Proteome Analysis Reveals Phosphorylation of ATP Synthase β-Subunit in Human Skeletal Muscle and Proteins with Potential Roles in Type 2 Diabetes

Insulin resistance in skeletal muscle is a hallmark feature of type 2 diabetes. An increasing number of enzymes and metabolic pathways have been implicated in the development of insulin resistance. However, the primary cellular cause of insulin resistance remains uncertain. Proteome analysis can quantitate a large number of proteins and their post-translational modifications simultaneously and is a powerful tool to study polygenic diseases like type 2 diabetes. Using this approach on human skeletal muscle biopsies, we have identified eight potential protein markers for type 2 diabetes in the fasting state. The observed changes in protein expression indicate increased cellular stress, e.g. up-regulation of two heat shock proteins, and perturbations in ATP (re)synthesis and mitochondrial metabolism, e.g. down-regulation of ATP synthase β-subunit and creatine kinase B, in skeletal muscle of patients with type 2 diabetes. Phosphorylation appears to play a key, potentially coordinating role for most of the proteins identified in this study. In particular, we demonstrated that the catalytic β-subunit of ATP synthase is phosphorylated in vivo and that the levels of a down-regulated ATP synthase β-subunit phosphoisoform in diabetic muscle correlated inversely with fasting plasma glucose levels. These data suggest a role for phosphorylation of ATP synthase β-subunit in the regulation of ATP synthesis and that alterations in the regulation of ATP synthesis and cellular stress proteins may contribute to the pathogenesis of type 2 diabetes. Insulin resistance in skeletal muscle is a hallmark feature of type 2 diabetes. An increasing number of enzymes and metabolic pathways have been implicated in the development of insulin resistance. However, the primary cellular cause of insulin resistance remains uncertain. Proteome analysis can quantitate a large number of proteins and their post-translational modifications simultaneously and is a powerful tool to study polygenic diseases like type 2 diabetes. Using this approach on human skeletal muscle biopsies, we have identified eight potential protein markers for type 2 diabetes in the fasting state. The observed changes in protein expression indicate increased cellular stress, e.g. up-regulation of two heat shock proteins, and perturbations in ATP (re)synthesis and mitochondrial metabolism, e.g. down-regulation of ATP synthase β-subunit and creatine kinase B, in skeletal muscle of patients with type 2 diabetes. Phosphorylation appears to play a key, potentially coordinating role for most of the proteins identified in this study. In particular, we demonstrated that the catalytic β-subunit of ATP synthase is phosphorylated in vivo and that the levels of a down-regulated ATP synthase β-subunit phosphoisoform in diabetic muscle correlated inversely with fasting plasma glucose levels. These data suggest a role for phosphorylation of ATP synthase β-subunit in the regulation of ATP synthesis and that alterations in the regulation of ATP synthesis and cellular stress proteins may contribute to the pathogenesis of type 2 diabetes. type 2 diabetes mellitus AMP-activated protein kinase mass spectrometry percentage integrated optical density matrix-assisted laser desorption/ionization time-of-flight ATP synthase β-subunit creatine kinase, brain isoform phosphoglucomutase-1 heat shock protein 78-kDa glucose-regulated protein α-1 chain of type VI collagen myosin regulatory light chain 2 reactive oxygen species two-dimensional free fatty acids 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid Insulin resistance in skeletal muscle, defined as reduced insulin-stimulated glucose disposal, is a characteristic feature of type 2 diabetes mellitus (T2DM)1 and is believed to be largely accounted for by reduced non-oxidative glucose metabolism (1Beck-Nielsen H. J. 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Conversely, in the basal, fasting state increased glucose oxidation and reduced lipid oxidation is seen in skeletal muscle of insulin resistant subjects, whether caused by T2DM or obesity alone (4Kelley D.E. Mandarino L.J. Diabetes. 2000; 49: 677-683Crossref PubMed Scopus (779) Google Scholar). These defects suggest an impaired capacity to switch between carbohydrate and fat as oxidative energy sources in insulin-resistant subjects. Together with reports of reduced oxidative enzyme activity and dysfunction of mitochondria in skeletal muscle of patients with T2DM (4Kelley D.E. Mandarino L.J. Diabetes. 2000; 49: 677-683Crossref PubMed Scopus (779) Google Scholar, 5He J. Watkins S. Kelley D.E. Diabetes. 2001; 50: 817-823Crossref PubMed Scopus (399) Google Scholar, 6Kelley D.E. He J. Menshikova E.V. Ritov V.B. Diabetes. 2002; 51: 2944-2950Crossref PubMed Scopus (1782) Google Scholar) and the fact that mitochondrial DNA defects cause T2DM through impairment of oxidative phosphorylation (7Gebhart S. Shoffner J.M. Koontz D. Kaufmann A. Wallace D. Metabolism. 1996; 45: 526-531Abstract Full Text PDF PubMed Scopus (37) Google Scholar, 8Wallace D.C. Science. 1999; 283: 1482-1488Crossref PubMed Scopus (2630) Google Scholar), these abnormalities in fuel metabolism have led to the hypotheses that perturbations in skeletal muscle mitochondrial metabolism (6Kelley D.E. He J. Menshikova E.V. Ritov V.B. Diabetes. 2002; 51: 2944-2950Crossref PubMed Scopus (1782) Google Scholar, 9Anderson C.M. Drug. Dev. Res. 1999; 46: 67-79Crossref Scopus (16) Google Scholar, 10Antonetti D.A. Reynet C. Kahn C.R. J. Clin. Invest. 1995; 95: 1383-1388Crossref PubMed Scopus (115) Google Scholar) and defects in the signaling pathways of AMP-activated protein kinase (AMPK) are implicated in the pathogenesis of T2DM (11Winder W.W. Hardie D.G. Am. J. Physiol. Endocrinol. Metab. 1999; 277: E1-E10Crossref PubMed Google Scholar). That rates of fuel oxidation and mitochondrial function can affect glucose uptake and glycogen synthesis has been reported earlier (12Bashan N. Burdett E. Guma A. Sargeant R. Tumiati L. Liu Z. Klip A. Am. J. Physiol. 1993; 264: C430-C440Crossref PubMed Google Scholar, 13Katz A. Westerblad H. Biochim. Biophys. Acta. 1995; 1244: 229-232Crossref PubMed Scopus (12) Google Scholar). In addition, both chronic activation of AMPK and induced expression of the transcriptional co-activator of peroxisome proliferator-activated receptor γ, PGC-1, result in improved mitochondrial biogenesis concomitant with increases in GLUT4 protein content in skeletal muscle (14Winder W.W. Holmes B.F. Rubink D.S. Jensen E.B. Chen M. Holloszy J.O. J. Appl. Physiol. 2000; 88: 2219-2226Crossref PubMed Scopus (605) Google Scholar, 15Michael L.F. Wu Z. Cheatham R.B. Puigserver P. Adelmant G. Lehman J.J. Kelly D.P. Spiegelman B.M. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 3820-3825Crossref PubMed Scopus (542) Google Scholar, 16Wu Z. Puigserver P. Andersson U. Zhang C. Adelmant G. Mootha V. Troy A. Cinti S. Lowell B. Scarpulla R.C. Spiegelman B.M. Cell. 1999; 98: 115-124Abstract Full Text Full Text PDF PubMed Scopus (3220) Google Scholar). The reduction in oxidative enzyme capacity in patients with T2DM could be attributed to an increased proportion of glycolytic, type 2 muscle fibers (17Gaster M. Staehr P. Beck-Nielsen H. Schrøder H.D. Handberg A. Diabetes. 2001; 50: 1324-1329Crossref PubMed Scopus (215) Google Scholar). However, reduced oxidative enzyme capacity seems to be equally present in all muscle fiber types in patients with T2DM (5He J. Watkins S. Kelley D.E. Diabetes. 2001; 50: 817-823Crossref PubMed Scopus (399) Google Scholar). An increasing number of enzymes and metabolic pathways has been suggested to be involved in the development of skeletal muscle insulin resistance in T2DM. Therefore, there is a growing demand for techniques able to evaluate proteins from many signaling and metabolic pathways simultaneously. The recently introduced technique of proteome analysis, high resolution two-dimensional (2-D) gel electrophoresis followed by protein identification using mass spectrometry (MS) and data base searching, offers the possibility to study a large number of proteins and their post-translational modifications simultaneously (18Fey S.J. Larsen P.M. Curr. Opin. Chem. Biol. 2001; 5: 26-33Crossref PubMed Scopus (167) Google Scholar). In the past, primarily defects in the insulin-stimulated state have been studied. However, as discussed above, there are several reasons to look at T2DM as a phenotype described by poor adaptations to fasting conditions, as well (4Kelley D.E. Mandarino L.J. Diabetes. 2000; 49: 677-683Crossref PubMed Scopus (779) Google Scholar). To identify protein markers of T2DM in the fasting state, we compared the protein expression profile in skeletal muscle from obese patients with T2DM and age- and gender-matched healthy control subjects using the methods of quantitative proteome analysis. The study was approved by the Local Ethics Committee and was performed in accordance with the Helsinki Declaration. Muscle samples were obtained from 10 patients with T2DM and 10 healthy age- and gender-matched control subjects. Five patients with T2DM were treated by diet alone, and five patients were treated by low doses of either sulfonylurea or metformin. These drugs were withdrawn 1 week prior to the study. The patients were all GAD65 antibody-negative and without signs of diabetic retinopathy, nephropathy, neuropathy, or macrovascular complications. All subjects had normal results on screening blood tests of hepatic and renal function. The control subjects had normal glucose tolerance and no family history of diabetes. Participants were instructed to avoid vigorous exercise for 48 h before the study, which was carried out after a 10-h overnight fast. Fasting blood samples were analyzed for glucose (Glucose Analyzer II; Beckman Instruments, Fullerton, CA), free fatty acids (FFA) (Wako Chemicals Gmbh, Neuss, Germany), insulin, and C-peptide (Wallac Oy, Turku, Finland). Percutaneous needle biopsies were obtained from the vastus lateralis muscle under local anesthesia using a biopsy pistol, and the muscle specimens (∼25 mg) were immediately blotted free of blood, fat, and connective tissue and frozen in liquid nitrogen. The frozen muscle samples were homogenized for 25 min in 100 μl of ice-cold DNase/RNase buffer (20 mmol/liter Tris-HCl buffer, pH 7.5, containing 30 mmol/liter NaCl, 5 mmol/liter CaCl2, 5 mmol/liter MgCl2, and 25 μg/ml RNase A, 25 μg/ml DNase I (Worthington, Freehold, NJ)). After homogenization, the samples were lyophilized overnight and then dissolved in 120 μl of lysis buffer (7 m urea (ICN Biomedicals), 2 m thiourea (Fluka), 2% CHAPS (Sigma), 0.4% dithiothreitol (Sigma), 0.5% Pharmalyte 3–10, and 0.5% Pharmalyte 6–11 (Amersham Biosciences) by shaking overnight. The homogenization was carried out in a buffer without kinase and phosphatase inhibitors (salts) to avoid destruction of the first dimensional gels. As this step was carried out at 0–4 °C, we assumed the activity of potential kinases and phosphatases to be very low and that the phosphorylation state of most proteins was stable. To confirm this, we have, after the present study, performed similar 2-D gels, in which the muscle specimens were solubilized directly in the lysis buffer for running the first dimensional gel. The pattern of the proteins on these 2-D gels and in particular the relative abundance of the different phosphoisoforms identified by MS showed no difference from the 2-D gels presented in this study. The protein concentration in the muscle samples was determined using the Bradford method, which was adopted for use with lysis buffer as described before (19Fey S.J. Nawrocki A. Larsen M.R. Gorg A. Roepstorff P. Skews G.N. Williams R. Larsen P.M. Electrophoresis. 1997; 8: 1361-1372Crossref Scopus (71) Google Scholar). First dimension gel electrophoresis was performed on IPG covering the pH range from 4 to 7 (AmershamBiosciences). Rehydration buffer for IPG 4–7 strips was identical with lysis buffer used for sample preparation, and the sample was applied by in-gel rehydration. 400 μg of protein were loaded on each gel. Focusing was performed on a Multiphor II at 20 °C using a voltage/time profile linearly increasing from 0 to 600 V for 2.25 h, from 600 to 3500 V for 1 h, and 3500 V for 13.5 h. After focusing, strips were equilibrated twice, each for 15 min in equilibration buffer (6 m urea, 2% SDS, 30% glycerol, 50 mm Tris-HCl, pH 8.8, 1% dithiothreitol). Gels were frozen at −80 °C between the equilibration steps. SDS-PAGE second dimension was performed using the ProteanTM II Multi Cell 2-D electrophoresis system (Bio-Rad) and laboratory-made single percentage gels (12.5% acrylamide; acrylamide:N,N′-ethylene-bis-acrylamide ratio was 200:1). The gels were run overnight at 20 °C at constant current. Running buffer was recirculated to maintain pH, SDS, temperature, and salt concentrations. After the second dimension, skeletal muscle proteins were visualized using a silver-staining method as described (19Fey S.J. Nawrocki A. Larsen M.R. Gorg A. Roepstorff P. Skews G.N. Williams R. Larsen P.M. Electrophoresis. 1997; 8: 1361-1372Crossref Scopus (71) Google Scholar). All gel images were analyzed by the same person using a Bio Image computer program (version 6.1; Bob Luton, Ann Arbor, MI). For comparison we have used 2-D gels from six subjects in the control group and nine subjects in the diabetes group (Table I). The remaining gels revealed that some protein degradation had occurred, and therefore these gels were excluded from the analysis. The expression of each protein was measured and expressed as its percentage integrated optical density (%IOD) (a percentage of the sum of all the pixel gray level values on and within the boundary of the spot in question compared with that of all detected spots). Images from each group were then matched, edited, and compared statistically. The average value of spot %IOD and S.D. were calculated for each protein in each group and then compared using the two-sided Student's t test. Protein spots whose expression was found different between the two groups at the significance level of p < 0.05 were selected for further analysis. For correlation analysis Spearman's rho was used.Table IFasting characteristics of study subjectsControl groupDiabetic grouppn69Duration of diabetes1.1 ± 0.4Age (years)46.1 ± 1.544.8 ± 1.5NSBody mass index (kg/m2)25.7 ± 1.233.3 ± 1.90.01Male/female3/35/4NSCholesterol (mmol/l)5.1 ± 0.45.9 ± 0.4NSHbA1c5.3 ± 0.16.6 ± 0.40.02Glucose (mmol/l)5.3 ± 0.18.5 ± 0.70.002FFA (mmol/l)0.28 ± 0.040.44 ± 0.08NSInsulin (pmol/l)37 ± 1193 ± 120.005C-peptide (pmol/l)575 ± 1201195 ± 930.001 Open table in a new tab Proteins of interest were cut out from the gels and, after in-gel digestion, analyzed by mass spectrometry using a Bruker REFLEX matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometer (19Fey S.J. Nawrocki A. Larsen M.R. Gorg A. Roepstorff P. Skews G.N. Williams R. Larsen P.M. Electrophoresis. 1997; 8: 1361-1372Crossref Scopus (71) Google Scholar, 20Jensen O.N. Larsen M.R. Roepstorff P. Proteins. 1998; 2: 74-89Crossref PubMed Google Scholar). The mass spectra obtained were internally calibrated using trypsin autodigestion peptides, and the masses were used to search the NCBI data base using the ProFound, FindPept, and FindMod programs (www.proteometrics.com). Data base searches were performed using the following attributes with minor modification needed for each program: all species, no restrictions for molecular weight and protein pI, trypsin digest, one missed cleavage allowed, cysteines modified by acrylamide, and oxidation of methionines possible, mass tolerance between 0.1 and 0.5 Da. Identification was considered positive when at least five peptides matched the protein with no sequence overlap. Human skeletal muscle cell cultures were established as described previously (21Hurel S.J. Rochford J.J. Borthwick A.C. Wells A.M. Vandenheede J.R. Turnbull D.M. Yeamna S.J. Biochem. J. 1996; 320: 871-877Crossref PubMed Scopus (83) Google Scholar). Myoblasts were grown in 12-wells plates, and growing cell medium was changed to Dulbecco's modified Eagle's medium containing 5 mm glucose and supplemented with 10% fetal calf serum 1 day before the labeling experiment. Prior to labeling, cells were incubated in serum-free Dulbecco's modified Eagle's medium containing 0.2% bovine serum albumin for 2.5 h. Phosphate groups in proteins of human myoblasts were labeled biosynthetically by incubating them in 300 μl of serum-free phosphate-free Dulbecco's modified Eagle's medium (ICN Biomedicals) supplemented with 2 mml-glutamine (Invitrogen), 0.2% bovine serum albumin, and 300 μCi of [32P]orthophosphate (AmershamBiosciences) for 2.5 h. Immediately after, labeling medium was removed, and cells were lysed in 400 μl of lysis buffer as described above. Determination of [32P]orthophosphate incorporation into myoblast proteins was performed using trichloroacetic acid precipitation as described (19Fey S.J. Nawrocki A. Larsen M.R. Gorg A. Roepstorff P. Skews G.N. Williams R. Larsen P.M. Electrophoresis. 1997; 8: 1361-1372Crossref Scopus (71) Google Scholar). 2-D gel electrophoresis was run as described above loading a cell lysate volume corresponding to 4 × 105 cpm on the gel. [32P]labeled proteins of myoblasts were visualized by exposing dried gels to phosphorimaging plates (AGFA). Using computerized image analysis 489 spots in each gel image were matched and quantitated. Fifteen protein spots were expressed at statistically significant different levels in the two groups (Fig.1). These potential protein markers of T2DM were excised from the 2-D gels for identification by MALDI-TOF-MS analysis. Eleven of these protein spots were positively identified: three metabolic enzymes, two heat shock proteins, and different isoforms of three structural proteins (TableII). Of the metabolic enzymes, ATP synthase β-subunit (ATPsynβ) and creatine kinase, brain isoform (CK-B) were significantly down-regulated, whereas phosphoglucomutase-1 (PGM-1) was significantly up-regulated in skeletal muscle of patients with T2DM. Heat shock protein (HSP) 90β and 78-kDa glucose-regulated protein (GRP78) were both significantly up-regulated in diabetic muscle. Of the structural proteins, four isoforms of α-1 chain of type VI collagen (α1(VI) collagen) and one isoenzyme of myosin regulatory light chain 2 (MRLC2-B) were significantly up-regulated, whereas another isoenzyme of myosin regulatory light chain 2 (MRLC2-A) was significantly down-regulated in diabetic muscle.Table IIProtein markers of type 2 diabetes in skeletal muscle identified by MALDI-TOF MS analysisSpot no.ProteinDatabase accession no.Theoretical pl/mWSequence coverageMatched peptidesControl groupDiabetic groupp%%IOD ± S.E.180ATP synthase β-subunitP065765.0/5256190.56 ± 0.050.41 ± 0.030.01295Myosin regulatory light chain 2 (A)P109164.9/195891.26 ± 0.100.92 ± 0.100.03541Creatine kinase BP122775.3/4342120.32 ± 0.040.20 ± 0.020.0410Collagen α1(VI) chainP121095.3/11010100.06 ± 0.010.09 ± 0.010.0411Collagen α1(VI) chainP121095.3/110550.04 ± 0.000.06 ± 0.010.02407Collagen α1(VI) chainP121095.3/110660.05 ± 0.010.08 ± 0.010.03520Collagen α1(VI) chainP121095.3/110650.06 ± 0.010.09 ± 0.010.0281Heat shock protein 90βP082385.0/8415110.03 ± 0.010.05 ± 0.000.02375Glucose-regulated protein 78P110214.9/7828120.06 ± 0.010.08 ± 0.010.02445Myosin regulatory light chain 2 (B)AAK527974.9/195270.30 ± 0.080.69 ± 0.120.03456Phosphoglucomutase 1P368716.2/6225120.14 ± 0.040.28 ± 0.040.03 Open table in a new tab In the diabetic group the levels of ATPsynβ (spot 180) and CK-B (spot 541) correlated negatively with fasting plasma glucose values (r = −0.75; p < 0.05) and (r = −0.82; p < 0.01), respectively (Fig. 2, A and B), whereas the levels of α1(VI) collagen (spot 11) correlated positively with fasting plasma glucose values (r = 0.67;p < 0.05). Furthermore, the levels of ATPsynβ (spot 180) correlated positively with the levels of CK-B (spot 541) (r = 0.71; p < 0.05) (Fig.2 C) and negatively with the levels of α1(VI) collagen (spot 11) (r = −0.72; p < 0.05) in the diabetic group. In contrast, in the control group the levels of ATPsynβ (spot 180) correlated negatively with fasting FFA values (r = −0.81; p < 0.05) rather than fasting plasma glucose values (Fig. 2 D), and the levels of PGM-1 (spot 456) correlated positively with insulin (r = 0.95; p < 0.01). No other correlations among glucose, FFA, insulin, or C-peptide levels and the identified potential protein markers of T2DM were observed in the two groups. In particular, none of these protein markers correlated with body mass index either in the entire study population or within the study groups. CK-B, HSP 90β, GRP78, PGM-1, and MRLC2 have been reported previously (22Hemmer W. Skarli M. Perriard J.C. Wallimann T. FEBS Lett. 1993; 327: 35-40Crossref PubMed Scopus (20) Google Scholar, 23Joshi J.G. Zimmerman A. Toxicology. 1988; 48: 21-29Crossref PubMed Scopus (36) Google Scholar, 24Zhao Y. Gilmore R. Leone G. Coffey M.C. Weber B. Lee P.W.K. J. Biol. Chem. 2001; 276: 32822-32827Abstract Full Text Full Text PDF PubMed Scopus (91) Google Scholar, 25Hendershot L.M. Ting J. Lee A.S. Mol. Cell. Biol. 1988; 8: 4250-4256Crossref PubMed Scopus (162) Google Scholar, 26Manning D.R. Stull S. Am. J. Physiol. 1982; 242: C234-C241Crossref PubMed Google Scholar) to be phosphorylated. Several of the identified protein spots were located within a row of three-six protein spots with the same molecular weight indicating multiple post-translational modifications (possibly phosphorylation) of the same protein (Fig. 1). These protein spots were excised and subjected to MALDI-MS analysis. Two additional isoforms of HSP 90β, GRP78, PGM-1, α1(VI) collagen, MRLC2-A, and MRLC2-B and three additional isoforms of ATPsynβ were positively identified (Fig.3). Calculation of sums of expression of the different isoforms of each protein marker showed that the sum of expression of HSP 90β, GRP78, α1(VI) collagen, and ATPsynβ were significantly up- or down-regulated in the same direction as the protein markers (Table III). This was also the case with the sum of expression of MRLC2-A and MRLC2-B, although these differences did not reach statistical significance (Table III). The relative abundance of the down-regulated ATPsynβ isoform (spot 180) was significantly reduced in muscle of patients with T2DM, as well (p = 0.03), indicating a role for post-translational modification of this protein. Interestingly, in the diabetic group the sum of expression of ATPsynβ and α1(VI) collagen correlated significantly with fasting plasma glucose values (r = −0.76 and r = 0.70;p < 0.05, respectively), and the sum of expression of ATPsynβ correlated significantly with the sum of expression of α1(VI) collagen (r = −0.87; p < 0.01).Table IIISum of expression of protein markers isoformsProteinControl groupDiabetic group%IOD ± S.E.ATP synthase β-subunit2.28 ± 0.161.86 ± 0.10ap < 0.05 vs. control.Myosin regulatory light chain 2 (A)3.67 ± 0.213.16 ± 0.29Collagen α1(VI) chain0.29 ± 0.040.43 ± 0.03ap < 0.05 vs. control.Heat shock protein 90β0.09 ± 0.010.13 ± 0.01ap < 0.05 vs. control.Glucose-regulated protein 780.13 ± 0.010.17 ± 0.01ap < 0.05 vs. control.Myosin regulatory light chain 2 (B)3.64 ± 0.214.77 ± 0.67a p < 0.05 vs. control. Open table in a new tab The identification of four isoforms of ATPsynβ with identical molecular weights but different pI values, suggested that ATPsynβ is also phosphorylated in vivo (Fig. 3). To further characterize the modification of ATP synthase β-subunit, we carried out 2-D gel electrophoresis of [32P]labeled human skeletal muscle cells (myoblasts) in culture. These 2-D gels revealed that all four identified ATPsynβ isoforms are in fact phosphorylated isoforms (Fig.4 A) and that a putative non-modified variant was below the level of detection by silver staining. MALDI-MS analysis and data base searching for phosphopeptides from the tryptic digest of the three most basic phosphoisoforms of ATPsynβ, including the down-regulated ATPsynβ spot (180), demonstrated the presence of a phosphorylated residue most likely at position Thr-213 in the nucleotide-binding domain of ATPsynβ (Fig. 4,B and C). Tyrosine sulfation may give rise to the same pattern on 2-D gels and the same increase in peptide mass (80 Da) as phosphorylation. However, according to the consensus features of a tyrosine sulfation site, such sites are not present in the sequence of ATPsynβ. The [32P]labeling of all four ATPsynβ isoforms and the MS data therefore indicate that ATPsynβ is regulated by multi-site phosphorylation. Using the methods of proteome analysis we have identified and characterized eight potential protein markers of T2DM in skeletal muscle in the fasting state. Although proteome analysis is somewhat limited by narrow dynamic ranges of silver staining and difficulties in MS identification of low abundant proteins recovered from fixed and or stained gels, it is the only technique that provides the possibility to quantitatively study global changes in expression profile of proteins, as well as certain post-translational protein modifications in a given cell or tissue. Because of the method of sample preparation used in this study the interpretation of the relative abundance of the specific protein marker phosphoisoforms should be done with some caution. However, in the present study MS identification of ∼75% of the protein markers and several additional isoforms demonstrates that combined 2-D gel and MS technology can be powerful tools for the study of molecular processes underlying a complex disease such as T2DM. Our findings indicate that the catalytic β-subunit of F1-ATP synthase is regulated by phosphorylation and that the expression and phosphorylation of ATPsynβ might be altered in skeletal muscle of patients with T2DM. Because there is no report to date on the phosphorylation of human ATPsynβ it is uncertain how such post-translational modification interacts with the proposed binding-change model for ATP synthesis, in which conformational changes in the nucleotide-binding sites of the three β-subunits are coupled to the catalytic activity of F1-ATP synthase (27Menz R.I. Walker J.E. Leslie A.G.W. Cell. 2001; 106: 331-341Abstract Full Text Full Text PDF PubMed Scopus (398) Google Scholar). That phosphorylation of human ATPsynβ may play a role for the catalytic activity of F1-ATP synthase is suggested by the recent observation that phosphoserine/phosphothreonine-binding 14–3-3 proteins were found to be associated with mitochondrial ATP synthase in plants in a phosphorylation-dependent manner through direct interaction with the β-subunit of F1-ATP synthase and that the activity of ATP synthase was reduced by recombinant 14–3-3 protein (28Bunney T.D. van Walraven H.S. de Boer A.H. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 4249-4254Crossref PubMed Scopus (158) Google Scholar). Consistent with the reduced expression of ATPsynβ in our study, several other studies argue for a lower ATP synthase activity in diabetic muscle. Thus, gene expression of several subunits from the other four complexes of the mitochondrial electron transport chain were found to be decreased in skeletal muscle of streptozotocin-diabetic mice (29Yechoor V.K. Patti M. Saccone R. Kahn C.R. Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 10587-10592Crossref PubMed Scopus (140) Google Scholar), and during fasting conditions reduced activity of cytochrome c oxidase and citrate synthase activity, as well as reduced overall activity of the respiratory chain, have been demonstrated in skeletal muscle of patients with T2DM (4Kelley D.E. Mandarino L.J. Diabetes. 2000; 49: 677-683Crossref PubMed Scopus (779) Google Scholar, 5He J. Watkins S. Kelley D.E. Diabetes. 2001; 50: 817-823Crossref PubMed Scopus (399) Google Scholar, 6Kelley D.E. He J. Menshikova E.V. Ritov V.B. Diabetes. 2002; 51: 2944-2950Crossref PubMed Scopus (1782) Google Scholar). Moreover, mutations in mtDNA can, via impaired oxidative phosphorylation, cause T2DM, with both muscle insulin resistance and impaired insulin secretion (7Gebhart S. Shoffner J.M. Koontz D. Kaufmann A. Wallace D. Metabolism. 1996; 45: 526-531Abstract Full Text PDF PubMed Scopus (37) Google Scholar, 8Wallace D.C. Science. 1999; 283: 1482-1488Crossref PubMed Scopus (2630) Google Scholar). Indeed, decreased ATP synthase activity and increased formation of reactive oxygen species (ROS) have been reported in hybrids constructed from mitochondria of patients with T2DM (9Anderson C.M. Drug. Dev. Res. 1999; 46: 67-79Crossref Scopus (16) Google Scholar). However, whether the down-regulation of a specific ATPsynβ phosphoisoform is a mechanism to maintain normal activity despite reduced total levels of ATPsynβ or whether it reflects an altered activity of ATPsynβ per se warrants further studies. Endurance training up-regulates ATPsynβ protein levels in soleus muscle of rats (30González B. Hernando R.