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Endocrinology, doi:10.1210/en.2007-0183
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Endocrinology Vol. 148, No. 12 5696-5705
Copyright © 2007 by The Endocrine Society

Evidence for Adipose-Muscle Cross Talk: Opposing Regulation of Muscle Proteolysis by Adiponectin and Fatty Acids

Qiugen Zhou, Jie Du, Zhaoyong Hu, Kenneth Walsh and Xiaonan H. Wang

Renal Division (Q.Z., X.H.W.), Department of Medicine, Emory University, Atlanta, Georgia 30322; Nephrology Division (J.D., Z.H.), Baylor College of Medicine, Houston, Texas 77030; and Boston University School of Medicine (K.W.), Boston, Massachusetts 02118

Address all correspondence and requests for reprints to: Dr. Xiaonan Wang, Renal Division, Emory University, School of Medicine, M/S 1930/001/1AG, 1639 Pierce Drive, WMB 338, Atlanta, Georgia 30322. E-mail: xwang03{at}emory.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Illnesses associated with insulin resistance exhibit increases in whole-body protein degradation and amino acid oxidation. However, the mechanisms stimulating muscle catabolism under these conditions are not clear. Because insulin resistance is associated with accumulation of lipids in muscle, we measured protein degradation in muscles of mice fed a high-fat diet. Muscle protein catabolism was accelerated on the high-fat diet, and this was associated with an increase in plasma free fatty acid and a decrease in plasma levels of the adipocyte-derived cytokine adiponectin. To evaluate how free fatty acids influence adiponectin-mediated changes in muscle protein breakdown we examined C2C12 skeletal muscle cells exposed to free fatty acids. Both saturated fatty acids (palmitate) and unsaturated fatty acids (oleate) increased protein degradation (25 and 18%, respectively) in part by activating the E3 ubiquitin ligases. Adenovirus-mediated overexpression of adiponectin blocked fatty acid-induced protein degradation in C2C12 cells. Palmitate activated the E3 ubiquitin ligases by suppressing insulin receptor substrate-1/Akt signaling in the C2C12 muscle cells, whereas adiponectin attenuated the E3 ubiquitin ligase activation by increasing both insulin receptor substrate-1 tyrosine phosphorylation and Akt Ser473 phosphorylation. In related experiments, adiponectin overexpression decreased TNF{alpha} and IL-6 expression in 3T3-L1 adipocytes, whereas exposure to free fatty acids had the opposite effect. We conclude that the balance between free fatty acids and adiponectin impacts muscle proteolysis in insulin-resistant conditions and suggest a role for adipose tissue-muscle cross talk in diabetes and obesity.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
MUSCLE WASTING IS a serious complication of several conditions that are associated with insulin resistance including uremia, heart failure, and diabetes mellitus (1, 2, 3, 4). The loss of muscle mass is an independent predictor of mortality in the elderly and in patients with chronic diseases (5, 6). Both obesity and type 2 diabetes are associated with an increase in whole-body protein catabolism (1, 7, 8). For example, it has been reported that whole-body protein degradation is 12–24% greater in type 2 diabetic patients compared with nondiabetic obese subjects in the isoenergetic fed state (9, 10, 11). In evaluating mechanisms for the proteolytic responses, we recently found that obesity-related insulin resistance is associated with an increase in muscle protein loss due to down-regulation of the insulin receptor substrate-1 (IRS-1)/phosphatidylinositol-3-kinase (PI3K) Akt pathway (1).

IRS-1/PI3K/Akt is an intracellular signaling pathway that is activated by insulin, and it regulates cell growth and apoptosis (12, 13, 14). In skeletal muscle, an increase in IRS-1/PI3K/Akt signaling stimulates myofiber hypertrophy and counteracts the loss of muscle mass that occurs in animals with catabolic disorders. This mechanism involves an increase in muscle protein synthesis and a decrease in protein degradation that is associated with activation of Akt (phosphorylation of Akt) (15, 16, 17). Akt phosphorylates members of the forkhead (FOXO) family, and this results in decrease the transcription of E3 ubiquitin ligases like atrogin-1/muscle atrophy F-box (MAFbx) and muscle RING finger-1 (MuRF-1) (18, 19, 20). A decrease of phosphorylated Akt (p-Akt) also causes the activation of caspase-3, which breaks down the complex myofibrillar structure to form substrates that are further degraded by the ubiquitin-proteasome system (15, 21, 22).

Obesity-linked metabolic changes are associated with a decrease in circulating adiponectin levels. Adiponectin functions as an endogenous, insulin-sensitizing factor (23, 24). Adiponectin is an abundant gene product (2–10 µg/ml in plasma) of adipose tissue and accounts for 0.01% of total plasma protein (25). Adiponectin levels are high in healthy lean individuals, but its levels are low in obese individuals. In addition, the plasma adiponectin concentration correlates negatively with glucose, insulin, triglyceride levels, and body mass index (BMI) (26, 27). Adiponectin enhances insulin sensitivity by regulating multiple signaling pathways including AMP-activated protein kinase (AMPK) (28). AMPK phosphorylates key regulatory enzymes and is considered to be a regulator of energy balance at both the cellular and whole-body levels. We previously found that plasma adiponectin concentration is low in obese db/db mice with accelerated muscle protein degradation, whereas increased adiponectin is associated with a decrease in muscle proteolysis in these mice (1).

The signaling mechanisms connecting insulin resistance, adiponectin levels, and muscle proteolysis have not been previously analyzed. In this study, protein degradation rates were examined in vivo and in vitro to investigate the impact of adiponectin and free fatty acids (FFA) on muscle protein metabolism. First, we examined the effect of a high-fat (HF) diet on the rate of protein degradation in muscles of C57BL/6JLer mice. Next, we examined the effect of adiponectin on cytokine production in 3T3-L1 adipocytes. Finally, we examined the effect of adiponectin on protein degradation in C2C12 skeletal muscle cells in response to treatment with FFA. We found that both HF feeding in vivo and FFA treatment in vitro reduced IRS-1/Akt signaling in muscle. FFA treatment led to accelerated protein degradation, and this effect was reversed by adiponectin. Our results provide evidence for a mechanism that controls muscle mass via adipose-muscle cross talk involving adiponectin.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
C57BL/6JLer mice (Gene Cdh23; Jackson Laboratories, Bar Harbor, ME) were fed a control or a HF diet. Four-week-old mice were randomly assigned to normal rodent chow (23% protein, 10% fat, 49% carbohydrate) or to a HF diet containing 23% protein, 35.8% fat, and 35.5% carbohydrate (Research Diets, New Brunswick, NJ) provided ad libitum for 5 months. Mice were housed in a 12-h light, 12-h dark animal care facility. At the end of experiments, HF-fed and birth-matched control mice were anesthetized by 12 mg/kg xylazine and 60 mg /kg ketamine, and the soleus (predominately oxidative, red fibers) and extensor digitorum longus (EDL, predominately glycolytic, white fibers) muscles were removed, and protein degradation was measured as the release of tyrosine. The hind-limb muscles were excised, flash-frozen in liquid nitrogen, and stored at –80 C for further experiments. The experiments were approved by the institutional animal care and use committee of the Emory University.

Plasma insulin was measured using 1-2-3 ultrasensitive mouse insulin enzyme immunoassay kit, (American Laboratories, Windham, NH). The coefficient of variation (Cv) of this assay is equal to 0.1%. Blood glucose was measured by Accu-CHEK Advantage blood glucose meter (Roche, Indianapolis, IN; Cv = 0.4%). Mouse plasma adiponectin concentration was measured by ELISA using a mouse adiponectin ELISA kit (ALPCO, Windham, NH; Cv = 0.1%). Serum FFA concentrations were measured by the colorimetric determination of nonesterified fatty acids (NEFA) using the NEFA kit (Wako Chemicals, Richmond, VA; Cv = 0.3%). Plasma cytokine levels were measured by the cytokine protein microarray quantification assay (Allied Biotech Inc., Ijamsville, MD; Cv = 0.2%).

Cell culture and adenovirus transduction
Mice skeletal muscle C2C12 myoblast cells (American Type Culture Collection, Rockville, MD) were cultured in DMEM with 10% fetal bovine serum (or 2% horse serum for differentiation), 4.5 g/liter glucose, 100 U/ml penicillin, 100 µg/ml streptomycin, and 4 mM glutamine and studied between passages 3 and 8. HEK 293A cells (Invitrogen, Carlsbad, CA) were cultured in DMEM and studied between passages 3 and 20. Mouse 3T3-L1 preadipocytes, purchased from American Type Culture Collection, were studied between passages 3 and 8. Protocols for maintaining and differentiating cells were described previously (1).

The two adenoviruses (Ad-ctrl and Ad-adipo) were used in this study; Ad-adipo contains full-length adiponectin cDNA (29), and Ad-ctrl contains a nonspecific gene. Virus was amplified in HEK 293A cells and purified by adeno-X virus purification kit (Clontech, Inc., Mountain View, CA). Virus titer was detected by series dilution and calculated as transduction units per milliliter as follows: (infected cells per field) x (fields per well)/volume virus (milliliters) x dilution factor. The conditions (virus titer, infection time, and transduction efficiency) of Ad-ctrl and Ad-adipo transduction were identical. The transduced cells showed greater than 95% infection.

Protein degradation
In isolated muscle.
We measured protein degradation as the rate of tyrosine release into the media as muscle neither synthesizes nor degrades tyrosine, and tyrosine does not accumulate in the intracellular pool of muscle (30). Soleus and EDL muscles (10–20 mg) were fixed at resting length on a plastic support and incubated in Krebs-Ringer bicarbonate (KRB) buffer (135.5 mM NaCl, 4.7 mM KCl, 24.8 mM NaHCO3, 1 mM MgSO4, 1 mM KH2PO4, 2.5 mM CaCl2, 10 mM glucose) containing 0.5 mM cycloheximide to block tyrosine reutilization. The muscles of the size we studied (10–20 mg) is adequate for the diffusion of glucose and oxygen. Thus, measurements of protein metabolism can be made reliably (31). Muscles incubated at resting length maintain high levels of ATP and phosphocreatine and achieve almost neutral protein balance when studied for 2 h (32). KRB buffer was pregassed for 30 min using 5% CO2/95% oxygen. After a 30-min preincubation, each muscle was transferred to a flask containing fresh medium and incubated at 37 C for 2 h. Tyrosine released into incubation medium was measured by a fluorometric technique and calculated for protein degradation (Cv = 0.1%) (30).

In cultured cells.
Protein degradation in cultured myotubes was measured using cells prelabeled with L-[U-14C]phenylalanine as described (33). Briefly, C2C12 cells were cultured in six-well plates with normal growth medium. The cells were transduced with Ad-adipo or Ad-ctrl (109 transduction units) in 2% horse serum DMEM (differentiation medium). After 48 h, L-[U-14C]phenylalanine was added to the media for 3 d to metabolically label intracellular proteins. Phenylalanine was used because it is neither synthesized nor degraded in muscle cells and does not accumulate in myocytes (34). To minimize reutilization of L-[U-14C]phenylalanine, nonradioactive L-phenylalanine was added into experimental medium (35). The myotubes were incubated in fresh experimental medium for 48 h. Aliquots were removed at four different time points, and protein was precipitated by adding 10% trichloroacetic acid. The amount of released free L-[U-14C]phenylalanine in the supernatant fractions was used for calculation of the rate of protein degradation (35). To determine whether changes of glucose concentration affected proteolysis in cultured cells, we used media with 4.5 g/liter glucose along with FFA because in a pilot study, we found that different concentrations of glucose (4.5 g/liter vs. 1.0 g/liter) did not change the rate of protein degradation in C2C12 cells.

Western blot analysis and immunoprecipitation
Western blot.
Hind-limb muscles from control and HF-fed mice were homogenized in radioimmunoprecipitation assay buffer. C2C12 cells were transduced with Ad-adipo or Ad-ctrl in differentiation medium for 48 h. Cells were treated with BSA-bound FFA or BSA alone (control) for 16 h. Cells were routinely harvested in radioimmunoprecipitation assay buffer except for analysis of actin cleavage. These cells were harvested in hypotonic buffer as described (21). Protein concentration was assayed using the RC-PC protein assay kit (Bio-Rad, Hercules, CA). The primary antibodies that we used included anti-IRS-1 (1:1000 dilution; Upstate, Lake Placid, NY), anti-pFOXO1/FOXO3-Thr32 antibody (1:500 dilution; Upstate), anti-pFOXO1-Ser253 antibody (1:500 dilution; Upstate), anti Ser307 IRS-1 (1:1000 dilution; Cell Signaling Technology, Beverly, MA), anti-Akt1 and Akt2 (1:1000 dilution; Cell Signaling), anti-p-Akt (Ser473) (1:1000 dilution; Cell Signaling), anti-AMPK (1:1000 dilution; Cell Signaling), anti-p-AMPK (Thr172) (1:1000 dilution; Cell Signaling).

Immunoprecipitation.
C2C12 skeletal muscle total cell lysate was prepared in immunoprecipitation buffer [20 mM Tris-HCl (pH 7.6), 5 mM EDTA, 0.5% Nonidet P-40, 10% glycerol, 0.1% SDS, 50 mM NaF, 150 mM NaCl, 2.8 mM ß-glycerophosphate, 1 mM Na3VO4, 10 mM Na pyrophosphate, 1 mM dithiothreitol, and one tablet protease inhibitor cocktail (Roche) per 10 ml buffer). Phosphotyrosine-containing proteins from C2C12 cells were immunoprecipitated from equal amounts of cellular protein (0.5 mg) with anti-phosphotyrosine PY-20 (10 µg; BD Transduction Laboratories, San Jose, CA) overnight. Phosphotyrosine-containing IRS-1 was determined by Western blotting of these proteins followed by probing with anti-IRS-1 antibody.

Muscle histology
Gastrocnemius muscles were processed for cryosectioning after placing them in cryo-molds containing embedding medium. The samples were frozen in isopentane cooled in dry ice, and the sections were stained with oil red-O in propylene glycol (Poly Scientific R & D Corp., Bay Shore, NY). Images were visualized with an Olympus 1 x 51 inverted microscope and captured by SIS-CC12 CLR camera.

Northern blots and RT-PCR
Muscle was removed and immediately freeze-clamped in liquid nitrogen. Total RNA was isolated using TriReagent (Molecular Research Inc., Cincinnati, OH) and used in Northern blots for atrogin-1/MAFbx and MuRF-1 as described (22). Autoradiographic signals were quantified by densitometry (Bio-Rad) and expressed relative to the corresponding ß-actin to correct for variations in RNA loading and transfer. For measuring RT-PCR assays, we isolated total RNA from C2C12 myotubes or from differentiated 3T3-L1 cells and performed RT using the GeneAmp RNA PCR Core kit (Applied Biosystems, Foster City, CA). Primers for adiponectin, TNF{alpha}, and IL-6 genes were designed as described (1). Primers for atrogin-1/MAFbx and MuRF-1 were designed to cross intron-exon boundaries. They were used to generate amplicons in their linear ranges as follows: atrogin-1/MAFbx (NM_026346; 382-bp product), forward 5'-AGC TTC GTG AGC GAC CTC AG-3' and reverse 5'-TGC CCA CCA GCA CAG ACT TG-3', and MuRF-1 (DQ229108; 405-bp product), forward 5'-ATC TTC CAG GCT GCG AAT CC-3' and reverse 5'-TGG CGT AGA GGG TGT CAA AC-3'. For each sample, 18S rRNA was used as an internal control using QuantumRNA 18S primers (Ambion, Austin, TX).

Albumin-bound fatty acids
Stock solutions of fatty acids with BSA were prepared by dissolving at 37 C for 8 h under a nitrogen atmosphere in KRB buffer containing 5% fatty acid-free BSA. We used a filtered (0.2-µm filter) 5 mmol/liter fatty acid stock solution (pH 7.4) with a molar ratio of fatty acid to BSA of 6:1. The solution of fatty acid bound to BSA was stored at –20 C under nitrogen to prevent oxidation and diluted in culture medium to obtain concentrations of fatty acid ranging from 0.25–1 mM. The fatty acid concentrations in the medium were verified using the NEFA kit (Wako).

Statistical analysis
Results are presented as mean ± the SEM. Densitometric data for blots are expressed as a percentage of the control mean density after normalization to loading controls. To identify significant differences between two groups, comparisons were made by using the Student’s t test. When multiple treatments were compared, ANOVA was performed. Differences with P values < 0.05 were considered significant. The correlation coefficient was calculated by simple regression using GB-stat V10 software (Dynamic Microsystems, Inc., Silver Spring, MD). The coefficient of variation (Cv) was calculated by SD/mean x 100.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
HF diet induces muscle wasting by increasing muscle protein degradation
C57BL/6JLer mice were fed a HF diet, and control mice were fed normal rodent chow for 5 months. As shown in Table 1Go, blood glucose was 1.7-fold (P < 0.01) higher, plasma insulin was 9.9-fold (P < 0.001) higher, serum FFA was 2.6-fold (P < 0.001) higher, plasma adiponectin concentration was 36% (P < 0.001) lower, and body mass index (BMI) was 1.8-fold (P < 0.001) higher in HF-fed mice compared with control mice. The increased levels of blood glucose and plasma insulin were indicative of insulin resistance and type 2 diabetes in the HF-fed mice.


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  		TABLE 1. Physiological parameters

 
The body weights of HF-fed mice were 1.83-fold greater than the control-diet-fed mice at 5 months (Table 2Go, P < 0.001). However, there was a significant decrease by 11% in soleus and by 12% in EDL muscle mass after 5 months of HF feeding (P < 0.05). Likewise, the ratio of muscle weight to body weight was decreased by 51% in soleus and by 53% in EDL muscle of HF-fed mice compared with controls (P < 0.001). To avoid the confounding effects of extracellular volume and FFA deposits in muscle on muscle mass measurements, we also measured muscle protein content by precipitation of soluble protein from muscle lysate. These measurements revealed that muscle soluble protein was decreased 17% in soleus and 23% in EDL muscle in HF-fed mice compared with control mice (P < 0.01).


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  		TABLE 2. Characteristics of HF-fed and control mice

 
Muscle was excised, and the release of tyrosine into the media was measured to determine whether muscle wasting in the HF-fed mice was due to accelerated muscle protein degradation. Based upon these measurements, muscle protein degradation was increased 18% in soleus and 25% in EDL muscles (Fig. 1Go, P < 0.05).


Figure 1
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  		FIG. 1. Protein degradation is elevated in the skeletal muscle of HF-fed mice. Protein degradation was measured by tyrosine release from isolated muscle incubated in standard KRB buffer containing 0.5 mM cycloheximide to block tyrosine reutilization. Values of bar graph are mean ± SE; n = 12 pairs. *, P < 0.05 vs. control.

 
Muscle wasting is associated with low plasma adiponectin levels and high serum FFA concentrations in HF-fed mice
Circulating levels of adiponectin and FFA were determined in HF-fed and control-diet-fed mice. A decrease in plasma adiponectin levels correlated with an increase in serum FFA in HF-fed mice (Fig. 2AGo, r = –0.77; P < 0.001). Unutilized dietary fat can be deposited into nonadipose tissues, causing dysfunction and possible cell death (36). Thus, gastrocnemius muscle samples were stained with oil red-O to examine whether the excess in circulating lipids was deposited in skeletal muscle. As shown in Fig. 2BGo, lipid accumulation was detected in the muscle of the HF-fed mouse but not in control mouse muscle. An increase in BMI correlated with a decrease in muscle weight after 4 months of HF feeding (Fig. 3CGo, r = –0.69; P < 0.01).


Figure 2
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  		FIG. 2. A, Low plasma adiponectin levels correlate with high serum FFA concentrations. Each point represents one animal. Plasma adiponectin (µg/ml) is plotted against Serum FFA (µmol) and analyzed by linear regression analysis. They were significantly correlated (r = –0.77; P < 0.01). B, Fat accumulated in skeletal muscle of HF-fed mice. Gastrocnemius muscles were placed in cryo-mold for embedding and freezing isopentane in dry ice. Fat staining was determined by oil red-O in propylene glycol staining. Skeletal muscle accumulates lipid deposits in HF-fed mice. C, The increase of BMI correlates with the decrease of muscle mass after 4–6 months of HF feeding. The percent increase of BMI and the percent decrease of muscle mass were examined by linear regression and found to be significantly correlated (r = –0.69; P < 0.01). Each point represents one pair of animals (HF-fed and birth-matched control) where the percent change was calculated as the difference between these two animals.

 

Figure 3
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  		FIG. 3. A, Plasma cytokine concentrations are elevated in HF-fed mice. Plasma cytokine levels were measured by the cytokine protein microarray quantification assay in HF-fed and control mice. The bar graph provides the levels of the eight cytokines that increased with HF-fed (black bar) compared with control (white bar) mice. The values of the bar graph show mean ± SE; n = 6. *, P < 0.05 vs. control. GM, Granulocyte-macrophage colony-stimulating factor; MCP, monocyte chemoattractant protein; VEGF, vascular endothelial growth factor. B, The effect of FFA on IL-6 and TNF{alpha} expression in adiponectin-treated adipocytes. 3T3-L1 adipocytes were differentiated and transduced either with adenovirus encoding full-length mouse adiponectin (Adipo) or control virus (Ctrl). Cells were treated with 0.5 mM palmitate (Palm) for 16 h. RT-PCR was used to detect adiponectin, IL-6, and TNF{alpha} expression. The values of the bar graph show mean ± SE; n = 6. *, P < 0.01 vs. Ctrl or Adipo+Palm. Adiponectin and FFA had opposite effects on TNF{alpha} and IL-6 expression.

 
Muscle wasting is associated with increased plasma cytokines in HF-fed mice
Inflammation-induced cytokines have been implicated in muscle protein loss (37). Therefore, we employed a quantitative cytokine protein microarray assay to analyze levels of 15 plasma cytokines in mice. HF-fed mice showed significant increases in eight of the 15 cytokines (Fig. 3AGo, P < 0.05). Of these, TNF{alpha}, IL-6, and interferon-{gamma} (IFN{gamma}) have previously described roles in muscle catabolism (38, 39, 40). Because adiponectin can suppress production of inflammatory cytokines in adipocytes (41), we cultured 3T3-L1 adipocytes and transduced the cells with a recombinant adenovirus (Ad-adipo) that encodes a full-length mouse adiponectin (29) or a control virus (Ad-ctrl). 3T3-L1 cells were treated with FFA (0.5 mM palmitate), and RT-PCR was performed to detect adiponectin, TNF{alpha}, and IL-6 expression. As shown in Fig. 3BGo, expression of adiponectin decreased TNF{alpha} and IL-6 expression in 3T3-L1 adipocytes, whereas exposure to FFA had the opposite effect.

Muscle wasting is associated with down-regulation of p-Akt signaling, resulting in increasing E3 ubiquitin ligase expression in HF-fed mice
In muscle of HF-fed mice, we found that the level of Akt phosphorylation was reduced (Fig. 4AGo). This decrease in Akt signaling corresponds to a decrease in the content of p-FOXO1 Ser253 and Thr32 in the same HF-fed mice. Consistent with the activation of FOXO1 by dephosphorylation, we observed increased expression of the E3 ubiquitin ligases, atrogin-1/MAFbx and MuRF-1 in the muscle of HF-fed mice (Fig. 4BGo). Although AMPK can be a signaling target of adiponectin, the abundance of AMPK and p-AMPK Thr172 did not significantly change in HF-fed mice compared with controls (Fig. 4AGo).


Figure 4
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  		FIG. 4. A, The phosphorylation of Akt and FOXO are reduced in the muscle of HF-fed mice. Akt phosphorylation was measured by a phosphospecific Ser473 polyclonal antibody in hind-limb muscle extracts of HF-fed and control mice. The FOXO phosphorylation was measured using phosphospecific anti-pFOXO1/FOXO3-Thr32 antibody or phosphospecific anti-pFOXO1-Ser253 antibody. The AMPK phosphorylation was measured by phosphospecific Thr172 antibody. The bar graphs of the Western band densities provide means ± SE; n = 6. *, P < 0.01 vs. control. HF feeding reduced Akt phosphorylation and FOXO phosphorylation in skeletal muscle of mice. AMPK phosphorylation was unchanged by HF feeding. B, E3 ubiquitin ligase expressions are elevated in the muscle of HF-fed mice. Atrogin-1 and MuRF-1 expression was measured by Northern blot in the muscle of HF-fed and control mice. The values of the bar graph show mean ± SE; n = 6 pairs. *, P < 0.01 vs. control. These E3 ubiquitin ligating enzymes, atrogin-1 and MuRF-1, increased in HF-fed mice.

 
Adiponectin blocks accelerated protein degradation by up-regulating the IRS-1/Akt signaling pathway in skeletal muscle cells
To determine the effect of adiponectin on protein degradation and IRS-1/Akt signaling, cultured skeletal muscle C2C12 cells were transduced with either Ad-adipo or Ad-ctrl and treated with 0.5 mM BSA-bound palmitate (saturated fatty acid) or oleate (unsaturated fatty acid). Both palmitate and oleate increased the rate of protein degradation (25% by palmitate and 18% by oleate). Adenovirus-mediated gene transfer of adiponectin prevented the accelerated protein degradation in both FFA-treated cells (Fig. 5AGo, P < 0.05). Levels of the 14-kDa actin fragment in muscle can be used as an index of skeletal muscle protein degradation (21). FFA-treated cells have higher levels of the 14-kDa actin fragment, and adiponectin was found to blunt this response to FFA (Fig. 5BGo, P < 0.01).


Figure 5
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  		FIG. 5. A, The effect of FFAs and adiponectin on protein degradation in C2C12 skeletal muscle cells. Cells were transduced with Ad-adipo or Ad-ctrl virus. The rate of protein degradation was measured as the remaining radiolabeled [14C]phenylalanine (log percent) in C2C12 cells incubated for 48 h with 0.5 mM palmitate (Palm) and oleate (Ole). Values are mean ± SE; n = 6 pairs. *, P < 0.05 vs. control. Palmitate and oleate increase protein degradation, and adiponectin attenuated protein breakdown induced by FFAs in skeletal muscle cells. B, The effect of FFA and adiponectin on actin cleavage in C2C12 skeletal muscle cells. Cells were transduced with Ad-adipo or Ad-ctrl for 48 h and treated with 0.5 mM palmitate (Palm) for 16 h. Protein was harvested in hypotonic buffer for Western blots to detect 14-kDa actin fragments. The bar graph shows that the quantity of actin fragments calculated by the density of 14-kDa bands divided by the density of GAPDH bands as a percentage of control values (mean ± SE, n = 6). *, P < 0.01 vs. control or Palm+adipo. Palmitate (Palm)-treated cells had substantially higher levels of the actin fragment compared with control group (Ctrl). Adiponectin (Adipo) attenuated myofibril protein cleavage induced by palmitate.

 
IRS-1/Akt cellular signaling was reduced in muscle cells treated with palmitate (Fig. 6AGo). IRS-1 serine phosphorylation was significantly increased in palmitate-treated cells. Phosphorylation of Ser307 in IRS-1 corresponded to a decrease of total IRS-1 and a reduction in insulin signaling, as indicated by a decrease in p-Akt. Adiponectin overexpression reduced the FFA-mediated Ser307 phosphorylation of IRS-1 and up-regulated tyrosine phosphorylation of IRS-1, leading to an increase in IRS-1/Akt signaling. Correspondingly, adiponectin significantly increased Ser473 phosphorylation of Akt. The effect of palmitate or adiponectin on Akt phosphorylation was dose dependent (Fig. 6BGo). The decrease in p-Akt by palmitate corresponds to an increase in the expression of the E3 ubiquitin ligases, atrogin-1/MAFbx, and MuRF-1 in muscle cells (Fig. 6CGo).


Figure 6
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  		FIG. 6. A, The effect of FFA and adiponectin on IRS-1/Akt signaling pathway in C2C12 cells. The components of IRS-1/Akt signaling pathway were measured in muscle cell extracts by immunoblotting for Akt-1, Akt-2, and p-Akt Ser473 or by immunoprecipitating with an antityrosine (PY-20) antibody followed by Western blotting for IRS-1. For serine 307 phosphorylation of IRS-1, we used a phosphospecific serine 307 antibody. The values of the bar graph show mean ± SE; n = 6 pairs. **, P < 0.001 vs. Ad-ctrl (white bar) and Ad-adipo+palmitate (cross-hatched bar); ##, P < 0.05 vs. Ad-ctrl and Ad-adipo+palmitate; *, P < 0.05 vs. Ad-ctrl; #, P < 0.0001 vs. Ad-adipo+palmitate. Palmitate down-regulated the IRS-1/Akt signaling. Adiponectin diminished the effect induced by palmitate in C2C12 cells. B, The dose-dependent changes of Akt phosphorylation in C2C12 cells. C2C12 cells were treated with different does of palmitate and different titers of Ad-adipo virus. Akt phosphorylation was measured by a phosphospecific Ser473 polyclonal antibody. Palmitate and adiponectin have opposite effects on Akt phosphorylation. C, The effect of FFA and adiponectin on E3 ubiquitin ligase expression in C2C12 skeletal muscle cells. C2C12 cells were treated with 0.5 mM palmitate with or without Ad-adipo virus. Atrogin-1 and MuRF-1 expression was measured by RT-PCR. The values of the bar graph show mean ± SE; n = 6 pairs. *, P < 0.01 vs. Ad-ctrl (white bar) and ad-adipo+palmitate (cross-hatched bar). These E3 ubiquitin ligating enzymes, atrogin-1 and MuRF-1, increased in palmitate-treated cells, and adiponectin diminished the effect induced by palmitate.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Muscle protein loss is a serious complication of several conditions that are associated with insulin resistance (1, 4, 42, 43). Abnormalities in muscle protein metabolism that accompany specific disorders such as obesity-induced insulin resistance has received little attention. Our results suggest that both an increase in FFA and a decrease in adiponectin can trigger muscle protein loss in obesity-induced insulin resistance. We provide evidence in support of the hypothesis that adiponectin inhibits accelerated muscle protein degradation through activation of the IRS-1/Akt signaling.

Patients with type 2 diabetes exhibit increased whole-body protein catabolism, and type 2 diabetic patients treated by hemodialysis have a 2-fold higher rate of protein breakdown compared with age-, gender-, and race-matched nondiabetic hemodialysis patients (9, 42). Whole-body protein catabolism also occurs in obese subjects without diabetes (8, 44) and is linked to increased insulin resistance. Our results suggest that muscle protein degradation occurs in HF-fed mice (present study) and in obese, insulin-resistant db/db mice (1) and that it can be controlled by the balance between adiponectin and FFA.

The perception that adipose tissue generally serves as a fuel reservoir supplying FFA for the energy requirements of other tissues including skeletal muscle has been modified with the realization that adipose tissue also functions as an endocrine organ. Adipose tissue releases adipocytokines, a series of adipocyte-derived biologically active molecules that influence the function of other tissues (45, 46, 47). This is a prime opportunity for adipose-muscle cross talk; i.e. fatty acids and adipocytokines from adipose tissue cause altered signaling pathways (PI3K/Akt) that result in altered muscle protein metabolism. This type of interaction could be important in diabetes or obesity. When visceral adipose mass is expanded, many products are secreted, including FFA, TNF{alpha}, ILs, and complement factors, which reduce insulin sensitivity (48). Our results indicate that serum FFA levels and plasma inflammatory cytokines are associated with an increase in muscle protein degradation (Table 1Go and Fig. 3AGo). These changes are accompanied by accumulation of fat in muscle fibers (Fig. 2BGo). Many studies show that TNF{alpha}, IL-6, and IFN{gamma} influence muscle catabolism (38, 39, 40). TNF{alpha} can up-regulate the expression of the E3 ubiquitin ligase atrogin-1/MAFbx in skeletal muscle (38). Modest levels of IL-6 infused into animals causes significant muscle atrophy (39). Administration of IFN{gamma} to normal animals increases ubiquitin mRNA expression (49) and the transcription of selected proteasome subunits in skeletal muscle (50). In contrast to the higher plasma level of these inflammatory cytokines, the plasma level of adiponectin is markedly reduced (Table 1Go).

Adiponectin enhances the insulin signal transduction cascade by increasing the tyrosine phosphorylation of IRS-1 in human skeletal muscle (51). Up-regulation of IRS-1 activates the Akt cascade, which phosphorylates members of the forkhead transcription factor family (52, 53). The FOXO proteins increase transcription of atrogin-1/MAFbx or MuRF-1, the E3 ubiquitin ligases that are closely associated with an increase in muscle protein degradation (19). Consequently, adiponectin can be considered as a candidate that limits protein losses from muscle. Alternatively, a decrease in IRS-1/Akt signaling such as the reduction in Akt-1 phosphorylation in response to a HF diet (Fig. 4AGo) or FFA treatment of cultured muscle cells (Fig. 6AGo) will be associated with an increased loss of muscle protein. We found that adiponectin slightly increased the amount of Akt-2, but not to a statistically significant level. There are three Akt isoforms (Akt-1, -2, and -3) encoded by distinct genes. In general, Akt-1 regulates growth and Akt-2 regulates glucose homeostasis. According to McCurdy and Cartee (54), Akt2 activation is tightly associated with insulin sensitivity. However, in skeletal muscle, Akt-1 is the most responsive to cellular stimulation in our studies and others (55). Thus, the differential regulation of Akt-1 and Akt-2 by adiponectin requires additional studies. A decrease in p-Akt resulted in augmentation of atrogen-1/MAFbx and MuRF-1 expression (Fig. 6CGo) and increased muscle protein degradation in HF-fed mice and cultured muscle cells. We hypothesize that adiponectin diminishes the accelerated protein degradation induced by FFA (Fig. 5AGo) due to the up-regulation of the IRS-1/Akt signaling pathway.

Adiponectin binding to its receptors activates multiple signaling pathways, including AMPK. The AMPK system is a regulator of energy balance at both the cellular and whole-body levels (56). AMPK is a critical regulator involved in initiating mitochondrial biogenesis (57, 58). Regarding protein metabolism, some studies show that activation of AMPK suppresses protein synthesis through down-regulation of the mammalian target of rapamycin (mTOR) signaling (59). Another study shows that AMPK is elevated after resistance exercise and associated with increasing p70S6K phosphorylation (60). There also are reports that activation of AMPK by activator of AMPK (AICAR) increases protein degradation (61). In our study, we did not find a change in the phosphorylation or protein level of AMPK in HF-fed mice compared with controls, suggested that HF feeding causes muscle protein loss through an adiponectin-dependent but most likely AMPK-independent pathway (62). A strong candidate for this mediator is IRS-1/Akt signaling.

Although FFA are known to play a key role in the etiology of insulin resistance, the molecular mechanisms by which high levels of FFA induce insulin resistance have remained obscure. Our data showing that increased FFA and inflammatory cytokines lead to insulin resistance and muscle wasting is consistent with a recent report that the increase in FFA levels can promote the toll-like receptor 4 (TLR4) inflammatory signaling pathway, which in turn contributes to obesity-dependent insulin resistance (63). TLR4 is the receptor for lipopolysaccharide. Stimulation of TLR4 activates proinflammatory pathways and induces cytokine expression in a variety of cell types, including muscle (64). Shi and his colleagues (63) show that increases in nutritional fatty acids activate TLR4 and induce inflammatory signaling in adipocytes and macrophages. The capacity of fatty acids to induce inflammatory signaling in adipose cells or tissue and macrophages is blunted in the absence of TLR4. What could be the underlying mechanism for adiponectin prevention of FFA-induced protein degradation? One potential mechanism is that adiponectin may inhibit FFA-induced TLR4 activation. Inhibition of the TLR4-dependent signaling pathway could reduce inflammatory cytokine production and limit or reverse the inactivation of the IRS-1/Akt signaling pathway. The measurement of protein degradation used in this study is very sensitive to the presence of an adequate energy source in the assay system. In the isolated muscle studies, we used glucose rather than exogenous FFA as the energy source because of concerns of inadequate transfer of FFA from carrier proteins to muscle or diffusion of FFA into muscle cells. It has been shown that rates of protein synthesis and degradation in muscles incubated for 2 h with glucose as the energy source were almost identical to rates measured over a similar period in muscle perfused with red blood cells, an artificial plasma, and glucose (65). These data suggest that the technique employed in our study accurately reflects changes in protein metabolism that are stimulated by in vivo conditions such as feeding lipids. The evidence that isolated muscles retain the biochemical responsiveness that has been established by in vivo metabolic conditions (memory) has been documented in a number of studies of muscle catabolism including acute uremia (66), chronic uremia (30), acute diabetes (67), and type 2 diabetes (1).

In conclusion, we found that HF feeding increased muscle protein catabolism, decreased plasma adiponectin levels, and down-regulated IRS-1/Akt signaling. Conversely, an elevation of adiponectin blocked the accelerated muscle protein degradation, and this was associated with the up-regulation of IRS-1/Akt signaling in cultured myotubes. Increasing adiponectin also blocked inflammatory cytokine expression in 3T3-L1 adipocytes. Our results suggest that muscle protein degradation present in HF-fed mice (present study) and in obese, insulin-resistant (db/db) mice (1) is impacted by the balance between adiponectin, an adipocyte-derived plasma protein, and fatty acids. These findings provide additional evidence of adipose tissue and muscle cross talk.


   Footnotes
 
This study was supported in part by the University Research Committee of Emory University, Norman S. Coplon Extramural Research Grant from Satellite Health, American Diabetes Association Junior Faculty Award 1-04-JF-48, and National Institutes of Health (NIH) DK62796 to X.H.W. and NIH HL70762 to J.D.

Disclosure Statement: Q.Z., J.D., Z.H., and X.H.W. have nothing to declare. K.W. consults for, has received lecture fees from, and has equity ownership in Nave Bioscience, Inc. (less than U.S. $10,000).

First Published Online August 30, 2007

Abbreviations: AMPK, AMP-activated protein kinase; BMI, body mass index; Cv, coefficient of variation; EDL, extensor digitorum longus; FFA, free fatty acids; FOXO, forkhead; HF, high fat; IFN{gamma}, interferon-{gamma}; IRS-1, insulin receptor substrate-1; KRB, Krebs-Ringer bicarbonate; MAFbx, muscle atrophy F-box; MuRF-1, muscle RING finger-1; NEFA, nonesterified fatty acids; p-Akt, phosphorylated Akt; PI3K, phosphatidylinositol-3-kinase; TLR4, toll-like receptor 4.

Received February 8, 2007.

Accepted for publication August 21, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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