Excess Lipid Availability Increases Mitochondrial Fatty Acid Oxidative Capacity in Muscle: Evidence Against a Role for Reduced Fatty Acid Oxidation in Lipid-Induced Insulin Resistance in Rodents

Eight-week-old male C57BL/6J mice and male Wistar rats were purchased from the animal resources center (Perth, Australia). The animals were kept in a temperature-controlled room (22 ± 1°C) on a 12-h light/dark cycle with free access to food and water. Mice were fed ad libitum for a period of 5 or 20 weeks with a standard lab diet (8% calories from fat, 21% calories from protein, and 71% calories from carbohydrate, 2.6 kcal/g; Gordon's Specialty Stock Feeds, Yanderra, Australia) or with a high-fat diet (45% of calories from fat, 20% calories from protein, and 35% calories from carbohydrates, 4.7 kcal/g, based on rodent diet no. D12451; Research Diets, New Brunswick, NJ). Rats were fed the standard or high-fat diet ad libitum for a period of 4 weeks. Quadriceps muscles were also obtained from 10- to 12-week-old db/db mice and obese Zucker rats and their respective lean controls. All experiments were carried out with the approval of the Garvan Institute/St. Vincent's Hospital Animal Experimentation Ethics Committee, following guidelines issued by the National Health and Medical Research Council of Australia.

Glucose tolerance tests (2g/kg glucose i.p.) were performed in overnight-fasted mice. Blood samples were obtained from the tail tip at the indicated times, and glucose levels were measured using a glucometer (AccuCheck II; Roche, New South Wales, Castle Hill, Australia). [³H]-2-deoxyglucose ([³H]-2-DOG) uptake in isolated soleus muscles was determined in sealed flasks containing pregassed (95% O₂/5% CO₂) Krebs-Henseleit buffer, pH 7.3, supplemented with 0.01% BSA, 25 mmol/l HEPES, 8 mmol/l mannitol, and 2 mmol/l pyruvate. Muscles were allowed to recover for 20 min and were incubated with or without insulin (1 mU/ml) for 40 min at 30°C. [³H]-2-DOG uptake was then measured for 16 min using 0.25 μCi/ml [³H]-2-DOG as described previously (25). Serum measurements were performed on blood collected from the chest cavity and centrifuged at 14,000g for 10 min to obtain the serum. Insulin concentrations were measured using a sensitive radioimmunoassay kit (Linco Research, St. Louis, MO). The concentration of nonesterified fatty acids was determined using a colorimetric kit (Wako Pure Chemical Industries, Osaka, Japan).

Oxygen consumption rate (Vo₂) was measured using an eight-chamber indirect calorimeter (Oxymax series; Columbus Instruments, Columbus, OH) with an airflow of 0.6 l/min. Studies were commenced after 2 h of acclimation to the metabolic chamber (20 × 10 × 12.5 cm). Vo₂ was measured in individual mice at 27-min intervals over a 24-h period under a consistent environmental temperature (22°C). During the study, mice had ad libitum access to food and water.

Fat and lean body mass was measured using dual-energy X-ray absorptiometry (Lunar PIXImus2 mouse densitometer; GE Healthcare) in accordance with the manufacturer's instructions.

Palmitate and glutamate oxidation were measured in muscle homogenates using a modified method of that described by Kim et al. (26). Briefly, muscles were homogenized in 19 volumes of ice-cold 250 mmol/l sucrose, 10 mmol/l Tris-HCl, and 1 mmol/l EDTA, pH 7.4. For assessment of substrate oxidation, 50 μl of muscle homogenate was incubated with 450 μl reaction mixture (pH 7.4). Final concentrations of the reaction mixture were (in mmol/l): 100 sucrose, 80 KCl, 10 Tris-HCl, 5 KH₂PO₄, 1 MgCl₂, 2 malate, 2 ATP, 1 dithiothreitol, 0.2 EDTA, and 0.3% fatty acid–free BSA. Substrates were 0.2 mmol/l [1-¹⁴C]palmitate (0.5 μCi) plus 2 mmol/l l-carnitine and 0.05 mmol/l coenzyme A or 10 mmol/l [1-¹⁴C]glutamate (0.1 μCi). After 90 min of incubation at 30°C, the reaction was stopped by the addition of 100 μl of ice-cold 1 mol/l perchloric acid. CO₂ produced during the incubation was collected in 100 μl of 1 mol/l sodium hydroxide. For palmitate incubations, ¹⁴C counts present in the acid-soluble fraction were also measured and combined with the CO₂ values to give the total palmitate oxidation rate. Protein content in the homogenates were measured using the Bradford method (protein assay kit; Bio-Rad Laboratories, Regents Park, Australia).

Mitochondria were isolated from the quadriceps muscle as described by Bruce et al. (27). Oxygen consumption was measured at 30°C in a Clark-type oxygen electrode (Strathkelvin Instruments, Motherwell, Scotland). The respiration medium contained (in mmol/l): 225 mannitol, 75 sucrose, 10 Tris-HCl, 10 KH₂PO₄, 10 KCl, 0.8 MgCl₂, 0.1 EDTA, and 0.3% fatty acid–free BSA, pH 7.0. Substrates were 0.3 mmol/l palmitoyl-CoA, 2 mmol/l l-carnitine, and 2 mmol/l malate or 10 mmol/l glutamate and 2 mmol/l malate. For each assay 0.1–0.3 mg mitochondria were used and state III respiration was initiated by the addition of 0.2 mmol/l ADP. Protein content of the mitochondrial preparations were measured as described above.

Powdered muscle samples were homogenized 1:19 (wt/vol) in 50 mmol/l Tris-HCl, 1 mmol/l EDTA, and 0.1% Triton X-100, pH 7.2, using a Polytron instrument (Kinematica, Littau-Lucerne, Switzerland) and were subjected to three freeze-thaw cycles. Citrate synthase, β-hydroxyacyl CoA dehydrogenase (βHAD), and medium-chain acyl-CoA dehydrogenase (MCAD) were determined at 30°C, as described previously (28,29), using a Spectra Max 250 microplate spectrophotometer (Molecular Devices, Sunnyvale, CA). Carnitine palmitoyl-transferase (CPT)-1 activity was measured at 30°C in isolated mitochondria and corrected to muscle wet weight based on recovery rates of citrate synthase (27). Enzyme activities are presented as units per gram wet weight, where units are defined as micromoles per minute.

Powdered muscle samples were resuspended in radioimmunoprecipitation assay buffer (PBS, pH 7.5; 1% nonidet NP-40; 0.5% sodium deoxy-cholate; and 0.1% SDS), supplemented with protease and phosphatase inhibitors (10 μg/ml phenylmethylsulfonyl fluoride, 10 μg/ml aprotinin, 10 μg/ml leupeptin, 1 mmol/l Na₃VO₄, and 10 mmol/l NaF) and solubilized for 2 h at 4°C. Equal amounts of tissue lysates (10 μg protein) were resolved by SDS-PAGE and immunoblotted with appropriate antibodies against PGC-1α (Chemicon International, Temecula, CA), uncoupling protein (UCP) 3 (Affinity Bioreagents, Golden, CO), cytochrome oxidase (complex IV) subunit 1 (Invitrogen, Victoria, Australia), and an antibody cocktail that recognizes several subunits of the mitochondrial respiratory chain (MS601; Mitosciences, Eugene, OR). Immunolabeled bands were quantitated by densitometry.

Data are presented as means ± SE. An unpaired Student's t test was used for comparison of relevant groups. Differences at P < 0.05 were considered to be statistically significant.

Table 1 shows the body mass, fat mass, and oxygen consumption measured in 5 and 20 weeks fat-fed mice along with their standard diet controls. Mice fed a high-fat diet for 5 weeks weighed the same as their standard diet–fed controls; however, they displayed a 2.3-fold increase (P < 0.01) in the mass of the epididymal fat pad and an increase (P < 0.01) in the percentage of whole-body adiposity by dual-energy X-ray absorptiometry scanning (Table 1). At the 20-week time point, fat-fed mice weighed on average 4.2 g more than standard diet–fed animals (P < 0.01), and this was associated with a 3.7-fold increase (P < 0.01) in epididymal fat pad mass and an increased (P < 0.01) percentage of whole-body adiposity by dual-energy X-ray absorptiometry scanning (Table 1). Brown adipose tissue mass was not significantly different between standard diet–and fat-fed mice at the 5-week time point; however, following 20 weeks of fat feeding there was a 65% (P < 0.01) increase in the size of the brown adipose depot in fat-fed mice.

We measured energy expenditure and food intake, as changes in either of these parameters may have contributed to the increased adiposity observed in the fat-fed animals. Whole-body oxygen consumption was measured over 24 h using indirect calorimetry, and we observed no difference in oxygen consumption (expressed per gram of body mass) between standard diet–and fat-fed mice (Table 1); however, when expressed per gram of lean mass there was a 20 and 19% higher Vo₂ (P < 0.01) in animals fed the high-fat diet for 5 and 20 weeks, respectively. As expected, the respiratory exchange ratio was significantly lower in the fat-fed animals, reflecting the difference in diet between the two groups (Table 1). We also measured total cage food intake and calculated the average energy intake per animal and found that at the 5-week time point fat-fed animals were consuming 12% more calories than standard diet–fed controls (11.2 ± 0.2 vs. 12.6 ± 0.5 kcal/day, n = 21, P < 0.02), while at the 20-week time point fat-fed mice consumed 30% more calories than standard diet controls (10.1 ± 0.1 vs. 13.1 ± 0.3 kcal/day, n = 12, P < 0.001).

To determine whether the increased adiposity in the fat-fed mice was associated with reduced insulin action, we examined whole-body glucose clearance during an intraperitoneal glucose tolerance test (Fig. 1). At both the 5- and 20-week time point, high-fat feeding resulted in a significant impairment in glucose clearance (Fig. 1C). In a group of 5 weeks fat-fed mice, we also observed impaired insulin action at the level of skeletal muscle, with an approximate 40% reduction in insulin-stimulated [³H]-2-DOG uptake compared with standard diet–fed controls (Fig. 1D). Fat-fed mice displayed higher circulating insulin levels after both 5 weeks (0.48 ± 0.07 vs. 0.74 ± 0.08 ng/ml, n = 6, P < 0.05) and 20 weeks (0.65 ± 0.11 vs. 1.86 ± 0.36 ng/ml, n = 5, P < 0.05) of the high-fat diet. High-fat feeding also resulted in a significant increase in circulating nonesterified fatty acid levels compared with standard diet–fed controls at both the 5-week (0.68 ± 0.05 vs. 1.12 ± 0.16 mmol/l, n = 6, P < 0.05) and 20-week (0.71 ± 0.04 vs. 0.92 ± 0.04 mmol/l, n = 4–5, P < 0.01) time points.

To determine the effect of the high-fat regime on muscle fatty acid oxidative capacity, we measured the palmitate oxidation rate in tissue homogenates. Skeletal muscle from both 5 and 20 weeks fat-fed mice displayed a significantly increased capacity for fatty acid oxidation (P < 0.05) compared with muscle from standard diet controls (Fig. 2A). Recently, it has been suggested that a reduced ratio of complete (measured as CO₂ production) to incomplete (measured as acid-soluble metabolites) fatty acid oxidation may be important in high-fat diet–induced insulin resistance (23); however, we observed no difference in this ratio between standard diet–and fat-fed mice in our assays (5 weeks: 2.2 ± 0.3 vs. 2.2 ± 0.2; 20 weeks: 2.6 ± 0.3 vs. 2.4 ± 0.2). We also measured glutamate oxidation and found no significant difference between standard diet–and fat-fed animals at either the 5-week (96 ± 8 vs. 104 ± 10 nmol/h/mg protein; standard diet versus fat) or 20-week (102 ± 13 vs. 125 ± 11 nmol/h/mg protein; standard diet versus fat) time point.

As a further measure of fatty acid oxidative capacity, we measured oxygen consumption in isolated mitochondria, with palmitoyl-CoA as the substrate. In mitochondria from 20 weeks fat-fed mice, there was a significant increase (49%; P < 0.01) in ADP-stimulated respiration rate compared with standard diet controls, while for 5 weeks fat-fed mice respiration rate was 20% higher than standard diet controls; however, this difference did not reach statistical significance (Fig. 2B). Similar to the homogenate oxidations, there was no significant difference in mitochondrial respiration when glutamate was used as an alternative substrate (data not shown).

We also measured the activity of a range of enzymes associated with fatty acid utilization and oxidative capacity, including βHAD, MCAD, CPT-1, and citrate synthase. Consistent with the results observed in the homogenate oxidations and mitochondrial respiration measurements, there was a significant increase in the activity of all of these enzymes in muscle from 5 and 20 weeks fat-fed mice compared with standard diet controls (Table 2).

Our data suggested an increase in mitochondrial content in skeletal muscle of mice fed a high-fat diet. Accordingly, we examined the protein expression of several subunits of the respiratory chain, both nuclear (30-kDa subunit of complex II, core protein 2 subunit of complex III, and the α subunit of complex V) and mitochondrial (ND6 subunit of complex I and subunit 1 of complex IV) encoded, as well as the expression of PGC-1α, given its important role in the regulation of mitochondrial biogenesis and fatty acid oxidation (15). In muscle from mice fed the high-fat diet for 5 or 20 weeks, there was increased protein expression of all subunits from the respiratory chain compared with standard diet–fed controls (Fig. 3). Consistent with the increased level of mitochondrial proteins, we found increased PGC-1α protein content in both 5 weeks (64%, P < 0.01, n = 7) and 20 weeks (42%, P < 0.01, n = 8) fat-fed animals compared with standard diet controls (Fig. 3).

Additionally, we determined the protein expression of UCP3 because although its precise function has yet to be determined, it has been suggested to be a potentially important protein for the regulation of fatty acid transport, and metabolism and its expression is increased during periods of elevated fatty acid oxidation (30,31). There was a 1.8-fold (P < 0.01, n = 6) and 2.2-fold (P < 0.01, n = 6) increase in the expression of UCP3 in the 5 and 20 weeks fat-fed mice, respectively, compared with standard diet–fed controls (Fig. 3).

To determine whether similar patterns may also be present in other rodent models of insulin resistance, we examined enzyme activity (βHAD, MCAD, and citrate synthase) and PGC-1α and mitochondrial protein expression in skeletal muscle from fat-fed Wistar rats, obese Zucker rats, and db/db mice. Compared with control animals, enzyme activity was significantly increased in muscle from fat-fed rats (15–27%), obese Zucker rats (33–41%), and db/db mice (17–46%) (Table 3). We also examined protein expression of PGC-1α, UCP3, and respiratory chain subunits (complex I and complex III) and found elevated protein levels in muscle from the insulin-resistant animals (Fig. 4).

Inappropriate lipid deposition in skeletal muscle is recognized as an important factor associated with insulin resistance (1). Recent studies (7–9,12,13) in humans have suggested that aberrant mitochondrial fatty acid metabolism may be associated with intramuscular lipid accumulation in conditions of reduced insulin action. Many of these studies, however, have been conducted in subjects with well-established insulin resistance, and whether defects in muscle mitochondrial metabolism are a cause or correlate of insulin resistance remains to be clarified.

In the current study, we examined markers of mitochondrial fatty acid metabolism in skeletal muscle from rodents, in which insulin resistance is associated with an oversupply of lipids. In C57BL/6J mice fed a high-fat diet, we observed a significant impairment in glucose tolerance and a 40% reduction in insulin-stimulated glucose uptake in skeletal muscle. Despite this, we observed increased fatty acid oxidative capacity; higher activity of βHAD, MCAD, CPT1, and citrate synthase; and elevated protein expression of PGC-1α, UCP3, and mitochondrial respiratory chain subunits in skeletal muscle from these animals. Similar findings were also observed in muscle from fat-fed Wistar rats, obese Zucker rats, and db/db mice, in which there was increased βHAD, MCAD, and citrate synthase activity and elevated expression of PGC-1α, UCP3, and subunits of the mitochondrial respiratory chain compared with respective control animals. Collectively, our findings suggest that mitochondrial fatty acid oxidative capacity is increased in skeletal muscle from insulin-resistant rodents.

Insulin resistance is associated with elevated fatty acid levels in the circulation. The increased capacity for fatty acid oxidation observed in skeletal muscle of insulin-resistant rodents in the current study is potentially a compensatory response to elevated fatty acid substrate availability. Mice with muscle-specific overexpression of lipoprotein lipase, which increases fatty acid influx into skeletal muscle, display extensive mitochondrial proliferation (32). Fatty acids have also been shown to increase PGC-1α expression in muscle cells (33) and β-cells (34), and this is associated with increased mitochondrial metabolism (33,35). The fatty acid subtype appears to be important (33), as palmitate alone reduces PGC-1α expression (36); however, this may be related to its activation of inflammatory pathways that are known to impact on PGC-1α expression (37).

The coordinated increase in the activity of β-oxidation and trichloroacetic cycle enzymes, along with the increased expression of respiratory chain subunits observed in the current study, suggest that PGC-1α is in part mediating the increase in fatty acid oxidative capacity and mitochondrial content by coactivating its known binding partners estrogen-related receptor α, PPARα, PPARδ, and nuclear respiratory factor-1 (15). Mechanistically elevated fatty acid could stimulate PGC-1α expression and increase fatty acid oxidative capacity via a number of pathways. Fatty acids are known ligands for the PPAR family of nuclear hormone receptors, and part of the increase in fatty acid oxidative capacity in the current study may be related to direct activation of PPARα or PPARδ by fatty acids. In skeletal muscle, activation of PPARδ has been shown to increase expression of PGC-1α (38). PGC-1α is also known to coactivate PPARδ (39), resulting in a feed-forward loop that stabilizes PGC-1α protein expression and drives the transcription of genes associated with fatty acid metabolism (38).

Other studies in insulin-resistant rodents have reported reduced expression of PGC-1α and other markers of mitochondrial metabolism in muscle (21–24). The reason for the disparity in results is unclear but may be related to methodological factors such as diet composition, the length of high-fat feeding, or the particular muscle groups examined (i.e., quadriceps versus gastrocnemius bundle). Furthermore, it must also be noted that many studies in rodents and humans have only examined mRNA expression for PGC-1α, and as PGC-1α is known to be posttranslationally modified, its gene expression may not always correlate with protein levels (40).

Despite our findings of elevated fatty acid oxidative capacity in insulin-resistant rodents, increased intramuscular lipid is a characteristic feature of these animal models (1,4). Excess lipid storage may be in part related to a greater increase in the efficiency of fatty acid uptake, as it has been observed in insulin-resistant rodents that there is increased clearance of fatty acid into muscle (3,4). We did not directly measure fatty acid uptake in our animals; however, adipose mass and circulating nonesterified fatty acids levels were elevated in fat-fed mice (Table 1), and the increased UCP3 protein in all of our rodent models is consistent with an increased influx of fatty acid into muscle (30,31,41). Skeletal muscle is quantitatively an important tissue for whole-body fat oxidation, and lipid overload in muscle may be linked to the reduction in muscle mass observed in insulin resistance (Table 1; [42]). Another factor to be considered is that our ex vivo measurements represent the capacity of enzymes and fatty acid oxidation pathways under favorable conditions of substrate availability, and in vivo, regulatory factors such as elevated levels of malonyl-CoA (43) or reduced activity of adiponectin and leptin signaling pathways (44) might contribute to ectopic deposition of lipids in muscle.

The proposed causative role for mitochondrial dysfunction in the development of insulin resistance is yet to be definitively demonstrated. Several studies (7–9,12,13) have reported defects in various markers of mitochondrial metabolism and biogenesis in skeletal muscle from subjects with obesity, insulin resistance, and type 2 diabetes. These results may be confounded by various disease factors; however, investigations demonstrating mitochondrial defects in first-degree relatives of patients with type 2 diabetes (6,11,12) suggest that mitochondrial dysfunction may be among the earliest defects that predisposes these subjects to lipid accumulation and insulin resistance, as opposed to increased lipid availability leading to decreased mitochondrial function and then insulin resistance. Studies in humans in which lipid availability has been experimentally altered have provided inconclusive results in this respect. Acute oversupply of lipids, via lipid infusion, has been shown to reduce gene expression for PGC-1α and mitochondrial respiratory chain components in muscle (45,46). However, 1 week of pharmacological reduction of plasma free fatty acids (and subsequently intramuscular acyl-CoA concentrations) in insulin-resistant subjects also reduced gene expression of PGC-1α and other mitochondrial markers in muscle (47). Dietary studies in humans are also equivocal with high-fat feeding studies reporting increases (48), decreases (24), or no change (49) in various markers of muscle mitochondrial metabolism. Thus, the effect of elevated lipid availability on muscle mitochondrial oxidative capacity in humans remains to be clarified.

In summary, our study demonstrates that fatty acid oxidative capacity and protein expression of PGC-1α and mitochondrial respiratory chain subunits are upregulated in skeletal muscle of a variety of rodent models of insulin resistance. We suggest that these changes likely represent a homeostatic response to attempt to compensate for elevated availability of lipids in these animals. We therefore conclude that increased lipid availability is unlikely to lead to lipid accumulation and insulin resistance via a specific effect to diminish mitochondrial fatty acid oxidative capacity.

This work was supported by the National Health and Medical Research Council of Australia (NHMRC), the Diabetes Australia Research Trust (DART), and the Rebecca L. Cooper Medical Research Foundation. N.T. and C.R.B. are supported by Peter Doherty Fellowships and G.J.C. by a Research Fellowship from the NHMRC.

The contribution of Dr. Bronwyn Hegarty, Andrew Hoy, Donna Wilks, and Ron Enriquez to specific methodological and technical aspects of the study is gratefully acknowledged. We thank the Biological Testing Facility at the Garvan Institute for their help with animal care.