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• W2080954430 abstract "To examine the relationship between mitochondrial energy coupling in skeletal muscle and change in uncoupling protein 3 (UCP3) expression during the transition from the fed to fasted state, we used a novel noninvasive31P/13C NMR spectroscopic approach to measure the degree of mitochondrial energy coupling in the hind limb muscles of awake rats before and after a 48-h fast. Compared with fed levels, UCP3 mRNA and protein levels in the gastrocnemius increased 1.7- (p < 0.01) and 2.9-fold (p < 0.001), respectively, following a 48-h fast. Tricarboxylic acid cycle flux measured using 13C NMR as an index of mitochondrial substrate oxidation was 212 ± 23 and 173 ± 25 nmol/g/min (p not significant) in the fed and 48-h fasted groups, respectively. Unidirectional ATP synthesis flux measured using31P NMR was 79 ± 15 and 57 ± 9 nmol/g/s (p not significant) in the fed and 48-h fasted groups, respectively. Mitochondrial energy coupling as expressed by the ratio of ATP synthesis to tricarboxylic acid cycle flux was not different between the fed and fasted states. To test the hypothesis that UCP3 may be involved in the translocation of long chain free fatty acids (FFA) into the mitochondrial matrix under conditions of elevated FFA availability, [U-13C]palmitate/albumin was administered in a separate group of rats with (+) or without (−) etomoxir (an inhibitor of carnitine palmitoyltransferase I). The ratio of glutamate enrichment ((+) etomoxir/(−) etomoxir) in the hind limb muscles was the same between groups, indicating that UCP3 does not appear to function as a translocator for long chain FFA in skeletal muscle following a 48-h fast. In summary, these data demonstrate that despite a 2–3-fold increase in UCP3 mRNA and protein expression in skeletal muscle during the transition from the fed to fasted state, mitochondrial energy coupling does not change. Furthermore, UCP3 does not appear to have a major role in FFA translocation into the mitochondria. The physiological role of UCP3 following a 48-h fast in skeletal muscle remains to be elucidated. To examine the relationship between mitochondrial energy coupling in skeletal muscle and change in uncoupling protein 3 (UCP3) expression during the transition from the fed to fasted state, we used a novel noninvasive31P/13C NMR spectroscopic approach to measure the degree of mitochondrial energy coupling in the hind limb muscles of awake rats before and after a 48-h fast. Compared with fed levels, UCP3 mRNA and protein levels in the gastrocnemius increased 1.7- (p < 0.01) and 2.9-fold (p < 0.001), respectively, following a 48-h fast. Tricarboxylic acid cycle flux measured using 13C NMR as an index of mitochondrial substrate oxidation was 212 ± 23 and 173 ± 25 nmol/g/min (p not significant) in the fed and 48-h fasted groups, respectively. Unidirectional ATP synthesis flux measured using31P NMR was 79 ± 15 and 57 ± 9 nmol/g/s (p not significant) in the fed and 48-h fasted groups, respectively. Mitochondrial energy coupling as expressed by the ratio of ATP synthesis to tricarboxylic acid cycle flux was not different between the fed and fasted states. To test the hypothesis that UCP3 may be involved in the translocation of long chain free fatty acids (FFA) into the mitochondrial matrix under conditions of elevated FFA availability, [U-13C]palmitate/albumin was administered in a separate group of rats with (+) or without (−) etomoxir (an inhibitor of carnitine palmitoyltransferase I). The ratio of glutamate enrichment ((+) etomoxir/(−) etomoxir) in the hind limb muscles was the same between groups, indicating that UCP3 does not appear to function as a translocator for long chain FFA in skeletal muscle following a 48-h fast. In summary, these data demonstrate that despite a 2–3-fold increase in UCP3 mRNA and protein expression in skeletal muscle during the transition from the fed to fasted state, mitochondrial energy coupling does not change. Furthermore, UCP3 does not appear to have a major role in FFA translocation into the mitochondria. The physiological role of UCP3 following a 48-h fast in skeletal muscle remains to be elucidated. uncoupling protein free fatty acid reactive oxygen species not significant Mitochondrial uncoupling proteins (UCPs)1 play an integral role in regulating cellular energy consumption via nonshivering thermogenesis (1Himms-Hagen J. Prog. Lipid Res. 1989; 28: 67-115Crossref PubMed Scopus (277) Google Scholar). This is accomplished by diminishing the proton motive force across the inner mitochondrial membrane, which results in uncoupling of respiration from ATP synthesis. Unlike UCP1, which is expressed exclusively in brown adipose tissue, the recently discovered homolog UCP3 is expressed primarily in muscle (2Boss O. Samec S. Paoloni-Giacobino A. Rossier C. Dulloo A. Seydoux J. Muzzin P Giacobino J.P. FEBS Lett. 1997; 408: 39-42Crossref PubMed Scopus (998) Google Scholar, 3Vidal-Puig A. Solanes G. Grujic D. Flier J.S. Lowell B.B. Biochem. Biophys. Res. Commun. 1997; 235: 79-82Crossref PubMed Scopus (682) Google Scholar) and is encoded in a chromosomal region linked to hyperinsulinemia and obesity (4Gong D-W. He Y. Karas M. Reitman M. J. Biol. Chem. 1997; 272: 24129-24132Abstract Full Text Full Text PDF PubMed Scopus (739) Google Scholar). Because quiescent skeletal muscle utilizes approximately 33% of whole body oxygen consumption (5Field J. Beldings H.S. Martin A.W. J. Cell. Comp. Physiol. 1939; 14: 143-157Crossref Google Scholar), much attention has been given to the control and function of UCP3 as a means of regulating energy expenditure and body weight. Increased UCP3 mRNA expression results from a number of physiological perturbations including thyroid hormone (4Gong D-W. He Y. Karas M. Reitman M. J. Biol. Chem. 1997; 272: 24129-24132Abstract Full Text Full Text PDF PubMed Scopus (739) Google Scholar, 6Larkin S. Mull E. Miao W. Pittner R. Albrandt K. Moore C. Young A. Denaro M. Beaumont K. Biochem. Biophys. Res. Commun. 1997; 240: 222-227Crossref PubMed Scopus (172) Google Scholar) and leptin (4Gong D-W. He Y. Karas M. Reitman M. J. Biol. Chem. 1997; 272: 24129-24132Abstract Full Text Full Text PDF PubMed Scopus (739) Google Scholar, 7Cusin I. Zakrzreska K.E. Boss O. Muzzin P. Giacobino J.P. Ricquier D. Jeanrenaud B. Rohner-Jeanrenaud F. Diabetes. 1998; 47: 1014-1019Crossref PubMed Scopus (180) Google Scholar, 8Gómez-Ambrosi J. Frühbeck G. Martı́nez J.A. Cell. Mol. Life Sci. 1999; 55: 992-997PubMed Google Scholar, 9Sivitz W.I. Fink B.D. Donohoue P.A. Endocrinology. 1999; 140: 1511-1519Crossref PubMed Scopus (97) Google Scholar) in skeletal muscle and β-adrenoreceptor agonist (4Gong D-W. He Y. Karas M. Reitman M. J. Biol. Chem. 1997; 272: 24129-24132Abstract Full Text Full Text PDF PubMed Scopus (739) Google Scholar, 10Boss O. Bachman E. Vidal-Puig A. Zhang C-Y Peroni O. Lowell B.B. Biochem. Biophys. Res. Commun. 1999; 261: 870-876Crossref PubMed Scopus (98) Google Scholar) in white adipose tissue, which have all been implicated in increased regulatory thermogenesis. However, a paradoxical situation occurs in fasting when UCP3 mRNA expression in skeletal muscle has been shown to increase ∼2–12-fold (4Gong D-W. He Y. Karas M. Reitman M. J. Biol. Chem. 1997; 272: 24129-24132Abstract Full Text Full Text PDF PubMed Scopus (739) Google Scholar, 11Boss O. Samec S. Kuhne F. Bijlenga P. Assimacopoulos-Jeannet F. Seydoux J. Giacobino J.P. Muzzin P. J. Biol. Chem. 1998; 273: 5-8Abstract Full Text Full Text PDF PubMed Scopus (263) Google Scholar, 12Semec S. Seydoux J. Dulloo A.G. Eur. J. Physiol. 1999; 438: 452-457PubMed Google Scholar, 13Hwang C-S. Lane M.D. Biochem. Biophys. Res. Commun. 1999; 258: 464-469Crossref PubMed Scopus (64) Google Scholar, 14Semec S. Seydoux J. Dulloo A.G. Diabetes. 1998; 47: 1693-1698Crossref PubMed Scopus (130) Google Scholar, 15Weigle D.S. Selfridge L.E. Schwartz M.W. Seeley R.J. Cummings D.E. Havel P.J. Kuijper J.L. BeltrandelRio H. Diabetes. 1998; 47: 298-302Crossref PubMed Google Scholar, 16Cadenas S. Buckingham J.A. Samec S. Seydoux J. Din N. Dulloo A.G. Brand M.D. FEBS Lett. 1999; 462: 257-260Crossref PubMed Scopus (205) Google Scholar) at a time when mitochondrial energy coupling efficiency might be expected to remain constant or increase to conserve energy. This paradoxical relationship may be a result of UCP3 mRNA expression not correlating with protein levels or possibly a result of alterations in the concentration of an unknown allosteric regulator to UCP3. It has also been suggested that UCP3 may play a role in the regulation of lipids as fuel substrate rather than mediators of regulatory thermogenesis (17Semec S. Seydoux J. Dulloo A.G. FASEB J. 1998; 12: 715-724Crossref PubMed Scopus (299) Google Scholar). Therefore, the UCP3 protein may function specifically as a free fatty acid anion translocator rather than a direct proton shuttle or as part of a proton-FFA cycle under conditions of elevated plasma FFA following a 48-h fast. It is now generally accepted that fatty acids are necessary in activating uncoupling proteins via direct or indirect mechanisms (15Weigle D.S. Selfridge L.E. Schwartz M.W. Seeley R.J. Cummings D.E. Havel P.J. Kuijper J.L. BeltrandelRio H. Diabetes. 1998; 47: 298-302Crossref PubMed Google Scholar, 18Winkler E. Klingenberg M. J. Biol. Chem. 1994; 269: 2508-2515Abstract Full Text PDF PubMed Google Scholar, 19Skulachev V.P. FEBS Lett. 1991; 294: 158-162Crossref PubMed Scopus (394) Google Scholar, 20Garlid K.D. Orosz D.E. Modriansky M. Vassanelli S. Jezek P. J. Biol. Chem. 1996; 271: 2615-2620Abstract Full Text Full Text PDF PubMed Scopus (292) Google Scholar). Although indirect measurements of mitochondrial uncoupling have been made in UCP3-reconstituted vesicles, transfected yeast, or myoblasts coupled with electric potential measurements using an electrode method (18Winkler E. Klingenberg M. J. Biol. Chem. 1994; 269: 2508-2515Abstract Full Text PDF PubMed Google Scholar, 21Klingenberg M. Winkler E. EMBO J. 1985; 4: 3087-3092Crossref PubMed Scopus (135) Google Scholar) or coupled with fluorescence measurements using flow cytometric techniques (4Gong D-W. He Y. Karas M. Reitman M. J. Biol. Chem. 1997; 272: 24129-24132Abstract Full Text Full Text PDF PubMed Scopus (739) Google Scholar, 11Boss O. Samec S. Kuhne F. Bijlenga P. Assimacopoulos-Jeannet F. Seydoux J. Giacobino J.P. Muzzin P. J. Biol. Chem. 1998; 273: 5-8Abstract Full Text Full Text PDF PubMed Scopus (263) Google Scholar), these measurements have not been made in explicit tissues of interest in situ. Additionally, although electric potential measurements across the inner mitochondrial membrane provide an index for potential energy uncoupling, they do not measure functional energy uncoupling. This is because of the non-ohmic nature of proton conductance, which is not linear with the inner mitochondrial membrane potential (22Brand M.D. Brindle K.M. Buckingham J.A. Harper J.A. Rolfe D.F.S. Stuart J.A. Int. J. Obes. 1999; 23 Suppl. 6: 4-11Crossref Scopus (130) Google Scholar). We recently reported on the development of a novel NMR spectroscopic method used to assess mitochondrial energy coupling in skeletal muscle, noninvasively, by combining 13C NMR spectroscopy to measure rates of mitochondrial substrate oxidation along with 31P NMR spectroscopy to assess rates of ATP synthesis in chronic T3 (triiodothyronine) treated rats (a model of increased UCP3 expression) (23Jucker B.M. Dufour S. Ren J. Cao X. Previs S.F. Underhill B. Cadman K.S. Shulman G.I. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 6880-6884Crossref PubMed Scopus (95) Google Scholar). Therefore, in this study, we used this noninvasive NMR technique to determine if increased UCP3 mRNA and protein levels correlate with measurements of mitochondrial energy coupling in skeletal muscle of 48-h fasted rats. Additionally, we tested the hypothesis that UCP3 may function as a fatty acid translocator under fasting conditions during which levels of plasma FFA are increased. Harlan Sprague-Dawley rats (Charles River, Raleigh, NC) were individually housed in an environmentally controlled room with a 12-h light/dark cycle and maintained on standard rat chow (Ralston Purina Co., St. Louis, MO). Two groups of rats were used: fed (n = 17) and 48-h fasted (n = 19). When weighing ∼300 g, the 48-h fasted rats had cannulas inserted into both the right jugular vein and carotid artery (24Smith D. Rossetti L. Ferrannini E. Johnson C.M. Cobelli L. Toffolo G. Katz L.D. DeFronzo R.A. Metabolism. 1987; 36: 1167-1174Abstract Full Text PDF PubMed Scopus (90) Google Scholar) and were allowed to recuperate for 5 days prior to fasting. Following fasting, these rats weighed ∼270 g. The fed group weight was pair-matched to the 48-h fasted group at the time of the experiment. On the day of the NMR experiment, rats were placed in a customized restraining tube that allowed their left hind limb to be secured to the outside of the tube in a manner to limit free movement of the leg for NMR measurements. The rats were transiently anesthetized (<30 s) with a low dose (2.5–5.0 mg) of thiopental (Sigma) in order to place them in the restraining tube. This protocol was approved by the Yale University Animal Care Committee. The 31P NMR measurements preceded the 13C NMR measurements in each experiment. During the 31P measurements, unlabeled acetate was administered at a constant infusion rate (138 μmol/kg/min). During the 13C measurements, [2-13C]acetate (sodium salt, 99% enriched; Cambridge Isotope Laboratories, Cambridge, MA) was administered in order to study the glutamate labeling kinetics for tricarboxylic acid cycle flux measurements (Fig. 1). The [2-13C]acetate infusion consisted of a bolus (100 mg/kg of body weight) for 1 min followed by a 138 μmol/kg/min continuous infusion for 150 min (sufficient time required to achieve isotopic steady state). Blood samples were drawn during the base-line NMR measurement, at 7.5 min, 15 min, and every 15 min thereafter to assess the [2-13C]acetate enrichment time course. At the end of the in vivo NMR experiment, rats were anesthetized with thiopental (50 mg/kg). Superficial skin was rapidly removed from the left hind quarter followed by in situ freeze clamping of the gastrocnemius and biceps femoris muscles. Rats were euthanized with a lethal dose of thiopental. All in vivo NMR experiments were performed on a Bruker Biospec 7.0T system (horizontal/22-cm diameter bore magnet). Both13C observe/1H decouple and 31P observe NMR spectroscopy were performed using concentric surface coils (the outer 1H coil (30 mm) tuned to 300.54 MHz and the inner dual frequency 13C or 31P coil (18 mm) tuned to 75.59 and 121.66 MHz, respectively). The rat hind limb was positioned over the 13C/31P coil (vertical in plane) and placed in the magnet isocenter. Due to the13C/31P sensitivity in the hind limb experiments, it was necessary to measure the bulk signal from the larger tissue beds of mixed fiber type including the gastrocnemius and biceps femoris. Global 1H shimming was followed by localized shimming using a STEAM sequence over a 1 × 2 × 2-cm volume of the leg. Water line widths of 35–60 Hz, as a reflection of the average inhomogeneity caused by the subtle movements of the leg, were routinely achieved. The creatine/phosphocreatine peak (54.4 ppm) in the13C spectrum and β-ATP (−16.0 ppm) in the31P spectrum were used as an internal reference standard for movement of the leg. If severe movement compromised these peak integrals, the experiment was terminated. All spectra were processed off-line using Nuts NMR processing software (Acorn NMR Inc., Fremont, CA) with peak fitting capabilities. Base line-subtracted 13C NMR and 31P NMR spectra were processed using Gaussian filtering and a Gaussian weighted peak-fitting algorithm. The saturation transfer study requires that two sets of spectra be acquired: one with and one without steady state saturation. To measure the kinetics of Pi → ATP, a continuous wave selective saturation of the γ-ATP resonance was used. In the spectrum without γ-ATP saturation, the CW pulse was placed an equal frequency offset to the downfield side of Pi. The ratio of the resulting magnetization (Mz) to the equilibrium magnetization (M0) in the absence of γ-ATP saturation is given by the equation, Mz/M0=1/(1+kT1)Equation 1 where k is the first order rate constant describing the loss of magnetization from Pi, andT1 is the intrinsic spin lattice relaxation time for the Pi nucleus. A 90° pulse was optimized for the nonselective detection of both spectra (repetition time = 4.4 s, scans = 128, sweep width = 20 KHz, 4K data). The spin lattice relaxation time measured using an inversion recovery pulse sequence in the presence of continuous γ-ATP saturation is defined as the observed T1 (T1obs) and is related to T1 by the following equation.1/T1obs=1/T1+kEquation 2 An adiabatic half-passage pulse was used to invert all Pi spins in the inhomogenous volume of the surface coil (repetition time = 6.0 s, scans = 64, sweep width = 4 KHz, 4K data) for the 180° pulse of the inversion recovery sequence. The six variable delay lengths used in the inversion recovery experiment will be between 10 ms and 6 s. Solving the above two equations simultaneously (where ΔM =M0 − Mz) yields the following equation.k=1/T1obs×ΔM/M0Equation 3 Therefore, the rate constant (k) may be calculated after acquiring spectra for the two experiments described above (saturation transfer and inversion recovery). The unidirectional ATP synthesis flux was calculated as k × [Pi]. The Pi concentration was extrapolated from the baseline NMR spectrum (comparing peak integrals from Pi and γ-ATP) and [ATP] measurement using a biochemical assay. The NMR-measured unidirectional ATP synthesis flux results from flux through both the F1F0-ATP synthase enzyme and the coupled glyceraldehyde-3-phosphate dehydrogenase and phosphoglycerate kinase reactions. Although the net glycolytic contribution to the production of ATP (via glyceraldehyde-3-phosphate dehydrogenase and phosphoglycerate kinase) versus that of oxidative phosphorylation is small, these enzymes are at near equilibrium, and consequently the unidirectional production of ATP (measured using the 31P saturation transfer experiment) via these enzymes can be high. We previously described a technique where uniformly labeled glucose (1, 2, 3, 4, 5, 6, 6-D7) was administered to rats to determine whether there are differences in the glycolytic ATP synthesis flux contribution between groups (23Jucker B.M. Dufour S. Ren J. Cao X. Previs S.F. Underhill B. Cadman K.S. Shulman G.I. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 6880-6884Crossref PubMed Scopus (95) Google Scholar). In separate experiments, this technique was applied to both fed (n = 3) and 48-h fasted (n = 3) rats. The glyceraldehyde-3-phosphate M + 2/M + 4 (5.8 ± 1.1 versus 5.0 ± 1.3 in fedversus 48-h fasted, respectively) and M + 3/M + 4 (1.3 ± 0.7 versus 1.6 ± 0.5 in fed versus 48-h fasted, respectively) ratios were similar in both groups, suggesting no differences in the glycolytic ATP contribution. Observation of the 13C label turnover in the NMR-detectable glutamate pool in skeletal muscle was necessary for the tricarboxylic acid cycle flux calculation. Therefore, 1H-decoupled 13C NMR spectroscopy was performed in the following manner. An initial frequency-selective sinc pulse (20 ms) set on the low field side of the methylene carbon of lipids at 30 ppm was immediately followed by a nonselective hard pulse (approximately 70° flip angle, 5 mm from surface coil). The sinc pulse power was adjusted to eliminate most of the signal in that region. This method improved the base-line subtraction of lipid peaks between 22 and 30 ppm. Broadband1H Waltz-16 decoupling was applied during acquisition, and additional nuclear Overhauser effect was achieved using low power decoupling (0.4 W) during the relaxation delay (repetition time = 0.5 s, scans = 1800, sweep width = 20 KHz, 4K data). A 15-min base-line spectrum was followed by subsequent 15-min acquisitions throughout the duration of the experiment. Metabolic steady state equations were derived for isotopic mass flow into the tricarboxylic acid cycle as described previously in brain (25Fitzpatrick S.M. Hetherington H.P. Behar K.L. Shulman R.G. J. Cereb. Blood Flow Metab. 1990; 10: 170-179Crossref PubMed Scopus (215) Google Scholar, 26Mason G.F. Rothman D.L. Behar K.L. Shulman R.G. J. Cereb. Blood Flow Metab. 1992; 12: 434-447Crossref PubMed Scopus (221) Google Scholar), heart (27Yu X. White L.T. Doumen C. Damico L.A. LaNoue K.F. Alpert N.M. Lewandowski E.D. Biophys. J. 1995; 69: 2090-2102Abstract Full Text PDF PubMed Scopus (91) Google Scholar, 28Chance E.M. Seeholzer S.H. Kobayashi K. Williamson J.R. J. Biol. Chem. 1983; 258: 13785-13794Abstract Full Text PDF PubMed Google Scholar), and liver (29Jucker B.M. Lee J.Y. Shulman R.G. J. Biol. Chem. 1998; 273: 12187-12194Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar). Yale University CWave software (Dr. Graeme F. Mason) was used to calculate the tricarboxylic acid cycle fluxes. The mathematical modeling was based on nonlinear least squares fitting of the calculated parameters (e.g. 4-13C- and 2-13C-labeled citrate, α-ketoglutarate, glutamate, etc.) from the set of isotopic mass balance equations describing the label flow through the tricarboxylic acid cycle to the acquired NMR data using a Runge-Kutta algorithm with an adaptive step size. From the labeled mass flow schematic shown in Fig. 1, we obtain the following isotopic mass balance equations, which were used in the analyses, ∂C2AcCoA∂t=C3PyrPyrVpdh+C2AcetAcetVacet−C2AcCoAAcCoAVtcaEquation 4 ∂C4Cit∂t=C2AcCoAAcCoA−C4CitCitVtcaEquation 5 ∂C4α­KG∂t=C4CitCitVtca+C4GluGluVglu−1−C4α­KGα­KG(Vtca+Vglu)Equation 6 ∂C4Glu∂t=C4α­KGα­KGVglu+C4GlnGlnVgln−C4GluGlu(Vgln+Vgln−1)Equation 7 ∂C3Oaa∂t=0.5×C4α­KGα­KGVtca+0.5×C3α­KGα­KGVtca+C3AspAspVasp−1+C3PyrPyrVpc−C3OaaOaa(Vasp+Vtca+VOaa)Equation 8 ∂C2Cit∂t=C3OaaOaa−C2CitCitVtcaEquation 9 ∂C2α­KG∂t=C2CitCitVtca+C2GluGluVglu−1−C2α­KGα­KG(Vtca+Vglu)Equation 10 ∂C2Glu∂t=C2α­KGα­KGVglu+C2GlnGlnVgln−C2GluGlu(Vgln+Vgln−1)Equation 11 where AcCoA represents acetyl-CoA; Pyr is pyruvate; Acet is acetate; Cit is citrate; α-KG is α-ketoglutarate; Glu is glutamate; Gln is glutamine; Oaa is oxaloacetate; Asp is aspartate, and the C2, C3, or C4 prefix refers to 13C label at the respective carbon isotope positions. Vpdh represents pyruvate dehydrogenase flux; Vacet is acetate thiokinase flux; Vtca is tricarboxylic acid cycle flux; Vglu andVglu−1 are aminotransferase and glutamate dehydrogenase flux, respectively; Vgln and Vgln−1 are glutamine synthetase and glutaminase flux, respectively; Vasp is aspartate aminotransferase flux; and Voaa is efflux from oxaloacetate (necessary to maintain steady state) The α-KG ⇄ Glu exchange via glutamate dehydrogenase and/or aminotransferase reaction (Vglu,Vglu−1) is rapid with respect to Vtca in the brain (26Mason G.F. Rothman D.L. Behar K.L. Shulman R.G. J. Cereb. Blood Flow Metab. 1992; 12: 434-447Crossref PubMed Scopus (221) Google Scholar) but has been shown to be significantly slower in the heart (27Yu X. White L.T. Doumen C. Damico L.A. LaNoue K.F. Alpert N.M. Lewandowski E.D. Biophys. J. 1995; 69: 2090-2102Abstract Full Text PDF PubMed Scopus (91) Google Scholar, 28Chance E.M. Seeholzer S.H. Kobayashi K. Williamson J.R. J. Biol. Chem. 1983; 258: 13785-13794Abstract Full Text PDF PubMed Google Scholar). Therefore, it was necessary to include both 2-13C- and 4-13C-labeled glutamate turnover data in the mathematical analysis to discriminate between Vtca and Vglu. The glutamate pool concentration was determined in tissue extracts for use in the mathematical analysis, while all other intermediate pool concentrations were taken from the literature (27Yu X. White L.T. Doumen C. Damico L.A. LaNoue K.F. Alpert N.M. Lewandowski E.D. Biophys. J. 1995; 69: 2090-2102Abstract Full Text PDF PubMed Scopus (91) Google Scholar, 30Garland P.B. Randal P.J. Biochem. J. 1964; 93: 678-687Crossref PubMed Scopus (164) Google Scholar). In the model, [2-13C]acetyl-CoA pool turnover was described as rapid and set equal to the [2-13C]acetate precursor pool turnover that was measured in plasma and tissue extracts. To test this assumption (i.e. [2-13C]acetate turnover = [2-13C]acetyl-CoA turnover), we used a non-steady state isotopic analysis of glutamate labeling derived from [2-13C]acetate measured in rats at 15, 30, 60, and 150 min. The rapid turnover of [2-13C]acetyl-CoA enrichment (58.7 ± 0.3, 59.4 ± 2.6, 53.6 ± 0.6, and 56.3 ± 5.0 at the respective time points) suggests that this assumption is valid. It has been shown that acyl-CoA synthetase can function to a low degree in the mitochondrial matrix (31Trevisan C. DiMauro S. Neurochem. Res. 1983; 8: 551-561Crossref PubMed Scopus (5) Google Scholar). Therefore, we measured the capacity for UCP3 to function as a FFA translocator to increase β-oxidation in a separate group of rats. We measured relative rates of [U-13C]palmitate oxidation in skeletal muscle in vivo with (+) and without (−) carnitine palmitoyltransferase I inhibition by etomoxir in the fed and 48-h fasted groups. These measurements were performed in the fed rats to determine the relative decrease in palmitate oxidation resulting from carnitine palmitoyltransferase I inhibition. The control value (percentage decrease in palmitate oxidation in the fed group) was compared with the values obtained from the 48-h fasted group to determine whether or not UCP3 functions in part as a FFA translocator (i.e. percentage decrease is less than control percentage decrease). In the basal group ((−) etomoxir), a constant infusion (0.935 μmol/kg/min) of [U-13C]palmitate (Cambridge Isotope Laboratories, Andover, MA) bound to 12% bovine albumin (Sigma) was administered for 120 min. In the etomoxir group, etomoxir (10 μmol/kg, gift from Bristol-Myers Squibb) in 100 μl of sterile water was administered 60 min prior to [U-13C]palmitate infusion (32Oakes N.D. Cooney G.J. Camilleri S. Chisholm D.J. Kraegen E.W. Diabetes. 1997; 46: 1768-1774Crossref PubMed Google Scholar). A bolus administration (25 mg/kg) of nicotinic acid (Sigma) was given every hour to maintain basal FFA concentrations (33Sidossis L.S. Stuart C.A. Shulman G.I. Lopaschuk G.D. Wolfe R.R. J. Clin. Invest. 1996; 98: 2244-2250Crossref PubMed Scopus (201) Google Scholar). After 120 min, hind limb muscles were freeze-clamped in situ, and glutamate enrichments (index of relative palmitate oxidation) were measured in tissue extract using gas chromatography/mass spectrometry (34Beylot M. David F. Brunengraber H. Anal. Biochem. 1993; 212: 532-536Crossref PubMed Scopus (25) Google Scholar). The measured M + 2 glutamate enrichments (∼0.5–2%) were significantly greater than the 0.02% background enrichment. Muscle tissue extracts were prepared for high field NMR analysis by homogenizing approximately 1 g of combined quadriceps and gastrocnemius muscle as described previously (35Jucker B.M. Rennings A.J.M. Cline G.W. Shulman G.I. J. Biol. Chem. 1997; 272: 10464-10473Abstract Full Text Full Text PDF PubMed Scopus (71) Google Scholar). Glutamate (34Beylot M. David F. Brunengraber H. Anal. Biochem. 1993; 212: 532-536Crossref PubMed Scopus (25) Google Scholar) and acetate (36Powers L. Osborn M.K. Yang D. Kien C.L. Murray R.D. Beylot M. Brunengraber H. J. Mass Spectrom. 1995; 30: 747-754Crossref Scopus (39) Google Scholar) 13C enrichments in plasma and/or skeletal muscle were determined using a Hewlett-Packard 5890 gas chromatography (HP-1 capillary column; 12 m × 0.2 mm × 0.33 mm film thickness) interfaced to a Hewlett-Packard 5971A mass-selective detector operating in the electron impact ionization mode. The Pi concentration was extrapolated from the base-line NMR spectrum (comparing peak integrals from Pi and (γ-ATP) and measured ATP concentration (ATP assay kit 366, Sigma-modified for tissue analysis). Plasma free fatty acids were determined using an acyl-CoA oxidase-based colorimetric kit (WAKO NEFA-C; WAKO Pure Chemical Industries, Osaka, Japan). Tissue extract glutamate concentration was determined using a 2300 STAT PLUS biochemical analyzer (Yellow Springs Instrument Co., Yellow Springs, OH). Skeletal muscles were homogenized in the buffer containing 0.1 m KCl, 0.05m Tris-HCl, pH 7.4, 0.005 m MgCl2, 0.001 m EDTA, and freshly added protease inhibitor mixture (Roche Molecular Biochemicals). After brief homogenization, samples were spun at 650 × g for 10 min. Supernatant was collected and transferred to a new tube and spun at 14,000 ×g for 10 min. Pellets were collected and resuspended in 0.15m KCl. Total RNA (20 μg) was electrophoresed in a 1.5% agarose gel containing formaldehyde, as described by Lehrach et al. (37Lehrach H. Frischauf A.M. Philipson L. Prog. Clin. Biol. Res. 1985; 177: 7-15PubMed Google Scholar), and transferred to Hybond nylon membranes (Amersham Pharmacia Biotech) by capillary blotting. Probes were labeled by random priming with [α-32P]dCTP (PerkinElmer Life Sciences) to a specific radioactivity of approximately 2 × 108cpm/μg of DNA. RNA blots were hybridized in hybridization buffer containing 50% formamide, 0.25 mNa2HPO4, 0.25 m NaCl, and 1 mm EDTA at 65 °C overnight and then washed in a solution containing 0.25 m Na2HPO4, 0.5% SDS, and 1 mm EDTA at 65 °C for 20 min. Blots were exposed to scientific imaging film (PerkinElmer Life Sciences) at −80 °C with intensifying screens. The signals on the membrane were quantified by an Instant Imager (Canberra Company, Meriden, CT). Mitochondrial protein (25 μg) was loaded on SDS-polyacrylamide gel electrophoresis (4–15% Tris-HCl gradient gel; Bio-Rad) and transferred to nitrocellulose (Bio-Rad) at 100 V for 1 h. Nitrocellulose was blocked in a buffer containing 5% fetal bovine serum in 1× TBS and 0.2% Tween 20 for 1 h at room temperature. The membrane wa" @default.
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• W2080954430 cites W1970844785 @default.
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• W2080954430 cites W1987981231 @default.
• W2080954430 cites W1998854328 @default.
• W2080954430 cites W1999646017 @default.
• W2080954430 cites W2002461623 @default.
• W2080954430 cites W2002962229 @default.
• W2080954430 cites W2007819480 @default.
• W2080954430 cites W2013948514 @default.
• W2080954430 cites W2016070194 @default.
• W2080954430 cites W2020093222 @default.
• W2080954430 cites W2022628018 @default.
• W2080954430 cites W2023530279 @default.
• W2080954430 cites W2024337692 @default.
• W2080954430 cites W2027741102 @default.
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• W2080954430 cites W2041616502 @default.
• W2080954430 cites W2046812355 @default.
• W2080954430 cites W2052688011 @default.
• W2080954430 cites W2057498089 @default.
• W2080954430 cites W2063945875 @default.
• W2080954430 cites W2072444935 @default.
• W2080954430 cites W2072678001 @default.
• W2080954430 cites W2072919666 @default.
• W2080954430 cites W2077911717 @default.
• W2080954430 cites W2083385526 @default.
• W2080954430 cites W2089471108 @default.
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• W2080954430 cites W2102546287 @default.
• W2080954430 cites W2102547070 @default.
• W2080954430 cites W2104502652 @default.
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• W2080954430 cites W2226380358 @default.
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