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• W2169492015 abstract "Triacylglyceride stored in cytosolic lipid droplets (LDs) constitutes a major energy reservoir in most eukaryotes. The regulated turnover of triacylglyceride in LDs provides fatty acids for mitochondrial β-oxidation and ATP generation in physiological states of high demand for energy. The mechanisms for the formation of LDs in conditions of energy excess are not entirely understood. Fat storage-inducing transmembrane protein 2 (FIT2/FITM2) is the anciently conserved member of the fat storage-inducing transmembrane family of proteins implicated to be important in the formation of LDs, but its role in energy metabolism has not been tested. Here, we report that expression of FIT2 in mouse skeletal muscle had profound effects on muscle energy metabolism. Mice with skeletal muscle-specific overexpression of FIT2 (CKF2) had significantly increased intramyocellular triacylglyceride and complete protection from high fat diet-induced weight gain due to increased energy expenditure. Mass spectrometry-based metabolite profiling suggested that CKF2 skeletal muscle had increased oxidation of branched chain amino acids but decreased oxidation of fatty acids. Glucose was primarily utilized in CKF2 muscle for synthesis of the glycerol backbone of triacylglyceride and not for glycogen production. CKF2 muscle was ATP-deficient and had activated AMP kinase. Together, these studies indicate that FIT2 expression in skeletal muscle plays an unexpected function in regulating muscle energy metabolism and indicates an important role for lipid droplet formation in this process. Triacylglyceride stored in cytosolic lipid droplets (LDs) constitutes a major energy reservoir in most eukaryotes. The regulated turnover of triacylglyceride in LDs provides fatty acids for mitochondrial β-oxidation and ATP generation in physiological states of high demand for energy. The mechanisms for the formation of LDs in conditions of energy excess are not entirely understood. Fat storage-inducing transmembrane protein 2 (FIT2/FITM2) is the anciently conserved member of the fat storage-inducing transmembrane family of proteins implicated to be important in the formation of LDs, but its role in energy metabolism has not been tested. Here, we report that expression of FIT2 in mouse skeletal muscle had profound effects on muscle energy metabolism. Mice with skeletal muscle-specific overexpression of FIT2 (CKF2) had significantly increased intramyocellular triacylglyceride and complete protection from high fat diet-induced weight gain due to increased energy expenditure. Mass spectrometry-based metabolite profiling suggested that CKF2 skeletal muscle had increased oxidation of branched chain amino acids but decreased oxidation of fatty acids. Glucose was primarily utilized in CKF2 muscle for synthesis of the glycerol backbone of triacylglyceride and not for glycogen production. CKF2 muscle was ATP-deficient and had activated AMP kinase. Together, these studies indicate that FIT2 expression in skeletal muscle plays an unexpected function in regulating muscle energy metabolism and indicates an important role for lipid droplet formation in this process. The maintenance of energy expenditure in mammals involves the dynamic integration of two metabolically opposed states, fasting and feeding. Fasting involves the induction of catabolic, or ATP generating pathways, whereas feeding engages anabolic or ATP-consuming pathways that build carbon skeletons and store energy. The integration of these states is governed by endocrine signals, such as insulin and glucagon, nutrient signals such as long chain fatty acids and branched chain amino acids, adipose and gut-derived polypeptide hormones, such as leptin and Glp-1, and nutrient-sensing kinases, such as AMPK 2The abbreviations used are: AMPKAMP-activated protein kinaseLDlipid dropletBCAAbranched chain amino acidTAGtriacylglycerideATGLadipose triacylglyceride lipasePPARperoxisome proliferator-activated receptorGTTglucose tolerance testITTinsulin tolerance testSCstandard chowEDLextensor digitorum longusHFhigh fatRQrespiratory quotientIMTGintramyocellular triacylglycerideDGATacyl-CoA:diacylglycerol acyltransferaseACCacetyl-CoA carboxylase. and mammalian target of rapamycin. One peripheral tissue that plays an important role in energy metabolism is skeletal muscle, because it comprises a large percentage of the organism's mass and is the major site of glucose disposal and fatty acid and amino acid catabolism. Changes in the efficiency of substrate utilization or in the type of substrate used by skeletal muscle might affect energy balance. For example, enhancing the cataplerosis of the TCA cycle in skeletal muscle by overexpression of cytosolic phosphoenolpyruvate carboxykinase resulted in mice with increased mitochondria activity and leanness (1Hakimi P. Yang J. Casadesus G. Massillon D. Tolentino-Silva F. Nye C.K. Cabrera M.E. Hagen D.R. Utter C.B. Baghdy Y. Johnson D.H. Wilson D.L. Kirwan J.P. Kalhan S.C. Hanson R.W. J. Biol. Chem. 2007; 282: 32844-32855Abstract Full Text Full Text PDF PubMed Scopus (161) Google Scholar). Similarly, deficiency of the mitochondrial branched chain amino acid aminotransferase isozyme (BCAT2) resulted in enhanced energy expenditure (2She P. Reid T.M. Bronson S.K. Vary T.C. Hajnal A. Lynch C.J. Hutson S.M. Cell Metab. 2007; 6: 181-194Abstract Full Text Full Text PDF PubMed Scopus (278) Google Scholar). AMP-activated protein kinase lipid droplet branched chain amino acid triacylglyceride adipose triacylglyceride lipase peroxisome proliferator-activated receptor glucose tolerance test insulin tolerance test standard chow extensor digitorum longus high fat respiratory quotient intramyocellular triacylglyceride acyl-CoA:diacylglycerol acyltransferase acetyl-CoA carboxylase. An anabolic pathway important for the maintenance of energy balance is the storage of triacylglycerides (TAG) in cells. TAG is the primary energy reservoir in most eukaryotes that is stored in structures called cytosolic lipid droplets (LD) (3Wolins N.E. Brasaemle D.L. Bickel P.E. FEBS Lett. 2006; 580: 5484-5491Crossref PubMed Scopus (320) Google Scholar). Fatty acids stored as TAG are important for ATP production in many tissues, such as skeletal muscle, during physiological stresses such as exercise (4Kiens B. Physiol. Rev. 2006; 86: 205-243Crossref PubMed Scopus (338) Google Scholar, 5van Loon L.J. J. Appl. Physiol. 2004; 97: 1170-1187Crossref PubMed Scopus (150) Google Scholar, 6Watt M.J. Heigenhauser G.J. Dyck D.J. Spriet L.L. J. Physiol. 2002; 541: 969-978Crossref PubMed Scopus (144) Google Scholar, 7Thyfault J.P. Cree M.G. Tapscott E.B. Bell J.A. Koves T.R. Ilkayeva O. Wolfe R.R. Dohm G.L. Muoio D.M. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2010; 299: R926-R934Crossref PubMed Scopus (16) Google Scholar). However, excessive accumulation of TAG-rich lipid droplets in tissues such as adipose, skeletal muscle, liver, and pancreas is associated with insulin resistance and type 2 diabetes (8Savage D.B. Petersen K.F. Shulman G.I. Physiol. Rev. 2007; 87: 507-520Crossref PubMed Scopus (776) Google Scholar). The ability to store energy in the form of TAG-rich lipid droplets is evolutionarily conserved from Saccharomyces cerevisiae to humans (9Greenberg A.S. Coleman R.A. Kraemer F.B. McManaman J.L. Obin M.S. Puri V. Yan Q.W. Miyoshi H. Mashek D.G. J. Clin. Invest. 2011; 121: 2102-2110Crossref PubMed Scopus (463) Google Scholar). TAG-rich LDs are composed of a monolayer of phospholipids surrounding the TAG core and a unique proteome that associates with the surface of LDs (9Greenberg A.S. Coleman R.A. Kraemer F.B. McManaman J.L. Obin M.S. Puri V. Yan Q.W. Miyoshi H. Mashek D.G. J. Clin. Invest. 2011; 121: 2102-2110Crossref PubMed Scopus (463) Google Scholar, 10Brasaemle D.L. Dolios G. Shapiro L. Wang R. J. Biol. Chem. 2004; 279: 46835-46842Abstract Full Text Full Text PDF PubMed Scopus (634) Google Scholar). A common theme in the turnover of TAG from LD in many mammalian tissues is the regulation of lipase action involving perilipin proteins that is coordinated with the energy demands of the cell (9Greenberg A.S. Coleman R.A. Kraemer F.B. McManaman J.L. Obin M.S. Puri V. Yan Q.W. Miyoshi H. Mashek D.G. J. Clin. Invest. 2011; 121: 2102-2110Crossref PubMed Scopus (463) Google Scholar, 11Brasaemle D.L. J. Lipid Res. 2007; 48: 2547-2559Abstract Full Text Full Text PDF PubMed Scopus (768) Google Scholar, 12Zechner R. Kienesberger P.C. Haemmerle G. Zimmermann R. Lass A. J. Lipid Res. 2009; 50: 3-21Abstract Full Text Full Text PDF PubMed Scopus (412) Google Scholar). The formation of TAG LDs might also be coordinated with the energy demands of the cell. TAG-rich LDs form at the endoplasmic reticulum and require the enzymatic activity of the endoplasmic reticulum-resident membrane proteins diacylglycerol acyltransferase 1 and 2 in mammals (13Harris C.A. Haas J.T. Streeper R.S. Stone S.J. Kumari M. Yang K. Han X. Brownell N. Gross R.W. Zechner R. Farese Jr., R.V. J. Lipid Res. 2011; 52: 657-667Abstract Full Text Full Text PDF PubMed Scopus (220) Google Scholar). The process of LD formation and regulation of LD formation downstream of DGAT activity are poorly understood. Our laboratory has recently identified a two-gene family of evolutionarily conserved proteins that are important in the partitioning of newly synthesized triacylglyceride into lipid droplets, which we named fat storage-inducing transmembrane protein 1 and 2 (FITM1/FIT1 and FITM2/FIT2) (14Kadereit B. Kumar P. Wang W.J. Miranda D. Snapp E.L. Severina N. Torregroza I. Evans T. Silver D.L. Proc. Natl. Acad. Sci. U.S.A. 2008; 105: 94-99Crossref PubMed Scopus (189) Google Scholar). Fat storage-inducing transmembrane proteins have six transmembrane domains and reside exclusively in the endoplasmic reticulum (14Kadereit B. Kumar P. Wang W.J. Miranda D. Snapp E.L. Severina N. Torregroza I. Evans T. Silver D.L. Proc. Natl. Acad. Sci. U.S.A. 2008; 105: 94-99Crossref PubMed Scopus (189) Google Scholar, 15Gross D.A. Snapp E.L. Silver D.L. PLoS One. 2010; 5e10796Crossref PubMed Scopus (45) Google Scholar). FIT2 is the anciently conserved fat storage-inducing transmembrane family member and is ubiquitously expressed at low levels in mammalian tissues with highest expression in white adipose tissue. This expression profile of FIT2 raises the possibility that FIT2 may play a role in TAG storage in “long term” fat depots like adipose tissue, as opposed to the more rapidly turning over TAG-LDs in muscle (5van Loon L.J. J. Appl. Physiol. 2004; 97: 1170-1187Crossref PubMed Scopus (150) Google Scholar, 16Saddik M. Lopaschuk G.D. J. Biol. Chem. 1991; 266: 8162-8170Abstract Full Text PDF PubMed Google Scholar). Overexpression of FIT2 in mammalian cells in vitro and in vivo resulted in TAG LD accumulation. The process of TAG LD accumulation was found to be through the partitioning of newly synthesized TAG into LD without an increase in TAG biosynthesis. Conversely, knockdown of FIT2 in pre-adipocyte NIH 3T3-L1 cells reduced the size and number of LDs, indicating that FIT2 plays an important role in lipid droplet formation but not TAG biosynthesis. The distinct tissue distribution pattern of FIT1 and FIT2 raises the idea that FIT2, being highly expressed in adipose, is primarily involved in the generation of large lipid droplets for long term storage of triacylglycerides, whereas FIT1 is important for generating small lipid droplets involved in the rapid mobilization of triacylglycerides during increased ATP demand. We took advantage of the unique biochemical function of FIT2 in LD formation to examine the effects of FIT2 expression and TAG-LDs on muscle energy metabolism. Here, we report that muscle-specific overexpression of FIT2 resulted in robust reprogramming of skeletal muscle metabolism whereby reduced metabolic efficiency resulted in increased energy expenditure and decreased energy charge. The data indicate that FIT2 plays an important role in regulating energy expenditure through modulating TAG storage. The following rabbit polyclonal antibodies were used for these studies: p-AKT, p-AMPK, AMPK, p-glycogen synthase, glycogen synthase, p-GSK3B, GSK3B, p-ACC, ACC, LC3B (Cell Signaling), Glut1 (Abcam), myoglobin (Santa Cruz Biotechnology), OXPAT (Novus Biologicals), anti-calnexin (Sigma), and mouse monoclonal AKT (Cell Signaling). Mouse FIT2 cDNA was cloned downstream of the mouse creatine kinase promoter for use in transgenesis. CKF2 mice were backcrossed for 10 generations to C57BL/6J. All experiments were conducted using 2–3-month-old male transgenic and WT littermates. Study protocols were approved by the Institute of Animal Studies of the Albert Einstein College of Medicine. Mice were fed a standard chow (LabDiet number 5058) or high fat diet (Research Diets, Inc., number D12492) composed of 60 kcal % fat for the indicated amount of time. Body weight was monitored weekly during the study period. Prior to GTT or ITT, mice were fasted overnight for 12 h with ad libitum access to water and then administered glucose (1 g/kg) or insulin (1 unit/kg) intraperitoneally, respectively. Blood glucose was measured in blood samples collected from tail vain using a glucometer (Accu-Check Compact, Roche Applied Science). C2C12 myocytes were cultured in DMEM (Invitrogen) supplemented with 10% fetal bovine calf serum and penicillin/streptomycin. For differentiation, C2C12 cell were induced at 80–90% confluency by switching the cell media to DMEM supplemented with 2% horse serum (differentiation media). On day 5 of differentiation, C2C12 were infected for 12 h in differentiation media. All experiments were performed 36 h post-infection. C2C12 cells were differentiated in 6-well plates and infected with empty or C-terminal tagged FIT2 (FIT2-V5) adenovirus as described above. For fatty acid oxidation, differentiation media was then replaced with 1 ml of low glucose DMEM (Invitrogen) supplemented with 100 μm [3H]palmitic acid/BSA (0.4 μCi/ml) and 1 μm l-carnitine. After 4 h, media was collected and centrifuged at 16,000 × g for 15 min in a table top centrifuge to remove cellular debris. 100 μl of media was then treated with 900 μl of 20 mm Tris-HCl, 5% w/v activated charcoal (Sigma) for 30 min at room temperature to absorb fatty acids. Samples were then centrifuged at 16,000 × g for 15 min at room temperature. 200 μl of the supernatant was used to determine fatty acid oxidation by measuring tritiated water using a scintillation counter and normalized to total protein. BCAA oxidation was performed in C2C12 myotubes expressing FIT2-V5 cultured in 6-well plates using a pulse-chase method. C2C12 myotubes were pulsed for 30 min with DMEM 5% horse serum supplemented with [3H]leucine (1 μCi/ml). After 30 min, C2C12 myotubes were extensively washed with Krebs-Henseleit buffer and then chased for 4 h with Krebs-Henseleit buffer supplemented with 5 mm glucose, 10 mm leucine, and 0.2% (w/v) fatty acid-free BSA. After 4 h, media was collected and centrifuged at 16,000 × g for 15 min in a table top centrifuge to remove cellular debris. 500 μl of media was then treated with 500 μl of 6% perchloric acid for 30 min at room temperature to precipitate protein. Samples were then centrifuged at 16,000 × g for 15 min at room temperature. 200 μl of the supernatant was used to determine BCAA oxidation by measuring tritiated water using a scintillation counter and normalized to total protein. For glucose uptake, C2C12 myotubes were starved in low glucose DMEM for 2 h, then washed twice with KRPH buffer (5 mm NaH2PO4, 20 mm HEPES, 1 mm MgSO4, 1 mm CaCl2, 136 mm NaCl, 4.7 mm KCl), and incubated for 20 min in 2 ml of KRPH buffer supplemented with 5 μm 2-deoxy-d-glucose, 1 mCi/ml 2-[3H]deoxy-d-glucose, plus or minus 100 nm insulin. After 20 min, C2C12 myotubes were washed extensively with PBS buffer and lysed with 1 ml of RIPA buffer supplemented with EDTA-free protease inhibitors (Roche Applied Science) and phosphatase inhibitors (Calbiochem). Glucose uptake was determined by measuring 800 μl of lysate using a scintillation counter and normalized to total protein. C2C12 myocytes were differentiated in 25-mm glass bottom dishes (Wilco) and infected with empty or C-terminally tagged FIT2 (FIT2-V5) adenovirus as described above. Visualization of LDs in C2C12 cells were performed as described previously using BODIPY 493/503 (15Gross D.A. Snapp E.L. Silver D.L. PLoS One. 2010; 5e10796Crossref PubMed Scopus (45) Google Scholar). Metabolic measurements (oxygen consumption, carbon dioxide production, food intake, and physical activity) were obtained continuously using a CLAMS (Columbus Instruments) open-circuit indirect calorimetry system for 5 consecutive days and normalized to lean body mass. Mice were allowed to acclimate for 2 days in individual chambers prior to data collection. Mitochondria isolation and mitochondrial ATP production rate were performed as described previously with some modifications (17Wibom R. Lundin A. Hultman E. Scand. J. Clin. Lab. Invest. 1990; 50: 143-152Crossref PubMed Google Scholar). Briefly, 50 mg of quadriceps was homogenized in 800 μl of Buffer A (100 mm KCl, 50 mm Tris-HCl, pH 7.4, 5 mm MgCl2, 1.8 mm ATP, and 1 mm EDTA) plus EDTA-free protease inhibitors (Roche Applied Science) and spun down at 1,000 × g for 20 min at 4 °C. The supernatant was then spun down at 10,000 × g for 10 min at 4 °C. The resulting pellet was then washed in 800 μl of Buffer A and spun down at 9,000 × g for 10 min at 4 °C. The resulting pellet was resuspended in 50 μl of Buffer B (180 mm sucrose, 35 mm KH2PO4, 10 mm magnesium acetate, and 5 mm EDTA) plus protease inhibitors. Protein concentration was determined by BCA protein assay (Pierce). Mitochondria lysates were then diluted 1:500 in Buffer B and immediately used to determine maximal mitochondrial ATP production rate using the following substrates for oxidation: 10 mm glutamate plus 1 mm malate and 35 mm ADP (state 3 respiration). ATP production was monitored using the ENLITEN® ATP bioluminescence detection kit (Promega). All reactions were conducted simultaneously at 25 °C for 25 min in a 200-μl reaction volume using 10 μl of diluted mitochondria protein, 100 μl of the luciferase/luciferin reagent, and substrates for oxidation plus or minus ADP (state 2 respiration). Mitochondrial ATP production rate was normalized to total protein. Blood glucose was measured under fasting conditions in blood samples collected from tail vein using a glucometer (Accu-Check Compact, Roche Applied Science). Plasma insulin levels were measured under fasting conditions and after treating mice with 1 g/kg glucose intraperitoneally using ELISA (Millipore). Total serum ketone bodies and lactate were determined using a colorimetric assay (Stanbio and Bioassays). Total serum triacylglyceride and cholesterol levels were measured using a colorimetric assay (Infinity). DGAT activity was determined using microsomes isolated from quadriceps as described previously (18Levin M.C. Monetti M. Watt M.J. Sajan M.P. Stevens R.D. Bain J.R. Newgard C.B. Farese Sr., R.V. Farese Jr., R.V. Am. J. Physiol. Endocrinol. Metab. 2007; 293: E1772-E1781Crossref PubMed Scopus (79) Google Scholar). TAG was separated by TLC and quantified as described previously (15Gross D.A. Snapp E.L. Silver D.L. PLoS One. 2010; 5e10796Crossref PubMed Scopus (45) Google Scholar). Mice were fasted for 4 h and injected intraperitoneally with universally labeled d-[14C]glucose (310 mCi/mmol, PerkinElmer Life Sciences) and insulin at 2,500 μCi/kg and 1 unit/kg, respectively. Quadriceps were collected and freeze-clamped 15 min post-injection, homogenized in 1 ml of PBS, and spun down 16,000 × g at 4 °C. 50 μl of the resulting supernatant was used for scintillation counting to determine total glucose uptake. Protein concentration was determined by BCA protein assay (Pierce). Lipids were then extracted from 1 mg of protein and TAG separated by TLC as described previously (15Gross D.A. Snapp E.L. Silver D.L. PLoS One. 2010; 5e10796Crossref PubMed Scopus (45) Google Scholar). Radioactive TAGs were quantified by using PhosphorImager analysis and represented as a rate of d-[14C]glucose incorporated into TAG (arbitrary units obtained from PhosphorImager analysis) per mg of protein normalized to total glucose uptake. Muscle glycogen content was determined in WT and CKF2 quadriceps as described previously (19Vaitheesvaran B. LeRoith D. Kurland I.J. Diabetologia. 2010; 53: 2224-2232Crossref PubMed Scopus (22) Google Scholar). Ex vivo basal and insulin-stimulated glucose transport was performed using extensor digitorum longus (EDL) muscle as described previously (20Chadt A. Leicht K. Deshmukh A. Jiang L.Q. Scherneck S. Bernhardt U. Dreja T. Vogel H. Schmolz K. Kluge R. Zierath J.R. Hultschig C. Hoeben R.C. Schürmann A. Joost H.G. Al-Hasani H. Nat. Genet. 2008; 40: 1354-1359Crossref PubMed Scopus (168) Google Scholar). Quadriceps were elongated, placed on cork, covered with OCT, and frozen in pre-chilled 2-methylbutane. Samples were then sectioned and stained for neutral lipids using Oil Red-O. Lipids were extracted from 1 mg of protein isolated from WT and CKF2 quadriceps using chloroform/methanol (2:1). The organic phase was transferred to glass bottom tubes and dried down under a stream of nitrogen. Total TAG was then quantified using TLC as described previously (15Gross D.A. Snapp E.L. Silver D.L. PLoS One. 2010; 5e10796Crossref PubMed Scopus (45) Google Scholar). TAG was quantified from C2C12 myotubes cultured in 6-well plates as described previously (15Gross D.A. Snapp E.L. Silver D.L. PLoS One. 2010; 5e10796Crossref PubMed Scopus (45) Google Scholar). cDNA was synthesized from mouse quadriceps RNA using SuperScript III first strand cDNA synthesis kit and oligo(dT) primers (Invitrogen). Gene expression analysis was assessed by quantitative SYBER Green real time PCR using a 7300 real time PCR system (Applied Biosystems). Primers were obtained from Invitrogen; sequences are available upon request. Relative quantification of each target was normalized to hypoxanthine-guanine phosphoribosyltransferase. HPLC-based nucleotide analysis was performed on freeze-clamped quadriceps as described previously (21Ellis J.M. Mentock S.M. Depetrillo M.A. Koves T.R. Sen S. Watkins S.M. Muoio D.M. Cline G.W. Taegtmeyer H. Shulman G.I. Willis M.S. Coleman R.A. Mol. Cell. Biol. 2011; 31: 1252-1262Crossref PubMed Scopus (138) Google Scholar). Mass spectrometry-based metabolite profiling was performed following a 4-h fast as described previously (22Koves T.R. Ussher J.R. Noland R.C. Slentz D. Mosedale M. Ilkayeva O. Bain J. Stevens R. Dyck J.R. Newgard C.B. Lopaschuk G.D. Muoio D.M. Cell Metab. 2008; 7: 45-56Abstract Full Text Full Text PDF PubMed Scopus (1436) Google Scholar). Tissues were homogenized in RIPA buffer plus EDTA-free protease inhibitors (Roche Applied Science). Protein concentration was determined by BCA protein assay (Pierce), and 80 μg of protein were used per sample. Samples were separated by SDS-PAGE on 8, 12, or 15% polyacrylamide gels, transferred to nitrocellulose membranes (Bio-Rad), and incubated with indicated antibodies. Quantification of signals was performed using an Odyssey infrared scanner (Li-Cor). All quantitative data are represented as mean ± S.E. p values were generated by Student's t test. To examine the effects of FIT2 overexpression in skeletal muscle, we generated muscle-specific FIT2 transgenic mice (CKF2) by using the mouse creatine kinase promoter. We identified two founder lines, 130 and 137, that exhibited similar levels of skeletal muscle-specific expression without enhanced expression in heart or changes in FIT1 expression (Fig. 1A and data not shown). Because both lines exhibited similar phenotypes, we arbitrarily focused on line 137 throughout these studies. CKF2 mice were viable, reproduced normally, did not die prematurely, and had similar food intake as wild-type (WT) mice (see below). We have previously demonstrated that overexpression of FIT2 in mouse liver and in cells in culture resulted in the accumulation of TAG in lipid droplets (14Kadereit B. Kumar P. Wang W.J. Miranda D. Snapp E.L. Severina N. Torregroza I. Evans T. Silver D.L. Proc. Natl. Acad. Sci. U.S.A. 2008; 105: 94-99Crossref PubMed Scopus (189) Google Scholar, 15Gross D.A. Snapp E.L. Silver D.L. PLoS One. 2010; 5e10796Crossref PubMed Scopus (45) Google Scholar). As expected, overexpression of FIT2 resulted in a significant intramyocellular accumulation of neutral lipid as indicated by Oil-Red O staining of sections of quadriceps (Fig. 1B). To further characterize this phenotype, total lipids were extracted from quadriceps of 12-week-old WT and CKF2 mice fed a standard chow (SC) diet and analyzed using thin layer chromatography. CKF2 mice exhibited a 7-fold increase in total TAG (Fig. 1C). Diacylglycerol acyltransferase activity was unchanged in muscles from CKF2 mice relative to WT muscle (Fig. 1D), supporting previous findings that FIT2 does not mediate triacylglyceride biosynthesis (14Kadereit B. Kumar P. Wang W.J. Miranda D. Snapp E.L. Severina N. Torregroza I. Evans T. Silver D.L. Proc. Natl. Acad. Sci. U.S.A. 2008; 105: 94-99Crossref PubMed Scopus (189) Google Scholar). Increased expression of FIT2 in skeletal muscle had a profound effect on body weight. At 6 weeks of age CKF2 exhibited 20% less body weight as compared with their WT littermates (Fig. 2A, time point zero). EchoMRI analysis indicated that the difference in body weight was due to decreased fat mass and lean mass (Fig. 2B). Decreased lean mass was not due to increased activation of atrophy pathways because Murf1 and Atrogin-1/Fbox32 were not increased but were decreased in CKF2 muscle (Fig. 3A). However, CKF2 mice had increased levels of LC3-II compared with WT, potentially indicating an increase in autophagy (Fig. 3B). Increased autophagy may be an indication of decreased energy state in muscle (discussed below). We then challenged CKF2 mice with high fat (HF) diet beginning at 6 weeks of age for 12 weeks. Surprisingly, CKF2 mice were completely protected from HF diet-induced weight gain and maintained a similar body weight and fat mass as CKF2 mice on a SC diet (Fig. 2, A and B). These findings raised the possibility that FIT2 overexpression might be affecting energy expenditure.FIGURE 3No increase in muscle atrophy markers. A, real time PCR analysis of E3 ligases Atrogin-1 and Murf1 indicated that CKF2 mice did not have increased atrophy in quadriceps. B, CKF2 mice exhibited increased autophagy as indicated by increased levels of LC3B-I and LC3B-II; data are represented as mean ± S.E.; n = 4 mice per genotype; **, p < 0.01. Mb, myoglobin.View Large Image Figure ViewerDownload Hi-res image Download (PPT) Several possible mechanisms can contribute to the maintenance of body weight in CKF2 mice on SC and HF diets relative to their respective WT controls, namely decreased food intake, increased voluntary physical activity, and increased thermogenesis. Because CKF2 mice are leaner than WT mice even when fed a SC diet, we focused the remainder of our studies on mice fed a SC diet. CKF2 mice exhibited similar food intake as WT mice and had decreased physical activity compared with WT mice (Fig. 4, A and B). CKF2 mice had decreased body temperature compared with WT littermates that was correlated with significantly decreased UCP-1 protein levels in brown adipose tissue compared with WT mice (Fig. 4, C and D). In addition, UCP-3 expression in skeletal muscle was unchanged (Fig. 4E). An alternative explanation for the leanness of CKF2 mice and protection from HF diet-induced weight gain is that CKF2 mice have enhanced energy expenditure independent of enhanced thermogenesis. To determine the mechanisms by which CKF2 mice are lean and resistant to diet-induced weight gain, we measured energy expenditure using indirect calorimetry of mice fed a SC diet. We chose to examine SC-fed mice because CKF2 mice fed a SC diet were already protected from body weight gain over time (Fig. 2A), and thus we could avoid complicating effects of HF diet on our analysis of metabolism. CKF2 mice had increased oxygen consumption in both the light and dark cycles as compared with WT mice, indicating increased energy expenditure (Fig. 4F). CKF2 mice also exhibited a significant increase in carbon dioxide production compared with WT mice (Fig. 4F). Respiratory quotient (RQ) was calculated to determine whether CKF2 mice have a particular preference for carbohydrate or fatty acids in their substrate utilization (Fig. 4F). RQ in CKF2 and WT mice increased from the light phase to the dark phase indicating that CKF2 mice were similarly metabolically flexible as WT mice. CKF2 mice did show a slight but significant reduction in RQ in the dark phase, which reflects a selective decline in glucose oxidation relative to fatty acid and/or amino acid oxidation. However, given that CKF2 mice had significantly higher plasma ketone bodies (Table 1), oxidation of fatty acids by liver partly explains the small decrease in RQ. We observed a significant increase in serum triacylglyceride and free fatty acids and a small but significant decrease in cholesterol. Levels of serum lactate were unchanged (Table 1).TABLE 1CKF2 mice serum metabolitesMetaboliteWTCKF2Triglyceride (mg/dl)65.27 ± 4.6085.76 ± 11.63ap < 0.001.Cholesterol (mg/dl)82.22 ± 3.0173.11 ± 1.75ap < 0.001.FFA (nmol/ml)562.52 ± 30.04858.82 ± 66.24bp < 0.0001.Ketones (nmol/ml)314.84 ± 50.14700.21 ± 82.10bp < 0.0001.Lactate (nmol/ml)3.39 ± 0.6872.74 ± 0.306a p < 0.001.b p < 0.0001. Open table in a new tab Intramyocellular accumulation of lipids has been correlated with insulin resi" @default.
• W2169492015 created "2016-06-24" @default.
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• W2169492015 date "2011-12-01" @default.
• W2169492015 modified "2023-09-29" @default.
• W2169492015 title "Re-patterning of Skeletal Muscle Energy Metabolism by Fat Storage-inducing Transmembrane Protein 2" @default.
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