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Am J Physiol Endocrinol Metab 288: E347-E352, 2005. First published October 19, 2004; doi:10.1152/ajpendo.00238.2004
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Creatine feeding increases GLUT4 expression in rat skeletal muscle

Jeong-Sun Ju, Jill L. Smith, Peter J. Oppelt, and Jonathan S. Fisher

Department of Biology, Saint Louis University, St. Louis, Missouri

Submitted 3 June 2004 ; accepted in final form 18 October 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The purpose of this study was to investigate the potential role of creatine in GLUT4 gene expression in rat skeletal muscle. Female Wistar rats were fed normal rat chow (controls) or chow containing 2% creatine monohydrate ad libitum for 3 wk. GLUT4 protein levels of creatine-fed rats were significantly increased in extensor digitorum longus (EDL), triceps, and epitrochlearis muscles compared with muscles from controls (P < 0.05), and triceps GLUT4 mRNA levels were ~100% greater in triceps muscles from creatine-fed rats than in muscles from controls (P < 0.05). In epitrochlearis muscles from creatine-fed animals, glycogen content was ~40% greater (P < 0.05), and insulin-stimulated glucose transport rates were higher (P < 0.05) than in epitrochlearis muscles from controls. Despite no changes in [ATP], [creatine], [phosphocreatine], or [AMP], creatine feeding increased AMP-activated protein kinase (AMPK) phosphorylation by 50% in rat EDL muscle (P < 0.05). Creatinine content of EDL muscle was almost twofold higher for creatine-fed animals than for controls (P < 0.05). Creatine feeding increased protein levels of myocyte enhancer factor 2 (MEF2) isoforms MEF2A (~70%, P < 0.05), MEF2C (~60%, P < 0.05), and MEF2D (~90%, P < 0.05), which are transcription factors that regulate GLUT4 expression, in creatine-fed rat EDL muscle nuclear extracts. Electrophoretic mobility shift assay showed that DNA binding activity of MEF2 was increased by ~40% (P < 0.05) in creatine-fed rat EDL compared with controls. Our data suggest that creatine feeding enhances the nuclear content and DNA binding activity of MEF2 isoforms, which is concomitant with an increase in GLUT4 gene expression.

acetyl-coenzyme A carboxylase; adenosine monophosphate-activated protein kinase; myocyte enhancer factor 2; phosphocreatine; creatinine

A NUMBER OF STUDIES regarding beneficial effects of creatine supplementation on muscle glucose metabolism have been reported. For example, creatine supplementation improves impaired glucose tolerance (9) and increases glycogen content (18). Furthermore, combined creatine and carbohydrate supplements result in a greater postexercise muscle glycogen resynthesis than carbohydrate alone (23). However, few studies (17, 18, 28) have been done regarding the effects of creatine supplementation on changes in the expression level of the insulin- and contraction-regulated glucose transporter (GLUT4) that mediates glucose uptake in muscle tissue, and a clear picture has not emerged from the findings described in the literature. It has been demonstrated that creatine supplementation prevents a decrease in muscle GLUT4 protein content during 2 wk of immobilization and increases GLUT4 protein content during a subsequent 10 wk of rehabilitation training in healthy subjects (18). However, the physiological mechanisms supporting elevated GLUT4 expression by creatine supplementation were not addressed. A separate study of creatine supplementation in humans found ~30% greater (not statistically significant) GLUT4 protein in muscle from creatine-supplemented subjects than from control subjects (28). There is reportedly no effect of 5 days of creatine supplementation on GLUT4 expression in rats (17).

A recent study demonstrated an approximately twofold increase in AMP-activated protein kinase (AMPK) phosphorylation after 48 h of creatine supplementation in L6 myocytes (5). It has been shown that myocyte enhancer factors 2 (MEF2)A, -C and -D, transcription factors that regulate GLUT4 expression in muscle (27), increase in response to AMPK activation (16). We hypothesized that the beneficial effects on glucose metabolism related to creatine supplementation might occur concomitantly with increased GLUT4 expression. We have evaluated whether 3 wk of oral creatine supplementation enhances GLUT4 biogenesis concomitantly with increased AMPK phosphorylation and MEF2 DNA binding activity.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials. A polyclonal antibody specific for the GLUT4 glucose transporter was the generous gift of Dr. Mike Mueckler (Washington University, St. Louis, MO). The primers and the internal standard mRNA for competitive RT-PCR were kindly supplied by Dr. John Holloszy (Washington University). Horseradish peroxidase-conjugated goat anti-rabbit IgG was purchased from Pierce Biotechnology (Rockford, IL). MEF2A and -D antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). MEF2C, phospho-AMPK{alpha} (P-AMPK), AMPK{alpha}, and phospho-acetyl-CoA carboxylase (P-ACC) antibodies were purchased from Cell Signaling Technology (Beverly, MA). The antibodies against AMPK and P-AMPK are pan-{alpha} antibodies that bind to both isoforms, {alpha}1 and {alpha}2, of the catalytic subunit of AMPK. ATPlite and reagents for enhanced chemiluminescence (ECL) were obtained from Perkin-Elmer Life Sciences (Boston, MA). NE-PER Nuclear Extraction Reagent, Biotin 3'-End DNA Labeling Kit and LightShift Chemiluminescent EMSA (electrophoretic mobility shift assay) kit were purchased from Pierce Biotechnology (Rockford, IL). Creatinine amidohydrolase was obtained from MP Biomedicals (Philadelphia, PA). Radiolabeled 2-deoxyglucose and mannitol were obtained from American Radiolabeled Chemicals (St. Louis, MO). All other reagents were obtained from Sigma Chemical (St. Louis, MO).

Animal care. Fouteen female Wistar rats weighing ~90 g were obtained from Charles River Laboratories and divided into two groups: 1) creatine (Cr) supplementation [2% creatine monohydrate (Harlan Teklad, Madison, WI), n = 7 rats] and normal chow diet (n = 7 rats). The rats were provided with unrestricted access to water and diets for 3 wk. After an overnight fast, animals were anesthetized with an intraperitoneal injection of pentobarbital sodium (5 mg/100 g body wt), and the skeletal muscle tissues were rapidly dissected out. Epitrochlearis muscles were assayed for glucose transport activity, as described in Glucose transport and glycogen assays. Other muscles were immediately frozen with clamps cooled in liquid nitrogen. This research was approved by the Saint Louis University Animal Care Committee.

Metabolite analysis. Muscle samples were stored at –80°C until analyzed. For assay of ATP, phosphocreatine (PCr), creatine, and AMP, extensor digitorum longus (EDL) muscles were ground in 5.6% perchloric acid containing 40% ethanol at –17°C. The –17°C homogenization temperature was maintained by cooling grinding tubes in an ethanol-dry ice bath during homogenizations. Samples were spun and supernatants neutralized with buffer containing 3 M KOH, 0.5 M imidazole, and 0.4 M KCl. ATP was analyzed with the ATPlite Luminescence ATP Detection Assay System (Perkin-Elmer) according to the manufacturer's instructions. PCr, creatine, and AMP content were measured with spectrophotometric or fluorometric assays as previously described by Bergmeyer (2) and Passonneau and Lowry (20). Total creatine (TCr) content was obtained by adding PCr and creatine concentrations. Creatinine was assayed by an end point creatinine amidohydrolase method based on the kinetic assay described by Moss et al. (15).

Western blot analysis. Homogenates for Western blot analysis were made by grinding muscle samples with ice-cold buffer with grinding tubes resting in ice-water baths. Muscle extracts for GLUT4 analyses were prepared by homogenizing muscle in 250 mM sucrose containing 20 mM HEPES and 1 mM EDTA, pH 7.4 (22). For P-AMPK, AMPK, and P-ACC Western blots, muscle tissues were homogenized in a buffer suitable for preserving phosphorylation states of enzymes, containing 50 mM HEPES, pH 7.4, 150 mM NaCl, 10% glycerol, 1% Triton X-100, 1.5 mM MgCl2, 1 mM EDTA, 10 mM Na3PO4, 100 mM NaF, 2 mM Na3VO4, 10 µg/ml leupeptin, 10 µg/ml aprotinin, and 1 mM phenylmethylsulfonyl fluoride (24), and samples for Western blots were prepared from supernatants after centrifugation of homogenates for 10 min at 14,000 g. For evaluation of MEF2 isoform protein content, nuclear extract from EDL muscle was isolated with the NE-PER nuclear extraction reagent (Pierce). Protein concentrations in the samples were measured using a bicinchoninc acid protein assay reagent kit (Pierce). Aliquots of homogenates were solubilized in Laemmli sample buffer containing dithiothreitol and boiled for 5 min, except for samples for GLUT4 analysis, which did not contain dithiothreitol and were not boiled. Samples in Laemmli sample buffer were subjected to SDS-PAGE (10% resolving gels, 5% gels for P-ACC), Western blotting, and visualization with ECL. GLUT4, AMPK, P-AMPK, and P-ACC blots yielded single bands at the expected molecular masses. ACC{beta} (280 kDa) is the primary ACC isoform in muscle, but it is possible that if the {alpha}-isoform (257 kDa) was present, it could have run to the same position as the ACC{beta} band on P-ACC blots. The prominent band of the two shown on MEF2 blots is at the expected molecular mass.

Competitive RT-PCR. Triceps muscle was used for the GLUT4 mRNA analysis instead of EDL, because the mRNA extraction procedure required a larger amount of tissue than was available from EDL muscles. However, we believe that triceps is a good proxy for EDL for the mRNA analysis, because the fiber type distributions for the two muscles are similar (7). Total RNA from triceps muscles was isolated using TRIzol reagent (Invitrogen). Competitive RT-PCR was performed as described by Garcia-Roves et al. (12).

Glucose transport and glycogen assays. 2-Deoxyglucose (2-DG) glucose transport in the absence or presence of 60 µU/ml insulin was assayed as described by Young et al. (31). Epitrochlearis muscles were allowed to recover for 1 h after dissection at 35°C with gentle shaking in an incubation medium that consisted of 0.1% RIA-grade bovine serum albumin, 8 mM glucose, and 32 mM mannitol in Krebs-Henseleit bicarbonate buffer gassed with 5% CO2-95% O2. Muscles were then incubated for 30 min in the same medium in the absence or presence of 60 µU/ml insulin (Iletin II; Lilly, Indianapolis, IN) before being washed twice at 30°C for 10 min per wash in glucose-free medium containing 40 mM mannitol and insulin, if it had been present for the previous step. Finally, muscles were incubated at 30°C for 20 min in glucose-free medium containing 8 mM 2-DG (with 3 µCi/ml [3H]2-DG) and 32 mM mannitol (with 0.2 µCi/ml [14C]mannitol), with insulin present if it had been present in the previous incubations. Muscles were blotted and trimmed at 4°C and frozen with clamps cooled in liquid nitrogen. Muscle homogenates were made in 0.3 M perchloric acid, and intracellular 2-DG content was determined after scintillation counting, as described previously (31). Glycogen content for epitrochlearis muscles that were not exposed to insulin was assayed with the amyloglucosidase method described by Passoneau and Lauderdale (19).

EMSA. The MEF2 DNA binding activity assay was performed with nuclear extract from EDL muscle tissues prepared with the NE-PER nuclear extraction reagent (Pierce). Synthetic oligonucleotides were labeled with the biotin 3'-End DNA Labeling Kit (Pierce). This oligonucleotide probe contains a functional MEF2 binding site in the GLUT4 promoter (the recognition sequence for MEF2 is italicized): forward oligo, 5'-GAT CGC TCT AAA AAT AAC CCT GTC G-3'; reverse oligo, 5'-C GAC AGG GTT ATT TTT AGA GCG ATC-3' (14). Reactions of biotin end-labeled target DNA and nuclear extracts in the presence or absence or 200-fold excess unlabeled oligonucleotides and subsequent electrophoresis and ECL visualization were performed with an EMSA kit (LightShift, Pierce Biotechnology) according to the manufacturer's instructions.

Data analysis. Data are presented as means ± SE. Creatine effects (creatine vs. control) were evaluated by a univariate analysis of variance (ANOVA). Glucose transport data were analyzed with a 2 x 2 ANOVA (insulin absence/presence x control/creatine).


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Body weights and food intakes were measured every 2–3 days and did not differ between groups.

Muscle metabolites. EDL muscle high-energy phosphate metabolite contents were not altered by creatine feeding (Table 1). There were no significant differences in PCr, creatine, ATP, or AMP levels in rat muscle tissue between creatine-fed and chow-fed control animals (Table 1). The PCr-to-TCr concentration ([PCr]/[TCr]) ratio did not differ in EDL muscles from creatine-fed animals compared with muscles from controls (Table 1). However, creatinine levels were almost twofold greater in muscles from creatine-fed rats than in muscles from controls (Table 1). The creatinine concentrations for these muscles are in the ranges previously reported for rat skeletal muscle [~1 µmol/g, assayed after homogenization in water and mixing with trichloracetic acid (26)] and rat heart [~2.6 µmol/g, assayed after homogenization in perchloric acid (11)].


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  		Table 1. Muscle high-energy phosphate metabolite and creatine content

 
Muscle GLUT4 expression, glucose transport activity, and glycogen content. Creatine feeding induced an ~60% increase in GLUT4 protein in EDL muscle (Fig. 1A), an ~80% increase in GLUT content of triceps muscle (Fig. 1B), and a twofold increase of GLUT4 in epitrochlearis (Fig. 1C). As shown in Fig. 1D, triceps muscle GLUT4 mRNA levels increased by 100% in the creatine-fed group compared with chow-fed control animals, suggesting that increases in GLUT4 expression induced by creatine supplementation are mediated at the transcriptional level. Because of the similar creatine-related increase in GLUT4 in EDL and triceps and the fiber type similarity between the two (7), we believe that the mRNA data for triceps are generalizable to EDL. Insulin (60 µU/ml) stimulated an ~2.5-fold increase in glucose transport rate in epitrochlearis muscles for control animals and an ~5-fold increase in glucose transport for creatine-fed animals (Fig. 2A). The creatine x insulin interaction (chow/creatine x basal/insulin) was statistically significant (P < 0.05), and insulin-stimulated glucose transport was greater for creatine-fed animals than for controls (P < 0.05). The data in Fig. 2A represent values for paired muscles (for each animal, glucose transport was assayed in both the absence and the presence of insulin). These muscle pairs allow computation of the increase in glucose transport that is stimulated by insulin for each animal, and this increase is about twofold higher for creatine-fed animals than for controls (P < 0.005). Glycogen content of epitrochlearis muscles was 40% higher for creatine-fed animals than for controls (Fig. 2B).



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  		Fig. 1. Creatine feeding increases skeletal muscle GLUT4 expression. Muscle GLUT4 protein content was determined by Western blot analysis as described in MATERIALS AND METHODS for extensor digitorum longus (EDL; A), triceps (B), and epitrochlearis (EPI; C). D: intensities (arbitrary units) of triceps GLUT4 mRNA transcript from chow-fed control (CON) and creatine-supplemented (Cr) muscle samples. GLUT4 mRNA was measured in triceps muscles by competitive RT-PCR as described in MATERIALS AND METHODS. Values are means ± SE; n = 7/group for CON and Cr rats. *P < 0.05.

 


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  		Fig. 2. Epitrochlearis muscle glucose transport activity and glycogen content. A: isolated epitrochlearis muscles from CON (n = 7 rats) and Cr animals (n = 6 rats) were assayed for glucose transport activity in the absence or presence of 60 µU/ml insulin and the insulin-stimulated increase in glucose transport, calculated as the difference between basal glucose transport and insulin-stimulated glucose transport for each animal. B: glycogen content of epitrochlearis muscles that were not exposed to insulin (n = 7/group). Values are means ± SE. *P < 0.05 for the creatine x insulin interaction in A, greater insulin-stimulated glucose transport in creatine-fed animals than controls in A, and higher muscle glycogen content for creatine-fed animals than for controls in B. {dagger}P < 0.005 for greater insulin-stimulated increase in glucose transport for Cr animals than for CON.

 
AMPK and ACC phosphorylation. The phosphorylation of Thr172 of AMPK{alpha} is shown in Fig. 3, A and B. AMPK phosphorylation of creatine-fed rats was significantly increased by 50% (P < 0.05) in EDL muscle compared with muscles from regular chow-fed rats. Muscle content of nonphosphorylated AMPK was not affected by creatine feeding (control 1.00 ± 0.02, creatine-fed rats 0.94 ± 0.08 arbitrary units, n = 7/group) Phosphorylation levels of ACC were 46% higher in EDL muscles from creatine-fed animals compared with EDL muscles from controls (Fig. 3, C and D). The increase in phosphorylation (Ser79) of ACC, an AMPK substrate, is consistent with increased AMPK phosphorylation.



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  		Fig. 3. Effects of creatine feeding on phosphorylation of AMP-activated protein kinase (AMPK) and acetyl-CoA carboxylase (ACC) in EDL muscles. A: representative phospho-AMPK (P-AMPK, top) and AMPK (bottom) Western blot samples. B: densitometry of P-AMPK; n = 7/group for CON and Cr rats. C: representative sample of phospho-ACC (P-ACC) Western blots. D: densitometry of P-ACC; n = 7/group. *P < 0.05 vs. CON. Values are means ± SE.

 
MEF2 isoform content. Nuclear content of MEF2A and MEF2D, members of the MEF2 transcription factor family known to regulate GLUT4 expression, was increased in the muscle of the creatine-fed vs. the control group (Fig. 4). Seventy-two percent higher levels of MEF2A (Fig. 4A) and 90% higher levels of MEF2D (Fig. 4C) in creatine-fed compared with control muscle were found. Concomitantly, MEF2C, another MEF2 isoform known to participate in the activity of GLUT4 promoter, was also increased by 60% in muscle nuclear extracts of the creatine-fed vs. the control group (Fig. 4B).



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  		Fig. 4. Representative blots and their respective densitometric quantification of effects of creatine on myocyte enhancer factor 2 (MEF2) isoforms. Creatine feeding increased protein levels of MEF2 isoforms in Cr rat EDL muscle nuclear extracts. A: MEF2-A. B: MEF2-C. C: MEF2-D. Values are means ± SE for 7/group for CON and Cr rats. *P < 0.05.

 
MEF2-DNA binding activity. MEF2 EMSA was performed using a synthetic oligonucleotide probe containing a sequence encoding for the GLUT4 promoter known to have a functional binding site for MEF2 isoforms. Three weeks of creatine feeding caused a 44% increase in MEF2 binding activity in nuclear isolates of EDL muscle compared with the control group (Fig. 5).



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  		Fig. 5. MEF2 electrophoretic mobility shift assay from EDL muscle samples of rats fed either Cr or CON diet: DNA probe alone (lane 1), CON (lanes 2, 4, and 6) and vs. Cr group (lanes 3, 5, and 7), and an excess of unlabeled probe (lane 8). Values are means ± SE; n = 7/group. *P < 0.05.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The present study demonstrated that 3 wk of creatine supplementation increases the expression of the GLUT4 gene in rat skeletal muscle. Creatine supplementation was related to increased phosphorylation of AMPK, increased content of MEF2 protein in nuclear extracts, and increased DNA binding activity of MEF2 isoforms.

The present study showed that creatine feeding increases AMPK phosphorylation in rat skeletal muscle. A recent study also shows that 48-h treatment with creatine increased activating phosphorylation of both AMPK{alpha}1 and -{alpha}2 isoforms (5). Currently, the effects of creatine supplementation on AMPK activation have not been fully studied. There is considerable evidence that AMPK activation is regulated by phosphorylation of the AMPK {alpha}-subunit by upstream kinases (AMP kinase kinase) (13, 29, 30) and that decreasing [PCr]/[TCr] ratio causes allosteric activation of AMPK (21). In our hands, creatine and phosphocreatine concentrations were not altered by creatine feeding. Although this is not a common finding (see, e.g., Ref. 17), other investigators have found no changes in total creatine in skeletal muscle over 7 wk of creatine feeding (4). Interestingly, despite no changes in intracellular [creatine], increases in plasma [creatine], and no change in creatine transporter abundance, creatine uptake rates were lowered by ~50% in white muscle after the 7 wk of creatine feeding (4). This suggests that, during creatine feeding, some factor other than tissue total creatine content can affect muscle physiology. The present data demonstrating increased muscle creatinine concentrations after creatine feeding raise the question of whether muscle function can be altered by changes in the creatine plus creatinine pool, rather than creatine concentrations per se.

In disagreement with the present study and the work of others (4) demonstrating that creatine feeding does not alter creatine content in rat muscle tissue, creatine supplementation has been reported to decrease the [PCr]/[TCr] ratio (5, 25). Ceddia and Sweeney (5) suggested that a decrease in the [PCr]/[TCr] ratio after creatine supplementation might cause the phosphorylation of AMPK. However, there is currently no published information regarding whether a change in the [PCr]/[Cr] ratio regulates rates of AMPK phosphorylation or dephosphorylation. Although there are reports of stimulation of AMPK phosphorylation independent of changes in concentrations of adenine nucleotides, creatine, or phosphocreatine (6, 10), the mechanism mediating the increase in AMPK phosphorylation by creatine supplementation in the present study and as previously reported (5) is unclear at this time.

MEF2 is a transcription factor that plays a key role in specific skeletal muscle gene expression (3). The activation of MEF2 in skeletal muscle is regulated via parallel intracellular signaling pathways in response to cellular stress or activation of AMPK (1). It has been demonstrated that the activation of AMPK increases MEF2 DNA binding activity (1, 32), resulting in increased muscle GLUT4 protein levels (16, 32). In our study, creatine feeding increased protein levels of MEF2A, -C, and -D isoforms in nuclear extracts of skeletal muscle. We also found that DNA binding activity of MEF2 was increased by 44% in creatine-fed rat EDL muscle nuclear extracts.

Some studies have not found increased GLUT4 after creatine feeding. For example, 5 days of creatine supplementation did not alter muscle GLUT4 content in rat skeletal muscle (17), and creatine supplementation did not affect muscle GLUT4 expression in a 6-wk supplementation period in humans (28). It has been proposed that creatine supplementation per se has no effect on either GLUT4 expression or glycogen content in humans but that it is effective in combination with exercise (8). However, it has been reported that creatine supplementation prevented a decrease in muscle GLUT4 protein content during 2 wk of immobilization (18), suggesting that creatine supplementation itself (without dietary supplementation or exercise) is able to affect muscle GLUT4 expression. At the present time, we are unable to explain the reasons for discrepant findings about creatine effects on GLUT4 expression. More studies need to be done to elucidate reasons for varying results on this issue and to clarify mechanisms.

In summary, we found that creatine feeding increases AMPK phosphorylation in muscle through an unknown process that does not require changes in creatine or phosphocreatine levels in muscle. Creatine feeding was related to enhanced nuclear content of MEF2, increased DNA binding activity of MEF2, increased in GLUT4 gene expression, and increased glycogen content in skeletal muscle. Our findings elucidate some of the possible physiological factors that might explain how creatine supplementation regulates GLUT4 biogenesis in muscle tissue.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
J. Fisher is supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant K01-DK-066330.


   ACKNOWLEDGMENTS
 
We thank the Saint Louis University Department of Comparative Medicine for specialized animal care.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. Fisher, 3507 Laclede Ave., Dept. of Biology, Saint Louis University, St. Louis, MO 63103 (E-mail fisherjs{at}slu.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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