At the molecular and structural level, mitochondrial biogenesis and mitochondrial function are altered in diabetes, as well as in insulin-resistant relatives of type 2 diabetic subjects (1,2). At the ultra-structural level, a reduction in the number, location, and morphology of mitochondria is strongly associated with insulin resistance (1). Two recent microarray studies have shown that genes involved in oxidative phosphorylation (OXPHOS) exhibit reduced expression levels in the skeletal muscle of type 2 diabetic subjects and prediabetic subjects. These changes may be mediated by the peroxisome proliferator-activated receptor [gamma] coactivator-1 (PGC1) pathway. PGC1[alpha]- and PGC1[beta]-responsive OXPHOS genes show reduced expression in the muscle of patients with type 2 diabetes (3,4). In addition to the cellular energy sensor AMP kinase, the peroxisome proliferator-activated receptor cofactors PGC1[alpha] (5-7) and possibly PGC1[beta] (8) activate mitochondrial biogenesis and increase OXPHOS gene expression by increasing the transcription, translation, and activation of the transcription factors necessary for mitochondrial DNA (mtDNA) replication. Similarly, PGC1[alpha] increases the transcription of enzymes necessary for substrate oxidation, electron transport, and ATP synthesis. Morphological and functional studies (1,9,10), combined with the recent microarray data, indicate that PGC1 is important in the development of type 2 diabetes.
Rates of ATP synthesis, measured in situ with magnetic resonance spectroscopy, are decreased in subjects with a family history of diabetes before the onset of impaired glucose tolerance (2,10). Based on these results, the prevailing view is that these defects have a genetic origin (2). One common feature of diverse insulin-resistant states is an elevation in nonesterified fatty acids (11). This gave rise to the concept of "lipotoxicity" and "ectopic fat" (12) and shifted attention toward the adipose tissue and increased free fatty acid concentrations as a potential foundation for insulin resistance (11).
Excess dietary fat has also been implicated in the development of obesity and diabetes (13). At energy balance, high-fat diets (HFDs) increase the flux of fatty acids through skeletal muscle for oxidation. The purpose of these experiments was to identify the transcriptional responses of skeletal muscle to an isoenergetic HFD in healthy young men using oligonucleotide microarrays. We found a HFD downregulated PGC1 and PGC1[beta] mRNA, as well as genes encoding proteins in complexes I, II, III, and IV of the electron transport chain. These changes were recapitulated and amplified in a murine model after a 3-week HFD, along with decreases in PGC1[beta] and cytochrome C protein. These studies implicate increased dietary fat in the defects in OXPHOS genes observed in diabetes and the prediabetic/insulin-resistant state.
RESEARCH DESIGN AND METHODS
ADAPT is a short-term intervention study designed to examine inter-individual differences in the ability to increase fat oxidation after consuming an isoenergetic HFD (14,15). Ten healthy young men, aged 23.0 [+ or -] 3.1 years and with BMIs of 24.3 [+ or -] 3.0 kg/[m.sup.2] underwent physical examination, medical laboratory tests, and anthropometry. The subjects had high preference for dietary fat as indicated by food preference questionnaire (114.16 [+ or -] 18.14). This measure is highly correlated with serf-selected fat intake (16). Participants presented to Pennington inpatient unit on day -4 and ate a weight-maintaining (35% fat, 16% protein, and 49% carbohydrate) diet prepared by the metabolic kitchen. On day -3, a euglycemic-hyperinsulinemic clamp was performed, and this diet was continued on days -2, -1, and 1. On day 1, participants ate the same weight-maintaining 35% fat diet, and total dally energy expenditure, fat oxidation, protein oxidation, and carbohydrate oxidation were measured at energy balance as previously described in a wholeroom calorimeter (17,18). On days 2, 3, and 4 subjects ate a 50% fat, 16% protein, 34% carbohydrate diet with an isoenergetic energy clamp procedure. Each meal was served to the subjects and consumed within 20 min.
Euglycemic-hyperinsulinemic clamp, insulin sensitivity was measured only at baseline (day -3) by euglycemic-hyperinsulinemic clamp (19) before HFD. After an overnight fast, glucose and insulin (80 mIU/[m.sup.2] BSA) were administered. The glucose disposal rate (M value) was adjusted for kilograms of lean body mass.
Maximal aerobic capacity. Maximal oxygen uptake was determined by a progressive treadmill test to exhaustion (20). The volume of [O.sub.2] and C[O.sub.2] was measured continuously using a metabolic cart (V-Minx29 Series; SensorMedics, Yorba Linda, CA).
Body composition. Body fat mass and lean body mass were measured on a Hologic Dual Energy X-ray Absorptiometer (QDR 4500; Hologic, Waltham, MA).
Indirect calorimetry. Respiratory quotient and 24-h energy expenditure were determined in the whole-room calorimeter before and during 3 days of isocaloric HFD and confirmed an increase in fat oxidation (data not shown). Energy expenditure was set at 1.4 times the resting metabolic rate measured by metabolic cart and clamped across the 4-day chamber stay.
Animal study. Male C57BL/6J mice were housed at room temperature with a 12-h-light/12-h-dark cycle for 5 weeks. Six mice consumed control diet ad libitum (D12450B [Research Diets, New Brunswick, NJ]: 10% of energy from fat, 20% of energy from protein, and 70% of energy from carbohydrate), and seven mice consumed HFD (D12451 [Research Diets]: 45% of energy from fat, 20[degrees]/5 of energy from protein, and 35% of energy from carbohydrate). All animals ate the control diet ad libitum for 2 weeks, and seven were switched to HFD for 3 additional weeks. Gastrocnemius muscles were dissected and snap-frozen in liquid nitrogen.
RNA and DNA extraction. Human and mouse total RNA from 50-100 mg of vastus lateralis and gastrocnemius muscle, respectively, was isolated with Trizol reagent (Invitrogen, Carlsbad, CA). Gastrocnemius was digested overnight in proteinase K (FisherBiotech, Houston, TX) at 55[degrees]C. DNA was extracted with phenol-chloroform. The quantity and integrity of the RNA and DNA were confirmed by Agilent 2100 Bioanalyzer (Agilent Technologies, Palo Alto, CA).
Oligonucleotide microarrays. RNA sample pairs (2 [micro]g) from human subjects were labeled by reverse transcriptase with dCTP-Cy3 and dCTP-Cy5, respectively, and in inverse order (dye swap) using MICROMAX TSA Labeling and Detection kit (Perkin-Elmer, Wellesley, MA). Equal amounts of labeled cDNA probes were hybridized in duplicate to oligonucleotide slides containing 18,861 spots corresponding to 17,260 unique genes (Compugen, San Jose, CA) in hybridization chambers (GenomicSohitions, Ann Arbor, MI) for up to 72 h at 42[degrees]C. Detection and washing were performed at room temperature according to manufacturer's protocol (Perkin-Eimer). Oligonucleotide chips were spotted on to poly-L-lysine slides using a GeneMachine OmniGrid microarrayer (GenomicSolutions) equipped with a Stealth SPH32 printhead and Stealth SMP4 Micro Spotting Pins (Telechem International, Sunnyvale, CA). Oligonucleotides were stored in 384-well plates in 45% DMSO. Microarray slides were scanned using a GSI Lumonics ScanArray 5,000 scanner (PerkinElmer) at high intensities (~95% for Cy3, ~75% for Cy5) and low intensities (~55% for Cy3, ~35% for Cy5) applying ScanArray Express software and quantified using QuantArray (GenomicSolutions). All subsequent microarray analyses were performed using SAS version 8.2 (SAS, Cary, NC). A robust local regression procedure (LOWESS) was performed to remove the systematic variations in the measured gene expression levels so that differences in expression across the samples could be distinguished accurately and precisely (21). After normalization, gene shaving (22), bootstrapping (23), and cluster analysis were performed (24), and the slide effect, dye effect, variety effect, and duplicate design were taken into account in an ANOVA model (25). Resampling-based multiple pairwise comparison was used to identify the differentially expressed genes before versus after the HFD. Differentially expressed genes were identified based on a Bonferroni adjusted P < 0.01.
Real-time quantitative RT-PCR for RNA. RNA sample pairs (1 [micro]g) were reverse transcribed using iScript cDNA synthesis kit (BioRad, Hercules, CA). All primers and probes were designed using Primer Express version 2.1 (Applied Biosystems-Roche, Branchburg, NJ). Sequences of primers and probes are shown in Supplemental Table 1 (online appendix [available at http://diabetes.diabetesjournals.org]).
