 and repeated contractions.
|
|
|
The results thus far indicated that ROS/H2O2 (or their derivatives) produced during contraction resulted in an accelerated glucose transport that was presumably mediated by the Glut-4 transport protein. To investigate the pathway through which ROS were working, we measured AMPK activity following contraction in the absence and presence of NAC. N-Acetylcysteine had no significant effect on AMPK activity in resting muscle (Fig. 5). Contraction resulted in almost a 10-fold increase in AMPK activity and about 50% of the increase was blocked by NAC. Thus NAC inhibited contraction-mediated activation of AMPK and 2-DG uptake to the same relative extent. We verified that exogenous H2O2 activates AMPK in mouse EDL muscles (basal, 5.9 ± 1.6 nmol (g dry weight)–1 min–1; and in the presence of 3 mM H2O2, 18.2 ± 4.0 nmol (g dry weight)–1 min–1; n
= 6; P < 0.05). It is noteworthy that the increase in AMPK activity induced by H2O2 under basal conditions (12 nmol (g dry weight)–1 min–1) is similar to that blocked by NAC during contraction (15 nmol (g dry weight)–1 min–1; Fig. 5).
|
|
N-Acetylcysteine is a general antioxidant and thus does not indicate the species of ROS that may mediate glucose transport during contraction. However, the results thus far were consistent with the involvement of H2O2. To assess the role of H2O2 more directly, two strategies were employed: (1) use of another antioxidant, ebselen; and (2) use of muscles overexpressing SOD2. Ebselen is a glutathione peroxidase mimetic that removes H2O2 in the presence of GSH (Cotgreave et al. 1987). Thus ebselen should enhance H2O2 removal and would therefore be expected to inhibit contraction-mediated glucose uptake. With respect to SOD2 overexpression, we assumed that these muscles would exhibit increased H2O2 production (hence increased glucose uptake) during contraction, owing to increased conversion of superoxide to H2O2. Consistent with this assumption is the observation that when superoxide production in mitochondria isolated from skeletal muscle is accelerated, the accumulation of superoxide is decreased in mice overexpressing SOD2 versus wild type (Silva et al. 2005).
Ebselen significantly increased basal 2-DG uptake (Fig. 7A). Despite this increase, 2-DG uptake was still significantly lower after contraction in the presence of ebselen. Thus the contraction-mediated 2-DG uptake was diminished by ebselen by almost 60%, which is similar to the inhibition seen with NAC. Ebselen resulted in a small decrease in initial force (control, 15.6 ± 1.4 mN (mg wet weight)–1; ebselen, 13.0 ± 0.9 mN (mg wet weight)–1; P < 0.05). However, the fatigue profile was not significantly affected during the repeated contractions (Fig. 7B).
|
25%) after a series of repeated contractions in SOD2 overexpressing versus wild-type muscles. This increase was not associated with alterations in force production (Fig. 7D). It might be argued that the increase in contraction-mediated 2-DG uptake is a consequence of an adaptive increase in the capacity of the glucose transport system and thus all interventions that result in Glut-4 translocation will result in higher values in SOD2 overexpressing muscles. We therefore stimulated 2-DG uptake with insulin. In contrast to the effect of SOD2 overexpression on contraction-mediated 2-DG uptake, insulin-mediated 2-DG uptake was not significantly altered (WT, 2.72 ± 0.31 µmol (20 min)–1 (ml intracellular water)–1; SOD2, 2.51 ± 0.24 µmol (20 min)–1 (ml intracellular water)–1; n
= 6). The data from the NAC, ebselen and SOD2 experiments are consistent with the idea that H2O2, or its derivatives, plays a significant in role in contraction-mediated glucose transport. In line with this idea, we determined that exogenous H2O2 also stimulates 2-DG uptake in mouse EDL muscles (basal, 0.82 ± 0.31 µmol (20 min)–1 (ml intracellular water)–1; in presence of 3 mM H2O2, 2.10 ± 0.31 µmol (20 min)–1 (ml intracellular water)–1; P < 0.001).
|
Discussion |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
While this study was in progress, it was reported that exogenous H2O2, as well as a superoxide-generating system, accelerated glucose transport in isolated resting rat epitrochlearis muscles by a mechanism that involved the activation of the 1 isoform of AMPK (Toyoda et al. 2004). Moreover, the effects of H2O2 and the superoxide-generating system on glucose transport and AMPK activity, as well as glutathione status, were partly blocked by NAC. These results are consistent with the findings in the present study, which indicate that endogenous ROS production is involved in activating glucose transport during contraction. Thus, of the various ROS components, H2O2, or its derivatives, is most likely to be the activating factor during contraction.
The possibility that reactive nitrogen species (RNS) also participate in contraction-mediated glucose transport was not addressed in the present study. Indeed, there is considerable evidence that RNS are produced during muscle contraction (Murrant & Reid, 2001). Specifically, increased NO production has been proposed to mediate contraction-stimulated glucose transport (Balon & Nadler, 1997; Roberts et al. 1997). However, other studies indicate that NO-mediated glucose transport occurs via a pathway that is distinct from the contraction-mediated pathway (Etgen et al. 1997; Higaki et al. 2001).
The idea that ROS are involved in contraction-mediated glucose transport may not be intuitive. This is because oxidative stress may play a role in insulin resistance and the pathogenesis of diabetes (Bonnefont-Rousselot, 2002; Houstis et al. 2006). Indeed, it is well known that excessive levels of ROS have deleterious effects on cell function and viability, whereas low/physiological levels of ROS are requisite for various signalling pathways and optimal cell function (Murrant & Reid, 2001; Goldstein et al. 2005). The present results implicating endogenously produced ROS in contraction-mediated glucose transport support the idea that a limited ROS production is requisite for normal physiological function.
An interesting finding was that NAC significantly blocked contraction- but not hypoxia-mediated glucose transport. These two stimuli are believed to activate glucose transport by similar but not identical pathways (Wojtaszewski et al. 1998). It is generally believed that mitochondria are the primary source of ROS produced in eukaryotic cells and that 2–5% of electron flux through the respiratory chain escapes to produce superoxide anions (Murrant & Reid, 2001). Thus an increase in mitochondrial respiration would be expected to result in a proportional increase in superoxide production. There is also evidence that ROS can be produced in extramitochondrial sites and under anaerobic conditions (Murrant & Reid, 2001; Zuo & Clanton, 2005). Our results are consistent with the idea that more ROS are produced during conditions associated with high rates of mitochondrial respiration.
The findings that NAC partly inhibited contraction-mediated activation of AMPK and that exogenous H2O2 activated AMPK raise the question of the mechanism whereby ROS results in AMPK activation. AMPK is usually considered to be activated by increases in AMP and decreases in ATP and phosphocreatine (PCr; Ponticos et al. 1998). While this manuscript was under review, evidence was presented from vascular endothelium cells that mitochondrially derived ROS activated AMPK by a mechanism independent of nucleotide concentrations (Quintero et al. 2006).
Incubation of isolated rat skeletal muscle with ROS (1 mM H2O2 or a superoxide-generating system) also did not result in significant changes in the total tissue AMP/ATP ratio (Toyoda et al. 2004). Similarly, perfusion of isolated rat hearts with 300 µM H2O2, which was sufficient to significantly activate AMPK, did not significantly alter the AMP/ATP ratio (Leon et al. 2004). Furthermore, if oxidants induce decreases in high-energy phosphates, then antioxidants should block such decreases. However, antioxidants do not affect the changes in high-energy phosphates (ATP, PCr and inorganic phosphate) that occur in the isolated rat diaphragm during repeated contractions, hypoxia or contractions in the presence of hypoxia (Wright et al. 2005). Consistent with these data is the present finding that NAC does not interfere with AMP-mediated activation of AMPK (AICAR experiment).
It was recently demonstrated that an important upstream kinase in the AMPK cascade is LKB1 and, based on current evidence, LKB1-mediated phosphorylation and activation of AMPK are not sensitive to AMP. Moreover, blocking LKB1 activity resulted in an inability of H2O2 to activate AMPK (Woods et al. 2003). We therefore suggest that the activation of AMPK by endogenously produced ROS during contraction does not occur via alterations in high-energy phosphates. Other modes of activation of AMPK that are independent of changes in adenine nucleotides include hyperosmotic shock and exposure to metformin (Fryer et al. 2002). Thus, it appears that contraction-mediated activation of AMPK can occur by at least two mechanisms: one that is NAC insensitive and involves changes in high-energy phosphates and one that is NAC sensitive and independent of changes in high-energy phosphates.
A scheme describing the steps whereby we suggest that endogenously produced ROS result in the activation of AMPK and glucose transport is provided in Fig. 8. The scheme indicates that Ca2+, in addition to initiating contraction, can participate in ROS production at different sites in the cell. Hydrogen peroxide-mediated activation of AMPK probably occurs via LKB1 (see above). However, previously presented data indicate that H2O2 does not directly activate LKB1 (Woods et al. 2003). Rather, it appears that H2O2 enhances the substrate suitability of AMPK for LKB1.
|
A complete understanding of the mechanisms involved in contraction-mediated glucose transport is still lacking. To our knowledge, this study provides the first evidence in support of the idea that endogenously produced ROS play a significant role in contraction-mediated glucose transport in fast-twitch muscle.
|
References |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Belluardo N, Westerblad H, Mudo G, Casabona A, Bruton J, Caniglia G, Pastoris O, Grassi F & Ibanez CF (2001). Neuromuscular junction disassembly and muscle fatigue in mice lacking neurotrophin-4. Mol Cell Neurosci 18, 56–67.[CrossRef][Medline]
Bonnefont-Rousselot D (2002). Glucose and reactive oxygen species. Curr Opin Clin Nutr Metab Care 5, 561–568.[CrossRef][Medline]
Cartee
GD, Douen
AG, Ramlal
T, Klip
A
&
Holloszy
JO (1991). Stimulation of glucose transport in skeletal muscle by hypoxia. J Appl Physiol
70, 1593–1600.
Cartee GD & Holloszy JO (1990). Exercise increases susceptibility of muscle glucose transport to activation by various stimuli. Am J Physiol 258, E390–E393.[Medline]
Choi SL, Kim SJ, Lee KT, Kim J, Mu J, Birnbaum MJ, Soo KS & Ha J (2001). The regulation of AMP-activated protein kinase by H2O2. Biochem Biophys Res Commun 287, 92–97.[CrossRef][Medline]
Coderre
L, Kandror
KV, Vallega
G
&
Pilch
PF (1995). Identification and characterization of an exercise-sensitive pool of glucose transporters in skeletal muscle. J Biol Chem
270, 27584–27588.
Cotgreave IA, Sandy MS, Berggren M, Moldeus PW & Smith MT (1987). N-Acetylcysteine and glutathione-dependent protective effect of PZ51 (ebselen) against diquat-induced cytotoxicity in isolated hepatocytes. Biochem Pharmacol 36, 2899–2904.[CrossRef][Medline]
Derave W, Ai H, Ihlemann J, Witters LA, Kristiansen S, Richter EA & Ploug T (2000). Dissociation of AMP-activated protein kinase activation and glucose transport in contracting slow-twitch muscle. Diabetes 49, 1281–1287.[Abstract]
Etgen GJ Jr, Fryburg DA & Gibbs EM (1997). Nitric oxide stimulates skeletal muscle glucose transport through a calcium/contraction- and phosphatidylinositol-3-kinase-independent pathway. Diabetes 46, 1915–1919.[Abstract]
Fischer Y, Rose H, Thomas J, Deuticke B & Kammermeier H (1993). Phenylarsine oxide and hydrogen peroxide stimulate glucose transport via different pathways in isolated cardiac myocytes. Biochim Biophys Acta 1153, 97–104.[Medline]
Fryer
LG, Parbu-Patel
A
&
Carling
D (2002). The anti-diabetic drugs rosiglitazone and metformin stimulate AMP-activated protein kinase through distinct signaling pathways. J Biol Chem
277, 25226–25232.
Ghezzi P (2005). Regulation of protein function by glutathionylation. Free Radic Res 39, 573–580.[Medline]
Goldstein
BJ, Mahadev
K
&
Wu
X (2005). Redox paradox: insulin action is facilitated by insulin-stimulated reactive oxygen species with multiple potential signaling targets. Diabetes
54, 311–321.
Hancock
CR, Janssen
E
&
Terjung
RL (2006). Contraction-mediated phosphorylation of AMPK is lower in skeletal muscle of adenylate kinase-deficient mice. J Appl Physiol
100, 406–413.
Hayashi T, Hirshman MF, Kurth EJ, Winder WW & Goodyear LJ (1998). Evidence for 5'-AMP-activated protein kinase mediation of the effect of muscle contraction on glucose transport. Diabetes 47, 1369–1373.[Abstract]
Higaki
Y, Hirshman
MF, Fujii
N
&
Goodyear
LJ (2001). Nitric oxide increases glucose uptake through a mechanism that is distinct from the insulin and contraction pathways in rat skeletal muscle. Diabetes
50, 241–247.
Holloszy
JO (2003). A forty-year memoir of research on the regulation of glucose transport into muscle. Am J Physiol Endocrinol Metab
284, E453–E467.
Houstis N, Rosen ED & Lander ES (2006). Reactive oxygen species have a causal role in multiple forms of insulin resistance. Nature 440, 944–948.[CrossRef][Medline]
Jessen
N
&
Goodyear
LJ (2005). Contraction signaling to glucose transport in skeletal muscle. J Appl Physiol
99, 330–337.
Jorgensen
SB, Viollet
B, Andreelli
F, Frosig
C, Birk
JB, Schjerling
P, Vaulont
S, Richter
EA
&
Wojtaszewski
JF (2004). Knockout of the 2 but not 1 5'-AMP-activated protein kinase isoform abolishes 5-aminoimidazole-4-carboxamide-1-beta-4-ribofuranoside- but not contraction-induced glucose uptake in skeletal muscle. J Biol Chem
279, 1070–1079.
Kurth-Kraczek EJ, Hirshman MF, Goodyear LJ & Winder WW (1999). 5'-AMP-activated protein kinase activation causes GLUT4 translocation in skeletal muscle. Diabetes 48, 1667–1671.[Abstract]
Leon H, Atkinson LL, Sawicka J, Strynadka K, Lopaschuk GD & Schulz R (2004). Pyruvate prevents cardiac dysfunction and AMP-activated protein kinase activation by hydrogen peroxide in isolated rat hearts. Can J Physiol Pharmacol 82, 409–416.[CrossRef][Medline]
Livingston
JN, Gurny
PA
&
Lockwood
DH (1977). Insulin-like effects of polyamines in fat cells. Mediation by H2O2 formation. J Biol Chem
252, 560–562.
Lund
S, Holman
GD, Schmitz
O
&
Pedersen
O (1995). Contraction stimulates translocation of glucose transporter GLUT4 in skeletal muscle through a mechanism distinct from that of insulin. Proc Natl Acad Sci U S A
92, 5817–5821.
Mu J, Brozinick JT Jr, Valladares O, Bucan M & Birnbaum MJ (2001). A role for AMP-activated protein kinase in contraction- and hypoxia-regulated glucose transport in skeletal muscle. Mol Cell 7, 1085–1094.[CrossRef][Medline]
Mueckler M (1994). Facilitative glucose transporters. Eur J Biochem 219, 713–725.[Medline]
Murrant CL & Reid MB (2001). Detection of reactive oxygen and reactive nitrogen species in skeletal muscle. Microsc Res Tech 55, 236–248.[CrossRef][Medline]
Nagata
D, Takeda
R, Sata
M, Satonaka
H, Suzuki
E, Nagano
T
&
Hirata
Y (2004). AMP-activated protein kinase inhibits angiotensin II-stimulated vascular smooth muscle cell proliferation. Circulation
110, 444–451.
Park
SH, Gammon
SR, Knippers
JD, Paulsen
SR, Rubink
DS
&
Winder
WW (2002). Phosphorylation-activity relationships of AMPK and acetyl-CoA carboxylase in muscle. J Appl Physiol
92, 2475–2482.
Ponticos M, Lu QL, Morgan JE, Hardie DG, Partridge TA & Carling D (1998). Dual regulation of the AMP-activated protein kinase provides a novel mechanism for the control of creatine kinase in skeletal muscle. EMBO J 17, 1688–1699.[CrossRef][Medline]
Quintero
M, Colombo
SL, Godfrey
A
&
Moncada
S (2006). Mitochondria as signaling organelles in the vascular endothelium. Proc Natl Acad Sci U S A
103, 5379–5384.
Reid
MB, Haack
KE, Franchek
KM, Valberg
PA, Kobzik
L
&
West
MS (1992). Reactive oxygen in skeletal muscle. I. Intracellular oxidant kinetics and fatigue in vitro. J Appl Physiol
73, 1797–1804.
Roberts CK, Barnard RJ, Scheck SH & Balon TW (1997). Exercise-stimulated glucose transport in skeletal muscle is nitric oxide dependent. Am J Physiol 273, E220–E225.[Medline]
Rose
AJ
&
Richter
EA (2005). Skeletal muscle glucose uptake during exercise: how is it regulated?
Physiology
20, 260–270.
Sakamoto K, McCarthy A, Smith D, Green KA, Grahame HD, Ashworth A & Alessi DR (2005). Deficiency of LKB1 in skeletal muscle prevents AMPK activation and glucose uptake during contraction. EMBO J 24, 1810–1820.[CrossRef][Medline]
Sandström ME, Abbate F, Andersson DC, Zhang SJ, Westerblad H & Katz A (2004). Insulin-independent glycogen supercompensation in isolated mouse skeletal muscle: role of phosphorylase inactivation. Pflugers Arch 448, 533–538.[Medline]
Shashkin
PN, Koshkin
A, Langley
DR, Ren
J-M, Westerblad
H
&
Katz
A (1995). Effects of CGS 9343B (a putative calmodulin antagonist) on isolated skeletal muscle: dissociation of signaling pathways for insulin-mediated activation of glycogen synthase and hexose transport. J Biol Chem
270, 25613–25618.
Silva JP, Shabalina IG, Dufour E, Petrovic N, Backlund EC, Hultenby K, Wibom R, Nedergaard J, Cannon B & Larsson NG (2005). SOD2 overexpression: enhanced mitochondrial tolerance but absence of effect on UCP activity. EMBO J 24, 4061–4070.[CrossRef][Medline]
Toyoda
T, Hayashi
T, Miyamoto
L, Yonemitsu
S, Nakano
M, Tanaka
S, Ebihara
K, Masuzaki
H, Hosoda
K, Inoue
G, Otaka
A, Sato
K, Fushiki
T
&
Nakao
K (2004). Possible involvement of the alpha1 isoform of 5'AMP-activated protein kinase in oxidative stress-stimulated glucose transport in skeletal muscle. Am J Physiol Endocrinol Metab
287, E166–E173.
Winder WW & Hardie DG (1996). Inactivation of acetyl-CoA carboxylase and activation of AMP-activated protein kinase in muscle during exercise. Am J Physiol 270, E299–E304.[Medline]
Winder WW & Hardie DG (1999). AMP-activated protein kinase, a metabolic master switch: possible roles in type 2 diabetes. Am J Physiol 277, E1–E10.[Medline]
Wojtaszewski JF, Laustsen JL, Derave W & Richter EA (1998). Hypoxia and contractions do not utilize the same signaling mechanism in stimulating skeletal muscle glucose transport. Biochim Biophys Acta 1380, 396–404.[Medline]
Woods A, Johnstone SR, Dickerson K, Leiper FC, Fryer LG, Neumann D, Schlattner U, Wallimann T, Carlson M & Carling D (2003). LKB1 is the upstream kinase in the AMP-activated protein kinase cascade. Curr Biol 13, 2004–2008.[CrossRef][Medline]
Wright
VP, Klawitter
PF, Iscru
DF, Merola
AJ
&
Clanton
TL (2005). Superoxide scavengers augment contractile but not energetic responses to hypoxia in rat diaphragm. J Appl Physiol
98, 1753–1760.
Xie
Z, Kometiani
P, Liu
J, Li
J, Shapiro
JI
&
Askari
A (1999). Intracellular reactive oxygen species mediate the linkage of Na+/K+-ATPase to hypertrophy and its marker genes in cardiac myocytes. J Biol Chem
274, 19323–19328.
Young ME, Radda GK & Leighton B (1996). Activation of glycogen phosphorylase and glycogenolysis in rat skeletal muscle by AICAR – an activator of AMP-activated protein kinase. FEBS Lett 382, 43–47.[CrossRef][Medline]
Zhang
SJ, Bruton
JD, Katz
A
&
Westerblad
H (2006). Limited oxygen diffusion accelerates fatigue development in mouse skeletal muscle. J Physiol
572, 551–559.
Zuo
L
&
Clanton
TL (2005). Reactive oxygen species formation in the transition to hypoxia in skeletal muscle. Am J Physiol Cell Physiol
289, C207–C216.
|
Acknowledgements |
|---|
This article has been cited by other articles:
|
|
 |
|
  Y. Higaki, T. Mikami, N. Fujii, M. F. Hirshman, K. Koyama, T. Seino, K. Tanaka, and L. J. Goodyear Oxidative stress stimulates skeletal muscle glucose uptake through a phosphatidylinositol 3-kinase-dependent pathway Am J Physiol Endocrinol Metab, May 1, 2008; 294(5): E889 - E897. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 N. P. Whitehead, C. Pham, O. L. Gervasio, and D. G. Allen N-Acetylcysteine ameliorates skeletal muscle pathophysiology in mdx mice J. Physiol., April 1, 2008; 586(7): 2003 - 2014. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. D. Bruton, N. Place, T. Yamada, J. P. Silva, F. H. Andrade, A. J. Dahlstedt, S.-J. Zhang, A. Katz, N.-G. Larsson, and H. Westerblad Reactive oxygen species and fatigue-induced prolonged low-frequency force depression in skeletal muscle fibres of rats, mice and SOD2 overexpressing mice J. Physiol., January 1, 2008; 586(1): 175 - 184. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. M. Ross, G. D. Wadley, M. G. Clark, S. Rattigan, and G. K. McConell Local Nitric Oxide Synthase Inhibition Reduces Skeletal Muscle Glucose Uptake but Not Capillary Blood Flow During In Situ Muscle Contraction in Rats Diabetes, December 1, 2007; 56(12): 2885 - 2892. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S.-J. Zhang, M. E. Sandstrom, J. T. Lanner, A. Thorell, H. Westerblad, and A. Katz Activation of aconitase in mouse fast-twitch skeletal muscle during contraction-mediated oxidative stress Am J Physiol Cell Physiol, September 1, 2007; 293(3): C1154 - C1159. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. N. Edwards, W. A. Macdonald, C. van der Poel, and D. G. Stephenson O2bullet production at 37{degrees}C plays a critical role in depressing tetanic force of isolated rat and mouse skeletal muscle Am J Physiol Cell Physiol, August 1, 2007; 293(2): C650 - C660. |
, control; •, NAC). Initial force averaged 20.8 ± 1.8 mN (mg wet weight)–1 for control conditions and 23.0 ± 1.4 mN (mg wet weight)–1 for NAC; n.s.).
