AMP kinase activation with AICAR simultaneously increases fatty acid and glucose oxidation in resting rat soleus muscle
1 Department of Human Biology and Nutritional Sciences, University of Guelph, Guelph, Ontario, Canada
Abstract
5-Amino-4-imidazolecarboxamide riboside (AICAR), a pharmacological activator of AMP-activated protein kinase (AMPK), acutely stimulates glucose uptake and fatty acid (FA) oxidation in skeletal muscle. However, it is not fully understood whether AICAR-induced changes in glucose oxidation are secondary to changes in FA oxidation (i.e. glucose fatty acid cycle), or what role AMPK may be playing in the regulation of intramuscular triacylglycerol (TAG) esterification and hydrolysis. We examined the acute (60 min) effects of AICAR (2 mM) on FA metabolism, glucose oxidation and pyruvate dehydrogenase (PDH) activation in isolated resting rat soleus muscle strips exposed to two different FA concentrations (low fatty acid, LFA, 0.2 mM; high fatty acid, HFA, 1 mM). AICAR significantly increased AMPK 2 activity (+192%; P < 0.05) over 60 min, and simultaneously increased both FA (LFA: +33%, P < 0.05; HFA: +36%, P < 0.05) and glucose (LFA: +105%, P < 0.05; HFA: +170, P < 0.001) oxidation regardless of FA availability. While there were no changes in TAG esterification, AICAR did increase the ratio of FA partitioned to oxidation relative to TAG esterification (LFA: +15%, P < 0.05; HFA: +49%, P < 0.05). AICAR had no effect on endogenous TAG hydrolysis and oxidation in resting soleus. The stimulation of glucose oxidation with AICAR was associated with an increase in PDH activation (+126%; P < 0.05) but was without effect on pyruvate, an allosteric activator of the PDH complex, suggesting that AMPK may stimulate PDH directly. In conclusion, AMPK appears to be an important regulator of both FA metabolism and glucose oxidation in resting skeletal muscle.
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Introduction
AMP-activated protein kinase (AMPK) is an important energy sensor within skeletal muscle. AMPK is activated allosterically by an increase in the AMP: ATP and creatine (Cr): phosphocreatine (PCr) ratio, as well as covalently by AMPK kinase (Hardie et al. 1999). AMPK is activated by muscle contraction (Hayashi et al. 1998; Ihlemann et al. 2000; Ai et al. 2002), and pharmacologically by 5-amino-4-imidazolecarboxamide riboside (AICAR). AICAR is a cell-permeable compound which is phosphorylated to form ZMP, and mimics the effects of AMP on the AMPK signalling cascade. The metabolic effects of AICAR include an increase in glucose uptake (Merrill et al. 1997; Hayashi et al. 1998; Bergeron et al. 1999; Musi et al. 2001), and a repartitioning of fatty acids (FA) toward oxidation (Merrill et al. 1997; Alam & Saggerson, 1998; Muoio et al. 1999; Kaushik et al. 2001) and away from intramuscular triacylglycerol (TAG) esterification (Muoio et al. 1999). Thus, pharmacological activation of AMPK appears to result in an increase in ATP-producing pathways, as is also observed during contraction.
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Regulation of TAG hydrolysis in skeletal muscle is poorly understood. AICAR-stimulated AMPK activation would be expected to stimulate TAG hydrolysis and subsequent oxidation for energy production. Paradoxically, AICAR treatment has been shown to have an anti-lipolytic effect in rat soleus muscle (Alam & Saggerson, 1998) and C2C12 myotubes (Muoio et al. 1999). In adipose tissue, AMPK phosphorylates hormone-sensitive lipase, having no effect on the activity of the enzyme in its basal state, but inhibits further phosphorylation by protein kinase A (Garton et al. 1989), preventing isoprenaline-induced lipid hydrolysis (Sullivan et al. 1994; Corton et al. 1995).
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Another area of controversy is the effect of AICAR on glucose oxidation in skeletal muscle. Previous studies have measured AICAR's ability to increase glucose uptake (Merrill et al. 1997; Hayashi et al. 1998; Bergeron et al. 1999; Musi et al. 2001) and increase glycogen synthesis (Aschenbach et al. 2002). However, few studies have specifically examined the effect of AICAR on glucose oxidation. In isolated human endothelial cells, AICAR simultaneously increased FA oxidation and glucose oxidation, leading to an increase in ATP production (Dagher et al. 1999). The only study using isolated skeletal muscle showed that 60 min of AICAR exposure in oxidative rat soleus muscle decreased glucose oxidation secondary to an increase in FA oxidation (i.e. glucose–fatty acid cycle) (Kaushik et al. 2001). However, teleologically, one would expect that, in response to a decrease in cellular energy charge, AMPK activation would simultaneously increase ATP production from both FA and glucose oxidation.
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Therefore, in the present study we used palmitate and glucose tracers to examine AICAR's acute effects on FA and glucose metabolism, as well as pyruvate dehydrogenase activation (PDHa), a key regulatory site of glucose oxidation. We used high fatty acid (HFA, 1 mM) and low fatty acid (LFA, 0.2 mM) buffers to determine whether AICAR-induced changes in glucose oxidation were dependent on changes in FA oxidation. Thus, an AICAR-induced decrease in glucose oxidation that is secondary to a stimulation of FA oxidation should occur with adequate FA availability, but not lower FA levels, when rates of oxidation are low. Furthermore, we also wished to confirm that AICAR-induced AMPK stimulation results in a reduction in TAG hydrolysis. Specifically, we also examined whether any AICAR-induced reduction in hydrolysis was dependent on FA availability, as increased FA availability is known to reduce lipolysis (Dyck & Bonen, 1998), potentially due to AMPK activation.
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Methods
Animals and preparation of muscle strips
Female Sprague-Dawley rats (Charles River Laboratory, QC, Canada; weight: 215 ± 2 g) were used for all experiments. Animals were housed in a controlled environment on a 12 h: 12 h reversed light–dark cycle and fed Purina rat chow and water ad libitum. All procedures were approved by the Animal Care Committee at the University of Guelph. Animals were anaesthetized with an intraperitoneal injection of pentobarbital sodium (6 mg (100 g body mass)–1) prior to all experimental procedures. Longitudinal soleus muscle strips were carefully dissected with tendons intact using a 27-gauge needle. Each strip was sutured, removed and suspended on brass hooks in a 7 ml incubation reservoir in order to maintain resting tension. Seven millilitres of warmed (30°C), pre-gassed (95% O2–5% CO2) modified Krebs-Henseleit buffer (KHB) containing 4% FA-free bovine serum albumin (Boehringer, QC, Canada), 10 mM glucose and 1 mM palmitate was immediately added to the incubation reservoir. This was the base buffer for all experiments and was maintained at 30°C, and continuously gassed during all stages except the ‘Chase phase’ (see below). A layer of heavy mineral oil was placed on top of the incubation buffer at all stages of the procedure in order to maintain gassing pressures. At the termination of the dissection procedure, the rats were humanely killed with an intracardiac injection of pentobarbital sodium. Muscle viability was assessed by measuring phosphagen content (ATP, PCr and Cr) in a separate set of experiments.
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Time course for AMPK activation
Muscle incubations. AMPK 1 and 2 activities were examined after 30, 40 and 60 min of AICAR treatment (Fig. 1A). Briefly, muscle strips were incubated in KHB with or without the addition of 2 mM AICAR (Toronto Research Chemicals, Toronto, ON, Canada). At the end of the incubation period, the muscle strips were quickly freeze-clamped and stored in liquid nitrogen until further analysis.
For time course experiments (A) soleus strips were incubated in the presence or absence of AICAR (2 mM) and muscle strips were freezeclamped at the time points indicated for further analysis of AMPK 1 and 2 activities, PDHa and pyruvate concentration. For FA metabolism (B), rat soleus muscle strips were incubated (30°C) in modified Krebs-Henseleit buffer (KHB) through four stages: preincubation (PREINC), pulse ([9,10-3H]palmitate), wash and chase ([1-14C]palmitate) to monitor endogenous and exogenous FA metabolism. During wash and chase (last 60 min), soleus strips were incubated in the presence or absence of AICAR (2 mM), in KHB with low fatty acid (LFA, 0.2 mM) or high fatty acid (1 mM) concentration. For glucose oxidation (C), soleus strips were incubated in the presence or absence of AICAR (2 mM), in KHB with low fatty acid (LFA, 0.2 mM) or high fatty acid (1 mM) concentration throughout the protocol (60 min) and glucose oxidation was monitored during the last 30 min.
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Immunoprecipitation and AMPK activity. Muscle strips (25–30 mg) were homogenized in buffer (50 mM Tris-HCl, pH 7.5, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, 50 mM NaF, 5 mM sodium pyrophosphate, 10% glycerol, 1% Triton X-100, 10 μg ml–1 trypsin inhibitor, 2 μg ml–1 aprotinin, 1 mM benzamidine, 1 mM phenylmethylsulphonyl fluoride). The homogenates were incubated with AMPK 1 and 2 (Upstate, Charlottesville, VA, USA) antibody-bound protein A beads (Sigma, St Louis, MO, USA), each for 2 h at 4°C. Immunocomplexes were washed with PBS and suspended in 60 μl dilution buffer (50 mM Tris (pH 7.5), 1 mM dithiothreitol (DTT), 10% glycerol, 0.1% Trition-X) for AMPK activity assay (Chen et al. 2000). Briefly, 20 μl of sample was combined with 20 μl of reagent mixture (5 mM Hepes (pH 7.5), 1 mM MgCl2, 0.5% glycerol, 1 mM DTT, 100 μM SAMS peptide (Upstate Scientific), 250 μM ATP with [-32P]ATP (Amersham Biosciences, QC, Canada), 100 μM AMP). The reaction proceeded for 15 min, after which 23 μl of reaction mixture was spotted onto p81 filter paper (Upstate Scientific) and washed three times in 1% phosphoric acid. Filter papers were dried and placed in organic scintillant for counting.
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Lipid metabolism (‘pulse–chase’ experiments) (Fig. 1B)
Pulse and wash. After an initial preincubation period (30 min), the buffer was drained from the reservoir and 7 ml KHB with 2 μCi [9,10-3H]palmitate (Amersham Biosciences, QC, Canada) was added to the reservoir. Strips were pulsed for 30 min to prelabel the endogenous lipid pools (intramuscular diacylglycerol (DAG) and TAG). After the pulse phase, the buffer was drained and the muscles were washed for 30 min in the absence of radiolabelled palmitate to allow for the removal of non-incorporated [3H]palmitate. During the wash, the buffer either remained at 1 mM palmitate (HFA) or was decreased to 0.2 mM palmitate (LFA). During this period, some strips were exposed to 2 mM AICAR to ensure sufficient time for AICAR to diffuse into the muscle and activate AMPK prior to the chase phase. At the end of the wash phase, one strip from each pair was removed and extracted for endogenous lipids to determine prelabelling.
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Chase phase (‘experimental phase’). The remaining muscle strips continued to be incubated for 30 min in pre-gassed, modified KHB containing 0.5 μCi [1-14C]palmitate (Amersham Biosciences, QC, Canada). Strips were incubated in the presence or absence of 2 mM AICAR, either at HFA or LFA. During this phase the gas was turned off to prevent the escape of 14CO2. Exogenous palmitate oxidation and esterification were monitored by the production of 14CO2 and incorporation of [1-14C]palmitate into intramuscular lipids, respectively. Intramuscular lipid hydrolysis and oxidation were monitored simultaneously by measuring the net change in lipid [3H]palmitate content and the production of 3H2O, respectively.
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Palmitate oxidation. Exogenous [1-14C]palmitate oxidation was determined as outlined previously (Dyck et al. 2000) with minor modifications. At the end of the chase phase, a 3.5 ml aliquot of buffer was transferred to a 50 ml Erlenmeyer flask, which was quickly sealed with a rubber stopper fitted with a stopcock and needle. The buffer was acidified with 3 ml of 1 M H2SO4 and the 14CO2 was captured over 120 min in a 0.5 ml microcentrifuge tube containing 400 μl of 1 M benzethonium hydroxide (Sigma, Oakville, ON, Canada) suspended from the stopper. The microcentrifuge tube was placed in a scintillation vial and counted using standard liquid scintillation techniques. Loss of 14C through isotopic exchange at the level of the Krebs cycle was accounted for by collecting a 0.5 ml aliquot of the aqueous phase produced during the muscle lipid extraction as previously described (Dyck et al. 2000). Endogenous [9,10-3H]palmitate oxidation was quantified by measuring 3H2O production. Two millilitres of the chase incubation buffer was transferred to a 13 ml plastic centrifuge tube containing 5 ml of 2: 1 chloroform–methanol (v/v). Samples were shaken for 15 min before adding 2 ml of 2 M KCl–HCl and were shaken for an additional 15 min. The samples were then centrifuged at 5000 g, 4°C for 10 min. A 1 ml aliquot was removed from the upper aqueous phase and quantified by liquid scintillation counting.
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Extraction of muscle lipids. After incubation, the muscles were removed, blotted and weighed, and placed in 13 ml plastic centrifuge tubes containing 5 ml ice-cold 2: 1 chloroform–methanol (v/v) and homogenized using a Polytron homogeniser. Connective tissue was weighed and subtracted from the initial wet weight of the muscle. Samples were then centrifuged at 5000 g, 4°C for 10 min. The supernatant was removed and transferred to a clean centrifuge tube. Distilled water (2 ml) was added and the samples were shaken for 10 min and centrifuged at 5000 g, 4°C for 10 min to separate the aqueous and lipophilic phases.
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The chloroform phase was transferred to glass centrifuge tubes and gently evaporated under a stream of N2. The samples were redissolved in 100 μl 2: 1 chloroform–methanol containing internal standards of dipalmitin and tripalmitin (Sigma, Oakville, ON, Canada) to ensure proper lipid identification. Fifty microlitres of each sample was spotted on a silica gel plate that had been oven-dried overnight. Silica gel plates were placed in a sealed tank containing solvent (60: 40: 3 heptane–isopropyl ether–acetic acid) for 60 min. Plates were then removed, permitted to dry and sprayed with dichlorofluorescein dye (0.02% w/v in ethanol) and visualized using long-wave ultraviolet light. The DAG and TAG bands were scraped into scintillation vials for counting.
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Glucose oxidation
Glucose oxidation (Fig. 1C) was determined in a separate set of experiments. The preincubation period utilized 7 ml KHB (LFA or HFA, 10 mM glucose) as described in the pulse–chase experiments, either in the presence or absence of 2 mM AICAR. After the 30 min preincubation, the buffer was drained and 7 ml of pre-gassed, modified KHB containing 2 μCi of [U-14C]glucose was added to the reservoir. The muscle strips were incubated for an additional 30 min in the presence or absence of 2 mM AICAR. Gaseous 14CO2 was captured as previously described.
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Time course for PDH activation and pyruvate content
In a separate series of experiments, we investigated the effect of AICAR on PDH activation after 30, 40 and 60 min by incubating muscle strips in the absence or presence of AICAR (Fig. 1A). Freeze-clamped soleus (10–15 mg) was used for the determination of PDHa, as previously described (Putman et al. 1993). The remainder of the muscle was freeze-dried, powdered and dissected free of all visible blood and connective tissue, and analysed for pyruvate concentration. Briefly, freeze-dried muscle (5 mg) was extracted with 0.5 M perchloric acid containing 1 mM EDTA and neutralized with 2.2 M KHCO3. Pyruvate was analysed fluorometrically (Passoneau & Lowry, 1993).
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Calculations and statistics
AMPK activity was expressed as picomoles 32P transferred to SAMS peptide per minute per milligram of protein. To calculate palmitate (nmol (g wet weight)–1) oxidized or incorporated into lipid pools, the specific activity of the incubation buffer (d.p.m. radiolabelled palmitate (nmol total palmitate)–1) was used. Intramuscular TAG hydrolysis was calculated as the net loss of preloaded [3H]palmitate (nmol g–1) from lipid pools (DAG and TAG) between paired soleus strips. Glucose oxidation was calculated with the specific activity of labelled glucose in KHB in the same manner as palmitate.
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Results are presented as mean ± S.E.M. One-way ANOVA tests were used to analyse muscle viability data over the 2 h incubation. Two-way ANOVA followed by Student-Neulman-Keuls post hoc analyses were used to assess statistical significance between time points for AMPK activity, with and without AICAR. Student's unpaired t tests were used to analyse the effect of AICAR on FA and glucose metabolism. Two-way repeated measures ANOVA followed by Student-Neulman-Keuls post hoc analyses were used to assess statistical significance between time points for PDHa and pyruvate, with and without AICAR. Significance was accepted at P 0.05.
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Results
Viability of incubated muscle
There were no significant differences in ATP, PCr and Cr (Table 1), and most importantly, all measured parameters remained stable after the initial 30 min when measurements of glucose oxidation and lipid metabolism were made.
Time course for AMPK activation with AICAR
AICAR treatment had no effect on AMPK 1 activity (Fig. 2A). However, over 60 min, AICAR resulted in significant increases in AMPK 2 activity (Fig. 2B). By 60 min of incubation with AICAR, AMPK 2 activity in soleus strips was higher (+192%; P < 0.05) than strips not treated with AICAR.
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Values are means ± S.E.M., pmol min–1 (mg protein)–1, n = 5–8 per group. *Significantly different from 30 min of same condition (P < 0.05). Treatment effect of AICAR significantly different from No AICAR (P < 0.05).
Effects of AICAR on FA metabolism
Exogenous FA metabolism. AICAR treatment significantly increased exogenous FA oxidation in both LFA (+33%; P < 0.05; Fig. 3A) and HFA conditions (+36%; P < 0.05; Fig. 3B). Although incorporation into TAG was not significantly different with AICAR (Fig. 3C and D), there was an increase in the amount of FA partitioned toward oxidation relative to TAG esterification at both LFA (+15%; P < 0.05; Fig. 3E) and HFA (+49%; P < 0.05; Fig. 3F). There were no significant differences in palmitate incorporation into DAG with AICAR (Table 2).
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Effect of AICAR on fatty acid oxidation in low fatty acid (0.2 mM, LFA: A) or high fatty acid (1 mM, HFA: B) modified KHB, on TAG esterification (LFA: C; HFA: D) and on oxidation: TAG esterification ratio (LFA: E; HFA: F). Values are means ± S.E.M., nmol (g wet wt)–1, n = 7 per group. *Significantly different from No AICAR (P < 0.05).
Endogenous FA metabolism. There were no significant effects on DAG or TAG hydrolysis with AICAR treatment in the LFA condition. Correspondingly, there was also no effect on endogenous oxidation with AICAR. These results were not different at HFA with AICAR treatment (Table 2).
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Effects of AICAR on glucose oxidation
AICAR increased glucose oxidation, both in the LFA (+105%; P < 0.05; Fig. 4A) and HFA conditions (+170%; P < 0.0001; Fig. 4B).
Effect of AICAR on glucose oxidation in low fatty acid (0.2 mM: A) or high fatty acid (1 mM: B) modified KHB. Values are means ± S.E.M., nmol (g wet wt)–1, n =6–8 per group. *Significantly different from No AICAR (P < 0.05).
Changes in calculated ATP production with AICAR
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Calculated total ATP production (Table 3) increased with AICAR both with LFA (+89%; P < 0.05) and HFA (+131%; P < 0.001) as a result of increased ATP production from glucose (LFA: +104%; P < 0.05; HFA: +165%; P < 0.001) and FA oxidation (LFA: +17%; P < 0.05; HFA: +30%; P < 0.05).
Time course for PDHa and pyruvate content with AICAR
Over 60 min, AICAR treatment resulted in significant increases in PDHa (Fig. 5A). By 60 min of incubation with AICAR, PDHa was higher than in strips not treated with AICAR (+71%; P < 0.005), and also significantly different from AICAR treatment after 30 min (+126%; P < 0.05). Muscle pyruvate was not significantly different during 60 min of AICAR treatment (Fig. 5B).
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Time course for pyruvate dehydrogenase activation (PDHa, A) and pyruvate content (B) in the presence or absence of AICAR. Values are means ± S.E.M., n = 6–8 per group. *Significantly different from 30 min of same condition (P < 0.05); **Significantly different from 30 min of different condition (P < 0.05). Trial effect of AICAR being significantly different from No AICAR (P < 0.05).
Discussion
With the use of palmitate and glucose tracers, we were able to directly examine the acute effects of AICAR on FA and glucose metabolism in isolated rodent soleus muscle. We confirmed previous results that in resting muscle AICAR activates AMPK 2 activity (Jessen et al. 2003; Raney et al. 2005) leading to stimulation of FA oxidation (Merrill et al. 1997; Muoio et al. 1999; Raney et al. 2005). In addition, several novel observations were made in this investigation: (1) AICAR increased glucose oxidation regardless of the level of FA available to the muscle, indicating an increase in total energy provision; (2) PDHa increased during 60 min of AICAR treatment, supporting the observed increase in glucose oxidation; however, this was not due to an increase in pyruvate; and (3) AICAR had no independent effects on resting endogenous FA metabolism (TAG hydrolysis, oxidation).
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Effect of AICAR on skeletal muscle FA metabolism
Exogenous oxidation. In agreement with previous studies (Merrill et al. 1997; Muoio et al. 1999; Kaushik et al. 2001; Raney et al. 2005), we show that AICAR increased FA oxidation 40%, demonstrating that AMPK is a key regulator of FA oxidation in soleus. Indeed, 60 min of AICAR administration activated AMPK 2 2-fold, which has been previously demonstrated in this muscle (1.5- to 3-fold; Wojtaszewski et al. 2002; Jessen et al. 2003). Increases in FA oxidation are presumably due to the downstream effects of AMPK activation, including inhibition of acetyl coenzyme A (acetyl-CoA) carboxylase (Winder et al. 1997), increases in malonyl-CoA decarboxylase (Saha et al. 2000), ultimately decreasing malonyl-CoA levels (Alam & Saggerson, 1998) with subsequent relief of the inhibition on carnitine palmitoyl transferase I, a rate-limiting step in FA oxidation in muscle.
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TAG esterification and endogenous hydrolysis. The regulation of intramuscular lipid metabolism, both at rest and during contraction, is poorly understood. To our surprise, we failed to see a significant decrease in TAG esterification with AICAR in both fat conditions. These findings are in direct contrast to those of Muoio et al. (1999), who demonstrated a decrease in TAG esterification in mouse soleus. Mitochondrial sn-glycerol-3-phosphate acyltransferase (GPAT) is a rate-limiting step in TAG esterification and has been suggested to be the likely mechanism by which AMPK may be regulating TAG esterification (Muoio et al. 1999). However, the degree by which AMPK-mediated mitochondrial GPAT activity plays a role in altering TAG esterification is a matter of debate. Indeed, regulation of mitochondrial GPAT in skeletal muscle has been difficult to determine (Park et al. 2002; Watt et al. 2004) and the current study does not support an AMPK-mediated effect on GPAT for TAG esterification. Differences in TAG esterification may be due to the time course of exposure to AICAR (60 min (present study) versus 180 min (Muoio et al. 1999)), or a species difference. Murine muscle has a higher metabolic rate, is highly oxidative (Muoio et al. 1999) and may respond more quickly to AICAR. However, even with no changes observed in TAG esterification, we did observe an increase in the ratio of FA oxidized relative to TAG esterification, which has been suggested to play a role in increased insulin sensitivity in skeletal muscle (Perdomo et al. 2004).
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Intuitively, one would expect that AICAR-induced AMPK activation would stimulate TAG hydrolysis and subsequent oxidation in muscle to provide energy for ATP production, similar to that observed during contraction (Dyck & Bonen, 1998). Regulation of intramuscular TAG hydrolysis is poorly understood in skeletal muscle. Hormone-sensitive lipase, a key enzyme involved in TAG hydrolysis, has been identified in muscle (Langfort et al. 1998; Peters et al. 1998) and is stimulated by adrenaline (epinephrine) and contraction by at least partially distinct mechanisms (Langfort et al. 2003). However, TAG hydrolysis is inhibited by AICAR in soleus (Alam & Saggerson, 1998) and in C2C12 myotubes (Muoio et al. 1999), demonstrating a similar anti-lipolytic effect as observed in adipocytes (Sullivan et al. 1994; Corton et al. 1995). Therefore, we expected to observe a similar anti-lipolytic effect with AICAR in resting soleus. However, AICAR had no effect on resting TAG hydrolysis rates, probably because these were already very low. To our surprise, reducing the FA substrate was not a strong enough stimulus to drive TAG hydrolysis to investigate the potential AICAR mediation of this process. It is also possible that the duration of this protocol (AICAR treatment over 60 min) may have been too short to observe any differences in lipolytic rate with AICAR. It should also be noted that with HFA, there was a slight mismatch between TAG hydrolysis and endogenous oxidation. This discrepancy may be due to possible reincorporation (recycling) of a small amount of 3H tracer which may ultimately underestimate TAG hydrolysis, and also because calculated endogenous oxidation may be overestimated, as it is based on the specific activity of the TAG pool. In addition, the calculation of TAG hydrolysis is derived from paired muscle strips from one soleus muscle, which could increase variability. Nonetheless, AICAR had no effect on endogenous FA oxidation in resting soleus, which may be the more accurate measure of endogenous metabolism, as this is measured in a single muscle strip.
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Effects of AICAR on skeletal muscle glucose metabolism
To our knowledge, this is the first study in isolated rat skeletal muscle to show that glucose oxidation is increased in the presence of AICAR and that this effect is independent of the increase in FA availability and oxidation. In fact, we observe even higher rates of glucose oxidation at HFA than at LFA in the presence of AICAR, results that argue against a glucose–fatty acid cycle phenomenon and reinforce the idea that activation of FA oxidation through AMPK does not lead to concomitant decreases in glucose oxidation. Previous studies have shown conflicting results for the role that AICAR-stimulated AMPK activation plays in regulating glucose oxidation. It has been proposed that increased FA oxidation may inhibit the uptake and subsequent oxidation of glucose in resting muscle (i.e. the glucose–fatty acid cycle). Dagher et al. (1999) subjected isolated human endothelial cells to AICAR and found that AMPK activity was increased 5-fold by 30 min and remained elevated for the duration of the 2 h incubation period, which correlated with a significant increase in glucose oxidation. However, Kaushik et al. (2001) found that a 60 min incubation with AICAR decreased glucose oxidation by 44% in soleus muscle isolated from fed but not from fasted rats. In this study, FA oxidation increased 90% with AICAR, suggesting that the AICAR-inhibited glucose oxidation was secondary to an increase in FA oxidation (Kaushik et al. 2001). Surprisingly, increases in AMPK activity could not be demonstrated or linked to changes in FA and glucose metabolism over 60 min (Kaushik et al. 2001). However, in the present study there were significant increases in AMPK 2 activity and we did not observe a concomitant decrease in glucose oxidation secondary to the increase in FA oxidation. Thus, the current data indicate that due to increases in both exogenous FA and glucose substrate oxidation, there is an overall increase in ATP provision at rest with AICAR-mediated AMPK activation.
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The resulting increase in PDHa over 60 min of AICAR exposure supports AMPK as being a stimulator of glucose oxidation. However, we cannot establish an allosteric regulatory mechanism for the increase in PDHa, as pyruvate content was not different in the presence of AICAR. Pyruvate is both a substrate and a negative allosteric inhibitor of PDH kinase, which phosphorylates and inactivates PDH (Spriet & Heigenhauser, 2002). We did not observe any changes in the phosphagens involved in the regulation of AMPK (ATP, PCr, Cr) and due to the fact that AICAR is phosphorylated to ZMP, there are no expected changes in endogenous adenine nucleotides (ATP, ADP, AMP; Merrill et al. 1997), which are involved in allosteric regulation of PDH kinase (Spriet & Heigenhauser, 2002). It is then tempting to speculate that the PDH complex may be a direct target of AMPK which may be interacting to regulate PDH directly or to regulate the relative activities of the kinase and phosphatase that determine the amount of PDH in its active form. However, there are no known covalent regulators of the kinase and phosphatase in skeletal muscle (Spriet & Heigenhauser, 2002).
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Summary
The results from the present study demonstrate that AMPK 2 activation by AICAR is an important mediator of FA and glucose metabolism in skeletal muscle. There were simultaneous increases in FA and glucose oxidation, independent of FA availability. Taken together, AMPK activation increases exogenous substrate oxidation to increase energy provision in skeletal muscle. Stimulation of PDHa appears to be responsible for the AMPK-activated increase in glucose oxidation, but this is not due to increases in pyruvate.
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Abstract
5-Amino-4-imidazolecarboxamide riboside (AICAR), a pharmacological activator of AMP-activated protein kinase (AMPK), acutely stimulates glucose uptake and fatty acid (FA) oxidation in skeletal muscle. However, it is not fully understood whether AICAR-induced changes in glucose oxidation are secondary to changes in FA oxidation (i.e. glucose fatty acid cycle), or what role AMPK may be playing in the regulation of intramuscular triacylglycerol (TAG) esterification and hydrolysis. We examined the acute (60 min) effects of AICAR (2 mM) on FA metabolism, glucose oxidation and pyruvate dehydrogenase (PDH) activation in isolated resting rat soleus muscle strips exposed to two different FA concentrations (low fatty acid, LFA, 0.2 mM; high fatty acid, HFA, 1 mM). AICAR significantly increased AMPK 2 activity (+192%; P < 0.05) over 60 min, and simultaneously increased both FA (LFA: +33%, P < 0.05; HFA: +36%, P < 0.05) and glucose (LFA: +105%, P < 0.05; HFA: +170, P < 0.001) oxidation regardless of FA availability. While there were no changes in TAG esterification, AICAR did increase the ratio of FA partitioned to oxidation relative to TAG esterification (LFA: +15%, P < 0.05; HFA: +49%, P < 0.05). AICAR had no effect on endogenous TAG hydrolysis and oxidation in resting soleus. The stimulation of glucose oxidation with AICAR was associated with an increase in PDH activation (+126%; P < 0.05) but was without effect on pyruvate, an allosteric activator of the PDH complex, suggesting that AMPK may stimulate PDH directly. In conclusion, AMPK appears to be an important regulator of both FA metabolism and glucose oxidation in resting skeletal muscle.
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Introduction
AMP-activated protein kinase (AMPK) is an important energy sensor within skeletal muscle. AMPK is activated allosterically by an increase in the AMP: ATP and creatine (Cr): phosphocreatine (PCr) ratio, as well as covalently by AMPK kinase (Hardie et al. 1999). AMPK is activated by muscle contraction (Hayashi et al. 1998; Ihlemann et al. 2000; Ai et al. 2002), and pharmacologically by 5-amino-4-imidazolecarboxamide riboside (AICAR). AICAR is a cell-permeable compound which is phosphorylated to form ZMP, and mimics the effects of AMP on the AMPK signalling cascade. The metabolic effects of AICAR include an increase in glucose uptake (Merrill et al. 1997; Hayashi et al. 1998; Bergeron et al. 1999; Musi et al. 2001), and a repartitioning of fatty acids (FA) toward oxidation (Merrill et al. 1997; Alam & Saggerson, 1998; Muoio et al. 1999; Kaushik et al. 2001) and away from intramuscular triacylglycerol (TAG) esterification (Muoio et al. 1999). Thus, pharmacological activation of AMPK appears to result in an increase in ATP-producing pathways, as is also observed during contraction.
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Regulation of TAG hydrolysis in skeletal muscle is poorly understood. AICAR-stimulated AMPK activation would be expected to stimulate TAG hydrolysis and subsequent oxidation for energy production. Paradoxically, AICAR treatment has been shown to have an anti-lipolytic effect in rat soleus muscle (Alam & Saggerson, 1998) and C2C12 myotubes (Muoio et al. 1999). In adipose tissue, AMPK phosphorylates hormone-sensitive lipase, having no effect on the activity of the enzyme in its basal state, but inhibits further phosphorylation by protein kinase A (Garton et al. 1989), preventing isoprenaline-induced lipid hydrolysis (Sullivan et al. 1994; Corton et al. 1995).
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Another area of controversy is the effect of AICAR on glucose oxidation in skeletal muscle. Previous studies have measured AICAR's ability to increase glucose uptake (Merrill et al. 1997; Hayashi et al. 1998; Bergeron et al. 1999; Musi et al. 2001) and increase glycogen synthesis (Aschenbach et al. 2002). However, few studies have specifically examined the effect of AICAR on glucose oxidation. In isolated human endothelial cells, AICAR simultaneously increased FA oxidation and glucose oxidation, leading to an increase in ATP production (Dagher et al. 1999). The only study using isolated skeletal muscle showed that 60 min of AICAR exposure in oxidative rat soleus muscle decreased glucose oxidation secondary to an increase in FA oxidation (i.e. glucose–fatty acid cycle) (Kaushik et al. 2001). However, teleologically, one would expect that, in response to a decrease in cellular energy charge, AMPK activation would simultaneously increase ATP production from both FA and glucose oxidation.
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Therefore, in the present study we used palmitate and glucose tracers to examine AICAR's acute effects on FA and glucose metabolism, as well as pyruvate dehydrogenase activation (PDHa), a key regulatory site of glucose oxidation. We used high fatty acid (HFA, 1 mM) and low fatty acid (LFA, 0.2 mM) buffers to determine whether AICAR-induced changes in glucose oxidation were dependent on changes in FA oxidation. Thus, an AICAR-induced decrease in glucose oxidation that is secondary to a stimulation of FA oxidation should occur with adequate FA availability, but not lower FA levels, when rates of oxidation are low. Furthermore, we also wished to confirm that AICAR-induced AMPK stimulation results in a reduction in TAG hydrolysis. Specifically, we also examined whether any AICAR-induced reduction in hydrolysis was dependent on FA availability, as increased FA availability is known to reduce lipolysis (Dyck & Bonen, 1998), potentially due to AMPK activation.
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Methods
Animals and preparation of muscle strips
Female Sprague-Dawley rats (Charles River Laboratory, QC, Canada; weight: 215 ± 2 g) were used for all experiments. Animals were housed in a controlled environment on a 12 h: 12 h reversed light–dark cycle and fed Purina rat chow and water ad libitum. All procedures were approved by the Animal Care Committee at the University of Guelph. Animals were anaesthetized with an intraperitoneal injection of pentobarbital sodium (6 mg (100 g body mass)–1) prior to all experimental procedures. Longitudinal soleus muscle strips were carefully dissected with tendons intact using a 27-gauge needle. Each strip was sutured, removed and suspended on brass hooks in a 7 ml incubation reservoir in order to maintain resting tension. Seven millilitres of warmed (30°C), pre-gassed (95% O2–5% CO2) modified Krebs-Henseleit buffer (KHB) containing 4% FA-free bovine serum albumin (Boehringer, QC, Canada), 10 mM glucose and 1 mM palmitate was immediately added to the incubation reservoir. This was the base buffer for all experiments and was maintained at 30°C, and continuously gassed during all stages except the ‘Chase phase’ (see below). A layer of heavy mineral oil was placed on top of the incubation buffer at all stages of the procedure in order to maintain gassing pressures. At the termination of the dissection procedure, the rats were humanely killed with an intracardiac injection of pentobarbital sodium. Muscle viability was assessed by measuring phosphagen content (ATP, PCr and Cr) in a separate set of experiments.
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Time course for AMPK activation
Muscle incubations. AMPK 1 and 2 activities were examined after 30, 40 and 60 min of AICAR treatment (Fig. 1A). Briefly, muscle strips were incubated in KHB with or without the addition of 2 mM AICAR (Toronto Research Chemicals, Toronto, ON, Canada). At the end of the incubation period, the muscle strips were quickly freeze-clamped and stored in liquid nitrogen until further analysis.
For time course experiments (A) soleus strips were incubated in the presence or absence of AICAR (2 mM) and muscle strips were freezeclamped at the time points indicated for further analysis of AMPK 1 and 2 activities, PDHa and pyruvate concentration. For FA metabolism (B), rat soleus muscle strips were incubated (30°C) in modified Krebs-Henseleit buffer (KHB) through four stages: preincubation (PREINC), pulse ([9,10-3H]palmitate), wash and chase ([1-14C]palmitate) to monitor endogenous and exogenous FA metabolism. During wash and chase (last 60 min), soleus strips were incubated in the presence or absence of AICAR (2 mM), in KHB with low fatty acid (LFA, 0.2 mM) or high fatty acid (1 mM) concentration. For glucose oxidation (C), soleus strips were incubated in the presence or absence of AICAR (2 mM), in KHB with low fatty acid (LFA, 0.2 mM) or high fatty acid (1 mM) concentration throughout the protocol (60 min) and glucose oxidation was monitored during the last 30 min.
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Immunoprecipitation and AMPK activity. Muscle strips (25–30 mg) were homogenized in buffer (50 mM Tris-HCl, pH 7.5, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, 50 mM NaF, 5 mM sodium pyrophosphate, 10% glycerol, 1% Triton X-100, 10 μg ml–1 trypsin inhibitor, 2 μg ml–1 aprotinin, 1 mM benzamidine, 1 mM phenylmethylsulphonyl fluoride). The homogenates were incubated with AMPK 1 and 2 (Upstate, Charlottesville, VA, USA) antibody-bound protein A beads (Sigma, St Louis, MO, USA), each for 2 h at 4°C. Immunocomplexes were washed with PBS and suspended in 60 μl dilution buffer (50 mM Tris (pH 7.5), 1 mM dithiothreitol (DTT), 10% glycerol, 0.1% Trition-X) for AMPK activity assay (Chen et al. 2000). Briefly, 20 μl of sample was combined with 20 μl of reagent mixture (5 mM Hepes (pH 7.5), 1 mM MgCl2, 0.5% glycerol, 1 mM DTT, 100 μM SAMS peptide (Upstate Scientific), 250 μM ATP with [-32P]ATP (Amersham Biosciences, QC, Canada), 100 μM AMP). The reaction proceeded for 15 min, after which 23 μl of reaction mixture was spotted onto p81 filter paper (Upstate Scientific) and washed three times in 1% phosphoric acid. Filter papers were dried and placed in organic scintillant for counting.
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Lipid metabolism (‘pulse–chase’ experiments) (Fig. 1B)
Pulse and wash. After an initial preincubation period (30 min), the buffer was drained from the reservoir and 7 ml KHB with 2 μCi [9,10-3H]palmitate (Amersham Biosciences, QC, Canada) was added to the reservoir. Strips were pulsed for 30 min to prelabel the endogenous lipid pools (intramuscular diacylglycerol (DAG) and TAG). After the pulse phase, the buffer was drained and the muscles were washed for 30 min in the absence of radiolabelled palmitate to allow for the removal of non-incorporated [3H]palmitate. During the wash, the buffer either remained at 1 mM palmitate (HFA) or was decreased to 0.2 mM palmitate (LFA). During this period, some strips were exposed to 2 mM AICAR to ensure sufficient time for AICAR to diffuse into the muscle and activate AMPK prior to the chase phase. At the end of the wash phase, one strip from each pair was removed and extracted for endogenous lipids to determine prelabelling.
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Chase phase (‘experimental phase’). The remaining muscle strips continued to be incubated for 30 min in pre-gassed, modified KHB containing 0.5 μCi [1-14C]palmitate (Amersham Biosciences, QC, Canada). Strips were incubated in the presence or absence of 2 mM AICAR, either at HFA or LFA. During this phase the gas was turned off to prevent the escape of 14CO2. Exogenous palmitate oxidation and esterification were monitored by the production of 14CO2 and incorporation of [1-14C]palmitate into intramuscular lipids, respectively. Intramuscular lipid hydrolysis and oxidation were monitored simultaneously by measuring the net change in lipid [3H]palmitate content and the production of 3H2O, respectively.
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Palmitate oxidation. Exogenous [1-14C]palmitate oxidation was determined as outlined previously (Dyck et al. 2000) with minor modifications. At the end of the chase phase, a 3.5 ml aliquot of buffer was transferred to a 50 ml Erlenmeyer flask, which was quickly sealed with a rubber stopper fitted with a stopcock and needle. The buffer was acidified with 3 ml of 1 M H2SO4 and the 14CO2 was captured over 120 min in a 0.5 ml microcentrifuge tube containing 400 μl of 1 M benzethonium hydroxide (Sigma, Oakville, ON, Canada) suspended from the stopper. The microcentrifuge tube was placed in a scintillation vial and counted using standard liquid scintillation techniques. Loss of 14C through isotopic exchange at the level of the Krebs cycle was accounted for by collecting a 0.5 ml aliquot of the aqueous phase produced during the muscle lipid extraction as previously described (Dyck et al. 2000). Endogenous [9,10-3H]palmitate oxidation was quantified by measuring 3H2O production. Two millilitres of the chase incubation buffer was transferred to a 13 ml plastic centrifuge tube containing 5 ml of 2: 1 chloroform–methanol (v/v). Samples were shaken for 15 min before adding 2 ml of 2 M KCl–HCl and were shaken for an additional 15 min. The samples were then centrifuged at 5000 g, 4°C for 10 min. A 1 ml aliquot was removed from the upper aqueous phase and quantified by liquid scintillation counting.
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Extraction of muscle lipids. After incubation, the muscles were removed, blotted and weighed, and placed in 13 ml plastic centrifuge tubes containing 5 ml ice-cold 2: 1 chloroform–methanol (v/v) and homogenized using a Polytron homogeniser. Connective tissue was weighed and subtracted from the initial wet weight of the muscle. Samples were then centrifuged at 5000 g, 4°C for 10 min. The supernatant was removed and transferred to a clean centrifuge tube. Distilled water (2 ml) was added and the samples were shaken for 10 min and centrifuged at 5000 g, 4°C for 10 min to separate the aqueous and lipophilic phases.
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The chloroform phase was transferred to glass centrifuge tubes and gently evaporated under a stream of N2. The samples were redissolved in 100 μl 2: 1 chloroform–methanol containing internal standards of dipalmitin and tripalmitin (Sigma, Oakville, ON, Canada) to ensure proper lipid identification. Fifty microlitres of each sample was spotted on a silica gel plate that had been oven-dried overnight. Silica gel plates were placed in a sealed tank containing solvent (60: 40: 3 heptane–isopropyl ether–acetic acid) for 60 min. Plates were then removed, permitted to dry and sprayed with dichlorofluorescein dye (0.02% w/v in ethanol) and visualized using long-wave ultraviolet light. The DAG and TAG bands were scraped into scintillation vials for counting.
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Glucose oxidation
Glucose oxidation (Fig. 1C) was determined in a separate set of experiments. The preincubation period utilized 7 ml KHB (LFA or HFA, 10 mM glucose) as described in the pulse–chase experiments, either in the presence or absence of 2 mM AICAR. After the 30 min preincubation, the buffer was drained and 7 ml of pre-gassed, modified KHB containing 2 μCi of [U-14C]glucose was added to the reservoir. The muscle strips were incubated for an additional 30 min in the presence or absence of 2 mM AICAR. Gaseous 14CO2 was captured as previously described.
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Time course for PDH activation and pyruvate content
In a separate series of experiments, we investigated the effect of AICAR on PDH activation after 30, 40 and 60 min by incubating muscle strips in the absence or presence of AICAR (Fig. 1A). Freeze-clamped soleus (10–15 mg) was used for the determination of PDHa, as previously described (Putman et al. 1993). The remainder of the muscle was freeze-dried, powdered and dissected free of all visible blood and connective tissue, and analysed for pyruvate concentration. Briefly, freeze-dried muscle (5 mg) was extracted with 0.5 M perchloric acid containing 1 mM EDTA and neutralized with 2.2 M KHCO3. Pyruvate was analysed fluorometrically (Passoneau & Lowry, 1993).
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Calculations and statistics
AMPK activity was expressed as picomoles 32P transferred to SAMS peptide per minute per milligram of protein. To calculate palmitate (nmol (g wet weight)–1) oxidized or incorporated into lipid pools, the specific activity of the incubation buffer (d.p.m. radiolabelled palmitate (nmol total palmitate)–1) was used. Intramuscular TAG hydrolysis was calculated as the net loss of preloaded [3H]palmitate (nmol g–1) from lipid pools (DAG and TAG) between paired soleus strips. Glucose oxidation was calculated with the specific activity of labelled glucose in KHB in the same manner as palmitate.
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Results are presented as mean ± S.E.M. One-way ANOVA tests were used to analyse muscle viability data over the 2 h incubation. Two-way ANOVA followed by Student-Neulman-Keuls post hoc analyses were used to assess statistical significance between time points for AMPK activity, with and without AICAR. Student's unpaired t tests were used to analyse the effect of AICAR on FA and glucose metabolism. Two-way repeated measures ANOVA followed by Student-Neulman-Keuls post hoc analyses were used to assess statistical significance between time points for PDHa and pyruvate, with and without AICAR. Significance was accepted at P 0.05.
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Results
Viability of incubated muscle
There were no significant differences in ATP, PCr and Cr (Table 1), and most importantly, all measured parameters remained stable after the initial 30 min when measurements of glucose oxidation and lipid metabolism were made.
Time course for AMPK activation with AICAR
AICAR treatment had no effect on AMPK 1 activity (Fig. 2A). However, over 60 min, AICAR resulted in significant increases in AMPK 2 activity (Fig. 2B). By 60 min of incubation with AICAR, AMPK 2 activity in soleus strips was higher (+192%; P < 0.05) than strips not treated with AICAR.
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Values are means ± S.E.M., pmol min–1 (mg protein)–1, n = 5–8 per group. *Significantly different from 30 min of same condition (P < 0.05). Treatment effect of AICAR significantly different from No AICAR (P < 0.05).
Effects of AICAR on FA metabolism
Exogenous FA metabolism. AICAR treatment significantly increased exogenous FA oxidation in both LFA (+33%; P < 0.05; Fig. 3A) and HFA conditions (+36%; P < 0.05; Fig. 3B). Although incorporation into TAG was not significantly different with AICAR (Fig. 3C and D), there was an increase in the amount of FA partitioned toward oxidation relative to TAG esterification at both LFA (+15%; P < 0.05; Fig. 3E) and HFA (+49%; P < 0.05; Fig. 3F). There were no significant differences in palmitate incorporation into DAG with AICAR (Table 2).
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Effect of AICAR on fatty acid oxidation in low fatty acid (0.2 mM, LFA: A) or high fatty acid (1 mM, HFA: B) modified KHB, on TAG esterification (LFA: C; HFA: D) and on oxidation: TAG esterification ratio (LFA: E; HFA: F). Values are means ± S.E.M., nmol (g wet wt)–1, n = 7 per group. *Significantly different from No AICAR (P < 0.05).
Endogenous FA metabolism. There were no significant effects on DAG or TAG hydrolysis with AICAR treatment in the LFA condition. Correspondingly, there was also no effect on endogenous oxidation with AICAR. These results were not different at HFA with AICAR treatment (Table 2).
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Effects of AICAR on glucose oxidation
AICAR increased glucose oxidation, both in the LFA (+105%; P < 0.05; Fig. 4A) and HFA conditions (+170%; P < 0.0001; Fig. 4B).
Effect of AICAR on glucose oxidation in low fatty acid (0.2 mM: A) or high fatty acid (1 mM: B) modified KHB. Values are means ± S.E.M., nmol (g wet wt)–1, n =6–8 per group. *Significantly different from No AICAR (P < 0.05).
Changes in calculated ATP production with AICAR
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Calculated total ATP production (Table 3) increased with AICAR both with LFA (+89%; P < 0.05) and HFA (+131%; P < 0.001) as a result of increased ATP production from glucose (LFA: +104%; P < 0.05; HFA: +165%; P < 0.001) and FA oxidation (LFA: +17%; P < 0.05; HFA: +30%; P < 0.05).
Time course for PDHa and pyruvate content with AICAR
Over 60 min, AICAR treatment resulted in significant increases in PDHa (Fig. 5A). By 60 min of incubation with AICAR, PDHa was higher than in strips not treated with AICAR (+71%; P < 0.005), and also significantly different from AICAR treatment after 30 min (+126%; P < 0.05). Muscle pyruvate was not significantly different during 60 min of AICAR treatment (Fig. 5B).
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Time course for pyruvate dehydrogenase activation (PDHa, A) and pyruvate content (B) in the presence or absence of AICAR. Values are means ± S.E.M., n = 6–8 per group. *Significantly different from 30 min of same condition (P < 0.05); **Significantly different from 30 min of different condition (P < 0.05). Trial effect of AICAR being significantly different from No AICAR (P < 0.05).
Discussion
With the use of palmitate and glucose tracers, we were able to directly examine the acute effects of AICAR on FA and glucose metabolism in isolated rodent soleus muscle. We confirmed previous results that in resting muscle AICAR activates AMPK 2 activity (Jessen et al. 2003; Raney et al. 2005) leading to stimulation of FA oxidation (Merrill et al. 1997; Muoio et al. 1999; Raney et al. 2005). In addition, several novel observations were made in this investigation: (1) AICAR increased glucose oxidation regardless of the level of FA available to the muscle, indicating an increase in total energy provision; (2) PDHa increased during 60 min of AICAR treatment, supporting the observed increase in glucose oxidation; however, this was not due to an increase in pyruvate; and (3) AICAR had no independent effects on resting endogenous FA metabolism (TAG hydrolysis, oxidation).
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Effect of AICAR on skeletal muscle FA metabolism
Exogenous oxidation. In agreement with previous studies (Merrill et al. 1997; Muoio et al. 1999; Kaushik et al. 2001; Raney et al. 2005), we show that AICAR increased FA oxidation 40%, demonstrating that AMPK is a key regulator of FA oxidation in soleus. Indeed, 60 min of AICAR administration activated AMPK 2 2-fold, which has been previously demonstrated in this muscle (1.5- to 3-fold; Wojtaszewski et al. 2002; Jessen et al. 2003). Increases in FA oxidation are presumably due to the downstream effects of AMPK activation, including inhibition of acetyl coenzyme A (acetyl-CoA) carboxylase (Winder et al. 1997), increases in malonyl-CoA decarboxylase (Saha et al. 2000), ultimately decreasing malonyl-CoA levels (Alam & Saggerson, 1998) with subsequent relief of the inhibition on carnitine palmitoyl transferase I, a rate-limiting step in FA oxidation in muscle.
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TAG esterification and endogenous hydrolysis. The regulation of intramuscular lipid metabolism, both at rest and during contraction, is poorly understood. To our surprise, we failed to see a significant decrease in TAG esterification with AICAR in both fat conditions. These findings are in direct contrast to those of Muoio et al. (1999), who demonstrated a decrease in TAG esterification in mouse soleus. Mitochondrial sn-glycerol-3-phosphate acyltransferase (GPAT) is a rate-limiting step in TAG esterification and has been suggested to be the likely mechanism by which AMPK may be regulating TAG esterification (Muoio et al. 1999). However, the degree by which AMPK-mediated mitochondrial GPAT activity plays a role in altering TAG esterification is a matter of debate. Indeed, regulation of mitochondrial GPAT in skeletal muscle has been difficult to determine (Park et al. 2002; Watt et al. 2004) and the current study does not support an AMPK-mediated effect on GPAT for TAG esterification. Differences in TAG esterification may be due to the time course of exposure to AICAR (60 min (present study) versus 180 min (Muoio et al. 1999)), or a species difference. Murine muscle has a higher metabolic rate, is highly oxidative (Muoio et al. 1999) and may respond more quickly to AICAR. However, even with no changes observed in TAG esterification, we did observe an increase in the ratio of FA oxidized relative to TAG esterification, which has been suggested to play a role in increased insulin sensitivity in skeletal muscle (Perdomo et al. 2004).
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Intuitively, one would expect that AICAR-induced AMPK activation would stimulate TAG hydrolysis and subsequent oxidation in muscle to provide energy for ATP production, similar to that observed during contraction (Dyck & Bonen, 1998). Regulation of intramuscular TAG hydrolysis is poorly understood in skeletal muscle. Hormone-sensitive lipase, a key enzyme involved in TAG hydrolysis, has been identified in muscle (Langfort et al. 1998; Peters et al. 1998) and is stimulated by adrenaline (epinephrine) and contraction by at least partially distinct mechanisms (Langfort et al. 2003). However, TAG hydrolysis is inhibited by AICAR in soleus (Alam & Saggerson, 1998) and in C2C12 myotubes (Muoio et al. 1999), demonstrating a similar anti-lipolytic effect as observed in adipocytes (Sullivan et al. 1994; Corton et al. 1995). Therefore, we expected to observe a similar anti-lipolytic effect with AICAR in resting soleus. However, AICAR had no effect on resting TAG hydrolysis rates, probably because these were already very low. To our surprise, reducing the FA substrate was not a strong enough stimulus to drive TAG hydrolysis to investigate the potential AICAR mediation of this process. It is also possible that the duration of this protocol (AICAR treatment over 60 min) may have been too short to observe any differences in lipolytic rate with AICAR. It should also be noted that with HFA, there was a slight mismatch between TAG hydrolysis and endogenous oxidation. This discrepancy may be due to possible reincorporation (recycling) of a small amount of 3H tracer which may ultimately underestimate TAG hydrolysis, and also because calculated endogenous oxidation may be overestimated, as it is based on the specific activity of the TAG pool. In addition, the calculation of TAG hydrolysis is derived from paired muscle strips from one soleus muscle, which could increase variability. Nonetheless, AICAR had no effect on endogenous FA oxidation in resting soleus, which may be the more accurate measure of endogenous metabolism, as this is measured in a single muscle strip.
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Effects of AICAR on skeletal muscle glucose metabolism
To our knowledge, this is the first study in isolated rat skeletal muscle to show that glucose oxidation is increased in the presence of AICAR and that this effect is independent of the increase in FA availability and oxidation. In fact, we observe even higher rates of glucose oxidation at HFA than at LFA in the presence of AICAR, results that argue against a glucose–fatty acid cycle phenomenon and reinforce the idea that activation of FA oxidation through AMPK does not lead to concomitant decreases in glucose oxidation. Previous studies have shown conflicting results for the role that AICAR-stimulated AMPK activation plays in regulating glucose oxidation. It has been proposed that increased FA oxidation may inhibit the uptake and subsequent oxidation of glucose in resting muscle (i.e. the glucose–fatty acid cycle). Dagher et al. (1999) subjected isolated human endothelial cells to AICAR and found that AMPK activity was increased 5-fold by 30 min and remained elevated for the duration of the 2 h incubation period, which correlated with a significant increase in glucose oxidation. However, Kaushik et al. (2001) found that a 60 min incubation with AICAR decreased glucose oxidation by 44% in soleus muscle isolated from fed but not from fasted rats. In this study, FA oxidation increased 90% with AICAR, suggesting that the AICAR-inhibited glucose oxidation was secondary to an increase in FA oxidation (Kaushik et al. 2001). Surprisingly, increases in AMPK activity could not be demonstrated or linked to changes in FA and glucose metabolism over 60 min (Kaushik et al. 2001). However, in the present study there were significant increases in AMPK 2 activity and we did not observe a concomitant decrease in glucose oxidation secondary to the increase in FA oxidation. Thus, the current data indicate that due to increases in both exogenous FA and glucose substrate oxidation, there is an overall increase in ATP provision at rest with AICAR-mediated AMPK activation.
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The resulting increase in PDHa over 60 min of AICAR exposure supports AMPK as being a stimulator of glucose oxidation. However, we cannot establish an allosteric regulatory mechanism for the increase in PDHa, as pyruvate content was not different in the presence of AICAR. Pyruvate is both a substrate and a negative allosteric inhibitor of PDH kinase, which phosphorylates and inactivates PDH (Spriet & Heigenhauser, 2002). We did not observe any changes in the phosphagens involved in the regulation of AMPK (ATP, PCr, Cr) and due to the fact that AICAR is phosphorylated to ZMP, there are no expected changes in endogenous adenine nucleotides (ATP, ADP, AMP; Merrill et al. 1997), which are involved in allosteric regulation of PDH kinase (Spriet & Heigenhauser, 2002). It is then tempting to speculate that the PDH complex may be a direct target of AMPK which may be interacting to regulate PDH directly or to regulate the relative activities of the kinase and phosphatase that determine the amount of PDH in its active form. However, there are no known covalent regulators of the kinase and phosphatase in skeletal muscle (Spriet & Heigenhauser, 2002).
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Summary
The results from the present study demonstrate that AMPK 2 activation by AICAR is an important mediator of FA and glucose metabolism in skeletal muscle. There were simultaneous increases in FA and glucose oxidation, independent of FA availability. Taken together, AMPK activation increases exogenous substrate oxidation to increase energy provision in skeletal muscle. Stimulation of PDHa appears to be responsible for the AMPK-activated increase in glucose oxidation, but this is not due to increases in pyruvate.
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