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Physiol. Genomics 34: 149-161, 2008. First published April 29, 2008; doi:10.1152/physiolgenomics.00281.2007
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Received 6 December 2007; accepted in final form 25 April 2008.
Physiological Genomics 34:149-161 (2008)
1094-8341/08 $8.00 © 2008 American Physiological Society

Muscle unloading-induced metabolic remodeling is associated with acute alterations in PPAR{delta} and UCP-3 expression

Dawn J. Mazzatti1, Melissa A. Smith2, Radu C. Oita1, Fei-Ling Lim3, Andrew J. White3 and Michael B. Reid2

1 Unilever Corporate Research, Colworth Park, Sharnbrook, Bedfordshire, United Kingdom
2 Department of Physiology, University of Kentucky, Lexington, Kentucky
3 Unilever Measurement Sciences, Colworth Park, Sharnbrook, Bedfordshire, United Kingdom


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
A number of physiological changes follow prolonged skeletal muscle unloading as occurs in spaceflight, bed rest, and hindlimb suspension (HLS) and also in aging. These include muscle atrophy, fiber type switching, and loss of the ability to switch between lipid and glucose usage, or metabolic inflexibility. The signaling and genomic events that precede these physiological manifestations have not been investigated in detail, particularly in regard to loss of metabolic flexibility. Here we used gene arrays to determine the effects of 24-h HLS on metabolic remodeling in mouse muscle. Acute unloading resulted in differential expression of a number of transcripts in soleus and gastrocnemius muscle, including many involved in lipid and glucose metabolism. These include the peroxisome proliferator-activated receptors (PPARs). In contrast to Ppar-{alpha} and Ppar-{gamma}, which were downregulated by acute HLS, Ppar-{delta} was upregulated concomitant with increased expression of its downstream target, uncoupling protein-3 (Ucp-3). However, differential expression of Ppar-{delta} was both acute and transient in nature, suggesting that regulation of PPAR{delta} may represent an adaptive, compensatory response aimed at regulating fuel utilization and maintaining metabolic flexibility.

hindlimb suspension; metabolic flexibility; peroxisome proliferator-activated receptor; gene expression; uncoupling protein-3; fuel utilization; microarray


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
DECREASED PHYSICAL ACTIVITY associated with a sedentary lifestyle, bed rest, spaceflight, and hindlimb suspension (HLS) leads to many debilitating alterations in skeletal muscle including a dramatic reduction in soleus muscle mass and concomitant loss of function. These physiological changes manifest in microcirculatory disturbances, atrophy, protein loss, changes in contractile properties, and fiber type switching, all of which have been extensively investigated. In addition, several studies have attempted to characterize the biochemical and metabolic processes that underlie the physiological alterations associated with skeletal muscle unloading. Disturbances in inter- and intracellular signaling, transcriptional regulation, proteolysis, protein metabolism, molecular transport, and energy metabolism have all been described after HLS (reviewed in Refs. 31, 73).

In contrast to the widely investigated effects of HLS on muscle structure, less attention has been paid to understanding the metabolic consequences of unloading. Unloading-induced muscle fiber type switching is associated with an increased reliance on glucose and a corresponding decreased use of lipid as a fuel source. Loss of the ability to switch between lipid and carbohydrate usage in periods of fasting and after insulin stimulation (e.g., food intake), respectively—a condition termed metabolic inflexibility—is a key feature of the metabolic disturbance associated with bed rest (1, 57), aging (5, 6a), obesity (29), insulin resistance syndrome (38), and diabetes (37, 76). Furthermore, multiple studies have demonstrated increased substrate-level activation of glycolysis and inhibition of fatty acid oxidation in unloaded skeletal muscle (3, 30). These physiological changes were associated with decreased activity of cytochrome oxidase, β-hydroxyacyl CoA dehydrogenase, and citrate synthase, key components of oxidative metabolism, following unloading (53), concomitant with decreased mitochondrial respiration (81). Together, these data indicate that metabolic flexibility is lost in maladapted, unloaded skeletal and cardiac muscle.

The maladaptive metabolic responses following unloading occur because of alterations in signals that control transcription of key regulatory genes. These metabolic alterations include reduced expression of genes involved in fatty acid oxidation and decreased transcription of genes involved in lipid metabolism corresponding with increased expression of glycolytic enzymes (68, 80). These genomic changes result in increased glycogen synthesis concomitant with increased reliance on glucose—rather than lipid—as an energy source (30, 32, 43). An investigation of the temporal regulation of physiological changes following muscle unloading demonstrated that these metabolic changes preceded the reduction of muscle mass (43), suggesting that alterations in the regulation of fuel switching (or metabolic inflexibility) may underlie the muscle unloading phenotype. Similarly, it has been proposed that metabolic remodeling including loss of metabolic flexibility precedes the functional and structural remodeling of the unloaded heart (71). However, few studies have attempted to elucidate the genomic mechanisms that are involved in initiating the early changes in gene expression that may be responsible for subsequent disturbances in metabolic flexibility and fuel utilization and, ultimately, result in the unloading phenotype.

The peroxisome proliferator-activated receptors (PPARs) are nuclear receptors with pleiotropic biological functions that include regulation of inflammation, lipid metabolism, and whole body fuel usage. These nuclear receptors are widely studied in the context of energy metabolism because two of three family members (PPAR{alpha} and PPAR{gamma}) are molecular targets for pharmaceutical treatment of metabolic disease. Although PPAR{delta} (also referred to as PPARβ) is the most highly expressed PPAR in muscle (51), it initially attracted less interest because of its ubiquitous expression and lack of synthetic agonist (10). However, development of selective agonists of PPAR{delta} led to the discovery of broad functions in early development, wound healing, and regulation of signaling pathways that are fundamental in the development of insulin resistance and cardiovascular disease, including regulation of glucose and lipid metabolism. PPAR{delta} agonists are known to improve lipid profile and sensitivity to insulin as well as increase whole body fat burning, lipid oxidation, and energy dissipation. Because of its proposed role in regulating genes involved in lipid catabolism, energy uncoupling, and fuel switching (17, 27) we hypothesize that PPAR{delta} expression may be altered as an adaptive response following metabolic disturbances caused by unloading and that changes in its expression following long-term unloading may be an important contributor to loss of metabolic flexibility and the subsequent physiological changes observed.

In the present study, Agilent whole-genome arrays were utilized to examine the effects of acute 24-h mouse HLS on soleus and gastrocnemius mRNA expression profiles. The acute exposure of 24-h HLS was chosen because atrophy and fiber type switching mediated by acceleration in proteolytic processes occur after at least 3 days of HLS (3, 34, 73) and thus 24 h represents a time point prior to these physiological alterations. Additionally, it was demonstrated previously that after 24-h HLS there are no changes in the expression of genes defining fast and slow contractile muscle phenotypes (69). However, both glucose and lipid metabolism are altered at 24 h (43), suggesting that this is a reasonable time point at which to investigate the molecular events that coordinate later physiological events. This investigation aimed to determine coordinated regulation of gene expression in the two muscle types, thereby obtaining a more global view of the musculoskeletal aspects of acute HLS. Because of the proposed roles of nuclear hormone receptors—such as PPARs and in particular PPAR{delta}—in regulating lipid metabolism and oxidative capacity (35, 48), we further assessed whether expression of components of the PPAR signaling pathway and PPAR-interacting proteins are altered after acute HLS. It is envisioned that a greater understanding of the genes involved in the early adaptive metabolic responses to HLS, including the PPARs and specifically PPAR{delta}, may allow the discovery of novel targets for prevention of the adverse physiological effects of muscle unloading and physical inactivity.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Experimental Animals
All experiments in this study were in agreement with the guiding principles on care and use of laboratory animals of the American Physiological Society and the National Institutes of Health. Animals used for this study were approved in advance by the Institutional Animal Care and Use Committee of the University of Kentucky Medical Center. Male ICR mice (28- to 32-g body wt; Harlan, Indianapolis, IN) were maintained on a reversed 12:12-h light-dark cycle with controlled temperature (21 ± 1°C) and humidity. Food and water were provided ad libitum. One week after arrival, the animals were separated into individual cages for a 3-day acclimation period and then were randomly assigned to either of two experimental groups: freely ambulating control animals and hindlimb-unloaded animals.

Hindlimb Unloading
Hindlimb muscles were unloaded by methods described previously (47). Briefly, Elastoplast tape (Beiersdorf-Jobst, Rutherford College, NC) was used to wrap the tail of each animal. A metal clip on the tape was attached to a nylon monofilament line via a stainless steel swivel. The distal end of the nylon line was attached to an overhead support and shortened to suspend the animal in a 45° head-down tilt position. The swivel enabled the animal to explore the cage (360° range of motion) and obtain food and water freely. Animals were observed daily for changes in appearance and activity. Each animal was weighed, and the angle of HLS was adjusted if necessary. After 1 day or 12 days, each animal was deeply anesthetized in the hindlimb-unloaded condition. The animal was removed from the unloading device, the soleus and gastrocnemius muscles were excised with the animal under surgical anesthesia, and the animal was euthanized. Muscle was immediately placed in RNAlater (Ambion, Foster City, CA) for subsequent analyses.

Microarray Analysis
All products were purchased from Agilent Technologies UK (Wokingham, UK) unless otherwise noted.

Target RNA preparation.
Total RNA was isolated with the RNeasy kit (Qiagen, Crawley, UK). The integrity of the RNA was confirmed with analysis by the Agilent 2100 bioanalyzer (Palo Alto, CA) and the RNA 600 LabChip kit.

Labeling.
Six hundred nanograms of RNA and three microliters (1:2,500 dilution) of Agilent One-Color RNA Spike-In RNA were labeled with the Agilent Low RNA Input Linear Amplification Kit PLUS, One-Color according to the manufacturer's instructions as follows: 1.2 µl of T7 Promoter Primer was added to 600 ng of RNA and 3 µl of spike-in control (in a 11.5-µl volume) and denatured at 65°C. First-Strand Buffer (to 1x), DTT (to 10 mM), dNTP (to 0.5 mM), Moloney murine leukemia virus (MMLV) reverse transcriptase (1 µl of stock provided in the kit in a 20-µl reaction), and RNaseOut (0.5 µl of stock provided in the kit in a 20-µl reaction) were added. The cDNA was synthesized during the following incubation step (2 h at 40°C). After 10-min denaturation at 65°C and the addition of Cy-labeled CTP (to 0.3 mM), Transcription Buffer (to 1x), DTT (to 10 mM), NTP (8 µl of stock provided in the kit in a 80-µl reaction), polyethylene glycol (to 4%), RNaseOUT (0.5 µl of stock provided in the kit in a 80-µl reaction), inorganic phosphatase (0.6 µl of stock provided in the kit in a 80-µl reaction), and T7 RNA polymerase (0.8 µl of stock provided in the kit in a 80-µl reaction), the synthesis of the fluorescent labeled cRNA was performed during the second incubation step (2 h at 40°C). The labeled cRNA was purified with the RNeasy Mini Kit (Qiagen) according to the manufacturer's protocol.

Hybridization and scanning.
The Agilent Hybridization Kit (catalog no. 5188-5242) was used in conjunction with Agilent Mouse Oligo Arrays (catalog no. G4122F). Two micrograms of the labeled sample RNA was used for hybridization according to the Agilent One-Color Microarray-Based Gene Expression Analysis Protocol. The hybridization was performed for 17 h at 65°C and 10 rpm. Slides were them washed for 1 min at 22°C in Wash Solution 1 (catalog no. 5188-5325) and 1 min at 22°C in Wash Solution 2, prewarmed to 37°C (catalog no. 5188-5326). Slides were incubated for 30 s in Agilent Stabilization and Drying Solution (catalog no. 5185-5979). The slides were scanned with the Agilent G2565BA Microarray Scanner System.

Data extraction and deposition into Gene Expression Omnibus.
For data extraction and quality control, Agilent G2567AA Feature Extraction Software (v.9.1) was used. Data files were deposited into the NCBI Gene Expression Omnibus (GEO) in order to comply with MIAME requirements. The following link was created to allow review of these data: http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?token=hfsfbmsqsgwievs&acc=GSE9802.

Bioinformatics analysis.
Extracted data were analyzed with GeneSpring GX 7.3.1 (Silicon Genetics). Agilent standard scenario normalizations for FE1-color arrays were applied to all data sets. A subset of genes for data interrogation was generated that excluded controls, spots of poor quality, and gene probes that were present in <50% of samples. From these selected genes, relative expression in hindlimb-unloaded samples compared with ambulatory samples was determined. Genes differentially regulated by >1.5-fold were selected. One-way parametric ANOVA tests were performed, followed by Benjamini and Hochberg multiple test correction with a false discovery rate of 0.05. Microsoft Excel templates were prepared containing genes that were over- and underexpressed in soleus and gastrocnemius muscle after hindlimb suspension. Ingenuity Pathway Analysis 3.0 was utilized to assemble functional networks of altered hindlimb unloading (Ingenuity Systems).

Real-Time Quantitative Reverse-Transcription PCR
RNA was prepared from cells as described above, and cDNA synthesis was performed with 1 µg of total RNA and random hexamer primers (Invitrogen, Paisley, UK). Taqman (Applied Biosystems, Applera) probes were used for RT-PCR: peroxisome proliferator activator receptor delta: Mm00803186_g1; peroxisome proliferator activated receptor gamma: Mm00440945_m1; peroxisome proliferator activated receptor alpha: Mm00440939_m1; peroxisome proliferator activated receptor, gamma, coactivator 1 beta: Mm01258518_m1; peroxisome proliferator activated receptor, gamma, coactivator 1 alpha: Mm00447183_m1; retinol binding protein 7, cellular: Mm00458145_m1; retinoic acid receptor, alpha: Mm00436264_m1; uncoupling protein 3 (mitochondrial, proton carrier): Mm00494074_m1; YKT6 homolog (Saccharomyces cerevisiae) snare protein: Mm00457727_m1. The Bio-Rad I-Cycler (Bio-Rad, Hercules, CA) with FAM-490 system detection was used for real-time RT-PCR. PCR thermocycler conditions were 50°C for 2 min, 90°C for 2 min, followed by 45 cycles of 95°C for 15 s and 60°C for 60 s. All samples were run in triplicate with both primer sets and the control gene mouse snare to control for differences in amount of starting material. A standard curve was created for each PCR reaction in order to determine amplification efficiency. Fold changes were calculated with the 2Formula method. Statistical analyses were performed with Student's t-test. A P value of <0.05 was considered significant.

Protein Analysis
For Western blot analysis, soleus and gastrocnemius tissue samples were homogenized and lysed in sample buffer [50 mM Tris, pH 8.0, 120 mM NaCl, 0.5% NP-40, 10 µg/ml phenylmethylsulfonyl fluoride (PMSF), and 1x protease inhibitor cocktail (Sigma, St. Louis, MO)]. A total of 30 µg of protein was suspended in Laemmli buffer (50 mM Tris, pH 6.8, 1% β-mercaptoethanol, 2% sodium dodecyl sulfate, 0.1% bromophenol blue, and 10% glycerol). Protein samples were boiled, loaded on 10% NuPAGE Bis-Tris gels (Invitrogen) with MES running buffer, run according to the instructions of the manufacturer, and transferred to nitrocellulose membranes (Schleicher and Schuell, Dassel, Germany). UCP-3, PPAR{delta}, and β-actin proteins were detected with polyclonal antibodies (Abcam, Cambridge, UK). Proteins were visualized by enhanced chemiluminescence (GE Healthcare, Little Chalfont, UK).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Identification of HLS-Induced Early Responsive Genes and Ontologies
The primary aim of this study was to investigate the effects of acute mouse HLS on genes and proteins involved in regulation of glucose and lipid metabolism. Our previous investigations (2, 24, 47) demonstrated that mice are a reasonable laboratory model to investigate the effects of HLS because they display a phenotype similar to other animal species after muscle unloading (reviewed in Ref. 74) and they additionally can be genetically manipulated. To examine the effect of acute HLS on gene expression profiles, RNA samples from gastrocnemius and soleus muscle from five control and five 24-h HLS animals were subjected to whole-genome microarray analysis. Stringent data analysis was performed in order to ensure that the genes identified through microarray analysis were truly differentially expressed. After exclusion of absent calls and probes that were expressed in less than half of experimental samples, 35,282 of an initial 41,267 probes (85%) were considered for analysis. Only probes with a differential expression of at least 1.5-fold and that passed ANOVA testing at P < 0.05 were considered to be truly differentially expressed after HLS. Gene expression profiling of control and unloaded muscle demonstrated that a total of 1,451 and 1,065 genes were differentially expressed in soleus and gastrocnemius muscle, respectively, after HLS. Of these probes, in soleus muscle 417 were upregulated and 1,034 were downregulated, while in gastrocnemius muscle 318 were upregulated and 747 were downregulated by hindlimb unloading. Figure 1A shows the number of genes up- and downregulated in gastrocnemius and soleus muscle by Venn diagram. Although 1,451 and 1,065 probes were differentially regulated in soleus and gastrocnemius muscle after unloading, only 600 were regulated in both muscle types, while 851 and 465 were regulated in a tissue-specific manner, respectively. The 600 commonly significantly regulated probes were subjected to gene cluster analysis using a Pearson correlation. A heat map was generated in which each row represents 1 of 600 probes and each column represents 1 sample. Expression of the 600 probes in 5 soleus and 5 gastrocnemius muscle samples after hindlimb unloading is shown in Fig. 1B (normalized to the average expression of ambulatory controls). The probes are ordered by tree clustering and significance, where average expression of each probe does not correlate between gastrocnemius and soleus samples (Fig. 1B, top), average expression is downregulated in both gastrocnemius and soleus (middle), and average expression is upregulated in both gastrocnemius and soleus muscle after unloading (bottom). The intensity of color in a cell represents the normalized expression of the probe, while green and red coloring depict low and high expression, respectively.


Figure 1
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  		Fig. 1. Venn analysis and hierarchical clustering of hindlimb suspension (HLS)-sensitive gene probes. Gene expression profiling of control and unloaded muscle demonstrated that a total of 1,451 and 1,065 genes were differentially expressed in soleus and gastrocnemius muscle, respectively, after HLS (>1.5-fold, P < 0.05). Of these probes, in soleus muscle 417 were upregulated and 1,034 were downregulated while in gastrocnemius muscle 318 were downregulated and 747 were upregulated by hindlimb unloading. A: Venn analysis found that although 1,451 and 1,065 probes were differentially regulated in soleus and gastrocnemius muscle after unloading, 600 were regulated in both muscle types, while 851 and 465 were differentially regulated in a tissue-specific manner, respectively. B: the 600 gene probes differentially regulated in both soleus and gastrocnemius muscles were subjected to gene cluster analysis using a Pearson correlation. A heat map was generated in which each row represents 1 of 600 probe sets and each column represents 1 sample. Expression of the 600 probes in soleus and gastrocnemius muscle samples (n = 5 animals for each group and tissue type) after hindlimb unloading compared with control is shown. The probes are ordered by correlation and significance. The intensity of color in a cell represents the normalized expression of the probe, where green and red depict low and high expression, respectively.

 
Biological processes affected by unloading were investigated initially with Gene Ontology (GO) analysis of the genes significantly regulated by HLS in both soleus and gastrocnemius muscle. Table 1 shows the 10 most significantly regulated GO biological processes in muscle ordered by P value. The most significant processes affected by HLS are muscle and musculoskeletal function and protein synthesis. Of the entire list of ontologies significantly altered by unloading, the 18 selected ontologies functionally regulating glucose and lipid metabolism that were altered by HLS are depicted in Table 2, with corresponding P values. Multiple lipid and glucose metabolic functions were altered in muscle after HLS. Of interest, one of the main metabolism-related ontologies affected by HLS was the gluconeogenesis pathway. On further inspection, it appeared that the expression of three enzymes that are involved in the synthesis of 2-phosphoglycerate, a substrate for gluconeogenesis, was found to be downregulated in HLS. The enzyme that subsequently converts 2-phosphoglycerate to 3-phosphoglycerate, phosphoglycerate mutase, was also found to be decreased. However, paradoxically, we observed increased expression of one of the rate-limiting enzymes of gluconeogenesis, fructose-1,6-bisphosphatase, while the expression of two other critical enzymes, pyruvate carboxylase and phosphoenolpyruvate carboxykinase, was unchanged by HLS. Since the liver is the major organ that contributes to whole body gluconeogenesis, the relevance of these changes in gene expression in skeletal muscle—in the absence of correlated changes in liver—is unclear. It is of course important to note that many of the same metabolic genes appear in multiple ontologies. For example, both PPAR{alpha} and peroxisome proliferator-activated receptor {gamma} coactivator (PGC)-1{alpha} appear in the multiple glucose-related and lipid ontologies. Together, these data suggest that HLS-dependent regulation of metabolic pathways is complex and that changes observed 24 h after HLS may differ from those observed at later time points, at which time substantial metabolic remodeling and adaptation has occurred.


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  		Table 1. Top 10 most significantly regulated ontologies after unloading (sorted by P value)

 

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  		Table 2. Significant glucose and lipid metabolic ontologies regulated by HLS in soleus muscle

 
Signaling Pathways and Cellular Functions Altered by Acute HLS
Biological processes are not equivalent to molecular pathways; at present, GO does not represent the dynamics or interactions that would be required to fully describe a biological pathway. Therefore, to gain an understanding of which functional cellular pathways and processes may be coordinately regulated during acute HLS, the 600 genes that were commonly significantly altered in both soleus and gastrocnemius muscle after unloading were visualized by Ingenuity Pathway Analysis (IPA). The Ingenuity entry tool systematically encodes findings presented in peer-reviewed scientific publications into ontologies, or groups of genes/proteins related by common function. Molecular networks of direct physical, transcriptional, and enzymatic interactions were computed from this knowledge base with the 600 genes differentially regulated after HLS. The resulting networks contain molecular relationships with a high degree of connectivity, and every gene in the network is supported by published literature. A total of 19 networks were significantly coregulated in soleus and gastrocnemius muscle after HLS (P < 0.001). Table 3 shows the score (related to the probability of each network being assembled by chance alone), number of HLS-regulated genes (focus genes), and main cellular functions or conditions/diseases in which these genes play a role for each of these 19 networks.


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  		Table 3. Networks significantly regulated in soleus and gastrocnemius muscle after HLS (P < 0.001) as determined by Ingenuity Pathway Analysis tool

 
From the 19 networks significantly affected by HLS, the 1 highly statistically significant molecular network involved in carbohydrate and lipid metabolism was visualized with the Ingenuity tool. This network assembled with IPA profiling of genes differentially expressed in soleus and gastrocnemius muscle after HLS, including interacting partners, is depicted in Fig. 2, A (soleus) and B (gastrocnemius). Genes demonstrated to be differentially upregulated or downregulated after microarray analysis are colored in red and green, respectively. Genes that are uncolored were not found to be significantly altered in gastrocnemius and soleus muscle after HLS. The class of function for each of the genes, including enzymes, growth factors, and receptors, is depicted by nodal shape. The fold change in gene expression compared with ambulatory animals is shown below each colored node in Fig. 2. Central nodes in this metabolic pathway include the retinoid X receptor (RXR), which heterodimerizes with PPARs and is required for transcriptional regulation of PPAR target genes, in addition to PPAR{alpha}, and PPAR-regulated proteins involved in mitochondrial biogenesis and metabolic regulation: PGC-1{alpha} and PGC-1β. While RXR was not differentially regulated by unloading, PPAR{alpha}, PGC-1{alpha}, and PGC-1β were all downregulated in both soleus and gastrocnemius muscle after HLS. These data suggested that the PPAR signaling cascade was affected by unloading.


Figure 2
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  		Fig. 2. Analysis of HLS-regulated lipid and glucose metabolism molecular network. Differentially expressed genes from analysis of soleus and gastrocnemius muscle after HLS were analyzed by the Ingenuity Pathway Analysis (IPA) tool. The most significant network involved in glucose and lipid metabolism is shown as regulated by HLS in soleus (A) and gastrocnemius (B) muscle. Differentially regulated genes are shown in red and green, depicting up- and downregulation after HLS (compared to ambulatory control), respectively. Bold color indicates a high degree of regulation (>2.5-fold), while pale color indicates 1.5- to 2.5-fold differential regulation. Noncolored genes directly or indirectly associated with the differentially expressed genes were not found to be differentially expressed after treatment. Positive regulatory interactions are marked by solid arrows (direct interactions) or dashed arrows (indirect interactions). Negative interactions are shown by inhibitory arrows.

 
Because of the proposed roles of PPARs, in particular PPAR{delta}, in regulating lipid metabolism and oxidative capacity (35, 48) we focused our analysis on the regulation of the PPAR signaling pathway after acute HLS. Several PPAR family members were found to represent nodal points in the networks identified as altered after HLS. Because of their important role in lipid homeostasis, we further investigated the affect of HLS on the PPAR signaling pathway in detail, using the Ingenuity Canonical Pathway tool to better understand the biological significance of the altered individual genes. The canonical PPAR signaling pathway is depicted in Fig. 3. Genes demonstrated to be significantly upregulated or downregulated after microarray analysis of soleus (Fig. 3A) or gastrocnemius (Fig. 3B) muscle are colored in red and green, respectively. Genes that are uncolored were not found to be significantly altered in gastrocnemius or soleus muscle after HLS. The classes of function for each of the genes, including enzymes, growth factors, and receptors, are depicted by nodal shape. Interestingly, while Ppar-{alpha} is downregulated by HLS in both soleus and gastrocnemius muscle, several other global regulatory genes, namely, signal transducer and activator of transcription 5 (Stat5), inhibitor of NF{kappa}B (I{kappa}B), and inhibitor of I{kappa}B (I{kappa}K) are differentially regulated in soleus and gastrocnemius muscle, demonstrating tissue-specific effects of unloading.


Figure 3
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  		Fig. 3. Peroxisome proliferator-activated receptor (PPAR) canonical pathway in soleus and gastrocnemius muscle after HLS. Genes that were differentially expressed in soleus and gastrocnemius muscle after HLS were clustered into canonical pathways with the Ingenuity Canonical Pathway tool. The PPAR signaling pathway was significantly regulated by HLS in both soleus (A) and gastrocnemius (B) muscle (P < 0.001). Colored genes were identified by microarray analysis as differentially upregulated or downregulated in HLS samples compared with control (red and green, respectively). Other uncolored nodal genes are directly or indirectly associated with the differentially expressed genes but were not found to be significantly regulated by HLS. Genes are linked by their subcellular location (extracellular, cytoplasmic, nuclear; from top to bottom, respectively). Main processes modulated by transcriptional regulation are shown.

 
Expression of PPARs and PPAR-Interacting and PPAR-Regulated Genes
To validate results obtained with the microarrays and to further assess the extent to which the PPARs and PPAR-interacting proteins are altered after HLS, we assessed mRNA expression by real-time RT-PCR. Figure 4A shows the average mRNA expression of Ppars after unloading of gastrocnemius and soleus muscle, compared with control. While both Ppar-{alpha} and Ppar-{gamma} were downregulated after HLS, RT-PCR analysis revealed that Ppar-{delta} was upregulated by HLS.


Figure 4
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  		Fig. 4. mRNA expression of selected genes involved in metabolic flexibility and energy metabolism. RNA was prepared, and in vitro transcription was performed with 1 µg of total RNA and random hexamer primers (Invitrogen, Paisley, UK). The Bio-Rad I-Cycler (Bio-Rad, Hercules, CA) with FAM-490 system detection was used for real-time RT-PCR. Taqman (Applied Biosystems, Applera) probes were used as detailed in MATERIALS AND METHODS. Animals were grouped into ambulatory and HLS conditions, and mRNA expression for each animal was averaged for each gene analyzed for both soleus and gastrocnemius muscle. All samples were run in triplicate with test probes and the control gene mouse SNARE to control for differences in amount of starting material. Fold change in expression was calculated by normalizing the test gene crossing threshold (Ct) with the SNARE amplified control and then comparing to gene expression in ambulatory control animals. A: mRNA expression of differential regulation of PPAR{alpha}, -{gamma}, -β/{delta} in soleus and gastrocnemius muscle after HLS. B: mRNA expression of PPAR-regulated genes: peroxisome proliferator-activated receptor {gamma} coactivator (PGC)-1{alpha}, PGC-1β, muscle-specific carnitine palmitoyl transferase (CPT)-1b, CPT-2, and UCP-3 in soleus and gastrocnemius muscle after HLS. C: mRNA expression of acetyl-CoA carboxylate (ACC; {alpha} and β isoforms) and AMP-activated protein kinase (AMPK; {alpha}2, β2 subunits) in soleus and gastrocnemius muscle after HLS. n = 5 animals for each group and tissue type. *P < 0.05, **P < 0.01 by ANOVA.

 
PPAR{delta} was shown previously to directly and/or indirectly modulate the expression of proteins involved in fatty acid oxidation [carnitine palmitoyl transferase (CPT)1 and 2], lipid utilization [pyruvate dehydrogenase kinase 4 (PDK4)], lipid efflux (ABCA1), and energy expenditure (UCP-1, -2, -3) (23). We next assessed the expression of PPAR{delta}-regulated genes that may impact on metabolic flexibility by RT-PCR: Pgc-1{alpha}, Pgc-1β, muscle-specific CPT-1b, CPT-2, and UCP-3 (Fig. 4B). In agreement with microarray data, we observed decreased expression of Pgc-1{alpha} and Pgc-1β after unloading. This is not surprising, because these two genes are regulated by PPAR{gamma}—which was significantly downregulated by HLS—as well as by PPAR{delta}. Interestingly, we observed increased expression of the three genes specifically regulated by PPAR{delta}, Cpt-1b, Cpt-2, and Ucp-3, in soleus muscle after HLS. However, only Cpt-2 was significantly regulated by HLS in gastrocnemius muscle, while the other genes are significantly regulated by HLS only in soleus muscle, again demonstrating the tissue-specific effects of unloading.

We further assessed the expression of several genes that are important in lipid and glucose metabolism. Krämer et al. (42) recently demonstrated that the effects of PPAR{delta} on metabolic responses including regulation of glucose and lipid utilization may involve the AMP-activated protein kinase (AMPK) or, alternatively, acetyl-CoA-carboxylase (ACC). Figure 4C shows the expression of Acc ({alpha} and β isoforms) and Ampk ({alpha}2 and β2 subunits) in soleus and gastrocnemius muscle after HLS, relative to ambulatory control. HLS significantly increased expression of Acc-β and the {alpha}2 catalytic subunit of Ampk, while expression of Ampk noncatalytic subunit β2 and Acc-{alpha} were unchanged in soleus muscle after unloading. In contrast to soleus muscle, Ampk-β2 was downregulated in gastrocnemius muscle after HLS.

Uncoupling proteins are thought to be involved in mediation of cellular thermogenesis (7, 25, 64), modulation of mitochondrial hydrogen peroxide generation (52), and regulation of lipid as a fuel substrate (59). In agreement with our results demonstrating that Ucp-3 mRNA expression is increased in soleus muscle after short-term HLS, long-term unloading was previously associated with increased UCP-3 expression in skeletal muscle (20). To investigate whether increased mRNA expression was associated with enhanced protein production, Western blot analysis was performed on soleus muscle only. Gastrocnemius muscle was not investigated because Ucp-3 was not found to be differentially expressed in this tissue. Because UCP-3 is thought to be regulated by PPAR{delta} (56) and is found primarily in muscle (11, 23), we assessed expression of both proteins in soleus muscle after 24-h unloading. A representative Western blot of UCP-3 and PPAR{delta} expression in soleus muscle of three ambulatory and five HLS animals is shown in Fig. 5A. Figure 5B depicts average UCP-3 and PPAR{delta} expression in ambulatory and unloaded soleus muscle samples after normalization to β-actin expression, which was not found to be altered as a consequence of unloading (data not shown). Western blot analysis demonstrated that HLS resulted in significantly enhanced expression of both PPAR{delta} and UCP-3 compared with control.


Figure 5
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  		Fig. 5. Analysis of PPAR{delta} and UCP-3 expression in soleus muscle of ambulatory and 24-h HLS mice. For Western blot analysis, tissue was lysed in sample buffer [50 mM Tris, pH 8.0, 120 mM NaCl, 0.5% NP-40, 10 µg/ml PMSF, and 1x protease inhibitor cocktail]. A total of 50 µg of protein was suspended in Laemmli buffer (50 mM Tris, pH 6.8, 1% β-mercaptoethanol, 2% sodium dodecyl sulfate, 0.1% bromophenol blue, and 10% glycerol). Samples were boiled and subject to Western blot analysis on NuPAGE Bis-Tris gels with MES running buffer. Mouse PPAR{delta}, UCP-3, and β-actin proteins were detected by antibodies as detailed in MATERIALS AND METHODS and visualized by enhanced chemiluminescence. A: representative blot of PPAR{delta} and UCP-3 expression in soleus muscle of ambulatory (n = 3) and 1-day HLS (n = 5) mice. B: average PPAR{delta} and UCP-3 expression in soleus muscle of ambulatory and HLS soleus depicted graphically after normalization to β-actin expression in each sample. Expression in soleus muscle isolated from HLS mice is displayed relative to ambulatory control. Student's t-test was performed, and P < 0.05 was considered statistically significant.

 
PPAR{delta} Expression After Long-Term Exposure to HLS
Increased expression of UCP-3 after long-term unloading and in cachexia-induced muscle wasting has been reported previously (4, 20, 50). However, in these studies of long-term unloading, PPAR{delta} expression was not investigated. Figure 6 shows mRNA expression of Ppar-{delta} in soleus muscle isolated from ambulatory animals and after 12-day HLS. In contrast to Fig. 5, in which a significant increase in PPAR{delta} RNA and protein expression was observed at 1 day of HLS, we observed no significant difference in Ppar-{delta} expression after 12-day unloading, suggesting that HLS-induced upregulation of PPAR{delta} is both acute and transient. In contrast, in agreement with previous studies (20), UCP-3 expressed is increased after long-term HLS. These data suggest that UCP-3 expression is increased in a Ppar{delta}-independent manner in the long-term unloading condition.


Figure 6
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  		Fig. 6. Analysis of UCP-3 and PPAR{delta} expression in ambulatory and 12-day unloaded soleus muscle. Soleus tissue was isolated from control animals and animals after 12-day HLS. RNA was extracted and mRNA expression of UCP-3 and PPAR{delta} was assessed as previously described in MATERIALS AND METHODS. Animals were grouped into ambulatory (n = 4) and 12-day HLS (n = 4) conditions, and mRNA expression for each animal was averaged for each gene analyzed. All samples were run in triplicate with test probes and the control gene mouse SNARE to control for differences in amount of starting material. Fold change in expression was calculated by normalizing the test gene crossing threshold (Ct) with the SNARE-amplified control and then comparing to gene expression in ambulatory control animals. P < 0.05. n.s., Not significant.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The goal of the present study was to determine the effects of acute, short-term HLS on metabolic genome remodeling. In contrast to the other Ppars and Ppar-regulated genes that were downregulated by HLS, Ppar-{delta} was upregulated concomitant with increased UCP-3 expression. Because of its role in regulating oxidative capacity and fuel switching in addition to its known interaction with UCP-3, we hypothesize that acute upregulation of Ppar-{delta} following short-term HLS may represent an adaptive, compensatory response aimed at regulating fuel utilization and maintaining metabolic flexibility. This hypothesis is supported by the finding that Ppar-{delta} expression is restored to basal levels at 12 days of HLS, at the same time at which muscle atrophy and loss of lipid oxidative capacity are observed. From these data, we hypothesize that loss of PPAR{delta} expression may contribute to metabolic inflexibility and adverse physiological alterations observed after long-term unloading. Thus improving metabolic flexibility via PPAR{delta} regulation may represent a novel intervention strategy to prevent the adverse metabolic and physiological effects of muscle unloading. Of course, examining metabolic fluxes would provide evidence in support of this hypothesis. However, these studies ideally would be performed with isotopes in unstressed animals, which would be technically challenging in mice that have undergone a stress-inducing procedure like HLS. Thus comparing changes in gene expression with the use of microarrays coupled with known metabolic alterations in long-term studies of unloading provides a complementary means to understand the genomic mechanisms that underlie the unloaded phenotype.

In recent years, several groups have used varying approaches to study gene expression after inactivity. These include analysis of gene expression after 12 h (4), 12 days (66), 14 days (18), or 35 days (80). Furthermore, temporal transcriptional profiling of rat soleus muscle after 1, 4, 7, and 14 days (69) or 1 and 35 days (43) of unloading has demonstrated that in addition to delayed activation of genes involved in muscle contraction, glycolysis, and catabolism, a vast number of regulatory genes are acutely affected—in a transient or sustained manner—by HLS. The conclusions of these studies conclusively suggest that muscle unloading is characterized by reduced fatty acid oxidation, decreased transcription of genes involved in lipid metabolism, and increased expression of glycolytic enzymes and genes involved in glycogen synthesis, all of which suggest an increased reliance on glucose as an energy source or a loss of metabolic flexibility (30, 32, 43). In corroboration with previous studies of acute HLS (43, 69) we observed many similar changes in genes involved in glucose and lipid regulation and energy homeostasis. These rapidly responding genes may be involved in the adaptive responses to HLS or may contribute to the physiological alterations associated with long-term unloading.

In the present study we focused our attempts on identifying early-responding signaling and regulatory genes that might correspond with the loss of metabolic flexibility in muscle. Here, we demonstrated a reduction in many genes involved in the PPAR signaling pathway including Ppar-{alpha}, Ppar-{gamma}, Lxr-{alpha}, and Pgc-1{alpha}, and Pgc-1β after acute HLS. However, we do not observe profound decreases in other genes involved in fatty acid transport, degradation, or oxidation, suggesting that alternative regulatory mechanisms compensate for loss of PPAR{alpha}/{gamma}-mediated control of lipid metabolism. In support of this hypothesis, PPAR{delta} has been demonstrated to compensate for loss of PPAR{alpha} (51). This may occur through PPAR{alpha}/{gamma}-independent heterodimerization with RXR and transcriptional regulation of genes involved in fatty acid oxidation and lipid homeostasis such as fatty acid binding protein 3 (FABP3), a cytosolic protein involved in uptake and transport of fatty acids (82), CPT1, a rate-limiting enzyme in mitochondrial fatty acid oxidation, and PDK4. Gene arrays demonstrated no significant change in Pdk-4 or Fabp-3 expression in soleus or gastrocnemius muscle after 24-h HLS. However, we did observe a significant increase in expression of Cpt-1b and Cpt-2, proteins involved in fatty acid transport into mitochondria, in soleus muscle after unloading. In contrast to these acute observations, in long-term exposure to HLS, there is decreased expression of genes involved in lipid metabolism including Cpt-1 and Cpt-2 concomitant with increased expression of glycolytic genes (hexokinase, phosphofructokinase, and pyruvate kinase) (68), suggesting that HLS-induced regulation of these genes is delayed.

These seemingly contradictory data further imply that a very early adaptation to acute unloading-induced metabolic deregulation occurs via increasing lipid utilization, while later time points indicate a reversal of lipid utilization and an increased reliance on glucose concomitant with loss of metabolic flexibility. It is possible that this switch is mediated by PPAR{delta}. Several pieces of evidence suggest that PPAR{delta} may respond to—and attempt to compensate for—global metabolic deregulation. PPAR{delta} is involved in remodeling of skeletal muscle, increasing oxidative fibers, and reducing body fat content. Increased PPAR{delta} has previously been implicated in the metabolic and structural adaptations to long-term fasting and endurance exercise (reviewed in Ref. 46a) and increased expression is an indicator of improvements in diabetes on training Ref. 78a. These data suggest that increased expression of PPAR{delta} following unloading may represent an adaptive, stress-induced response to prevent further metabolic consequences. However, we hypothesize that after longer-term exposure to unloading metabolic flexibility is lost, resulting in enhanced reliance on glucose utilization concomitant with increased lipid accumulation in tissue, thus contributing to muscle wasting (atrophy) and slow-to-fast fiber transformation.

In addition to its interactions with other nuclear receptors such as PPAR{alpha}, PPAR{gamma}, and RXR, and through its transcriptional targets, PPAR{delta} also interacts with AMPK and may influence metabolic regulation through this route (42). Krämer et al. (42) recently demonstrated that the effects of PPAR{delta} on metabolic responses including regulation of glucose and lipid utilization may involve AMPK or, alternatively, ACC. In the present study, both Acc-β isoform and Ampk-{alpha}2 subunit were upregulated in soleus muscle by HLS. AMPK is a heterotrimeric protein kinase that participates in cellular energy homeostasis (46). Once activated, AMPK enhances cellular nutrient uptake, activates ATP-producing catabolic processes, and downregulates energy-consuming processes (46). In addition, AMPK-dependent effects on energy homeostasis may partly involve activation of PPARs (44, 45, 55). In fact, PPAR{delta} agonist treatment results in AMPK activation through an unknown mechanism (41). Therefore, it remains probable that the beneficial effects of PPAR{delta} on metabolic flexibility may be mediated, at least in part, through its interactions with both AMPK and ACC.

Alterations in the ratio of AMP to ATP also regulate AMPK activity; increased AMP:ATP concentration and activation of AMPK can occur via mitochondrial uncoupling (22, 40). In the present study we observed a significant increase in muscle-specific UCP-3 expression in soleus muscle after HLS. These data suggest that in addition to interactions with PPAR{delta} after unloading, it is also possible that HLS-induced mitochondrial uncoupling via increased expression of UCP-3 may activate AMPK. Alternatively, AMPK activation has been shown to result in upregulation of UCP-3 in muscle (70), demonstrating the complexity of the relationship between energy-sensing and/or responsive genes in skeletal muscle.

On the basis of its homology with UCP-1, UCP-3 has been implicated in the regulation of energy expenditure and has recently been studied as a potential contributor to muscle wasting under cachectic conditions (6, 16, 58). In addition to cachexia, it has been shown that UCP-3 expression is modulated in soleus muscle by unloading and fasting (20, 59). It appears that UCP-3 is involved in the regulation of lipids as a fuel substrate rather than simply as a mediator of thermogenesis as previously believed, because of the demonstrated relationship between free fatty acid flux and muscle UCP-3 expression (60, 62). The data we have obtained in the present study are consistent with this role, particularly as unloading is associated with altered free fatty acid levels in muscle (3). We would further hypothesize that UCP-3 functions may be multifaceted and differ in the context of acute and chronic activation of the protein.

Although the loss of UCP-3 function appears not to be involved in the pathogenesis of obesity-related conditions, it is possible that upregulation of UCP-3 may improve metabolic parameters such as metabolic flexibility. Clapham et al. (15) reported that UCP-3 transgenic mice were more insulin sensitive and have lower fasting plasma glucose and insulin, suggesting a possible beneficial effect of UCP-3 on metabolic regulation. Several reports have proposed that fatty acids induce UCP-3 gene expression in skeletal muscle (8, 79). In addition, increased UCP-3 protected against lipotoxicity in a cachexia model (50). During cancer cachexia, the expression of UCP-3 is also upregulated in skeletal muscle (12, 61) and is dependent on rise in circulating levels of free fatty acids. When UCP-3 levels increased, free fatty acids decreased concomitant with increased lipid peroxidation markers, suggesting a switch to fat oxidation. The cachectic state is often accompanied by increased rates of adipose tissue lipolysis, reduced mitochondrial volume, and changes in mitochondrial protein synthesis rate, and UCP-3 is thought to protect mitochondria against the oxidative damage induced by nonesterified fatty acids under these conditions (28, 33, 63). Further evidence connecting UCP expression to metabolic flexibility includes the demonstration that both UCP-2 and UCP-3 are important in resting metabolic rate, resting and basal energy expenditure, and anorexia nervosa (9, 13, 77) through linkage analysis. Ukkola et al. (75) demonstrated a role for UCPs in the adaptive response to long-term overfeeding by regulating substrate oxidation.

Several investigations link upregulated PPAR{delta} with high levels of UCP-3 expression. (19, 26, 65). Direct evidence of an interaction between PPAR{delta} and UCP-3 is due to the finding that the UCP-3 gene promoter contains a PPAR response element (56). Specific PPAR{delta} agonist treatment resulted in upregulation of UCP-3 that was mediated by fatty acids (65). This same study reported that modulation of UCP-3 by PPARs appeared specific to PPAR{delta}, because neither PPAR{alpha} nor PPAR{gamma} was demonstrated to transcriptionally regulate UCP-3. In addition to modulation by fatty acids, Fritz et al. (26) have demonstrated that low-intensity exercise increases skeletal muscle protein expression of PPAR{delta} and UCP-3 in type 2 diabetic patients, suggesting that regulation of these two proteins may represent an early adaptation that may ultimately serve to improve clinical outcome.

Improving metabolic flexibility may represent a novel intervention strategy to prevent the adverse physiological effects of muscle unloading. This concept is gaining momentum particularly in the cardiac field, where alterations in myocardial energy substrate metabolism are thought to contribute to heart failure (reviewed in Refs. 39, 67, 72). Of interest, it appears that metabolic maladaptation in the stressed heart occurs as the result of a return to a pattern of fetal metabolism including predominance of the use of carbohydrate as the major fuel substrate (reviewed in Ref. 54). Furthermore, Jucker et al. (36) elegantly showed recently that PPAR{delta} activation in congestive heart failure normalizes cardiac substrate metabolism and improves hypertrophy and pulmonary congestion. These concepts mirror the findings and hypotheses set forth in the present study, suggesting that the metabolic disturbances observed in the muscle disuse phenotype may be similar to those demonstrated in cardiohypertrophy and heart failure.

In summary, in this report we have identified several proteins, namely, PPAR{delta}, UCP-3, AMPK, and CPT1/2, that have putative roles in the maintenance of metabolic flexibility and represent potential targets for intervention. Because of its interactions with the other putative targets, PPAR{delta} appears particularly promising. In addition, previous investigations have demonstrated that transgenic mice expressing an activated form of PPAR{delta} have enhanced fatty acid utilization and are protected against high-fat diet-induced obesity (78). Furthermore, PPAR{delta} agonism induces changes in muscle fuel metabolism (increases lipid oxidation, decreases carbohydrate utilization) (17, 48). Since enhanced UCP-3 expression is associated with improvements in glycemic regulation (49), we hypothesize that promoting PPAR{delta}-dependent upregulation of UCP-3 and other transcriptional targets might offer additional benefits to prevent metabolic derangement and physiological alterations associated with muscle unloading. Further studies are in progress to test this hypothesis.


   GRANTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
R. C. Oita is supported by the NucSys Marie Curie Research Training Network funded by the European Union (contract number MRTN-CT-019496).


   ACKNOWLEDGMENTS
 
We thank Ann Scarborough for technical assistance and Jonathan Powell for critical review of the manuscript.


   FOOTNOTES
 
Address for reprint requests and other correspondence: D. J. Mazzatti, Unilever Corporate Research, Colworth Park, Sharnbrook, Bedfordshire MK44 1LQ, UK (e-mail: dawn.mazzatti{at}unilever.com).

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
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Acheson KJ, Decombaz J, Piguet-Welsch C, Montigon F, Decarli B, Bartholdi I, Fern EB. Energy, protein, and substrate metabolism in simulated microgravity. Am J Physiol Regul Integr Comp Physiol 269: R252–R260, 1995.[Abstract/Free Full Text]
  2. Arbogast S, Smith J, Matuszczak Y, Hardin BJ, Moylan JS, Smith JD, Ware J, Kennedy AR, Reid MB. Bowman-Birk inhibitor concentrate prevents atrophy, weakness, and oxidative stress in soleus muscle of hindlimb-unloaded mice. J Appl Physiol 102: 956–964, 2007.[Abstract/Free Full Text]
  3. Baldwin KM, Herrick RE, McCue SA. Substrate oxidation capacity in rodent skeletal muscle: effects of exposure to zero gravity. J Appl Physiol 75: 2466–2470, 1993.[Abstract/Free Full Text]
  4. Bey L, Akunuri N, Zhao P, Hoffman EP, Hamilton DG, Hamilton MT. Patterns of global gene expression in rat skeletal muscle during unloading and low-intensity ambulatory activity. Physiol Genomics 13: 157–167, 2003.[Abstract/Free Full Text]
  5. Beyer RE, Fattore JE. The influence of age and endurance exercise on the myoglobin concentration of skeletal muscle of the rat. J Gerontol 39: 525–530, 1984.
  6. Bing C, Brown M, King P, Collins P, Tisdale MJ, Williams G. Increased gene expression of brown fat uncoupling protein (UCP)1 and skeletal muscle UCP2 and UCP3 in MAC16-induced cancer cachexia. Cancer Res 60: 2405–2410, 2000.[Abstract/Free Full Text]
  7. Bonadonna RC, Groop LC, Simonson DC, DeFronzo RA. Free fatty acid and glucose metabolism in human aging: evidence for the operation of the Randle cycle. Am J Physiol Endocrinol Metab 266: E501–E509, 1994.[Abstract/Free Full Text]
  8. Boss O, Samec S, Dulloo A, Seydoux J, Muzzin P, Giacobino JP. Tissue-dependent upregulation of rat uncoupling protein-2 expression in response to fasting or cold. FEBS Lett 412: 111–114, 1997.[CrossRef][Web of Science][Medline]
  9. Boss O, Bobbioni-Harsch E, Assimacopoulos-Jeannet F, Muzzin P, Munger R, Giacobino JP, Golay A. Uncoupling protein-3 expression in skeletal muscle and free fatty acids in obesity. Lancet 351: 1933, 1998.[Web of Science][Medline]
  10. Bouchard C, Perusse L, Chagnon YC, Warden C, Ricquier D. Linkage between markers in the vicinity of the uncoupling protein 2 gene and resting metabolic rate in humans. Hum Mol Genet 6: 1887–1889, 1997.[Abstract/Free Full Text]
  11. Braissant O, Foufelle F, Scotto C, Dauca M, Wahli W. Differential expression of peroxisome proliferator-activated receptors (PPARs): tissue distribution of PPAR-alpha, -beta, and -gamma in the adult rat. Endocrinology 137: 354–366, 1996.[Abstract]
  12. Brunmair B, Gras F, Wagner L, Artwohl M, Zierhut B, Waldhausl W, Furnsinn C. Expression of uncoupling protein-3 mRNA in rat skeletal muscle is acutely stimulated by thiazolidinediones: an exercise-like effect? Diabetologia 47: 1611–1614, 2004.[CrossRef][Web of Science][Medline]
  13. Busquets S, Carbo N, Almendro V, Figueras M, Lopez-Soriano FJ, Argiles JM. Hyperlipemia: a role in regulating UCP3 gene expression in skeletal muscle during cancer cachexia? FEBS Lett 505: 255–258, 2001.[CrossRef][Web of Science][Medline]
  14. Campbell DA, Sundaramurthy D, Gordon D, Markham AF, Pieri LF. Association between a marker in the UCP-2/UCP3 gene cluster and genetic susceptibility to anorexia nervosa. Mol Psychiatry 4: 68–70, 1999.[CrossRef][Web of Science][Medline]
  15. Clapham JC, Arch JR, Chapman H, Haynes A, Lister C, Moore GB, Piercy V, Carter SA, Lehner I, Smith SA, Beeley LJ, Godden RJ, Herrity N, Skehel M, Changani KK, Hockings PD, Reid DG, Squires SM, Hatcher J, Trail B, Latcham J, Rastan S, Harper AJ, Cadenas S, Buckingham JA, Brand MD, Abuin A. Mice overexpressing human uncoupling protein-3 in skeletal muscle are hyperphagic and lean. Nature 406: 415–418, 2000.[CrossRef][Medline]
  16. Collins P, Bing C, McCulloch P, Williams G. Muscle UCP3 mRNA levels are elevated in weight loss associated with gastrointestinal adenocarcinoma in humans. Br J Cancer 86: 372–375, 2002.[CrossRef][Web of Science][Medline]
  17. Constantin D, Constantin-Teodosiu D, Layfield R, Tsintzas K, Bennett AJ, Greenhaff PL. PPAR{delta} agonism induces a change in muscle fuel metabolism and activation of an atrophy programme, but does not impair mitochondrial function in rat skeletal muscle. J Physiol 583: 381–390, 2007.[Abstract/Free Full Text]
  18. Cros N, Tkatchenko AV, Pisani DF, Leclerc L, Leger JJ, Marini JF, Dechesne CA. Analysis of altered gene expression in rat soleus muscle atrophied by disuse. J Cell Biochem 83: 508–519, 2001.[CrossRef][Web of Science][Medline]
  19. de Lange P, Feola A, Ragni M, Senese R, Moreno M, Lombardi A, Silvestri E, Amat R, Villarroya F, Goglia F, Lanni A. Differential 3,5,3'-triiodothyronine-mediated regulation of uncoupling protein 3 transcription: role of fatty acids. Endocrinology 148: 4064–4072, 2007.[Abstract/Free Full Text]
  20. Denjean F, Desplanches D, Lachuer J, Cohen-Adad F, Mayet MH, Duchamp C. Muscle-specific up-regulation of rat UCP3 mRNA expression by long-term hindlimb unloading. Biochem Biophys Res Commun 266: 518–522, 1999.[CrossRef][Web of Science][Medline]
  21. Doenst T, Goodwin GW, Cedars AM, Wang M, Stepkowski S, Taegtmeyer H. Load-induced changes in vivo alter substrate fluxes and insulin responsiveness of rat heart in vitro. Metabolism 50: 1083–1090, 2001.[CrossRef][Web of Science][Medline]
  22. Dokladda K, Green KA, Pan DA, Hardie DG. PD98059 and U0126 activate AMP-activated protein kinase by increasing the cellular AMP:ATP ratio and not via inhibition of the MAP kinase pathway. FEBS Lett 579: 236–240, 2005.[CrossRef][Web of Science][Medline]
  23. Dressel U, Allen TL, Pippal JB, Rohde PR, Lau P, Muscat GE. The peroxisome proliferator-activated receptor beta/delta agonist, GW501516, regulates the expression of genes involved in lipid catabolism and energy uncoupling in skeletal muscle cells. Mol Endocrinol 17: 2477–2493, 2003.[Abstract/Free Full Text]
  24. Farid M, Reid MB, Li YP, Gerken E, Durham WJ. Effects of dietary curcumin or N-acetylcysteine on NF-kappaB activity and contractile performance in ambulatory and unloaded murine soleus. Nutr Metab (Lond) 2: 20, 2005.[CrossRef][Medline]
  25. Fleury C, Neverova M, Collins S, Raimbault S, Champigny O, Levi-Meyrueis C, Bouillaud F, Seldin MF, Surwit RS, Ricquier D, Warden CH. Uncoupling protein-2: a novel gene linked to obesity and hyperinsulinemia. Nat Genet 15: 269–272, 1997.[CrossRef][Web of Science][Medline]
  26. Fritz T, Kramer DK, Karlsson HK, Galuska D, Engfeldt P, Zierath JR, Krook A. Low-intensity exercise increases skeletal muscle protein expression of PPARdelta and UCP3 in type 2 diabetic patients. Diabetes Metab Res Rev 22: 492–498, 2006.[CrossRef][Web of Science][Medline]
  27. Fürnsinn C, Willson TM, Brunmair B. Peroxisome proliferator-activated receptor-delta, a regulator of oxidative capacity, fuel switching and cholesterol transport. Diabetologia 50: 8–17, 2007.[Web of Science][Medline]
  28. Goglia F, Skulachev VP. A function for novel uncoupling proteins: antioxidant defense of mitochondrial matrix by translocating fatty acid peroxides from the inner to the outer membrane leaflet. FASEB J 17: 1585–1591, 2003.[Abstract/Free Full Text]
  29. Goodpaster BH, Wolfe RR, Kelley DE. Effects of obesity on substrate utilization during exercise. Obes Res 10: 575–584, 2002.[Web of Science][Medline]
  30. Grichko VP, Heywood-Cooksey A, Kidd KR, Fitts RH. Substrate profile in rat soleus muscle fibers after hindlimb unloading and fatigue. J Appl Physiol 88: 473–478, 2000.[Abstract/Free Full Text]
  31. Hamilton MT, Booth FW. Skeletal muscle adaptation to exercise: a century of progress. J Appl Physiol 88: 327–331, 2000.[Abstract/Free Full Text]
  32. Henriksen EJ, Tischler ME. Glucose uptake in rat soleus: effect of acute unloading and subsequent reloading. J Appl Physiol 64: 1428–1432, 1988.[Abstract/Free Full Text]
  33. Himms-Hagen J, Harper ME. Physiological role of UCP3 may be export of fatty acids from mitochondria when fatty acid oxidation predominates: a hypothesis. Exp Biol Med (Maywood) 226: 78–84, 2001.[Abstract/Free Full Text]
  34. Huey KA, Roy RR, Baldwin KM, Edgerton VR. Temporal effects of inactivity on myosin heavy chain gene expression in rat slow muscle. Muscle Nerve 24: 517–526, 2001.[CrossRef][Web of Science][Medline]
  35. Huss JM, Kelly DP. Nuclear receptor signaling and cardiac energetics. Circ Res 95: 568–578, 2004.[Abstract/Free Full Text]
  36. Jucker BM, Doe CP, Schnackenberg CG, Olzinski AR, Maniscalco K, Williams C, Hu TC, Lenhard SC, Costell M, Bernard R, Sarov-Blat L, Steplewski K, Willette RN. PPARdelta activation normalizes cardiac substrate metabolism and reduces right ventricular hypertrophy in congestive heart failure. J Cardiovasc Pharmacol 50: 25–34, 2007.[CrossRef][Web of Science][Medline]
  37. Kang J, Kelley DE, Robertson RJ, Goss FL, Suminski RR, Utter AC, Dasilva SG. Substrate utilization and glucose turnover during exercise of varying intensities in individuals with NIDDM. Med Sci Sports Exerc 31: 82–89, 1999.[Web of Science][Medline]
  38. Kelley DE, Mandarino LJ. Fuel selection in human skeletal muscle in insulin resistance: a reexamination. Diabetes 49: 677–683, 2000.[Abstract]
  39. Kodde IF, van der Stok J, Smolenski RT, de Jong JW. Metabolic and genetic regulation of cardiac energy substrate preference. Comp Biochem Physiol A 146: 26–39, 2007.[CrossRef][Medline]
  40. Konrad D, Rudich A, Bilan PJ, Patel N, Richardson C, Witters LA, Klip A. Troglitazone causes acute mitochondrial membrane depolarisation and an AMPK-mediated increase in glucose phosphorylation in muscle cells. Diabetologia 48: 954–966, 2005.[CrossRef][Web of Science][Medline]
  41. Krämer DK, Al-Khalili L, Perrini S, Skogsberg J, Wretenberg P, Kannisto K, Wallberg-Henriksson H, Ehrenborg E, Zierath JR, Krook A. Direct activation of glucose transport in primary human myotubes after activation of peroxisome proliferator-activated receptor delta. Diabetes 54: 1157–1163, 2005.[Abstract/Free Full Text]
  42. Krämer DK, Al-Khalili L, Guigas B, Leng Y, Garcia-Roves PM, Krook A. Role of AMP kinase and PPARdelta in the regulation of lipid and glucose metabolism in human skeletal muscle. J Biol Chem 282: 19313–19320, 2007.[Abstract/Free Full Text]
  43. Langfort J, Zernicka E, Mayet-Sornay MH, Dubaniewicz A, Desplanches D. Effects of acute and chronic hindlimb suspension on sensitivity and responsiveness to insulin in the rat soleus muscle. Biochem Cell Biol 75: 41–44, 1997.[CrossRef][Web of Science][Medline]
  44. Lee WJ, Kim M, Park HS, Kim HS, Jeon MJ, Oh KS, Koh EH, Won JC, Kim MS, Oh GT, Yoon M, Lee KU, Park JY. AMPK activation increases fatty acid oxidation in skeletal muscle by activating PPARalpha and PGC-1. Biochem Biophys Res Commun 340: 291–295, 2006.[CrossRef][Web of Science][Medline]
  45. Leff T. AMP-activated protein kinase regulates gene expression by direct phosphorylation of nuclear proteins. Biochem Soc Trans 31: 224–227, 2003.[Web of Science][Medline]
  46. Long YC, Zierath JR. AMP-activated protein kinase signaling in metabolic regulation. J Clin Invest 116: 1776–1783, 2006.[CrossRef][Web of Science][Medline]
  47. Luquet S, Lopez-Soriano J, Holst D, Gaudel C, Jehl-Pietri C, Fredenrich A, Grimaldi PA. Roles of peroxisome proliferator-activated receptor delta (PPARdelta) in the control of fatty acid catabolism. A new target for the treatment of metabolic syndrome. Biochimie 86: 833–837, 2004.[Medline]
  48. Matuszczak Y, Arbogast S, Reid MB. Allopurinol mitigates muscle contractile dysfunction caused by hindlimb unloading in mice. Aviat Space Environ Med 75: 581–588, 2004.[Medline]
  49. Mensink M, Hesselink MK, Russell AP, Schaart G, Sels JP, Schrauwen P. Improved skeletal muscle oxidative enzyme activity and restoration of PGC-1alpha and PPARbeta/delta gene expression upon rosiglitazone treatment in obese patients with type 2 diabetes mellitus. Int J Obes (Lond) 31: 1302–1310, 2007.[CrossRef][Medline]
  50. Mensink M, Hesselink MK, Borghouts LB, Keizer H, Moonen- Kornips E, Schaart G, Blaak EE, Schrauwen P. Skeletal muscle uncoupling protein-3 restores upon intervention in the prediabetic and diabetic state: implications for diabetes pathogenesis? Diabetes Obes Metab 9: 594–596, 2007.[CrossRef][Web of Science][Medline]
  51. Minnaard R, Schrauwen P, Schaart G, Hesselink MK. UCP3 in muscle wasting, a role in modulating lipotoxicity? FEBS Lett 580: 5172–5176, 2006.[CrossRef][Web of Science][Medline]
  52. Muoio DM, MacLean PS, Lang DB, Li S, Houmard JA, Way JM, Winegar DA, Corton JC, Dohm GL, Kraus WE. Fatty acid homeostasis and induction of lipid regulatory genes in skeletal muscles of peroxisome proliferator-activated receptor (PPAR) alpha knock-out mice. J Biol Chem 277: 26089–26097, 2002.[Abstract/Free Full Text]
  53. Negre-Salvayre A, Hirtz C, Carrera G, Cazenave R, Troly M, Salvayre R, Penicaud L, Casteilla L. A role for uncoupling protein-2 as a regulator of mitochondrial hydrogen peroxide generation. FASEB J 11: 809–815, 1997.[Abstract]
  54. Ohira Y, Yasui W, Kariya F, Wakatsuki T, Nakamura K, Asakura T, Edgerton VR. Metabolic adaptation of skeletal muscles to gravitational unloading. Acta Astronaut 33: 113–117, 1994.[CrossRef][Web of Science][Medline]
  55. Rajabi M, Kassiotis C, Razeghi P, Taegtmeyer H. Return to the fetal gene program protects the stressed heart: a strong hypothesis. Heart Fail Rev 12: 331–343, 2007.[CrossRef][Web of Science][Medline]
  56. Ravnskjaer K, Boergesen M, Dalgaard LT, Mandrup S. Glucose-induced repression of PPARalpha gene expression in pancreatic beta-cells involves PP2A activation and AMPK inactivation. J Mol Endocrinol 36: 289–299, 2006.[Abstract/Free Full Text]
  57. Riquet FB, Rodriguez M, Guigal N, Dromaint S, Naime I, Boutin JA, Galizzi JP. In vivo characterisation of the human UCP3 gene minimal promoter in mice tibialis anterior muscles. Biochem Biophys Res Commun 311: 583–591, 2003.[CrossRef][Web of Science][Medline]
  58. Ritz P, Acheson KJ, Gachon P, Vico L, Bernard JJ, Alexandre C, Beaufrere B. Energy and substrate metabolism during a 42-day bed-rest in a head-down tilt position in humans. Eur J Appl Physiol Occup Physiol 78: 308–314, 1998.[CrossRef][Medline]
  59. Roubenoff R, Roubenoff RA, Cannon JG, Kehayias JJ, Zhuang H, Dawson-Hughes B, Dinarello CA, Rosenberg IH. Rheumatoid cachexia: cytokine-driven hypermetabolism accompanying reduced body cell mass in chronic inflammation. J Clin Invest 93: 2379–2386, 1994.[Web of Science][Medline]
  60. Samec S, Seydoux J, Dulloo AG. Role of UCP homologues in skeletal muscles and brown adipose tissue: mediators of thermogenesis or regulators of lipids as fuel substrate? FASEB J 12: 715–724, 1998.[Abstract/Free Full Text]
  61. Samec S, Seydoux J, Dulloo AG. Skeletal muscle UCP3 and UCP2 gene expression in response to inhibition of free fatty acid flux through mitochondrial beta-oxidation. Pflügers Arch 438: 452–457, 1999.[CrossRef][Web of Science][Medline]
  62. Sanchis D, Busquets S, Alvarez B, Ricquier D, Lopez-Soriano FJ, Argiles JM. Skeletal muscle UCP2 and UCP3 gene expression in a rat cancer cachexia model. FEBS Lett 436: 415–418, 1998.[CrossRef][Web of Science][Medline]
  63. Sbraccia P, D'Adamo M, Leonetti F, Buongiorno A, Silecchia G, Basso MS, Tamburrano G, Lauro D, Federici M, Di Daniele N, Lauro R. Relationship between plasma free fatty acids and uncoupling protein-3 gene expression in skeletal muscle of obese subjects: in vitro evidence of a causal link. Clin Endocrinol (Oxf) 57: 199–207, 2002.[CrossRef][Medline]
  64. Schrauwen P, Saris WH, Hesselink MK. An alternative function for human uncoupling protein 3: protection of mitochondria against accumulation of nonesterified fatty acids inside the mitochondrial matrix. FASEB J 15: 2497–2502, 2001.[Abstract/Free Full Text]
  65. Skulachev VP. Uncoupling: new approaches to an old problem of bioenergetics. Biochim Biophys Acta 1363: 100–124, 1998.[Medline]
  66. Son C, Hosoda K, Matsuda J, Fujikura J, Yonemitsu S, Iwakura H, Masuzaki H, Ogawa Y, Hayashi T, Itoh H, Nishimura H, Inoue G, Yoshimasa Y, Yamori Y, Nakao K. Up-regulation of uncoupling protein 3 gene expression by fatty acids and agonists for PPARs in L6 myotubes. Endocrinology 142: 4189–4194, 2001.[Abstract/Free Full Text]
  67. St-Amand J, Okamura K, Matsumoto K, Shimizu S, Sogawa Y. Characterization of control and immobilized skeletal muscle: an overview from genetic engineering. FASEB J 15: 684–692, 2001.[Abstract/Free Full Text]
  68. Stanley WC, Recchia FA, Lopaschuk GD. Myocardial substrate metabolism in the normal and failing heart. Physiol Rev 85: 1093–1129, 2005.[Abstract/Free Full Text]
  69. Stein TP, Schluter MD, Galante AT, Soteropoulos P, Tolias PP, Grindeland RE, Moran MM, Want TJ, Polansky M, Wade CE. Energy metabolism pathways in rat muscle under conditions of simulated microgravity. J Nutr Biochem 13: 471–478, 2002.[CrossRef][Web of Science][Medline]
  70. Stevenson EJ, Giresi PG, Koncarevic A, Kandarian SC. Global analysis of gene expression patterns during disuse atrophy in rat skeletal muscle. J Physiol 551: 33–48, 2003.[Abstract/Free Full Text]
  71. Stoppani J, Hildebrandt AL, Sakamoto K, Cameron-Smith D, Goodyear LJ, Neufer PD. AMP-activated protein kinase activates transcription of the UCP3 and HKII genes in rat skeletal muscle. Am J Physiol Endocrinol Metab 283: E1239–E1248, 2002.[Abstract/Free Full Text]
  72. Taegtmeyer H, Golfman L, Sharma S, Razeghi P, van Arsdall M. Linking gene expression to function: metabolic flexibility in the normal and diseased heart. Ann NY Acad Sci 1015: 202–213, 2004.[CrossRef][Web of Science][Medline]
  73. Taegtmeyer H, Wilson CR, Razeghi P, Sharma S. Metabolic energetics and genetics in the heart. Ann NY Acad Sci 1047: 208–218, 2005.[CrossRef][Web of Science][Medline]
  74. Thomason DB, Booth FW. Atrophy of the soleus muscle by hindlimb unweighting. J Appl Physiol 68: 1–12, 1990.[Abstract/Free Full Text]
  75. Tischler ME, Jaspers SR, Henriksen EJ, Jacob S. Responses of skeletal muscle to unloading—a review. Physiologist 28, Suppl: S13–S16, 1985.[Medline]
  76. Ukkola O, Tremblay A, Sun G, Chagnon YC, Bouchard C. Genetic variation at the uncoupling protein 1, 2 and 3 loci and the response to long-term overfeeding. Eur J Clin Nutr 55: 1008–1015, 2001.[CrossRef][Web of Science][Medline]
  77. Ukropcova B, Sereda O, de Jonge L, Bogacka I, Nguyen T, Xie H, Bray GA, Smith SR. Family history of diabetes links impaired substrate switching and reduced mitochondrial content in skeletal muscle. Diabetes 56: 720–727, 2007.[Abstract/Free Full Text]
  78. Walder K, Norman RA, Hanson RL, Schrauwen P, Neverova M, Jenkinson CP, Easlick J, Warden CH, Pecqueur C, Raimbault S, Ricquier D, Silver MH, Shuldiner AR, Solanes G, Lowell BB, Chung WK, Leibel RL, Pratley R, Ravussin E. Association between uncoupling protein polymorphisms (UCP2-UCP3) and energy metabolism/obesity in Pima indians. Hum Mol Genet 7: 1431–1435, 1998.[Abstract/Free Full Text]
  79. Wang YX, Lee CH, Tiep S, Yu RT, Ham J, Kang H, Evans RM. Peroxisome-proliferator-activated receptor {delta} activates fat metabolism to prevent obesity. Cell 113: 159–170, 2003.[CrossRef][Web of Science][Medline]
  80. Wang YX, Zhang CL, Yu RT, Cho HK, Nelson MC, Bayuga-Ocampo CR, Ham J, Kang H, Evans RM. Regulation of muscle fiber type and running endurance by PPARdelta. PLoS Biol 2: e294, 2004.[CrossRef][Medline]
  81. Weigle DS, Selfridge LE, Schwartz MW, Seeley RJ, Cummings DE, Havel PJ, Kuijper JL, BeltrandelRio H. Elevated free fatty acids induce uncoupling protein 3 expression in muscle: a potential explanation for the effect of fasting. Diabetes 47: 298–302, 1998.[Abstract]
  82. Wittwer M, Fluck M, Hoppeler H, Muller S, Desplanches D, Billeter R. Prolonged unloading of rat soleus muscle causes distinct adaptations of the gene profile. FASEB J 16: 884–886, 2002.[Abstract/Free Full Text]
  83. Yajid F, Mercier JG, Mercier BM, Dubouchaud H, Préfaut C. Effects of 4 wk of hindlimb suspension on skeletal muscle mitochondrial respiration in rats. J Appl Physiol 84: 479–485, 1998.[Abstract/Free Full Text]
  84. Zimmerman AW, Veerkamp JH. Fatty-acid-binding proteins do not protect against induced cytotoxicity in a kidney cell model. Biochem J 360: 159–165, 2001.[CrossRef][Web of Science][Medline]




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