J. Anim Sci. 2008. 86:1296-1305. doi:10.2527/jas.2007-0794
© 2008 American Society of Animal Science
ANIMAL GROWTH, PHYSIOLOGY, AND REPRODUCTION |
AMP-activated protein kinase and adipogenesis in sheep fetal skeletal muscle and 3T3-L1 cells1
J. Tong2,
M. J. Zhu2,
K. R. Underwood,
B. W. Hess,
S. P. Ford and
M. Du3
Department of Animal Science and Interdepartmental Molecular and Cellular Life Sciences Program, University of Wyoming, Laramie 82071
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Abstract
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Marbling, or i.m. fat, is an important factor determining beef quality. Both adipogenesis and hypertrophy of existing adipocytes contribute to enhanced marbling. We hypothesized that the fetal stage is important for the formation of i.m. adipocytes and that AMP-activated protein kinase (AMPK) has a key role in adipogenesis during this stage. The objective of this study was to assess the role of AMPK in adipogenesis in fetal sheep muscle and 3T3-L1 cells. Nonpregnant ewes were randomly assigned to a control (Con, 100% of NRC recommendations, n = 7) or overfed (OF, 150% of NRC, n = 7) diet from 60 d before to 75 d after conception, when the ewes were killed. The fetal LM was collected at necropsy for biochemical analyses. The activity of AMPK was less in the fetal muscle of OF sheep. The expression of peroxisome proliferator-activated receptor (PPAR)
, a marker of adipogenesis, was greater in OF fetal muscle compared with Con fetal muscle. To further show the role of AMPK in adipogenesis, we used 3T3-L1 cells. The 3T3-L1 cells were incubated in a standard adipogenic medium for 24 h and 10 d. Activation of AMPK by 5-aminoimidazole-4-car-boxamide-1-β-d-ribonucleoside dramatically inhibited the expression of PPAR and reduced the presence of adipocytes after 10 d of differentiation. Inhibition of AMPK by compound C enhanced the expression of PPAR. In conclusion, these data show that AMPK activity is inversely related to adipogenesis in fetal sheep muscle and 3T3-L1 cells.
Key Words: 3T3-L1 cells • adipogenesis • AMP-activated protein kinase • fetus • sheep • skeletal muscle
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INTRODUCTION
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Marbling (i.m. fat) is a primary criterion for beef quality grading, as well as for the eating quality of pork and lamb (Hausman et al., 2007
; Underwood et al., 2007b). Marbling is correlated with the number of adipocytes in skeletal muscle; however, mechanisms controlling adipogenesis in skeletal muscle remain poorly defined (Hausman and Poulos, 2004; Hausman et al., 2007). Understanding such mechanisms will allow us to develop practical strategies to enhance i.m. fat accumulation.
Skeletal muscle cells and adipocytes are both derived from mesenchymal pluripotent cells, which are abundant in fetal muscle of meat animals around mid-gestation (Yamanouchi et al., 2007). Adipogenesis starts with the formation of preadipocytes, the majority of which differentiate into mature adipocytes in late gestation (Feve, 2005; Gnanalingham et al., 2005; Muhlhausler et al., 2006). Adipose tissue growth in later life is due to hypertrophy of existing adipocytes and hyperplasia due to the generation of new adipocytes from pluripotent cells or preadipocytes (Feve, 2005). However, new fat cells generated later in life are mainly located in visceral, retroperitoneal, and subcutaneous fat depots, with few located in the i.m. fat depot (Faust et al., 1978; Miller et al., 1984; Valet et al., 2002). Thus, i.m. adipogenesis during the fetal stage is anticipated to have a dominant effect on the number of adipocytes existing within skeletal muscle.
Adenosine monophosphate-activated protein kinase (AMPK) has a central role in energy metabolism (Hardie, 2007). Once activated, AMPK promotes fatty acid oxidation and inhibits lipid synthesis in cells through phosphorylation and inhibition of acetyl-CoA carboxylase (ACC) activity (Carey et al., 2006; Ravnskjaer et al., 2006; Yoon et al., 2006). Thus, it is likely that AMPK is a key player in adipogenesis during fetal muscle development.
The objective of this study was to assess the role of AMPK in adipogenesis of fetal sheep muscle and 3T3-L1 cells.
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MATERIALS AND METHODS
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All animal procedures were approved by the University of Wyoming Animal Care and Use Committee.
Care and Use of Animals
All ewes (n = 14) used in current study were mated with a single ram. Beginning 60 d before conception and continuing to d 75 of gestation (day of mating = d 0), multiparous Rambouillet/Columbia ewes (parity of ewes was balanced between treatments) were individually fed a highly palatable concentrated diet (Table 1) at 100% (control, Con) of the NRC recommendations for energy (NRC, 1985) or 150% (OF) of the recommended energy requirements for early gestation. Ewes were housed in individual pens within a temperature-controlled (20°C) room. All ewes were weighed at weekly intervals, and the rations were adjusted for weekly changes in metabolic BW (BW0.75). Body condition was scored at monthly intervals to evaluate changes in fatness. A BCS of 1 (emaciated) to 9 (obese) was assigned by 2 trained observers after palpation of the transverse and vertical processes of the lumbar vertebrae (L2 through L5) and the region around the tail head (Sanson et al., 1993).
Immediately before necropsy, on d 75, 14 pregnant ewes (7 Con and 7 OF) were weighed. Sedation was induced by i.v. ketamine (10 mg/kg of BW, Sigma, St. Louis, MO) and anesthesia was induced and maintained by 1 to 2% isoflurane inhalation. Fetal blood (10 mL) was collected from the umbilical vein via a 20-ga, 3.8-cm needle and 3-mL syringe. Serum and plasma were collected and stored for another study. After the blood collection, fetuses were quickly removed and exsanguinated through the umbilical vein. Fetal LM samples were collected from 5 ewes carrying twin pregnancies in each group. After trimming the surface tissues, a small piece of muscle (1 g) was sampled at the anatomical center of the muscle and snap frozen in liquid nitrogen for biological analyses. No difference in BW was observed between twins and, thus, 1 fetus of each twin pregnancy was randomly selected for analyses. Although no difference in weight was observed among fetuses of different sexes, the sex of the fetuses in each group was balanced. Ewes were killed by exsanguination.
The 3T3-L1 Cell Culture
All chemicals for cell culture were bought from Sigma-Aldrich (St. Louis, MO) unless noted otherwise. Four independent studies were conducted. Preadipocyte 3T3-L1 cells (ATCC, Manassas, VA) were cultured in Dulbecco’s modified Eagle medium containing 10% (vol/vol) fetal bovine serum (FBS), 100 units/mL of penicillin, and 100 µg/mL of streptomycin in a humidified atmosphere of 95% air and 5% CO2 at 37°C. Medium was changed every other day. The 3T3-L1 cells were seeded into a 10-cm diam. dish and allotted to treatments. At 30% confluence, 3T3-L1 cells were incubated in a standard adipogenic medium composed of 10% FBS/Dulbecco’s modified Eagle medium supplemented with insulin (20 mIU/mL), dexamethasone (0.1 µM), 3-isobutyl-1-methylxanthine (IBMX, 0.5 mM), and troglitazone (10 µM; (Klemm et al., 2001; Kim and Chen, 2004; Yada et al., 2006; Yamanouchi et al., 2007). The 5-aminoimidazole-4-carboxamide-ribonucleoside (0.1 mM and 1.0 mM AICAR, Calbiochem, San Diego, CA) and compound C (1 and 10 µM, prepared using 1,000 x stock solution in 100 mM HCl, Calbiochem) were used to activate or inhibit AMPK in the cells, respectively. The cells were collected at 24 h for immunoblotting analyses and at 10 d (duration needed for adipocyte differentiation) for Oil Red O staining directly. Cells were collected after a brief rinse with PBS and were harvested in 0.5 mL of ice-cold lysis buffer [50 mM Tris-HCl (pH 7.4), 137 mM NaCl, 1 mM CaCl2, 1 mM MgCl, 10% glycerol (vol/vol), 2% SDS (wt/vol), 1% Triton X-100 (vol/vol), 2.5 mM EDTA, 100 mM NaF, 2 mM Na3VO4, and 1% proteinase inhibitor cocktail (Shen et al., 2007)].
Antibodies
Antibodies against phospho-AMPK at Thr 172, phospho-ACC at Ser 79, peroxisome proliferator-activated receptor (PPAR) , and horseradish peroxidase-linked secondary antibody were purchased from Cell Signaling (Danvers, MA). Anti-β-actin antibody was obtained from Developmental Studies Hybridoma Bank (DSHB, Iowa City, IA).
Immunoblotting Analysis
Muscle (0.1 g) powdered in liquid nitrogen was used for immunoblotting analyses as described previously (Zhu et al., 2004, 2006). Briefly, the muscle sample (0.1 g) was homogenized in 400 µL of ice-cold buffer containing 137 mM NaCl, 50 mM HEPES, 2% SDS, 1% NP-40, 10% glycerol, 2 mM phenylmethylsulfonyl fluoride, 10 mM sodium pyrophosphate, 10 µg/mL of aprotinin, 10 µg/mL of leupeptin, 2 mM Na3VO4, and 100 mM NaF (pH 7.4). The protein content of the lysates was determined by the Bradford method (Bio-Rad Laboratories, Hercules, CA; Zhu et al., 2006).
Muscle homogenate (30 µg of protein) was mixed with an equal volume of 2 x standard SDS sample loading buffer containing 137 mM Tris-HCl (pH 6.8), 4.0% SDS, 20% glycerol, and 0.002% bromophenol blue for SDS-PAGE separation of proteins. Proteins on the gels were transferred to nitrocellulose membranes in a transfer buffer containing 20 mM Tris-base, 192 mM glycine, 0.1% SDS, and 20% methanol. Membranes were incubated in a blocking solution consisting of 5% nonfat dry milk in TBS/T [0.1% Tween-20, 50 mM Tris-HCl (pH7.6), and 150 mM NaCl] for 1 h. Membranes were incubated overnight in a 1:1,000 dilution (vol/vol) of primary antibodies in TBS/T with 2% BSA (SeraCare Diagnostics, Millford, MA). Membranes were then incubated with horseradish peroxidase-conjugated secondary antibodies at a 1:1,000 dilution (vol/vol) for 1 h in TBS/T with gentle agitation. Membranes were visualized using Enhanced Chemiluminescence Western blotting reagents (Amersham Bioscience, Piscataway, NJ) and exposure to film (MR, Kodak, Rochester, NY). Density of the bands was quantified by using an Imager Scanner II and ImageQuant TL software (National Institutes of Health, Bethesda, MD). Band density was normalized according to the β-actin content (Zhu et al., 2004).
Real-Time Quantitative PCR
The mRNA was extracted from the fetal muscle using TRI Reagent (Sigma, St. Louis, MO) and reverse transcribed into cDNA using a kit (Qiagen, Valencia, CA). Reverse transcribed cDNA were used for real-time quantitative PCR analyses by using SYBR Green RT-PCR kit from Bio-Rad (Hercules, CA). Primer sets used were: PPAR forward, 5'-CCGCATCTTCCAGGGGTG TC-3', and reverse, 5'-CAAGGAGGCCAGCATCGT-GAAAT-3'; PPAR
forward, 5'-GCGCCGTGTGATT-TACGTT-3', and reverse, 5'-GAAGGGCGGATTGTTG TTGGTCT-3'; PPAR coactivator 1- (PGC-1) forward, 5'-GCGCCGTGTGATTTACGTT-3' and reverse, 5'-AAAACTTCAAAGCGGTCTCTCAA-3'; uncoupling protein 1 (UCP1) forward, 5'-GCTAGTTTAGGAAG-CAAGTC-3', and reverse, 5'-GCCCCGTCAAGCCTTC TGTTGTTG-3'. The 18S RNA was used as a control, forward, 5'-GTAACCCGTTGAACCCCATT-3', and reverse, 5'-CCATCCAATCGGTAGTAGCG-3' (Lomax et al., 2007). Each reaction yielded amplicons between 80 and 200 bp. The PCR conditions were as follows: 20 s at 95°C, 20 s at 56°C, and 20 s at 72°C for 35 cycles. After amplification, a melting curve (0.01 °C/sec) was used to confirm product purity. Results are expressed relative to 18S rRNA (Lomax et al., 2007).
Oil Red O Staining of Intramuscular Triacylglycerols
After 10 d of differentiation, muscle sections and 3T3-L1 cells were stained with Oil Red O working solution [a mixture of 0.5% (wt/vol) Oil Red O in 2-propanol and distilled water at a 3:2 ratio] for 7 min, rinsed with PBS to remove excessive Oil Red O dye (Kim and Chen, 2004), and then subjected to microscopic observation at 200x magnification. Two images were captured from each section, and 5 sections were examined for each muscle sample. The total area of Oil Red O staining for each image was quantified by using Image J software (National Institutes of Health, Washington, DC) and expressed as the percentage of total image area.
Statistical Analysis
Statistical analyses were conducted according to our previous studies in sheep (Zhu et al., 2004, 2006). Briefly, each animal or a separate cell culture experiment was considered as an experimental unit. Dietary treatments or chemical treatments of cells were considered as the main effect. Data were analyzed as a completely randomized design using PROC GLM (SAS Inst. Inc., Cary, NC). The differences in the mean values were compared by the Tukey’s multiple comparison, and mean ± SE were reported. Statistical significance was considered as P < 0.05.
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RESULTS AND DISCUSSION
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Skeletal muscle cells and adipocytes are both derived from mesenchymal pluripotent cells (Artaza et al., 2005; Poulos and Hausman, 2006). In approximately mid-gestation, fetal skeletal muscle has a large number of pluripotent cells which can differentiate into myogenic cells or adipogenic cells (Feve, 2005; Gnanalingham et al., 2005; Muhlhausler et al., 2006). Enhancing adipogenesis in fetal muscle is expected to provide sites for fat accumulation in later life, increasing marbling. Adenosine monophosphate-activated protein kinase has a central role in controlling energy metabolism (Hardie, 2007). Activated AMPK phosphorylates and inhibits the activity of ACC, a key enzyme in lipid synthesis. Activation of AMPK accelerates fatty acid oxidation due to a reduction in malonyl-CoA content (Merrill et al., 1997). We previously demonstrated that AMPK was negatively associated with marbling in beef cattle (Underwood et al., 2007a). Therefore, it is highly possible that AMPK activity is associated with adipogenesis in fetal muscle. Because obesity leads to the reduction of AMPK activity in skeletal muscle (Bandyopadhyay et al., 2006; Sriwijitkamol et al., 2006), we hypothesized that maternal obesity would inhibit AMPK and increase adipogenesis in fetal muscle. In this study, OF ewes developed severe obesity (Table 2). The BW of OF fetuses was greater than the BW of Con fetuses (374 ± 10 g and 268 ± 12 g, respectively; P < 0.05). These data are consistent with the macrosomal fetuses frequently observed in obese pregnant women (Sahu et al., 2007).
Sheep and cattle are both ruminant animals, which makes them physiologically similar. Because the small size of sheep provides convenience and also dramatically reduces experimental costs, we used sheep to study adipogenesis in fetal muscle as affected by maternal nutrition. Marbling is one of the most important traits for beef, but also has importance for lamb and pork (Hausman et al., 2007; Underwood et al., 2007b). Therefore, this study has implications for several meat animal species.
Downregulation of AMPK and ACC Activity in Fetal Muscle
Acetyl-CoA carboxylase is a key enzyme regulating lipid metabolism. Its activity is negatively controlled by AMPK through phosphorylation at Ser 79 (Horman et al., 2005; Takekoshi et al., 2006). Therefore, AMPK controls lipid metabolism in cells through phosphorylation of ACC. In addition, AMPK regulates adipogenesis though the exact mechanism is vague (Dagon et al., 2006). Thus, it is likely that AMPK and ACC are involved in adipogenesis within fetal muscle. Phosphorylation of AMPK at Thr 172 was downregulated in the skeletal muscle of OF sheep compared with Con sheep (25.4 ± 6.6%, P < 0.05; Figure 1A). Phosphorylation of ACC was also reduced (36.2 ± 8.1%, P < 0.05) in the muscle of OF sheep (Figure 1B). These data clearly show AMPK activity was downregulated in OF fetal muscle compared with Con muscle. The activation of ACC due to inhibition of AMPK should promote lipid accumulation in fetal muscle.
Adipogenesis in Fetal Muscle
In fetal muscle through late gestation, there are no mature adipocytes (Casteilla et al., 1987; Lomax et al., 2007). However, the lack of mature adipocytes does not exclude the cells which have progressed through differentiation to the point of being equipped at the molecular level to accumulate triacylglycerols. Indeed, the initial events in adipogenesis start in mid-gestation (Feve, 2005; Gnanalingham et al., 2005; Muhlhausler et al., 2006). Peroxisome proliferator-activated receptor- is a key regulator of adipogenesis. The expression of PPAR leads to adipogenic differentiation from pluripotent cells, and PPAR is highly expressed in adipose tissue (Spiegelman et al., 2000). Hence, we measured the PPAR mRNA expression to show whether there was enhanced adipogenesis. Indeed, PPAR mRNA expression levels were much greater in OF vs. Con fetal muscle (Figure 2A). Furthermore, we also analyzed that PPAR content by immunoblotting. Two bands of PPAR were detected, which might correspond to the 2 isoforms PPAR1 and PPAR2 (Tontonoz et al., 1994). Again, the PPAR content (2 bands combined) was greater in OF fetal muscle (Figure 2B). Because PPAR is a marker of adipocyte differentiation, these data indicated enhanced adipogenesis in OF fetal muscle. Of course, PPAR is also expressed in skeletal muscle, but the level of expression is very low (Vidal-Puig et al., 1996; Verma et al., 2004). To further evaluate adipogenesis in fetal muscle, the content of a preadipocyte marker, preadipocyte factor-1 was analyzed (Smas and Sul, 1993; Kim et al., 2007). Preadipocyte factor-1 is exclusively expressed in preadipocytes, not mature adipocytes (Fahrenkrug et al., 1999; Mei et al., 2002). Its expression was greater in OF fetal skeletal muscle compared with Con fetal skeletal muscle, showing a greater number of pluripotent cells had committed to adipogenesis in OF fetuses (Figure 3). Therefore, these data indicated that the adipogenesis was enhanced in OF fetal muscle. Previous studies in pigs and cattle indicate that fetal stage is important for the regulation of genes involved in i.m. fat accumulation (Cagnazzo et al., 2006; Lehnert et al., 2007). However, to the knowledge of authors, this is the first report showing that overfeeding dams enhances adipogenesis in fetal muscle in important livestock species.
In ruminant animal fetuses, brown adipose tissue is dominant, which rapidly transforms into white adipose tissue within the first week of life (Casteilla et al., 1987; Lomax et al., 2007). Peroxisome proliferators-activated receptor is preferably expressed in brown adipose tissue (Lomax et al., 2007). Its expression induces the expression of PPAR coactivator 1, which further induces the expression of uncoupling protein-1, a protein conferring the thermogenic function of brown adipose tissue (Lomax et al., 2007). Our data show that the mRNA expression for PPAR and PPAR coactivator-1 were greater in OF fetal muscle than Con fetal muscle (Figure 4A, B), although no difference was observed for uncoupling protein-1 mRNA expression. These data further confirmed the enhancement of adipogenesis in OF fetal muscle. These data also show that a portion of those developing adipocytes in fetal muscle might be destined to brown adipocytes.
During mid-gestation, adipogenesis in fetal muscle has just been initiated; thus, there are no mature adipocytes available at this stage (Casteilla et al., 1987). Therefore, we did not detect mature adipocytes in fetal muscle. However, the accumulation of lipids in OF fetal muscle was greater than Con muscle (Figure 5) in agreement with the enhanced adipogenesis in OF fetal muscle.
Association Between AMPK and Adipogenesis
Our data showed that AMPK activity was inhibited and adipogenesis was enhanced in fetal muscle. How- ever, these data do not establish the cause-effect relationship between AMPK and adipogenesis. To answer this question, we used 3T3-L1 cells. The 3T3-L1 cells were cultured in an adipogenic medium and treated with AICAR, a specific activator of AMPK, for 24 h and 10 d. Cells treated for both 24 h and 10 d were used for the detection of PPAR expression and phosphorylation of AMPK and ACC, whereas cells differentiated for 10 d were used for Oil Red O staining of adipocytes. A 10-d period is necessary for the differentiation of 3T3-L1 cells into mature adipocytes.
At 24 h after differentiation, activation of AMPK by AICAR induced a dose-dependent inhibition of PPAR, eliciting that activation of AMPK inhibited adipogenesis in 3T3-L1 cells (Figure 6). Compound C is a specific inhibitor of AMPK (Hong-Brown et al., 2007). Applying compound C induced a greater expression of PPAR (Figure 6). We further analyzed the activation of AMPK in 3T3-L1 cells due to AICAR and compound C treatments. Addition of AICAR to culture media induced a dose-dependent activation of AMPK phosphorylation at Thr 172 and ACC phosphorylation at Ser 79, demonstrating that AMPK was activated by AICAR (Figure 7A, B), except for ACC phosphorylation at 0.1 mM AICAR treatment where ACC phosphorylation was inhibited (Figure 7B). The reason for this inhibition was unclear, but may be associated with the differential effect of low and high AMPK activation on downstream signaling, as indicated in a recent study in colon cancer cells (Park et al., 2006). In contrast, phosphorylation of both AMPK and ACC was inhibited by compound C (Figure 7A, B). These data were in agreement with previous reports that AMPK activation inhibited the expression of PPAR and C/EBP in preadipocytes (Giri et al., 2006). After 10 d of differentiation, AICAR treatment induced similar changes in AMPK and ACC phosphorylation, as well as PPAR expression (Figure 8). After 10 d of differentiation, activation of AMPK dramatically reduced the number of adipocytes in 3T3-L1 cells (Figure 9). However, incubation with compound C for more than 5 d led to cell death. At d 10 of incubation, extensive cell detachment and death (more than 80% cell death) resulted, demonstrating the essential role of AMPK for cell survival (data not shown). These data in 3T3-L1 cells clearly demonstrate AMPK has a crucial regulatory role in adipogenesis, which is in agreement with previous reports (Habinowski and Witters, 2001; Giri et al., 2006).
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Figure 7. Phosphorylation of AMP-activated protein kinase (AMPK) and phosphorylation of acetyl-CoA carboxylase (ACC) in 3T3-L1 cells at 24 h of differentiation. Panel A shows representative phospho-AMPK immunoblots and mean ± SEM; panel B shows representative phospho-ACC immunoblots and mean ± SEM. AICAR = 5-Aminoimidazole-4-carboxamide ribonucleoside; and Comp. C = compound C. *Control vs. treatments, P < 0.05; **P < 0.01 (n = 4 per treatment).
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Figure 9. Presence of adipocytes in 3T3-L1 cells after 10 d of differentiation as affected by 5-Aminoimidazole-4-carboxamide ribonucleoside (AICAR) treatment. By Oil Red O staining, adipocytes appear dark in the micrographs.
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It is well established that PPAR expression induces adipogenic differentiation. This receptor functions as an obligate heterodimer with retinoid X receptor and binds to DNA sequences called direct repeat-1 elements, which induces the expression of adipocyte-specific genes (Spiegelman et al., 2000). However, it remains largely unclear how AMPK regulates PPAR expression and adipogenesis. Few reports examined the possible mechanisms associated with AMPK in adipogenesis (Habinowski and Witters, 2001; Giri et al., 2006). Activation of AMPK by AICAR induced downregulation of key adipogenic genes and suggested that this effect was mediated by phosphorylation of eukaryotic initiation factor 2 (Dagon et al., 2006). The effect of AMPK on adipogenesis in fetal muscle has not been previously evaluated.
In conclusion, AMPK was inhibited in fetal muscle of overfed ewes, and this inhibition was associated with enhanced adipogenesis in fetal muscle. Therefore, adipogenesis in sheep fetal muscle can be enhanced by inhibition of AMPK. These data indicate that maternal nutrient supplementation can be utilized to inhibit AMPK in fetal muscle, which may be a strategy to enhance marbling in beef cattle and other meat animals.
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Footnotes
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1 This work was supported by Research Initiative Grant 2006-55618-16914, 2007-35203-18065, and 2008-35206-18826 from the USDA Cooperative State Research, Education and Extension Service, and University of Wyoming INBRE P20 RR016474-04. The monoclonal antibody of actin developed by Jim Jung-Ching Lin was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by the University of Iowa, Department of Biological Sciences, Iowa City, IA 52242. 
2 These two authors made an equal contribution.
3 Corresponding author: mindu{at}uwyo.edu
Received for publication December 11, 2007.
Accepted for publication March 1, 2008.
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LITERATURE CITED
|
|---|
Artaza, J. N., S. Bhasin, T. R. Magee, S. Reisz-Porszasz, R. Shen, N. P. Groome, M. F. Meerasahib, and N. F. Gonzalez-Cadavid. 2005. Myostatin inhibits myogenesis and promotes adipogenesis in C3H 10T(1/2) mesenchymal multipotent cells. Endocrinology 146:3547–3557.[Abstract/Free Full Text]
Bandyopadhyay, G. K., J. G. Yu, J. Ofrecio, and J. M. Olefsky. 2006. Increased malonyl-CoA levels in muscle from obese and type 2 diabetic subjects lead to decreased fatty acid oxidation and increased lipogenesis; thiazolidinedione treatment reverses these defects. Diabetes 55:2277–2285.[Abstract/Free Full Text]
Cagnazzo, M., M. F. te Pas, J. Priem, A. A. de Wit, M. H. Pool, R. Davoli, and V. Russo. 2006. Comparison of prenatal muscle tissue expression profiles of two pig breeds differing in muscle characteristics. J. Anim. Sci. 84:1–10.[Abstract/Free Full Text]
Carey, A. L., G. R. Steinberg, S. L. Macaulay, W. G. Thomas, A. G. Holmes, G. Ramm, O. Prelovsek, C. Hohnen-Behrens, M. J. Watt, D. E. James, B. E. Kemp, B. K. Pedersen, and M. A. Febbraio. 2006. Interleukin-6 increases insulin-stimulated glucose disposal in humans and glucose uptake and fatty acid oxidation in vitro via AMP-activated protein kinase. Diabetes 55:2688–2697.[Abstract/Free Full Text]
Casteilla, L., C. Forest, J. Robelin, D. Ricquier, A. Lombet, and G. Ailhaud. 1987. Characterization of mitochondrial-uncoupling protein in bovine fetus and newborn calf. Am. J. Physiol. 252:E627–E636.[Medline]
Dagon, Y., Y. Avraham, and E. M. Berry. 2006. AMPK activation regulates apoptosis, adipogenesis, and lipolysis by eIF2alpha in adipocytes. Biochem. Biophys. Res. Commun. 340:43–47.[Medline]
Fahrenkrug, S. C., B. A. Freking, and T. P. Smith. 1999. Genomic organization and genetic mapping of the bovine PREF-1 gene. Biochem. Biophys. Res. Commun. 264:662–667.[CrossRef][Medline]
Faust, I. M., P. R. Johnson, J. S. Stern, and J. Hirsch. 1978. Diet-induced adipocyte number increase in adult rats: A new model of obesity. Am. J. Physiol. 235:E279–E286.[Medline]
Feve, B. 2005. Adipogenesis: Cellular and molecular aspects. Best Pract. Res. Clin. Endocrinol. Metab. 19:483–499.[CrossRef][Medline]
Giri, S., R. Rattan, E. Haq, M. Khan, R. Yasmin, J. S. Won, L. Key, A. K. Singh, and I. Singh. 2006. AICAR inhibits adipocyte differentiation in 3T3L1 and restores metabolic alterations in diet-induced obesity mice model. Nutr. Metab. (Lond.) 3:31–51.[CrossRef][Medline]
Gnanalingham, M. G., A. Mostyn, M. E. Symonds, and T. Stephenson. 2005. Ontogeny and nutritional programming of adiposity in sheep: Potential role of glucocorticoid action and uncoupling protein-2. Am. J. Physiol. Regul. Integr. Comp. Physiol. 289:R1407–R1415.[Abstract/Free Full Text]
Habinowski, S. A., and L. A. Witters. 2001. The effects of AICAR on adipocyte differentiation of 3T3-L1 cells. Biochem. Biophys. Res. Commun. 286:852–856.[CrossRef][Medline]
Hardie, D. G. 2007. Biochemistry. Balancing cellular energy. Science 315:1671–1672.
Hausman, G. J., and S. Poulos. 2004. Recruitment and differentiation of intramuscular preadipocytes in stromal-vascular cell cultures derived from neonatal pig semitendinosus muscles. J. Anim. Sci. 82:429–437.[Abstract/Free Full Text]
Hausman, G. J., S. P. Poulos, T. D. Pringle, and M. J. Azain. 2007. The influence of thiazolidinediones on adipogenesis in vitro and in vivo: Potential modifiers of intramuscular adipose tissue deposition in meat animals. J. Anim. Sci. 86 (E. Suppl.):E236–E243.[CrossRef][Medline]
Hong-Brown, L. Q., C. R. Brown, D. S. Huber, and C. H. Lang. 2007. Alcohol regulates eukaryotic elongation factor 2 phosphorylation via an AMP-activated protein kinase-dependent mechanism in C2C12 skeletal myocytes. J. Biol. Chem. 282:3702–3712.[Abstract/Free Full Text]
Horman, S., N. Hussain, S. M. Dilworth, K. B. Storey, and M. H. Rider. 2005. Evaluation of the role of AMP-activated protein kinase and its downstream targets in mammalian hibernation. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 142:374–382.[CrossRef][Medline]
Kim, J. E., and J. Chen. 2004. regulation of peroxisome proliferator-activated receptor-gamma activity by mammalian target of rapamycin and amino acids in adipogenesis. Diabetes 53:2748–2756.[Abstract/Free Full Text]
Kim, K. A., J. H. Kim, Y. Wang, and H. S. Sul. 2007. Pref-1 (preadipocyte factor 1) activates the MEK/extracellular signal-regulated kinase pathway to inhibit adipocyte differentiation. Mol. Cell. Biol. 27:2294–2308.[Abstract/Free Full Text]
Klemm, D. J., J. W. Leitner, P. Watson, A. Nesterova, J. E. Reusch, M. L. Goalstone, and B. Draznin. 2001. Insulin-induced adipocyte differentiation. Activation of CREB rescues adipogenesis from the arrest caused by inhibition of prenylation. J. Biol. Chem. 276:28430–28435.[Abstract/Free Full Text]
Lehnert, S. A., A. Reverter, K. A. Byrne, Y. Wang, G. S. Nattrass, N. J. Hudson, and P. L. Greenwood. 2007. Gene expression studies of developing bovine longissimus muscle from two different beef cattle breeds. BMC Dev. Biol. 7:95.[CrossRef][Medline]
Lomax, M. A., F. Sadiq, G. Karamanlidis, A. Karamitri, P. Trayhurn, and D. G. Hazlerigg. 2007. Ontogenic loss of brown adipose tissue sensitivity to beta-adrenergic stimulation in the ovine. Endocrinology 148:461–468.[Abstract/Free Full Text]
Mei, B., L. Zhao, L. Chen, and H. S. Sul. 2002. Only the large soluble form of preadipocyte factor-1 (Pref-1), but not the small soluble and membrane forms, inhibits adipocyte differentiation: role of alternative splicing. Biochem. J. 364:137–144.[Medline]
Merrill, G. F., E. J. Kurth, D. G. Hardie, and W. W. Winder. 1997. AICA riboside increases AMP-activated protein kinase, fatty acid oxidation, and glucose uptake in rat muscle. Am. J. Physiol. 273:E1107–E1112.[Medline]
Miller, W. H., Jr., I. M. Faust, and J. Hirsch. 1984. Demonstration of de novo production of adipocytes in adult rats by biochemical and radioautographic techniques. J. Lipid Res. 25:336–347.[Abstract]
Muhlhausler, B. S., J. A. Duffield, and I. C. McMillen. 2006. Increased maternal nutrition stimulates Peroxisome Proliferator Activated Receptor- (PPAR), adiponectin and leptin mRNA expression in adipose tissue before birth. Endocrinology 148:878–885.[CrossRef][Medline]
NRC. 1985. Nutrient Requirements of Sheep. 6th ed. National Academy Press, Washington, DC.
Park, I. J., J. T. Hwang, Y. M. Kim, J. Ha, and O. J. Park. 2006. Differential modulation of AMPK signaling pathways by low or high levels of exogenous reactive oxygen species in colon cancer cells. Ann. N. Y. Acad. Sci. 1091:102–109.[CrossRef][Medline]
Poulos, S. P., and G. J. Hausman. 2006. A comparison of thiazolidinedione-induced adipogenesis and myogenesis in stromal-vascular cells from subcutaneous adipose tissue or semitendinosus muscle of postnatal pigs. J. Anim. Sci. 84:1076–1082.[Abstract/Free Full Text]
Ravnskjaer, K., M. Boergesen, L. T. Dalgaard, and S. Mandrup. 2006. Glucose-induced repression of PPARalpha gene expression in pancreatic beta-cells involves PP2A activation and AMPK inactivation. J. Mol. Endocrinol. 36:289–299.[Abstract/Free Full Text]
Sahu, M. T., A. Agarwal, V. Das, and A. Pandey. 2007. Impact of maternal body mass index on obstetric outcome. J. Obstet. Gynaecol. Res. 33:655–659.[CrossRef][Medline]
Sanson, D. W., T. R. West, W. R. Tatman, M. L. Riley, M. B. Judkins, and G. E. Moss. 1993. Relationship of body composition of mature ewes with condition score and body weight. J. Anim. Sci. 71:1112–1116.[Abstract]
Shen, Q. W., M. J. Zhu, J. Tong, J. Ren, and M. Du. 2007. Ca2+/ calmodulin-dependent protein kinase kinase is involved in AMP-activated protein kinase activation by alpha-lipoic acid in C2C12 myotubes. Am. J. Physiol. Cell Physiol. 293:C1395–C1403.[Abstract/Free Full Text]
Smas, C. M., and H. S. Sul. 1993. Pref-1, a protein containing EGF-like repeats, inhibits adipocyte differentiation. Cell 73:725–734.[CrossRef][Medline]
Spiegelman, B. M., P. Puigserver, and Z. Wu. 2000. Regulation of adipogenesis and energy balance by PPARgamma and PGC-1. Int. J. Obes. Relat. Metab. Disord. 24(Suppl. 4):S8–S10.
Sriwijitkamol, A., J. L. Ivy, C. Christ-Roberts, R. A. DeFronzo, L. J. Mandarino, and N. Musi. 2006. LKB1-AMPK signaling in muscle from obese insulin-resistant Zucker rats and effects of training. Am. J. Physiol. Endocrinol. Metab. 290:E925–E932.[Abstract/Free Full Text]
Takekoshi, K., M. Fukuhara, Z. Quin, S. Nissato, K. Isobe, Y. Kawakami, and H. Ohmori. 2006. Long-term exercise stimulates adenosine monophosphate-activated protein kinase activity and subunit expression in rat visceral adipose tissue and liver. Metabolism 55:1122–1128.[CrossRef][Medline]
Tontonoz, P., E. Hu, and B. M. Spiegelman. 1994. Stimulation of adipogenesis in fibroblasts by PPAR gamma 2, a lipid-activated transcription factor. Cell 79:1147–1156.[CrossRef][Medline]
Underwood, K. R., W. J. Means, M. J. Zhu, S. P. Ford, B. W. Hess, and M. Du. 2007a. AMP-activated protein kinase is negatively associated with intramuscular fat content in longissimus dorsi muscle of beef cattle. Meat Sci. In press.
Underwood, K. R., J. Tong, M. J. Zhu, Q. W. Shen, W. J. Means, S. P. Ford, S. I. Paisley, B. W. Hess, and M. Du. 2007b. Relationship between kinase phosphorylation, muscle fiber typing, and glycogen accumulation in Longissimus muscle of beef cattle with high and low intramuscular fat. J. Agric. Food Chem. 55:9698–9703.[CrossRef][Medline]
Valet, P., G. Tavernier, I. Castan-Laurell, J. S. Saulnier-Blache, and D. Langin. 2002. Understanding adipose tissue development from transgenic animal models. J. Lipid Res. 43:835–860.[Abstract/Free Full Text]
Verma, N. K., J. Singh, and C. S. Dey. 2004. PPAR-gamma expression modulates insulin sensitivity in C2C12 skeletal muscle cells. Br. J. Pharmacol. 143:1006–1013.[CrossRef][Medline]
Vidal-Puig, A., M. Jimenez-Linan, B. B. Lowell, A. Hamann, E. Hu, B. Spiegelman, J. S. Flier, and D. E. Moller. 1996. Regulation of PPAR gamma gene expression by nutrition and obesity in rodents. J. Clin. Invest. 97:2553–2561.[Medline]
Yada, E., K. Yamanouchi, and M. Nishihara. 2006. Adipogenic potential of satellite cells from distinct skeletal muscle origins in the rat. J. Vet. Med. Sci. 68:479–486.[CrossRef][Medline]
Yamanouchi, K., T. Hosoyama, Y. Murakami, and M. Nishihara. 2007. Myogenic and adipogenic properties of goat skeletal muscle stem cells. J. Reprod. Dev. 53:51–58.[CrossRef][Medline]
Yoon, M. J., G. Y. Lee, J. J. Chung, Y. H. Ahn, S. H. Hong, and J. B. Kim. 2006. Adiponectin increases fatty acid oxidation in skeletal muscle cells by sequential activation of AMP-activated protein kinase, p38 mitogen-activated protein kinase, and peroxisome proliferator-activated receptor alpha. Diabetes 55:2562–2570.[Abstract/Free Full Text]
Zhu, M. J., S. P. Ford, W. J. Means, B. W. Hess, P. W. Nathanielsz, and M. Du. 2006. Maternal nutrient restriction affects properties of skeletal muscle in offspring. J. Physiol. 575:241–250.[Abstract/Free Full Text]
Zhu, M. J., S. P. Ford, P. W. Nathanielsz, and M. Du. 2004. Effect of maternal nutrient restriction in sheep on the development of fetal skeletal muscle. Biol. Reprod. 71:1968–1973.[Abstract/Free Full Text]
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Abstract
INTRODUCTION
)] or 150% [overfed (OF, 
)] of the NRC requirements for energy. Panel A shows representative phospho-AMPK immunoblots and mean ± SEM; and panel B shows representative phospho-ACC immunoblots and mean ± SEM. *Con vs. OF, P < 0.05 (n = 5 per group).
