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. 2006 Apr 27;574(Pt 1):139–147. doi: 10.1113/jphysiol.2006.107318

Fatty acids stimulate AMP-activated protein kinase and enhance fatty acid oxidation in L6 myotubes

Matthew J Watt 1,2, Gregory R Steinberg 2, Zhi-Ping Chen 2, Bruce E Kemp 2,3, Mark A Febbraio 1
PMCID: PMC1817791  PMID: 16644805

Abstract

We investigated the role of fatty acid availability on AMPK signalling and fatty acid oxidation in skeletal muscle. Incubating L6 skeletal muscle myotubes with palmitate (a saturated fatty acid) or linoleate (a polyunsaturated fatty acid) increased AMPK activity by 56 and 38%, respectively, compared with untreated cells. Consistent with these changes, AMPK Thr172 and acetyl-CoA carboxylase β Ser218 phosphorylation were increased in fatty acid treated cells. Pre-incubating cells with palmitate or linoleate increased subsequent fatty acid oxidation by 86 and 92%, respectively. The enhanced AMPK signalling occurred in the absence of detectable changes in free AMP and glycogen content. The activity of the upstream kinase LKB1 was decreased by fatty acid treatment indicating that AMPK activation was not a consequence of LKB1 activation. Instead, fatty acids enhanced LKB1 phosphorylation of AMPK. Fatty acids did not alter LKB1 activity when either synthetic peptide or AMPK α(1–312) catalytic fragment was used as substrate indicating that the βγ subunits were required for the fatty acid activation. Infection of cells with a dominant-negative AMPK adenovirus reduced basal fatty acid oxidation and inhibited the stimulatory effects of fatty acid pretreatment on fatty acid oxidation. These results indicate that increasing fatty acid availability increases AMPK activity independent of changes in the cellular energy charge and support the view that fatty acids may modulate AMPK allosterically, making it a better substrate for LKB1.

The Randle glucose–fatty acid cycle hypothesis states that elevating exogenous fatty acid levels increases fatty acid oxidation and suppresses glucose uptake and metabolism (Randle et al. 1963). The biochemical basis of this cycle is the finding that increasing fatty acid metabolism increases citrate and acetyl-CoA accumulation, thereby resulting in allosteric inhibition of key carbohydrate enzymes phosphofructokinase and pyruvate dehydrogenase, respectively. An unresolved question is how increased extracellular fatty acid availability leads to a ‘feed forward’ activation of fatty acid metabolism (Clark et al. 2004). A key regulatory enzyme of fuel selection is carnitine palmitoyl transferase (CPT) 1, which permits the entry of fatty acyl-CoAs into the mitochondria. CPT1 is potently inhibited by malonyl-CoA, generated by the mitochondrial associated acetyl-CoA carboxylase β (ACCβ). ACC also acts as a metabolic sensor, itself being regulated allosterically (via citrate activation and fatty acid inhibition) and by phosphorylation at Ser218 (rat), which inactivates the enzyme by increasing citrate dependence. Thus fatty acids may promote fatty acid oxidation by inactivating ACC, thereby releasing CPT1 from allosteric inhibition.

The AMP-activated protein kinase (AMPK) is an αβγ heterotrimeric enzyme that is activated by phosphorylation of Thr172 within the α-subunit activation loop by the constitutively active upstream AMPK kinase (AMPKK), LKB1 (Woods et al. 2003) and allosteric activation by increasing cellular AMP (Kemp et al. 2003). AMPK is described as a cellular ‘energy sensor’ because its activity is increased when AMP levels increase, resulting in increased catabolism and ATP regeneration. AMPK exerts an acute regulatory role on numerous metabolic processes, including fatty acid oxidation (Merrill et al. 1997).

Recent studies in heart and liver have revealed that AMPK may be sensitive to the ‘lipid status’ of the cell and activation may be influenced by intracellular fatty acid availability independent of the cellular AMP levels. Clark et al. (2004) reported AMPK Thr172 phosphorylation in rat hearts perfused with saturated fatty acids in the absence of changes in the cellular energy charge. AMPK activation also occurs in cultured rat cardiomyocytes with short-term palmitate and oleate treatment (Hickson-Bick et al. 2000). Studies in rodents show that dietary polyunsaturated fatty acids attenuate the decrease in hepatic AMPK activity observed with fat-free feeding (Suchankova et al. 2005). These data raise the possibility that fatty acids promote AMPK activation, and also support the concept of feed forward activation of fatty acid oxidation in response to increased fatty acid availability. This has not been directly tested in skeletal muscle, which is a quantitatively important tissue for free fatty acid uptake and oxidation (van Hall et al. 2002). The aim of the present study was to test the hypothesis that increasing fatty acid availability increases AMPK activation in skeletal muscle, which is essential for feed forward activation of fatty acid oxidation.

Methods

Cell culture

L6 myoblasts were maintained at 37°C (95% O2−5% CO2) in α-modified Eagle's medium (α-MEM) containing 10% fetal bovine serum (FBS) and 1% penicillin–streptomycin. Differentiation was induced by switching to medium containing 2% horse serum when the myoblasts were ∼90% confluent. Differentiated L6 myotubes were infected with adenovirus containing AdGo (Control) or a dominant negative (DN) AMPK mutant as previously described (Woods et al. 2000). Experimental treatments were started after 4 days, by which time nearly all of the myoblasts had fused to form myotubes. Cells were serum starved for 4 h prior to experiments. Stock solutions of palmitate (16 : 0) and linoleate (18 : 2) (Sigma Aldrich, Castle Hill, NSW, Australia) were made in the presence of 2% bovine serum albumin (BSA). Cells were incubated at 37°C in the presence of vehicle, 0.075 mm palmitate, 0.25 mm palmitate or 0.25 mm linoleate. After 60 min, cells were lysed for later analysis.

Fatty acid oxidation

To examine the effects of fatty acid pretreatment on fat oxidation, cells were grown in six-well plates and were incubated for 1 h as described above. Fatty acids in the medium were then removed by washing twice with phosphate buffered saline (PBS), and fresh medium (3 ml) consisting of α-MEM, 2% BSA and 14C-radiolabelled palmitate (0.5 mm, 1 μCi ml−1) (Amersham Biosciences, Buckinghamshire, UK) was added to the cells. The incubation was stopped after 1 h. Two millilitres of medium was added to an equal volume of 1 m acetic acid and the released 14CO2 was trapped in benzothonium hydroxide. Methanol was added to the plate and cells were scraped for subsequent analysis of the acid soluble metabolite (ASM) fraction. Oxidation rates (CO2 + ASMs) were determined after scintillation counting. We also assessed muscle lipid contents after 1 h pretreatment according to previously described methods (Lessard et al. 2005). The 1 h pretreatment with palmitate or linoleate did not effect L6 myotube ceramide (percentage change from untreated = 9 ± 1), diacylglycerol (−13 ± 6%) or triacylglycerol (9 ± 2%) contents. These experiments were performed in triplicate on three separate occasions.

Fatty acid esterification into cellular lipid pools

To examine the effects of fatty acid pretreatment on fatty acid esterification, cells lysed in methanol were exposed to chloroform: methanol (2 : 1) extraction. The organic phase containing triacylglycerol (TAG) and diacylglycerol (DAG) was removed and dried under vacuum. The lipids were reconstituted in chloroform and loaded onto silica gel plates, and the lipid fractions were separated by thin layer chromatography and the bands corresponding to TAG and DAG were scraped for liquid scintillation counting.

Fatty acid effects on purified LKB1 phosphorylation of AMPK

LKB1 was partially purified from bovine testes as reported for liver (Hawley et al. 2003). Recombinant GST tagged AMPK α1β1γ1 was expressed in insect cells using baculoviral driven expression and purified by glutathione affinity chromatography. Palmitate and linoleate were dissolved in ethanol as described and were diluted in 2% fat-free BSA. The LKB1 activity assay was performed as a two-step measurement. First, LKB1 was incubated with AMPK in 30 μl of buffer containing 20 mm Tris.HCl, pH 7.5; 400 μm ATP; 4 mm MgCl2; 0.1% Tween-20 and 1 mm DTT in the presence or absence of 120 μm AMP, plus fatty acid (500 μm) or vehicle (BSA alone) at 30°C for 20 min. Second, 10 μl of LKB1 reaction solution was added to the AMPK assay buffer which contained 100 μm SAMS peptide, 200 μm AMP and 250 μm ATP (500–1000 c.p.m. (pmol 32P-ATP)−1) in a total volume of 40 μl. At end of the incubation, 25 μl of the reaction buffer was spotted on P81 paper, washed in 75 mm phosphoric acid and counted in a scintillation counter (Glass et al. 1978). These experiments were also performed using bacterial expressed MBP tagged AMPKα1(1–312) fragment (Hamilton et al. 2002), instead of baculoviral AMPK α1β1γ1, as described above. LKB1 activity was assayed with NUAK2 peptide as previously described (Lizcano et al. 2004). LKB1 apparent activity was calculated as picomoles of phosphate transferred to the SAMS peptide per minute per microgram partially purified LKB1 protein.

AMPK activity assays

Briefly, lysed cells were centrifuged at 14 000 g for 25 min and the supernatant was incubated with AMPK α1 and AMPK β2 antibody-bound protein A agarose beads for 2 h. Immunocomplexes were washed and suspended in 50 mm Tris.HCl (pH 7.5) buffer for the AMPK assay in the presence of 200 μm AMP. Activities were calculated as picomoles of phosphate incorporated into the SAMS peptide per minute per milligram lysate protein subjected to immunoprecipitation (Chen et al. 1999).

LKB1 and AMPKK activity in L6 myotubes

Anti-LKB1 antibody (Upstate) prebound to Protein A beads were incubated with myotube lysates (500 μg) for 2 h at 4°C. AMPKK assays were performed on the beads using a two-step reaction with bacterially expressed human AMPK (α1β1γ1) as substrate as described (Chen et al. 2005). The AMPKK buffer contained 20 mm Tris.HCl, pH 7.5, 0.1% Tween-20, 10 mm DTT, 8 mm MgCl2 with 0.4 mm ATP, 0.12 mm AMP and AMPK (α1β1γ1) (3 μm) in 18 μl and was incubated with 12 μl of immunoprecipitated LKB1 at 30°C for 30 min. The AMPK (α1,β1,γ1) activity was then determined using the AMPK SAMS peptide assay, a 20 μl aliquot of the AMPKK reaction mixture was added to the peptide phosphorylation reaction to give a final volume of 40 μl and assayed as described above. To compare the activity of AMPKK to LKB1, AMPKK assays were performed on whole cell lysates (6 μg of protein) that were incubated before and after immunoprecipitation with LKB1 or an unrelated antibody (insulin receptor substrate-2).

Immunoblot analysis

AMPK and ACCβ expression and phosphorylation were determined from aliquots of the AMPK lysates. Protein content was determined (BCA kit, Pierce, Progen Industries, Darra, QLD, Australia) and equal amounts of protein were resolved by SDS-PAGE. Proteins were transferred to nitrocellulose membranes, blocked and immunoblotted. After incubation with horseradish peroxidase-conjugated secondary antibody (Amersham Biosciences, Castle Hill, NSW, Australia), the immunoreactive proteins were detected with enhanced chemiluminescence (Perkin Elmer, Rowville, VIC, Australia) and quantified by densitometry (ChemiDoc XRS, Bio-Rad Laboratories, Regents Park, NSW, Australia). Membranes were stripped (50 mm Tris.HCl, pH 6.8, 2% SDS, β-mercaptoethanol, 45 min at 55°C), washed and re-probed for total protein content where appropriate.

Nucleotides and metabolite contents

Cells were lysed in a volume of 0.5 m perchloric acid (1 mm EDTA) and neutralized with 2.2 m KHCO3. This extract was used for the determination of adenosine triphosphate (ATP), phosphocreatine (PCr), creatine and lactate by enzymatic fluorometric assays (Bergmeyer, 1974). Free ADP and AMP concentrations were calculated as described (Dudley et al. 1987). In separate experiments, cells were extracted for analysis of muscle glycogen content (Passoneau & Lowry, 1993).

Statistical analysis

Results are presented as the means ± standard error of the mean (s.e.m.). Data were analysed for differences by one-way analysis of variance (ANOVA) with specific differences located with a Student-Newman-Keuls post hoc test, or Student's t test for unpaired samples where appropriate. P < 0.05 was considered statistically significant.

Results

Fatty acids increase AMPK activity in L6 myotubes

To evaluate whether increasing fatty acid availability increases AMPK activity, L6 myotubes were incubated with the saturated fatty acid palmitate. Palmitate at low concentrations (0.075 mm) did not increase AMPK activity above basal levels but higher concentrations (0.25 mm) increased AMPK activity by 56 ± 11% (P < 0.05, Fig. 1A). Because palmitate treatment can result in the cellular accumulation of bioactive lipid species such as ceramide (Chavez et al. 2003), cells were also incubated in the presence of the polyunsaturated fatty acid linoleate. Linoleate treatment (0.25 mm) gave a modest increase in AMPK activity (38 ± 9% (P < 0.05) above basal). Consistent with the increased AMPK activity, phosphorylation of AMPK Thr172 (Fig. 1B) and ACCβ (Fig. 1C), a downstream AMPK target, was elevated (P < 0.05) in fatty acid treated cells. Fatty acid treatment over this time period did not affect cell viability as assessed by LDH release into the culture medium and trypan blue exclusion (data not shown).

Figure 1. Activation of AMPK in L6 myotubes with fatty acid treatment.

Figure 1

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A, AMPK activity in AMPK immunocomplexes from L6 myotubes after treatment for 1 h with the saturated fatty acid palmitate (PAL) or the polyunsaturated fatty acid linoleate (LIN). AMPK Thr172 phosphorylation (B) and ACCβ Ser219 phosphorylation (C) were determined by immunoblot analysis. D, LKB1 activity was determined after fatty acid treatment in L6 myotubes. Results are the mean (+ s.e.m.) of six independent experiments and are plotted relative to control activity. Western blot analysis is plotted as arbitrary units. *P < 0.05 versus CON.

AMPK is activated in the absence of changes in adenine nucleotide and metabolite content

Adenine nucleotides and creatine phosphate were measured in cell extracts to determine whether the increased AMPK activity was attributable to changes in the cellular energy charge. Treatment of cells with fatty acids did not alter the content of ATP, AMP or phosphocreatine (Table 1). Because AMPK can bind to glycogen (Hudson et al. 2003; Polekhina et al. 2003) and low cellular glycogen is associated with increased AMPK activity (Wojtaszewski et al. 2003; Watt et al. 2004) we determined cellular glycogen content and found no differences between treatments (data not shown).

Table 1. Effects of fatty acids on L6 myotube adenine nucleotide contents.

VEH PAL LIN
ATP (nmol (mg protein)−1) 22.4 ± 2.0 23.2 ± 1.7 21.5 ± 0.7
fAMP (pmol (mg protein)−1) 0.044 ± 0.012 0.043 ± 0.011 0.065 ± 0.011
fADP (pmol (mg protein)−1) 31.8 ± 5.0 31.8 ± 5.7 29.8 ± 4.6
Phosphocreatine (nmol (mg protein)−1) 45.2 ± 6.6 51.0 ± 2.8 46.5 ± 4.2
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L6 myotubes were incubated in the presence of BSA (VEH), 0.25 mm palmitate (PAL) or 0.25 mm linoleate (LIN) for 1 h. Cells were lysed in 1 mm PCA for later analysis by enzymatic fluorometric methods. Values are means ± s.e.m. (n = 5). f, free.

LKB1 activity is decreased by fatty acids

The activity of the upstream AMPK kinase, LKB1, was assessed in L6 myotubes. LKB1 activity was decreased by 50% following palmitate and linoleate treatment (Fig. 1D). Immunoprecipitation, with anti-LKB1 antibody removed all AMPKK activity present in cell lysates, indicating that LKB1 is the principal AMPKK regulating AMPK activity in L6 myotubes.

Fatty acids make AMPK a better substrate for LKB1

We next determined whether fatty acids affect the ability of LKB1 to phosphorylate AMPK holoenzyme. AMPK activity was increased 4-fold in the presence of LKB1 and the addition of either 0.50 mm palmitate or linoleate further increased AMPK activity (Fig. 2A). No activation was observed at 0.05 and 0.15 mm (data not shown), which is consistent with the lack of stimulation on AMPK activity with 0.075 mm palmitate in L6 myotubes. Similar results were obtained in the presence of AMP (Fig. 2B). To determine whether the fatty acid stimulatory effects on AMPK activity require all three subunits, we repeated the aforementioned experiments using the purified MBP tagged AMPK α1(1–312) catalytic fragment. The α1(1–312) catalytic fragment is inactive in the basal state but is activated on incubation with AMPKK (Hamilton et al. 2002). There was increased AMPK activity after incubation with LKB1 and no detectable effects of fatty acids or AMP (Fig. 2C and D). These data indicate that the fatty acid effects observed in the α1β1γ1 holoenzyme are likely to be dependent on the β or γ subunits of AMPK. Furthermore, we assessed whether the observed fatty acid effects were AMPK dependent by assaying LKB1 with the NUAK2 peptide substrate and found this was not enhanced by palmitate or linoleate (Fig. 2E).

Figure 2. Fatty acids facilitate phosphorylation of AMPK by LKB1.

Figure 2

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A, purified LKB1 was incubated in the presence of recombinant AMPK α1β1γ1 without or with 500 μm palmitate (PAL) or 500 μm linoleate (LIN) and AMPK activity was assessed. B, these experiments were repeated with the addition of AMP. C and D, purified LKB1 was incubated in the presence of recombinant AMPK α1(1–312) without or with 500 μm palmitate (PAL) or 500 μm linoleate (LIN) and AMPK activity was assessed in the absence (C) or presence (D) of AMP. E, purified LKB1 was incubated in the presence of NUAK2 peptide without or with 500 μm palmitate (PAL) or 500 μm linoleate (LIN). Results are presented as means ± s.e.m. of assays conducted in duplicate and representative of three independent experiments. *P < 0.05 versus AMPK, †different versus LKB1 + AMPK.

Fatty acid oxidation

Increases in AMPK activity result in enhanced fatty acid oxidation. We assessed whether pretreating cells with fatty acids results in a ‘feed forward’ activation of fatty acid oxidation. Palmitate (0.25 mm) and linoleate (0.25 mm) pretreatment of L6 myotubes increased fatty acid oxidation above untreated cells by 86 and 92%, respectively (Fig. 3A). To determine whether the increase in fatty acid oxidation was AMPK dependent, fatty acid oxidation was assessed in L6 myotubes infected with a dominant-negative (DN) AMPK adenovirus. Fatty acid oxidation was decreased (P < 0.05) in DN-AMPK cells compared with control infected cells. Treatment of control cells with linoleate and palmitate (P = 0.07) increased subsequent fatty acid oxidation. Fatty acid oxidation was decreased in DN-AMPK cells preincubated with palmitate (P < 0.05), whereas linoleate pretreatment did not affect fatty acid oxidation in these cells. These data support the conclusion that AMPK mediates the stimulatory effect of fatty acid pretreatment on increased fatty acid oxidation (Fig. 3B).

Figure 3. AMPK is essential for palmitate and linoleate effects on fatty acid oxidation.

Figure 3

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A, L6 myotubes were pretreated with palmitate or linoleate for 1 h and FA oxidation was subsequently determined. B, L6 myotubes were infected with adenovirus containing AdGo (CON) or a dominant negative (DN) AMPK mutant and FA oxidation was determined after treatment for 1 h with the saturated fatty acid palmitate (PAL) or the polyunsaturated fatty acid linoleate (LIN). Results are the mean (± s.e.m.) of two independent experiments (n = 8 total determinations). Fatty acid oxidation was greater (main effect, P < 0.05) in CON versus DN-AMPK. *P < 0.05 versus CON and PAL, †P < 0.05 versus CON and LIN within the same treatment.

Fatty acid esterification into cellular lipid pools

Fatty acid esterification into DAG was not affected by fatty acid pretreatment and was not different between CON and DN-AMPK cells. The incorporation of fatty acids into TAG was increased in CON compared with DN-AMPK (P < 0.05) and appeared to be a function of increased fatty acid uptake into the cell (average total fatty acid uptake across all treatments: DN-AMPK, 28.4 ± 2.5 nmol h−1; CON, 48.9 ± 3.8 nmol h−1; P < 0.05). The esterification: oxidation ratio, an index of fat storage compared with fat oxidation capacity, was increased (P < 0.05) in DN-AMPK but not CON after fatty acid pretreatment (Fig. 4).

Figure 4. AMPK inhibition increases the fatty acid esterification: oxidation ratio.

Figure 4

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L6 myotubes were infected with adenovirus containing AdGo (CON) or a dominant negative (DN) AMPK mutant and FA oxidation was determined after treatment for 1 h with the saturated fatty acid palmitate (PAL) or the polyunsaturated fatty acid linoleate (LIN). Results are the mean (± s.e.m.) of two independent experiments (n = 8 total determinations). *P < 0.05 versus CON within the same group.

Discussion

The present studies demonstrate that fatty acids increase AMPK Thr172 phosphorylation and AMPK activity without detectable changes in the cellular contents of AMP, ATP and phosphocreatine. The increased AMPK activity resulted in enhanced fatty acid oxidation, which was inhibited by the expression of a dominant negative AMPK. These results indicate that increasing fatty acid availability increases AMPK activity independent of changes in the cellular energy charge and support the idea that increasing fatty acid availability enhances fatty acid oxidation by an AMPK-mediated mechanism.

AMPK is a highly conserved heterotrimeric serine/threonine enzyme and functions as a cellular fuel sensor. AMPK is activated allosterically by AMP and is phosphorylated at Thr172 on the α subunit and activated by the upstream AMPK kinase, LKB1. Although increases in AMP were thought to be essential for increased AMPK activity, several studies demonstrate activation without changes in the cellular energy status (Fryer et al. 2002; Hawley et al. 2002; Minokoshi et al. 2002). In the present study, we demonstrate that acutely increasing fatty acid availability in vitro increases AMPK activation via Thr172 phosphorylation independent from increases in cellular AMP. This response was observed at 250 μm and not at low (75 μm) palmitate concentrations, indicating that these effects are relevant at physiological fatty acid levels. The experiments conducted in purified LKB1 indicate that fatty acids enhance AMPK phosphorylation by LKB1 in an AMP independent manner. The fatty acid effect appears to be specific to AMPK and may be due to fatty acid interacting with AMPK on the β or γ subunit, although the underlying mechanisms are not yet clear. In agreement with a previous report (Sakamoto et al. 2005), we confirm that LKB1 is the principal AMPKK present in skeletal muscle cells. LKB1 activity was reduced following treatment with both palmitate and linoleate, an effect not observed following treatment with palmitoyl-CoA. Whilst this indicates a direct inhibitory effect of fatty acids and not fatty acyl-CoA on LKB1 activity, a role for other lipid metabolites cannot be discounted. Unsaturated fatty acids are reported to stimulate PP2C activity in vitro (Klumpp et al. 1998) indicating that the changes in AMPK activity in response to fatty acid treatment are not likely to be due to inhibition of PP2C inactivation of AMPK.

The physiological relevance of these findings is unclear. On the one hand, high circulating plasma fatty acid levels are a hallmark of obesity and poorly controlled type 2 diabetes, and fatty acid oversupply is associated with fatty acid metabolite (e.g. ceramide, diacylglycerol) accumulation in skeletal muscle, which causes insulin resistance (Shulman, 2000). On the other hand it seems reasonable that AMPK is activated by fatty acids to protect normal cellular function. AMPK inactivates glycerol 3-phosphate acyltransferase (Muoio et al. 1999) and hormone sensitive lipase (Garton et al. 1989; Watt et al. 2004; Watt et al. 2005), rate-limiting enzymes for triacylglycerol esterification and hydrolysis, respectively. AMPK also phosphorylates and inhibits ACCβ and has been reported to activate malonyl-CoA decarboxylase (Sambandam et al. 2004), both important for decreasing malonyl-CoA content and enhancing fatty acid oxidation (Abu-Elheiga et al. 2001). Combined, these events would be expected to result in preferential partitioning of exogenous fatty acids away from esterification and towards oxidation. Such regulation would also prevent futile cycles of triacylglycerol breakdown and resynthesis from glycerol and FFAs (Tagliaferro et al. 1990). In support of this concept, increasing fatty acid availability in isolated skeletal muscle resulted in up-regulation of fatty acid oxidation while concomitantly reducing endogenous fatty acid oxidation (Dyck et al. 1997). In the present study AMPK-DN resulted in increased esterification of endogenous fatty acids into triacylglycerol and an increased esterification/oxidation ratio. Perhaps most significantly, AMPK-DN cells were unable to increase oxidation of fatty acids, resulting in greater fat storage in muscle cells. Collectively, our data in skeletal muscle cells demonstrate that AMPK is required to maintain basal rates of fatty acid oxidation, that AMPK is essential for the ‘feed forward’ activation of fatty acid oxidation and that AMPK is required to reduce fatty acid esterification in the face of a fat challenge.

Recently it was reported that palmitoyl-CoA (Ki 0.5–5 μm) was a strong inhibitor of AMPK activation by LKB1, and palmitate (400 μm) was without affect (Taylor et al. 2005). As mentioned above, we found no inhibition of AMPK activation by palmitoyl-CoA. While we do not know the reason for these differences there are several differences in the reagents used in the two studies; Taylor et al. used recombinant LKB1 and bacterially expressed AMPK holoenzyme α2β2γ2 whereas we used native bovine testis LKB1 and insect cell expressed AMPK α1β1γ1. We found no effect of fatty acids on LKB1 phosphorylation of the NUAC2 (LKB1tide) synthetic peptide substrate whereas Taylor et al. reported stimulation by 50 μm palmitoyl-CoA.

In conclusion, we provide evidence of a role for fatty acids in the control of AMPK in skeletal muscle. These data demonstrate that fatty acids activate AMPK by facilitating LKB1 phosphorylation of AMPK independent of changes in the cellular energy charge. In muscle cells, an increase in AMPK is required for acute fatty acid-induced increases in fat oxidation and prevention of excess intramyocellular fat storage.

Acknowledgments

M.J.W. is supported by a National Health and Medical Research Council (NHMRC) Peter Doherty postdoctoral fellowship, G.R.S. by a CIHR ‘Target Obesity’ research fellowship and B.E.K. by an Australian Research Council (ARC) Federation fellowship, and M.A.F. is a NHMRC senior research fellow. We thank the ARC, National Heart Foundation and NHMRC for support. GRS is supported by a ‘Target Obesity’ Research fellowship from the Canadian Institutes of Health Research, The Heart & Stroke Foundation of Canada and the Canadian Diabetes Association.

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