AJP - Endo Fuel your research with LabChart
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Endocrinol Metab 292: E1631-E1636, 2007. First published February 6, 2007; doi:10.1152/ajpendo.00702.2006
0193-1849/07 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow A corrigendum has been published
Right arrow All Versions of this Article:
292/6/E1631    most recent
00702.2006v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (2)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Spargo, F. J.
Right arrow Articles by Hawley, J. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Spargo, F. J.
Right arrow Articles by Hawley, J. A.

Dysregulation of muscle lipid metabolism in rats selectively bred for low aerobic running capacity

Fiona J. Spargo,1 Sean L. McGee,2 Nick Dzamko,3 Matthew J. Watt,3 Bruce E. Kemp,3 Steven L. Britton,4 Lauren G. Koch,4 Mark Hargreaves,2 and John A. Hawley1

1Exercise Metabolism Laboratory, RMIT University, Melbourne, 2Department of Physiology, University of Melbourne, Melbourne, and 3Department of Medicine, St. Vincent's Institute of Medical Research, Fitzroy, Australia; and 4Physical Medicine and Rehabilitation, University of Michigan, Ann Arbor, Michigan

Submitted 20 December 2006 ; accepted in final form 30 January 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
As substrate for evaluation of metabolic diseases, we developed novel rat models that contrast for endurance exercise capacity. Through two-way artificial selection, we created rodent phenotypes of intrinsically low-capacity runners (LCR) and high-capacity runners (HCR) that also differed markedly for cardiovascular and metabolic disease risk factors. Here, we determined skeletal muscle proteins with putative roles in lipid and carbohydrate metabolism to better understand the mechanisms underlying differences in whole body substrate handling between phenotypes. Animals (generation 16) differed for endurance running capacity by 295%. LCR animals had higher resting plasma glucose (6.58 ± 0.45 vs. 6.09 ± 0.45 mmol/l), insulin (0.48 ± 0.03 vs. 0.32 ± 0.02 ng/ml), nonesterified fatty acid (0.57 ± 0.14 v 0.35 ± 0.05 mM), and triglyceride (TG; 0.47 ± 0.11 vs. 0.25 ± 0.08 mmol/l) concentrations (all P < 0.05). Muscle TG (72.3 ± 14.7 vs. 38.9 ± 6.2 mmol/kg dry muscle wt; P < 0.05) and diacylglycerol (96 ± 28 vs. 42 ± 8 pmol/mg dry muscle wt; P < 0.05) contents were elevated in LCR vs. HCR rats. Accompanying the greater lipid accretion in LCR was increased fatty acid translocase/CD36 content (1,014 ± 80 vs. 781 ± 70 arbitrary units; P < 0.05) and reduced TG lipase activity (0.158 ± 0.0125 vs. 0.274 ± 0.018 mmol·min–1·kg dry muscle wt–1; P < 0.05). Muscle glycogen, GLUT4 protein, and basal phosphorylation states of AMP-activated protein kinase-{alpha}1, AMP-activated protein kinase-{alpha}2, and acetyl-CoA carboxylase were similar in LCR and HCR. In conclusion, rats with low intrinsic aerobic capacity demonstrate abnormalities in lipid-handling capacity. These disruptions may, in part, be responsible for the increased risk of metabolic disorders observed in this phenotype.

exercise capacity; insulin resistance; mitochondrial biogenesis; lipids

DURING THE PAST 50 YEARS, the prevalence of a cluster of interrelated chronic metabolic disease states including coronary heart disease, insulin resistance, type 2 diabetes mellitus, and obesity has reached epidemic proportions (5). The etiological basis of these disorders is both polygenic and dependent on environmental factors. However, because no new major human gene mutations have occurred in the latter half of the 20th century to cause this greater frequency of chronic metabolic diseases, the increased incidence must principally be due to alterations in environmental conditions.

One contemporary environmental factor that changed and is strongly associated with chronic metabolic disorders is the decline in physical activity (8). Indeed, the increased prevalence of coronary heart disease, insulin resistance, type 2 diabetes mellitus, and obesity and their strong associations with inactivity have resulted in a low aerobic phenotype in which individuals with a particular combination of disease-susceptible genes (i.e., intrinsic risk factors) interact with environmental conditions (e.g., level of physical activity) and cross a threshold of biological significance that results in overt clinical conditions (1). Evidence in support of this premise comes from studies in which multiple genes involved in aerobic metabolism are downregulated in several metabolic states and may be linked to the pathogenesis of these disorders (25, 28, 36, 44). Less clear is whether this breakdown in aerobic metabolism results mainly from the inactive state (i.e., environmental) or is derived from mostly intrinsic factors independent of activity.

To approach progressive metabolic disease states mechanistically, we recently developed novel rat models that contrast for aerobic phenotype (18). Through two-way artificial selection, we generated rodent models of low-capacity runners (LCR) and high-capacity runners (HCR) that originated from a population of genetically heterogeneous rats (18). Eleven generations of selection produced animals that differed markedly (374%) for intrinsic (i.e., untrained) aerobic treadmill running capacity. This selection also simultaneously generated a contrast for metabolic and cardiovascular disease risk factors, including insulin resistance and elevated blood lipid concentrations in LCR compared with HCR (43). LCR animals also had compromised mitochondrial oxidative function relative to HCR rats, as evidenced by a reduced cellular content of several proteins required for mitochondrial biogenesis and function (15, 38, 43).

The comparative evaluation of animal models divergently selected for a complex phenotype is useful for two reasons. First, contrasting allelic variation causative of phenotypic differences is concentrated at the extremes of the population at each generation. Second, both selected lines have shared similar historical and current environments. Given the central role of lipid metabolism in aerobic exercise capacity, we hypothesized that the low intrinsic exercise capacity phenotype of the LCR would be characterized by elevated muscle lipid content and alterations in lipid handling capacity relative to the HCR. A further aim was to test whether the downregulation of key proteins involved in cellular energy metabolism found at generation 11 was maintained at generation 16.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animal procedures. Twenty-four female rats (12 LCR and 12 HCR) derived from generation 16 of selection (12) were used for this study. Animals were phenotyped at the University of Michigan (Ann Arbor, MI) for running capacity at 11 wk of age and then transported to The University of Melbourne animal facility (Parkville, Victoria, Australia) for further evaluation. Animals were housed three per cage in an environmentally controlled facility (21°C, 50–60% relative humidity) with a 12:12-h light-dark cycle (0600–1800). Animals received standard rodent chow (Speciality Feeds, Glen Forest, Western Australia) and water ad libitum. At the time of death, animals were age ~30 wk. All animal experimentation procedures were carried out with the approval of The University of Melbourne Animal Ethics Committee.

Approximately 10–14 h before death, food was removed from animal cages, but animals had continued ad libitum water access. Approximately 60 min before surgical removal of tissues, animals were weighed and injected intraperitoneally with a 60–80 mg/kg body mass dose of pentobarbital sodium. Once the animals were fully anesthetized, blood (5–10 ml) was removed via cardiac puncture, one-half was transferred to EDTA-administered tubes for later recovery of plasma, and one-half remained in regular Eppendorf tubes, for later recovery of serum. Hindlimb muscles were removed and immediately freeze clamped in liquid nitrogen before removal for storage at –80°C.

Plasma analyses. Plasma glucose, triglycerides (TG), and total and high-density lipoprotein cholesterol analyses were performed at Melbourne Pathology, Collingwood, Victoria, Australia. Plasma glucose was assessed by the glucose-hexokinase method (35), whereas plasma TGs were measured with the colorimetric enzymatic GPO-PAP assay. Total cholesterol was also analyzed with the enzymatic colorimetric CHOD-PAP method, whereas high-density lipoprotein cholesterol was analyzed with a homogenous enzymatic colorimetric test. All tests were performed on a Modular Analytic SWA P module (Roche Diagnostics, Mannheim, Germany). Plasma nonesterified fatty acid and insulin analyses were performed with the use of commercially available kits (Wako nonesterified fatty acid C kit, Richmond, VA, and Linco RIA rat insulin kit, St. Charles, MO).

Muscle glycogen analysis. Muscle glycogen content was assayed as previously described (27). Briefly, ~3 mg of freeze-dried red gastrocnemius muscle were placed in 2 N HCl and incubated at 100°C for 2 h. After neutralization with 0.66 N NaOH, the liberated glucose units were assayed fluorometrically and glycogen content was expressed as millimoles of glucosyl units liberated per kilogram dry muscle weight (mmol/kg dry muscle wt).

Intramuscular TG analysis. Muscle triglyceride (TGm) content was determined as previously described (9). Briefly, ~7 mg of freeze-dried red gastrocnemius muscle were placed in 4 ml of chloroform-methanol (2:1) and left to rotate at room temperature for ~2 h to facilitate total lipid extraction. After lipid extraction, 0.6% NaCl was added to each sample tube before centrifugation at 2,000 rpm for 10 min to facilitate aqueous and organic phase separation. The TG-containing organic phase was then transferred to glass tubes and air dried. The isolated lipids were then resuspended in 250 µl of ethanol, and the TG concentration was determined spectrophotometrically at 490 nm with an enzymatic colorimetric test kit (triglycerides GPO-PAP, Boehringer Mannheim, Sydney, Australia).

Muscle diacylglycerol and ceramide analysis. Muscle diacylglycerol (DAG) and ceramide contents were determined as previously described (31). Briefly, ~5 mg of freeze-dried red gastrocnemius were placed in 3.8 ml of chloroform-methanol-PBS + 0.2% SDS (1:2:0.8) at room temperature for 3 h. DAG kinase and [{gamma}-32P]ATP (15 µCi/µmmol cold ATP) were added to the samples, and the reaction was stopped with 4 ml of chloroform-methanol (2:1). Samples were spotted onto TLC plates and developed to two-thirds of the total plate length. Bands containing DAG and ceramide were identified after phospho-imaging, dried, scraped from the TLC plate and counted in a liquid scintillation analyzer (Tri-Carb 2500TR; Packard, Canberra, Australia).

Western blot procedure. Red gastrocnemius muscle (~40–50 mg) was homogenized for ~30 s in either 1 ml of ice-cold buffer [for fatty acid translocase/CD36 (FAT/CD36)] [20 mM HEPES (pH 7.4), 2 mM EGTA, 50 mM beta-glycerophosphate, 1 mM DTT, 1 mM Na3VO4, 10% glycerol, 3 mM benzamidine, 10 µM leupeptin, 5 µM pepstatin A, Triton X-100, and 1 mM PMSF]; 10x vol ice-cold buffer [for cytochrome oxidase subunit IV (COX IV), PGC-1{alpha}, and GLUT4] [25 mM Tris (pH 6.8), 1% Triton X-100, 5 mM EGTA, 50 mM NaF, 1 mM sodium orthovanadate, 10% glycerol, 1% PMSF, 10 µg/ml leupeptin, 10 µg/ml aprotinin]; or ice-cold buffer (for total TG lipase) [50 mmol/l HEPES, 150 mmol/l NaCl, 10 mmol/l NaF, 1 mmol/l Na3VO4, 5 mmol/l EDTA, 0.5% Triton X-100, 10% glycerol (vol:vol), 2 µg/ml leupeptin, 100 µg/ml PMSF, and 2 µg/ml aprotinin]. Lysates were then spun at either 16,000 g at 4°C for 30 min (FAT/CD36), 13,000 g at 4°C for 5 min (COX IV, PGC-1{alpha}, GLUT4), or 16,000 g at 4°C for 60 min (total TG lipase). The supernatant was subsequently recovered and stored at –80°C before total protein content was analyzed with the use of the bicinchoninic acid method (Pierce Chemical, Rockford, IL). Muscle lysate containing either 50–60 µg protein (FAT/CD36, COX IV, PGC-1{alpha}, and GLUT4) or 120 µg protein (total TG lipase) was solubilized in 4x Laemmli buffer and heated at 95°C for 5 min before being resolved by SDS-PAGE on either 10% (total TG lipase) or 12% (FAT/CD36, COX IV, PGC-1{alpha}, GLUT4) polyacrylamide gels. Proteins were then transferred onto either activated polyvinylidene difluoride (PVDF) (FAT/CD36, COX IV, PGC-1{alpha}, GLUT4) or nitrocellulose (total TG lipase) membranes before either 1-h (COX IV, PGC-1{alpha}, GLUT4, total TG lipase) or overnight (FAT/CD36) blocking in 5% skim milk solution (in 1x Tris-buffered saline with 1% Tween 20). Membranes were then exposed overnight, at 4°C, to primary antibodies for FAT/CD36 (AbCam, Cambridge, UK), COX IV (Novus Biologicals, Littleton, UK), PGC-1{alpha} (Chemicon, Temecula, CA), GLUT4 (Biogenesis, Poole, UK), or hormone-sensitive lipase (HSL; produced in house; see Ref. 41). After primary antibody exposure, membranes were exposed to appropriate anti-species horseradish peroxidase-conjugated secondary antibodies for 45–60 min at room temperature. Antibody binding was then viewed after incubation in enhanced chemiluminescence substrate (Amersham BioSciences, Castle Hill, NSW, Australia, for COX IV, PGC-1{alpha}, GLUT4; Pierce SuperSignal chemiluminescent, Rockford, IL, for FAT/CD36; or Perkin-Elmer, Rowville, Victoria, Australia, for total TG lipase) and exposure to Chemidoc XRS (Bio-Rad Laboratories, Regents Park, NSW, Australia). Bands were identified and quantified with Quantity One one-dimensional image analysis software (Bio-Rad) (FAT/CD36, COX IV, PGC-1{alpha}, GLUT4) or Kodak one-dimensional image analysis software (total TG lipase).

PKC{theta} Western blot procedure. Approximately 75 mg of red gastrocnemius muscle were homogenized in 8x buffer (as described previously) excluding Triton X-100. The samples were then spun at 100,000 g for 30 min at 4°C. The supernatant (cytosolic fraction) was then removed for storage at –80°C, and the pellet was resuspended in 4x buffer containing all homogenization buffer ingredients, including Triton X-100. This membrane fraction was then spun at 16,000 g at 4°C for 10 min before the supernatant was removed for storage at –80°C. Both the cytosolic and membrane fractions were later assayed for total protein content as described previously. Lysate containing 100 µg of protein was then solubilized in 4x Laemmli buffer before samples were vortexed and heated at 95°C for 5 min. Samples were then loaded into 10% SDS-PAGE gels for separation by electrophoresis. From this point, all procedures were the same as those outlined above for FAT/CD36 analysis. The anti-PKC{theta} primary antibody was purchased from Cell Signal (Danvers, MA).

AMP-activated protein kinase and acetyl-CoA carboxylase Western blot procedure. Red gastrocnemius muscle (50 mg) was homogenized by pestle in 1:10 wt/vol lysis buffer A (50 mM Tris, pH 7.5, 1 mM EDTA, 1 mM EGTA, 50 mM NaF, 5 mM NaPO7, 10% glycerol, 1% Triton X-100, 10 µg/ml trypsin inhibitor, 2 µg/ml aprotinin, 1 mM benzamidine, and 1 mM PMSF). Proteins were extracted by rotating the homogenate at 4°C for 20 min followed by centrifugation at 14,000 rpm for 20 min and retention of the supernatant. Protein concentration was determined with the bicinchoninic acid method (Pierce). For the measurement of AMP-activated protein kinase (AMPK) subunits by Western blot, 75 µg of protein were separated on 10% SDS-PAGE gels and then transferred to Immobilon-FL membranes (Millipore, Billerica, MA). Primary antibodies to detect AMPK-{alpha}1, AMPK-{alpha}2, total AMPK-{alpha}, and the phosphorylation of AMPK at Thr172 were used at 1/2,000 dilution and have been described previously (24).

For the measurement of phosphorylated acetyl-CoA carboxylase (ACC), 75 µg of protein were separated on 7% SDS-PAGE gels and an ACC{alpha}-pSer79 phosphospecific antibody was used as described previously (7). Total ACC was measured with the use of a conjugated streptavidin antibody (Rockland Immunochemicals, Gilbertsville, PA). All immunoblots were analyzed with the Odyssey infrared imaging system (LI-COR Biosciences, Lincoln, NE).

TG lipase activity. TG lipase activity was determined as described previously (39, 42). Briefly, an aliquot of powdered skeletal muscle (~30 mg) was homogenized for 10 s on ice in 20 vol of homogenizing buffer using a Polytron homogenizer at full speed. After centrifugation (16,000 g at 4°C for 60 min), the supernatant was removed and stored on ice for immediate analysis. A substrate consisting of 5 mmol/l triolein, 14 x 106 dpm [9,10-3H]triolein, 0.6 mg phospholipid (phosphatidylcholine-phosphatidylinositol 3:1 wt/vol), 0.1 M potassium phosphate, and 20% BSA was emulsified by sonication. The supernatant was incubated at 37°C with enzyme dilution buffer and 100 µl of triolein substrate. The reaction was stopped after 1 h by the addition of 3.25 ml of a methanol-chloroform-heptane (10:9:7 vol/vol/vol) solution, and 0.1 M potassium carbonate-0.1 M boric acid (1.1 ml) was added to facilitate the separation of the organic and aqueous phases. The mixture was mixed on a vortex and spun in a centrifuge at 1,100 g for 20 min, and 1 ml of the upper phase containing the released fatty acids was removed for determination of radioactivity on a beta spectrometer (Tri-Carb 1500, Packard, Canberra, Australia). Activity was normalized to the total protein content of the homogenate.

Statistical analysis. All data are expressed as means ± SE. Statistical differences among groups were determined by one-way ANOVA. Where main effects were detected, a Student-Newman-Keuls post hoc test was used to probe for differences. Statistical significance was accepted at P ≤ 0.05.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Physiological characteristics and blood profiles. All rats were the same age (~7 mo) at time of death. LCR animals were heavier than HCR (275 ± 22 vs. 219 ± 21 g; P < 0.05). Table 1 shows plasma measurements.


View this table:
[in this window]
[in a new window]

  		Table 1. Plasma variables in animals selectively bred for low and high aerobic capacity

 
Intramuscular metabolites and TG lipase activity. LCR had significantly greater (47%) intramuscular TG (P < 0.05) and DAG (57%) (P < 0.05) percentages that HCR animals. There was no difference between phenotypes in intramuscular ceramide content (Fig. 1). Muscle glycogen (130 ± 16 vs. 126 ± 11 mmol/kg dry muscle wt) was also not different between phenotypes. LCR had reduced TG lipase activity (43%) (P < 0.01) when expressed relative to total protein content (see GoFig. 3). HSL is a major TG lipase in skeletal muscle, and consistent with the TG lipase activity measure its protein expression tended (P = 0.10) to be greater in HCR than in LCR animals. When animals from both phenotypes were combined in analyses, there was a strong negative correlation (R = –0.71, P < 0.05) between TGm and PGC-1{alpha} content (see Fig. 4).


Figure 1
View larger version (9K):
[in this window]
[in a new window]

  		Fig. 1. Muscle triglyceride (TGm), diacylglycerol (DAG), and ceramide content in animals selectively bred for low-capacity (LCR; black bars, n = 9) or high-capacity (HCR; white bars, n = 7) running. dm, dry muscle wt. Values are means ± SE. *P < 0.05.

 

Figure 2
View larger version (61K):
[in this window]
[in a new window]

  		Fig. 2. Representative Western blot images of selected proteins under investigation. ACC, acetyl-CoA carboxylase; AMPK, AMP-activated protein kinase; COX IV, cytochrome oxidase subunit IV; FAT/CD36, fatty acid translocase/CD36. Cyt, cytosolic; Mem, membrane. *P < 0.05 significantly different between phenotypes.

 

Figure 3
View larger version (9K):
[in this window]
[in a new window]

  		Fig. 3. Muscle FAT/CD36 content and triglyceride (TG) lipase activity in LCR (n = 7) and HCR (n = 7) animals. AU, arbitrary units. Values are means ± SE. *P < 0.05.

 

Figure 4
View larger version (7K):
[in this window]
[in a new window]

  		Fig. 4. Relationship between TGm and peroxisome proliferatory-activated receptor-{gamma} coactivator 1{alpha} (PGC-1{alpha}) in LCR (bullet; n = 6) and HCR ({circ}; n = 5) animals. R value = –0.71, R2 = 0.504, P < 0.05.

 
Skeletal muscle protein expression. Compared with HCR, LCR exhibited reduced protein content of both COX IV (P < 0.05) and PGC-1{alpha} (P < 0.05). LCR had greater muscle FAT/CD36 content than HCR animals (P < 0.05). There were no differences in GLUT4, cytosolic PKC-{theta}, membrane PKC-{theta}, AMPK-{alpha}1, AMPK-{alpha}2, AMPK Thr172 phosphorylation/total AMPK, or ACC{alpha} Ser79 phosphorylation/total ACC protein expression between groups (Fig. 2).


   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Low aerobic exercise capacity (16) and coordinated defects in oxidative metabolism (25, 28) are strongly linked to the pathogenesis of several chronic metabolic disease states, including insulin resistance, type 2 diabetes, and coronary heart disease (4). As such, it appears that the ability of an organism to utilize oxygen during exercise represents a point of divergence for health-disease outcomes (14). The results from the present study support the hypothesis that the low aerobic capacity phenotype of the LCR is related to alterations in lipid-handling capacity and that this increased metabolic load is associated with an elevated risk of metabolic diseases.

In agreement with Wisloff et al. (43) who studied animals from generation 11, we found that generation 16 LCR rats were insulin resistant, as demonstrated by higher fasting glucose, insulin, and lipid concentrations (Table 1). A novel finding from the present study was the elevated muscle lipid content in LCR compared with HCR animals (Fig. 1). Accumulation of TGm has been strongly associated with both whole body and skeletal muscle insulin resistance in a variety of animal models (17, 19, 26). However, the proposed relationship between TGm accumulation and skeletal muscle insulin resistance is equivocal, as endurance-trained humans are markedly insulin sensitive despite elevated TGm (10). The greater insulin sensitivity in the face of elevated TGm deposition in the endurance-trained state has been described as a "metabolic paradox" that represents an adaptive response to training, allowing a greater contribution of the TGm pool as a substrate source during exercise and thus "sparing" glycogen stores. In contrast, elevated TGm stores in the insulin-resistant state appear to be secondary to a structural imbalance between plasma free fatty acid availability, fatty acid uptake and storage, and oxidation.

In this context, it has been proposed that TGm merely acts as a marker for the presence of other, more metabolically active lipid intermediates, which are directly linked to defects in insulin signaling and may play causative roles in obesity-induced insulin resistance (34). One such lipid intermediate is DAG, which is elevated in both genetic and diet-induced insulin resistance (40). DAG is proposed to induce insulin resistance by activating DAG-sensitive PKC isoforms (33), which results in serine phosphorylation of insulin receptor substrate 1 (22). We found that, compared with HCR rats, muscle from LCR animals had a twofold increase in both TGm and DAG (Fig. 1). However, despite the marked elevation of these lipids in LCR rats, we did not find any differences in PKC{theta} protein content in muscle from either phenotype. In support of this anomaly, we have previously reported increased DAG content in muscle from obese Zucker rats independent of changes in membrane-associated PKC{theta}, PKC{alpha}/beta, or PKC{delta} content (21). Ceramide, a second messenger in the sphingomyelin signaling pathway, has also been shown to be elevated in the muscle of obese, insulin-resistant rodents (37) but was similar between LCR and HCR animals in the present investigation.

In an effort to determine the potential mechanism(s) mediating the elevations in TGm and DAG content observed in the LCR, we examined several steps regarded as important for the transport and/or degradation of triacylglycerol in skeletal muscle. We speculated that the observed increases in TGm and DAG content that the low-exercise capacity of the LCR may have resulted from alterations in regulatory enzymes of lipid turnover, such as the TG lipase (e.g., HSL). Because an increased rate of fatty acid transport into skeletal muscle has also been linked with the accumulation of intramuscular lipids and insulin resistance, we also measured the putative fatty acid transporter FAT/CD36. We show that, in muscle from LCR rats, there is a twofold reduction in TG lipase activity and a significant (23%) increase in FAT/CD36 protein content compared with HCR rats (Fig. 3). The lipid-regulatory enzymes play important roles in the response to various physiological conditions such as feeding/fasting and exercise and directly affect plasma free fatty acid concentrations (20), which in turn are known to determine carbohydrate and lipid utilization, storage, and synthesis in both skeletal muscle and liver. Any imbalances between lipid accumulation and fat oxidation and/or turnover in muscle due to the dysregulation of these enzymes might contribute to the development of obesity and related disorders. Indeed, in HSL–/– mice, there is marked DAG accumulation and insulin resistance (11). In the present study, the downregulation of HSL protein content and TG lipase activity is, in part, likely to be responsible for DAG accretion in the LCR rats. With regard to the protein levels of the putative fatty acid transporter FAT/CD36, Chabowski et al. (6) recently reported that, in muscle from obese, insulin-resistant Zucker rats, FAT/CD36 expression and plasmalemmal content are upregulated before the onset of diabetes. Thus a small but early increase in FAT/CD36 may precede the clinical staging of overt conditions such as Type 2 diabetes and may be reversible with interventions such as exercise training. Although Chabowski et al. found an inverse relationship between FAT/CD36 and GLUT4 protein expression in obese Zucker rats, we did not find any differences in either GLUT4 protein content or muscle glycogen levels between LCR and HCR phenotypes.

AMPK is an energy sensor that regulates cellular metabolism (for review, see Ref. 23). When activated by an energy charge disruption, AMPK stimulates glucose uptake and lipid oxidation to produce energy while turning off energy-consuming processes, including glucose and lipid production to restore energy balance (23). The AMPK pathway has profound effects on the regulation of lipid metabolism. Fatty acid oxidation in skeletal muscle involves a rate-controlling step that is regulated by carnitine palmitoyltransferase 1. Carnitine palmitoyltransferase 1 transfers long-chain acyl-CoA into the mitochondria, and this process is inhibited allosterically by malonyl-CoA, synthesized by ACC (13). The activity of ACC is regulated by reversible phosphorylation, and AMPK directly phosphorylates and inactivates this downstream target (23). The ability of AMPK to induce lipid oxidation and thus lower skeletal muscle and liver lipid deposition is considered an important feature for the insulin-sensitizing effect of AMPK activation (13). Hence, given the lipid accretion in LCR and the associated insulin resistance, it was surprising to find no differences in the basal phosphorylation state of both AMPK-{alpha}1 and AMPK-{alpha}2 isoforms between phenotypes. ACC phosphorylation state was also similar between LCR and HCR.

Wisloff et al. (43) first showed that LCR animals have compromised skeletal muscle mitochondrial function relative to HCR rats. These workers reported that HCR animals have higher muscle content of proteins known to be integral for mitochondrial function, including the peroxisome proliferatory-activated receptors and PGC-1{alpha}, both transcription regulators for the expression of metabolic and mitochondrial genes. In agreement, we report reduced protein content of both COX IV and PGC-1{alpha} in LCR compared with HCR rats. The compromised oxidative function in these rats is consistent with previous observations of mitochondrial dysfunction in several metabolic disease states, such as insulin resistance (30, 32). Such compromised oxidative function occurs in conjunction with reduced content of oxidative-type proteins in LCR rats. In this regard, Benton et al. (2) demonstrated a strong, negative relationship between TGm content and PGC-1{alpha} expression in FAT/CD36 knockout mice and concluded that PGC-1{alpha} expression is downregulated when triacylglycerol synthesis rates rise and, conversely, that PGC-1{alpha} expression is upregulated when triacylglycerol synthesis rates are reduced. They speculated that "muscle lipid sensing" may be involved in regulating the protein expression of PGC-1{alpha} in skeletal muscle. In agreement with this observation, we found a robust correlation between TGm and PGC-1{alpha} expression in LCR and HCR rats (Fig. 4).

In conclusion, we provide novel evidence that the LCR phenotype is associated with aberrant lipid handling in skeletal muscle and that these factors combine to create a differential load of metabolic risk factors previously shown to be strongly related with several chronic disease states (3, 29). Although our observations cannot prove a direct cause-effect relationship, they provide further support that low intrinsic aerobic capacity is associated with altered lipid homeostasis and that these factors may be mechanistically linked. The LCR/HCR model provides a unique platform for the discovery of the genetic and environmental causes of complex disease at all levels of biologic organization.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by a RMIT VRI II Grants Scheme (J. A. Hawley), National Heart, Lung, and Blood Institute Grant HL-64270, and National Center for Research Resources Grant RR-17718, a component of the National Institutes of Health.


   ACKNOWLEDGMENTS
 
The authors thank Dr. Sarah Lessard, Dr. Vernon Coffey, Lori Gilligan, and Ashley Duval for excellent technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. A. Hawley, Exercise Metabolism Group, School of Medical Sciences, RMIT Univ., PO Box 71, Plenty Road, Bundoora, Victoria 3083, Australia (e-mail: john.hawley{at}rmit.edu.au)

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
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Beaudet AL, Scriver CR, Sly WS, Valle D. Genetics, biochemistry, and molecular bases of variant human phenotypes. In: The Metabolic and Molecular Bases of Inherited Disease, edited by Scriver CR BA, Sly WS, Valle D, Stanbury JB, Wyngaarden JB, Fredrickson DS. New York: McGraw-Hill, 1995, p. 79.
  2. Benton CR, Han XX, Febbraio M, Graham TE, Bonen A. Inverse relationship between PGC-1{alpha} protein expression and triacylglycerol accumulation in rodent skeletal muscle. J Appl Physiol 100: 377–383, 2006.[Abstract/Free Full Text]
  3. Blaak EE. Metabolic fluxes in skeletal muscle in relation to obesity and insulin resistance. Best Pract Res Clin Endocrinol Metab 19: 391–403, 2005.[CrossRef][Medline]
  4. Booth FW, Gordon SE, Carlson CJ, Hamilton MT. Waging war on modern chronic diseases: primary prevention through exercise biology. J Appl Physiol 88: 774–787, 2000.[Abstract/Free Full Text]
  5. Centers for Disease Control and Prevention (CDC). State-specific prevalence of obesity among adults—United States, 2005. MMWR Morb Mortal Wkly Rep 55: 985–988, 2006.[Medline]
  6. Chabowski A, Chatham JC, Tandon NN, Calles-Escandon J, Glatz JFC, Luiken JJFP, Bonen A. Fatty acid transport and FAT/CD36 are increased in red but not white skeletal muscle of ZDF rats. Am J Physiol Endocrinol Metab 291: E675–E682, 2006.[Abstract/Free Full Text]
  7. Chen ZP, McConnell GK, Michell BJ, Snow RJ, Canny BJ, Kemp BE. AMPK signaling in contracting human skeletal muscle: acetyl-CoA carboxylase and NO synthase phosphorylation. Am J Physiol Endocrinol Metab 279: E1202–E1206, 2000.[Abstract/Free Full Text]
  8. Duncan GE. Exercise, fitness, and cardiovascular disease risk in type 2 diabetes and the metabolic syndrome. Curr Diab Rep 6: 29–35, 2006.[CrossRef][Medline]
  9. Frayn KN, Maycock PF. Skeletal muscle triacylglycerol in the rat: methods for sampling and measurement, and studies of biological variability. J Lipid Res 21: 139–144, 1980.[Abstract]
  10. Goodpaster BH, He J, Watkins S, Kelley DE. Skeletal muscle lipid content and insulin resistance: Evidence for a paradox in endurance-trained athletes. J Clin Endocrinol Metab 86: 5755–5761, 2001.[Abstract/Free Full Text]
  11. Haemmerle G, Zimmermann R, Hayn M, Theussl C, Waeg G, Wagner E, Sattler W, Magin TM, Wagner EF, Zechner R. Hormone-sensitive lipase deficiency in mice causes diglyceride accumulation in adipose tissue, muscle, and testis. J Biol Chem 277: 4806–4815, 2002.[Abstract/Free Full Text]
  12. Hansen C, Spuhler K. Development of the National Institutes of Health genetically heterogeneous rat stock. Alcohol Clin Exp Res 8: 477–479, 1984.[ISI][Medline]
  13. Hardie DG, Sakamoto K. AMPK: A key sensor of fuel and energy status in skeletal muscle. Physiology 21: 48–60, 2006.[Abstract/Free Full Text]
  14. Hawley JA, Spargo FJ. It's all in the genes, so pick your parents wisely. J Appl Physiol 100: 1751–1752, 2006.[Free Full Text]
  15. Howlett RA, Gonzalez NC, Wagner HE, Fu Z, Britton SL, Koch LG, Wagner PD. Selected contribution: skeletal muscle capillarity and enzyme activity in rats selectively bred for running endurance. J Appl Physiol 94: 1682–1688, 2003.[Abstract/Free Full Text]
  16. Kavanagh T, Mertens DJ, Hamm LF, Beyene J, Kennedy J, Corey P, Shepard RJ. Peak oxygen consumption and cardiac mortality in women referred for cardiac rehabilitation. J Am Coll Cardiol 42: 2139–2143, 2003.[Abstract/Free Full Text]
  17. Kelley DE, Goodpaster BH, Storlien L. Muscle triglyceride and insulin resistance. Annu Rev Nutr 22: 325–346, 2002.[CrossRef][ISI][Medline]
  18. Koch LG, Britton SL. Artificial selection for intrinsic aerobic endurance running capacity in rats. Physiol Genomics 5: 45–52, 2001.[Abstract/Free Full Text]
  19. Krssak M, Roden M. The role of lipid accumulation in liver and muscle for insulin resistance and type 2 diabetes mellitus in humans. Rev Endocr Metab Disord 5: 127–134, 2004.[CrossRef][ISI][Medline]
  20. Langfort J, Ploug T, Ihlemann J, Enevoldsen LH, Stallknecht B, Saldo M, Kjaer M, Holm C, Galbo H,. Hormone-sensitive lipase (HSL) expression and regulation in skeletal muscle. Adv Exp Med Biol 441: 219–228, 1998.[ISI][Medline]
  21. Lessard SJ, Lo Guidice SJ, Lau W, Reid J, Turner N, Febbraio MA, Hawley JA, Watt MJ. Rosiglitazone enhances glucose tolerance by mechanisms other than reduction of fatty acid accumulation within skeletal muscle. Endocrinology 145: 5665–5670, 2004.[Abstract/Free Full Text]
  22. Li Y, Soos TJ, Li X, Wu J, DeGennaro M, Sun X, Littman DR, Birnbaum MJ, Polakiewicz RD. Protein kinase C {theta}inhibits insulin signaling by phosphorylating IRS1 at Ser1101. J Biol Chem 279: 45304–45307, 2004.[Abstract/Free Full Text]
  23. Long YC, Zierath JR. AMP-activated protein kinase signaling in metabolic regulation. J Clin Invest 116: 1776–1783, 2006.[CrossRef][ISI][Medline]
  24. Michell BJ, Stapleton D, Mitchelhill KI, House CM, Katsis F, Witters LA, Kemp BE. Isoform-specific purification and substrate specificity of the 5'-AMP-activated protein kinase. J Biol Chem 271: 28445–28450, 1996.[Abstract/Free Full Text]
  25. Mootha VK, Lindgren CM, Eriksson KF, Subramanian A, Sihag S, Lehar J, Puigserver P, Carlsson E, Ridderstrale M, Laurila E, Houstis N, Daly MJ, Patterson N, Mesirov JP, Golub TR, Tamayo P, Spiegelman B, Lander ES, Hirschhorn JN, Altshuler D, Groop LC. PGC-1{alpha}-responsive genes involved in oxidative phosphorylation are coordinately downregulated in human diabetes. Nat Genet 34: 267–273, 2003.[CrossRef][ISI][Medline]
  26. Pan DA, Lillioja S, Kriketos AD, Milner MR, Baur LA, Bogardus C, Jenkins AB, Storlien LH. Skeletal muscle triglyceride levels are inversely related to insulin action. Diabetes 46: 983–988, 1997.[Abstract]
  27. Passoneau LA, Lowry OH. Enzymatic Analysis: A Practical Guide. Totawa, NJ: Humana, 1993.
  28. Patti ME, Butte AJ, Crunkhorn S, Cusi K, Berria R, Kashyap S, Miyazaki Y, Kohane I, Costello M, Saccone R, Landaker EJ, Goldfine AB, Mun E, DeFronzo R, Finlayson J, Kahn CR, Mandarino LJ. Coordinated reduction of genes of oxidative metabolism in humans with insulin resistance and diabetes: potential role of PGC1 and NRF1. Proc Natl Acad Sci USA 100: 8466–8471, 2003.[Abstract/Free Full Text]
  29. Perseghin G. Muscle lipid metabolism in the metabolic syndrome. Curr Opin Lipidol 16: 416–420, 2003.
  30. Petersen KF, Befroy D, Dufour S, Dziura J, Ariyan C, Rothman DL, DiPietro L, Cline GW, Shulman GI. Mitochondrial dysfunction in the elderly: possible role in insulin resistance. Science 300: 1140–1142, 2003.[Abstract/Free Full Text]
  31. Preiss J, Loomis CR, Bishop WR, Stein R, Niedel JE, Bell RM. Quantitative measurement of sn-1,2-diacylglycerols present in platelets, hepatocytes, and ras- and sis-transformed normal rat kidney cells. J Biol Chem 261: 8597–8600, 1986.[Abstract/Free Full Text]
  32. Reznick RM, Shulman GI. The role of AMP-activated protein kinase in mitochondrial biogenesis. J Physiol 574: 33–39, 2006.[Abstract/Free Full Text]
  33. Schmitz-Peiffer C, Browne CL, Oakes ND, Watkinson A, Chisholm DJ, Kraegen EW, Biden TJ. Alterations in the expression and cellular localisation of protein kinase C isozymes epsilon and theta are associated with insulin resistance in skeletal muscle of the high-fat-fed rat. Diabetes 46: 169–178, 1997.[Abstract]
  34. Shulman GI. Cellular mechanisms of insulin resistance. J Clin Invest 106: 171–176, 2000.[ISI][Medline]
  35. Slein MW. Methods of Enzymatic Analysis. New York: Academic, 1963.
  36. Sreekumar R, Halvatsiotis P, Schimke JC, Nair KS. Gene expression profile in skeletal muscle of type 2 diabetes and the effect of insulin treatment. Diabetes 51: 1913–1920, 2002.[Abstract/Free Full Text]
  37. Turinsky J, Bayly BP, O'Sullivan DM. 1,2-Diacylglycerol and ceramide levels in rat skeletal muscle and liver in vivo. J Biol Chem 265: 733–7938, 1990.
  38. Walsh B, Hooks RB, Hornyak JE, Koch LG, Britton SL, Hogan MC. Enhanced mitochondrial sensitivity to creatine in rats bred for high aerobic capacity. J Appl Physiol 100: 1751–1752, 2006.[Free Full Text]
  39. Watt MJ, Carey AL, Wolsk-Petersen E, Kraemer FB, Pedersen BK, Febbraio MA. Hormone-sensitive lipase is reduced in the adipose tissue of patients with type 2 diabetes mellitus: influence of IL-6 infusion. Diabetologia 48: 105–112, 2004.[CrossRef][ISI][Medline]
  40. Watt MJ, Hevener A, Lancaster GI, Febbraio MA. Ciliary neurotrophic factor prevents acute lipid-induced insulin resistance by attenuating ceramide accumulation and phosphorylation of c-Jun N-terminal kinase in peripheral tissues. Endocrinology 147: 2077–2085, 2006.[Abstract/Free Full Text]
  41. Watt MJ, Holmes AG, Pinnamaneni SK, Garnham AP, Steinberg GR, Kemp BE, Febbraio MA. Regulation of HSL serine phosphorylation in skeletal muscle and adipose tissue. Am J Physiol Endocrinol Metab 290: E500–E508, 2006.[Abstract/Free Full Text]
  42. Watt MJ, Stellingwerff T, Heigenhauser GJF, Spriet LL. Effects of plasma adrenaline on hormone-sensitive lipase at rest and during moderate exercise in human skeletal muscle. J Physiol 550: 325–332, 2003.[Abstract/Free Full Text]
  43. Wisloff U, Najjar SM, Ellingsen O, Haram PM, Swoap S, Al-Share Q, Fernstrom M, Rezaei K, Jun Lee S, Gerard Koch L, Britton SL. Cardiovascular risk factors emerge after artificial deletion for low aerobic capacity. Science 307: 418–420, 2005.[Abstract/Free Full Text]
  44. Yang X, Pratley RE, Tokraks S, Bogardus C, Permana PA. Microarray profiling of skeletal muscle tissues from equally obese, non-diabetic insulin-sensitive and insulin-resistant Pima Indians. Diabetologia 45: 1584–1593, 2002.[CrossRef][ISI][Medline]



This article has been cited by other articles:


Home page
 Am. J. Physiol. Endocrinol. Metab.Home page
J. A. Hawley
Retraction.
Am J Physiol Endocrinol Metab, December 1, 2007; 293(6): E1845 - E1845.
[Full Text] [PDF]

Home page
 Physiol. GenomicsHome page
L. G. Koch and S. L. Britton
Evolution, atmospheric oxygen, and complex disease
Physiol Genomics, August 20, 2007; 30(3): 205 - 208.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow A corrigendum has been published
Right arrow All Versions of this Article:
292/6/E1631    most recent
00702.2006v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (2)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Spargo, F. J.
Right arrow Articles by Hawley, J. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Spargo, F. J.
Right arrow Articles by Hawley, J. A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2007 by the American Physiological Society.