Compensatory Increases in Nuclear PGC1α Protein Are Primarily Associated With Subsarcolemmal Mitochondrial Adaptations in ZDF Rats Open Access

Male control (n = 5, weighing ∼400 g) and ZDF (n = 5, weighing ∼400 g) rats were purchased from Charles River. Animals were housed in a climate- and temperature-controlled room, on a 12:12-h light-dark cycle, with rat diet and water provided ad libitum. Twenty-four–week-old animals were anesthetized with an intraperitoneal injection of sodium pentobarbital (60 mg/kg), and subsequently blood was sampled from nonfasted animals using a cardiac puncture, and muscles were rapidly excised for various measurements (described below). For the PGC1α electotransfection experiments, Sprague-Dawley rats (∼300 g) from our breeding colony were used. All facets of this study were approved by the University of Guelph Animal Care Committee.

Serum samples were analyzed for glucose using a spectrophotometric method (Sigma, St. Louis), insulin by radioimmunoassay using a rat-specific antibody (Linco, St. Charles, MO), and fatty acid concentrations using a spectrophotometric procedure (Wako Chemicals, Richmond, VA).

Samples from the red and white portions of the tibialis anterior muscle were rapidly immersed in a fixing buffer (2.5% glutaraldehyde, 1.0% parafermaldehyde in PBS) and incubated at 4°C overnight. Tissue was then washed three times in 0.1 mol/l Hepes and subsequently suspended in 1.0% osmium tetroxide for 4 h. Thereafter, tissue was washed three times in 100 mmol/l Hepes and suspended in 2% uranyl acetate for 3 h, washed three times in 0.1% Hepes, and dehydrated by incubating in a graded ethanol series (i.e., 25, 50, 75, 95, and twice in 100% ethanol). Tissue was infiltrated with resin by suspending in 50/50 ethanol/resin (London Resin Company White) for 4 h on a rotating mixer and subsequently suspended in pure resin for 4 h on a rotating mixer. Tissue was then placed in an embedding capsule containing pure resin and incubated overnight at 60°C to polymerize. Sections (100 nm) were cut and laid onto 200 mesh formvar/carbon copper grids and then stained with 2% uranyl acetate and Reynold's lead citrate. Samples were viewed on a Philips CM 10 transmission electron microscope (TEM) at 80 kV, and images were obtained with an Olympus/SIS Morada CCD camera using the Olympus/SIS iTEM software. Images were analyzed using the measurement tools provided by this software. Individual lipid droplet and mitochondrial sizes were determined repeatedly and averaged for a given image taken at 25,000× magnification. Mitochondrial subpopulation densities were determined within a defined region (10-μm² area) at a minimum of three locations within an image taken at 5,800× magnification, similar to methodologies previously published (28). Several fibers (3,–5) were imaged for each animal. Mitochondrial number and density were not determined in white muscle because of their infrequent presence within our defined region.

Intracellular triacylglycerol concentrations were determined biochemically as previously described (29). Lipids were extracted in a standard Folch solution and subsequently separated by high-performance thin-layer chromatography and quantified against known standards.

Muscle samples were homogenized in 100 vol/wt of a 100 mmol/l potassium phosphate buffer and citrate synthase (CS) activity was assayed spectrophotometrically at 412 nm (37°C) (30).

mtDNA was determined using real-time PCR, as previously reported (10), using the following primers: NADH dehydrogenase subunit 5 forward, 5′-GCAGCCACAGGAAAATCCG-3′ and reverse, 5′-GTAGGGCAGAGACGGGAGTTG-3′; and the solute carrier family 16 member 1 forward, 5′-TAGCTGGATCCCTGATGCGA-3′ and reverse, 5′-GCATCAGACTTCCCAGCTTCC-3′.

Differential centrifugation was used to obtain both subsarcolemmal and intermyofibrillar mitochondrial fractions from the red and white portions of the tibialis anterior, as we have previously published (20,31). Mitochondria were further purified using a Percoll gradient for Western blotting analysis.

Whole-muscle homogenates were prepared as previously described (20) (n = 4). Isolated mitochondria (5 μg), whole muscle (30 μg), nuclear extract (45 μg), and cytosolic protein (25 μg) were analyzed. Samples were separated by electrophoresis by SDS-PAGE and transferred to polyvinylidene difluoride membranes. The MO-25 antibody has been used previously (10,12) to detect fatty acid translocase (FAT)/CD36 (a gift from Dr. N.N. Tandon), and commercially available antibodies were used to detect cytochrome c oxidase complex IV (COXIV; Invitrogen, Burlington, ON, Canada), PGC1α (Calbiochem, LA Jolla, CA), lactate dehydrogenase (LDH; Abcam; Cambridge, MA), and histone H2B (H2B; Abcam). Blots were visualized and quantified using chemiluminescence and the ChemiGenius 2 Bioimaging System (SynGene, Cambridge, U.K.).

The forward radioisotope assay was used for the determination of carnitine palmitoyltransferase I (CPTI) activity as described by McGarry et al. (32), with minor modifications as we have previously reported (20,33). Briefly, the assay was conducted in the presence of 75 μmol/l P-CoA (l-[³H]carnitine (Amersham Bioscience, Buckinghamshire, England). Palmitoyl-[³H] carnitine was extracted in water-saturated butanol and the radioactivity determined.

Palmitate oxidation was measured in the presence [1-¹⁴C] palmitate, as previously described (34). Briefly, gaseous ¹⁴CO₂ production and isotopic fixation were determined following a 30-min reaction at 37°C in the presence of 77 μmol/l palmitate.

PGC-1α mRNA was determined in the tibialis anterior muscle using real-time PCR as we have previously reported (20). The following primer sets were used: PGC-1α forward 5′-CAATGAGCCCGCGAACATAT-3′, PGC-1α reverse 5′-CAATCCGTCTTCATCCACCG-3′ and 18S forward 5′-GTTGGTTTTCGGAACTGAGGC-3′, 18S reverse ‘5-GTCGGCATCGTTTATGGTCG-3′.

Nuclear extraction was performed using a commercial kit (Pierce Biotechnology, Rockford, IL) according to the manufacturer's specifications, as done previously (26). Harvested muscles were immediately placed in 750 μl of PBS, minced, and briefly homogenized. Cytosolic and nuclear extraction was performed using supplied reagents supplemented with 1 mmol/l sodium orthovanadate, 1 mmol/l phenylmethylsulfonyl fluoride, and 10 μg/ml of pepstain A, aprotinin, and leupeptin. Isolated nuclei were washed 15 times in alternating PBS or PBS supplemented with 0.1% Nonidet P-40. To confirm the purity of nuclear extracts, both fractions were analyzed by Western blotting for cytosolic (LDH) and nuclear proteins (H2B).

Electrotransfection experiments were performed as described by us (20,35) and others (36,37), with minor modifications. The PGC1α expression construct (a gift from Dr. B. Spiegelman, Harvard University, Boston, MA) was produced by subcloning the PGC1α coding sequence into a mammalian expression vector (pcDNA 3.0) (Invitrogen, Burlington, ON, Canada). Plasmid stocks were produced by large-scale plasmid isolation from transformed Escherichia coli cells (One-Shot; Invitrogen) using commercially available kits (GIGA-prep kits; Invitrogen).

For electrotransfection, animals were anesthetized with isoflurane and the tibialis anterior muscle injected with 100 μl hyaluronidase (0.15 units/μl in 50% vol/vol saline). Two hours later, muscles were electrotransfected with 1) PGC1α-pcDNA plasmid (500 μg PGC1α in 50% vol/vol saline) or 2) empty pcDNA3.0 plasmid (500 μg pcDNA in 50% vol/vol saline) as described previously (20,38,39). Thereafter, rats were provided with an analgesic (Temgesic) and allowed to recover for 2 weeks (40).

All data are presented as means ± SE. Unpaired t tests, paired t tests, and two-way ANOVA were used where appropriate. When significance was obtained, a Fisher's least-significant-difference post hoc analysis was completed. Statistical significance was accepted at P < 0.05.

In ZDF animals, serum insulin concentrations were markedly lower (4.8 ± 0.8 vs. 1.5 ± 0.4 ng/ml; P < 0.05), while glucose (14 ± 1 vs. 37 ± 1 mmol/l) and fatty acid (0.3 ± 0.1 vs. 0.9 ± 0.2 mmol/l) concentrations were higher (P < 0.05).

In red ZDF muscle, the area of individual lipid droplets was +47% larger than in red control muscle (P < 0.05) (Fig. 1,A). In addition, TEM images revealed a large number of lipid droplets in red ZDF muscle, which were largely in contact with intermyofibrillar mitochondria, while the subsarcolemmal mitochondrial region was devoid of lipids (Figs. 2 and 3). Biochemical extraction revealed that triacylglycerol concentration was approximately threefold higher (P < 0.05) in ZDF muscle (Fig. 1 B). In white muscle, lipid droplets were not quantifiable.

In red and white ZDF muscles, subsarcolemmal mitochondrial size was not altered (Fig. 2,A and B). However, compared with control animals, red intermyofibrillar mitochondria in ZDF animals were +35% larger (P < 0.05) (Fig. 2,A), while white muscle intermyofibrillar mitochondria in ZDF animals were −37% smaller (P < 0.05) (Fig. 2 B).

In red muscle, the number of subsarcolemmal mitochondria in ZDF animals was markedly increased (+50%; P < 0.05) (Fig. 2,C), but the number of intermyofibrillar mitochondria was unaltered (Fig. 2,C). In white muscle, this parameter was not determined because of the scarcity and diffuse nature of mitochondria in this tissue (Figs. 2 and 4).

The width of the red muscle subsarcolemmal mitochondrial subpopulation was +69% larger (P < 0.05) (Fig. 3,A), and the density of red muscle subsarcolemmal mitochondria was +57% larger (P < 0.05) (Fig. 3,B) in ZDF animals. In contrast, the density of intermyofibrillar mitochondria was not different (P > 0.05) when analyzed in regions devoid (Fig. 3,C) or abundant (Fig. 3 D) in lipid droplets.

Whole-muscle COXIV protein (Fig. 5,A) and CS maximal activity (Fig. 5,B) were not different in ZDF animals. The amount of mitochondrial DNA (mtDNA) was also not altered in ZDF rats in either red (control: 2.23 ± 0.25 vs. ZDF: 2.13 ± 0.63 arbitrary units) or white (control: 1.31 ± 0.23 vs. ZDF: 0.91 ± 0.28 arbitrary units) muscles, suggesting that mitochondrial content was unaltered. In contrast, FAT/CD36 was increased +2.4-fold in red, and +50% in white, ZDF muscles (Fig. 5 C).

In isolated mitochondria, COXIV protein was constant across all mitochondrial subpopulations (Fig. 5,D and G), while in contrast, CPTI activity (Fig. 5,E and H) and FAT/CD36 protein (Fig. 5 F and I) were greater (P < 0.05) in all mitochondrial subpopulations in ZDF animals.

In both red and white muscle, ZDF animals displayed higher (P < 0.05) rates of palmitate oxidation in isolated subsarcolemmal (∼40%) and intermyofibrillar (∼40%) mitochondria (Fig. 6 A and B). Increased ratios of ¹⁴C -acid soluble intermediates (ASMs) to ¹⁴CO₂ have previously been used to infer incomplete oxidation of ¹⁴C-palmitate (41). In red intermyofibrillar mitochondria, the ASM/CO₂ ratio was unchanged. In contrast, this ratio was decreased (∼30%; P < 0.05) in red muscle subsarcolemmal mitochondria and in white muscle subsarcolemmal and intermyofibrillar mitochondria, indicating greater complete fatty acid oxidation in these mitochondria (data not shown).

In red muscle, PGC1α mRNA was not different (P > 0.05) between control and ZDF animals (Fig. 7,A). In control animals, PGC1α mRNA was −54% lower (P < 0.05) in white, compared with red, muscle (Fig. 7,A). In contrast, this fiber type difference was lost in ZDF animals as a result of the increase (P < 0.05) in white muscle PGC1α mRNA (Fig. 7 A).

Whole-muscle PGC1α protein was not altered in ZDF animals in either red or white muscles (Fig. 7,B). White muscle, compared with red muscle, had less (P < 0.05) PGC1α protein in both control (−19%) and ZDF (−23%) animals (Fig. 7 B).

Purified nuclear extracts were devoid of LDH and contained high levels of H2B (Fig. 7,D), indicating successful enrichment of nuclear proteins and the absence of cytosolic contamination. The amount of PGC1α protein located in the nuclei was higher (P < 0.05) in both red (+68%) and white (+67%) muscles of ZDF animals (Fig. 7,C). A fiber type difference existed in nuclear PGC1α protein, as white muscle contained less (P < 0.05) PGC1α nuclear protein in both control (−32%) and ZDF (−32%) animals (Fig. 7 C).

To test the notion that PGC1α preferentially targets subsarcolemmal mitochondria, we electrotransfected PGC1α cDNA into the tibialis anterior muscle of lean animals and determined the effect on FAT/CD36, a target protein. Transfection increased the mRNA (+29%) (Fig. 8,A), total protein (+22%) (Fig. 8,B), and the nuclear content of PGC1α (+15%) (Fig. 8,C). Subsequently, whole-muscle FAT/CD36 content was increased (P < 0.05) +30% (control: 100 ± 6.1 vs. transfected: 130 ± 18.4 arbitrary units), as was subsarcolemmal mitochondrial FAT/CD36 content (control: 100 ± 10.5 vs. transfected: 117 ± 15.5 arbitrary units) but not intermyofibrillar FAT/CD36 (control: 105.4 ± 6.3 vs. transfected: 98 ± 3.9 arbitrary units). Rates of palmitate oxidation increased (P < 0.05) +37% in subsarcolemmal mitochondria, but PGC1α transfection had no effect (P > 0.05) on intermyofibrillar mitochondria (Fig. 8 D). Around 30% of muscle fibers are transfected with our procedures (data not shown), and therefore we could not determine mitochondrial content in various subpopulations in transfected muscle fibers, as with TEM imaging one cannot determine which fibers have been affected.

The novel findings of the current study are that skeletal muscle from ZDF rats 1) have larger and more prevalent lipid droplets and 2) preferentially display compensatory increases in subsarcolemmal mitochondrial number, width, density, as well as fatty acid oxidation rates, which potentially result from 3) an increased nuclear content of PGC1α that appears to target subsarcolemmal mitochondria.

The original hypothesis of a mitochondrial dysfunction in fatty acid oxidation was partially based on observations of smaller subsarcolemmal mitochondria in insulin-resistant muscle (4). However, the current TEM images do not support the notion of smaller mitochondria with insulin resistance, as in red muscle subsarcolemmal mitochondrial size, was unchanged and intermyofibrillar mitochondrial size was actually increased in ZDF animals. In addition, it was not previously understood if mitochondrial size directly impacted mitochondrial oxidation rates, making the previous observations of reduced mitochondrial size (4) difficult to interpret. The current data suggests that mitochondrial size does not influence mitochondrial palmitate oxidation, as these rates were increased when the size of mitochondria were unaltered (red and white subsarcolemmal mitochondria), increased (red intermyofibrillar mitochondria), as well as decreased (white intermyofibrillar mitochondria).

Analysis of TEM images showed that the number (+50%), width (+69%), and density (+57%) of subsarcolemmal mitochondria were increased in ZDF animals. In contrast, ZDF intermyofibrillar mitochondrial number and density were not changed, nor were markers of mitochondrial content (mtDNA, CS, and COXIV). These data show that mitochondrial subpopulations can respond differently in insulin-resistant muscle, and whole-muscle measures of mitochondrial content cannot reveal the subtle differences of various signals that induce mitochondrial proliferation.

Previously, indirect assessments suggested that there was an intrinsic dysfunction in fatty acid oxidation within mitochondria (4). However, the current data do not support this notion, as subsarcolemmal and intermyofibrillar mitochondria, from both red and white muscles of ZDF animals, displayed ∼40% increased rates of palmitate oxidation. These data are consistent with more contemporary mitochondrial literature, as it has recently been shown that mitochondrial fatty acid oxidation, when measured in isolated mitochondria or permeabilized fibers, is not reduced in mitochondria of obese and type 2 diabetic individuals (12,–14). Rodent models of insulin resistance have also supported the notion that downregulation of mitochondrial fatty acid oxidation is not a requirement for lipid accumulation and impairments in insulin signaling. High-fat feeding has been shown to induce insulin resistance while increasing mitochondrial content (9,11). Magnetic resonance spectroscopy has suggested that fatty acid oxidation is not compromised in diabetic animals (42), and we have recently shown in both red and white muscles of obese Zucker rats that fatty acid oxidation was increased, not decreased, in subsarcolemmal mitochondria (10). Thus, our work (10,12) and that of others (9,13,14,43), indicates that in both human (12,–14) and animal models of insulin resistance (9,43) there is little evidence to support the view that an intrinsic impairment in the ability of mitochondria to oxidize fatty acids accounts for the increased intramuscular lipid accumulation associated with insulin resistance. Instead, evidence in both human (44) and rodent (10,45) models of insulin resistance suggests that plasma membrane fatty acid transport is increased as a result of an increased content of fatty acid transport proteins. Despite the potential compensatory adaptations in mitochondrial fatty acid oxidation, the large changes in plasma membrane fatty acid transport have been proposed to create an imbalance between delivery and utilization such that lipids accumulate (10).

In the current study, the increase in palmitate oxidation in ZDF mitochondria may result from the observed increase in 1) CPTI activity and/or 2) mitochondrial FAT/CD36 protein. The rate-limiting step in fatty acid transport/oxidation has long been attributed to CPTI activity, and the changes in CPTI activity mimicked the trends in mitochondrial fatty acid oxidation in both red and white muscle. However, the notion that CPTI activity represents the only regulatory site in mitochondrial fatty acid oxidation has been challenged, as several laboratories have recently found the fatty acid transport protein FAT/CD36 on mitochondrial membranes. Gain-of-function (46) and loss-of-function (34) molecular approaches, as well as physiological perturbations (31), have suggested that mitochondrial FAT/CD36 has a role in regulating mitochondrial fatty acid oxidation. While FAT/CD36 appears to have a role in regulating mitochondrial fatty acid oxidation, this is likely mediated in a concerted fashion with additional proteins, as mitochondrial FAT/CD36 and palmitate oxidation rates do not correlate under basal conditions (47) but rather multiple regression approaches that take into consideration both CPTI and mitochondrial FAT/CD36 highly correlate with mitochondrial fatty acid oxidation rates (47). Therefore, in the current study, the increase in both CPTI activity and mitochondrial FAT/CD36 may account for the observed changes in mitochondrial palmitate oxidation rates in ZDF animals, both of which are transcribed by PPARs and mediated by PGC1α.

It has been suggested that reductions in PGC1α expression has a role in the etiology of insulin resistance (21,22). However, in the current study we have found that PGC1α mRNA and total muscle protein were not reduced in ZDF muscles. However, the subcellular distribution of PGC1α did differ, as the nuclear content was markedly increased (∼70%). Others (6,23,24) have also now found that PGC1α is not reduced with insulin resistance in human skeletal muscle. The mechanism(s) triggering the PGC1α translocation to the nucleus are unknown. However, muscle contraction, through calcium-mediated signaling, has previously been shown to induce nuclear translocation of PGC1α (26), and therefore alterations in cytosolic calcium levels represent a potential mechanism of action.

Interestingly, the increase in nuclear PGC1α was associated with increased proliferation of the ZDF subsarcolemmal mitochondria not intermyofibrillar mitochondria. PGC1α overexpression experiments revealed that PGC1α targets subsarcolemmal mitochondria, as palmitate oxidation rates and FAT/CD36 content were only increased in subsarcolemmal, not intermyofibrillar, mitochondria. This was also observed previously in another study (20). However, in ZDF animals in the current study, intermyofibrillar mitochondrial CPTI, FAT/CD36, and rates of fatty acid oxidation were also all increased in conjunction with increased nuclear PGC1α. Proliferation of intermyofibrillar mitochondria is also possible in PGC1α transgenic animals (25). This may suggest that a greater increase in nuclear PGC1α is required to upregulate intermyofibrillar mitochondria, as transfection only had a modest affect on nuclear PGC1α (+15%) compared with the in vivo ZDF (+70%) and, arguably, the transgenic conditions. Alternatively, the apparent argeting of PGC1α to subsarcolemmal mitochondria may be a result of the proximity of nuclei to the subsarcolemmal subpopulation and differences in the rate of protein import into mitochondrial subpopulations (48). Regardless of the exact mechanism, more pronounced changes occur in subsarcolemmal mitochondria in response to increased nuclear PGC1α content in both ZDF and electrotransfected rats.

While ZDF animals share a number of similar traits with human type 2 diabetes, including hyperglycemia, insulin resistance (49), hyperlipidemia (50), increased plasmalemmal fatty acid transport, and intramuscular lipids (44), we recognize that the ZDF animal model is not fully representative of human type 2 diabetes. Therefore, it will be important to discern whether the current observation in ZDF animals extend to human type 2 diabetic individuals.

In summary, we show that in ZDF rats 1) intramuscular triacylglycerol accumulates, almost exclusively in the intermyofibrillar region, as a result of an increase in both the size and number of lipid droplets. In an attempt to compensate for the increased plasma membrane fatty acid transport and lipid delivery (as we have shown previously [45]), in ZDF muscle the there was an increase in 2) subsarcolemmal mitochondrial number, size, and density; 3) subsarcolemmal and intermyofibrillar CPTI activity and FAT/CD36 protein; and 4) mitochondrial palmitate oxidation rates in subsarcolemmal and intermyofibrillar mitochondria. The increase in ZDF mitochondrial oxidative capacity was greater in subsarcolemmal mitochondria, as the increase in mitochondrial fatty acid oxidation rates is amplified by the increase in subsarcolemmal mitochondrial number. These changes may result in part from 5) an increased nuclear translocation of PGC1α protein, which preferentially targets subsarcolemmal mitochondria.

This work was funded by the Canadian Institutes of Health Research (to A.B.) and the Natural Sciences and Engineering Research Council of Canada (to A.B. and G.H.).

No potential conflicts of interest relevant to this article were reported.

We express our appreciation to Robert Harris for his assistance with the TEM experiments.