Ubiquitin-Proteasome-mediated Degradation, Intracellular Localization, and Protein Synthesis of MyoD and Id1 during Muscle Differentiation
2005; Elsevier BV; Volume: 280; Issue: 28 Linguagem: Inglês
10.1074/jbc.m500373200
ISSN1083-351X
AutoresLiping Sun, Julie S. Trausch‐Azar, Aaron Ciechanover, Alan L. Schwartz,
Tópico(s)Cardiomyopathy and Myosin Studies
ResumoMammalian skeletal myogenesis results in the differentiation of myoblasts to mature syncytial myotubes, a process regulated by an intricate genetic network of at least three protein families: muscle regulatory factors, E proteins, and Id proteins. MyoD, a key muscle regulatory factor, and its negative regulator Id1 have both been shown to be degraded by the ubiquitin-proteasome system. Using C2C12 cells and confocal fluorescence microscopy, we showed that MyoD and Id1 co-localize within the nucleus in proliferating myoblasts. In mature myotubes, in contrast, they reside in distinctive subcellular compartments, with MyoD within the nucleus and Id1 exclusively in the cytoplasm. Cellular abundance of Id1 was markedly diminished from the very onset of muscle differentiation, whereas MyoD abundance was reduced to a much lesser extent and only at the later stages of differentiation. These reductions in MyoD and Id1 protein levels seem to result from a change in the rate of protein synthesis rather than the rate of degradation. In vivo protein stability studies revealed that the rates of ubiquitin-proteasome-mediated MyoD and Id1 degradation are independent of myogenic differentiation state. Id1 and MyoD were both rapidly degraded, each with a t½ ≃ 1 h in myoblasts and in myotubes. Furthermore, relative protein synthesis rates for MyoD and Id1 were significantly diminished during myoblast to myotube differentiation. These results provide insight as to the interaction between MyoD and Id1 in the process of muscle differentiation and have implications for the involvement of the ubiquitin-proteasome-mediated protein degradation and protein synthesis in muscle differentiation and metabolism under abnormal and pathological conditions. Mammalian skeletal myogenesis results in the differentiation of myoblasts to mature syncytial myotubes, a process regulated by an intricate genetic network of at least three protein families: muscle regulatory factors, E proteins, and Id proteins. MyoD, a key muscle regulatory factor, and its negative regulator Id1 have both been shown to be degraded by the ubiquitin-proteasome system. Using C2C12 cells and confocal fluorescence microscopy, we showed that MyoD and Id1 co-localize within the nucleus in proliferating myoblasts. In mature myotubes, in contrast, they reside in distinctive subcellular compartments, with MyoD within the nucleus and Id1 exclusively in the cytoplasm. Cellular abundance of Id1 was markedly diminished from the very onset of muscle differentiation, whereas MyoD abundance was reduced to a much lesser extent and only at the later stages of differentiation. These reductions in MyoD and Id1 protein levels seem to result from a change in the rate of protein synthesis rather than the rate of degradation. In vivo protein stability studies revealed that the rates of ubiquitin-proteasome-mediated MyoD and Id1 degradation are independent of myogenic differentiation state. Id1 and MyoD were both rapidly degraded, each with a t½ ≃ 1 h in myoblasts and in myotubes. Furthermore, relative protein synthesis rates for MyoD and Id1 were significantly diminished during myoblast to myotube differentiation. These results provide insight as to the interaction between MyoD and Id1 in the process of muscle differentiation and have implications for the involvement of the ubiquitin-proteasome-mediated protein degradation and protein synthesis in muscle differentiation and metabolism under abnormal and pathological conditions. Skeletal muscle differentiation is characterized by the terminal withdrawal of the myoblast from the cell cycle, activation of muscle-specific gene expression, and cell fusion into multinucleated myotubes. These events are coordinated by a family of four muscle-specific basic helix-loop-helix transcription factors, MyoD, Myf5, myogenin, and Mrf4, termed the muscle regulatory factors (1Puri P.L. Sartorelli V. J. Cell. Physiol. 2000; 185: 155-173Crossref PubMed Scopus (250) Google Scholar, 2Rudnicki M.A. Jaenisch R. BioEssays. 1995; 17: 203-209Crossref PubMed Scopus (365) Google Scholar, 3Yun K. Wold B. Curr. Opin. Cell Biol. 1996; 8: 877-889Crossref PubMed Scopus (318) Google Scholar, 4Pownall M.E. Gustafsson M.K. Emerson Jr., C.P. Annu. Rev. Cell Dev. Biol. 2002; 18: 747-783Crossref PubMed Scopus (457) Google Scholar). Mice lacking myogenin appropriately specify the skeletal muscle lineage but fail to terminally differentiate. Mrf4 is required for the maintenance of the differentiated myotubes. Although the specification of the myogenic lineage requires myoD and myf5, as double knock-out of both genes yields mice with no skeletal muscle (5Braun T. Arnold H.H. EMBO J. 1996; 15: 310-318Crossref PubMed Scopus (96) Google Scholar), MyoD is also required for healthy self-renewing proliferation of the adult skeletal muscle satellite cells (6Megeney L.A. Kablar B. Garrett K. Anderson J.E. Rudnicki M.A. Genes Dev. 1996; 10: 1173-1183Crossref PubMed Scopus (542) Google Scholar, 7Cooper R.N. Tajbakhsh S. Mouly V. Cossu G. Buckingham M. Butler-Browne G.S. J. Cell Sci. 1999; 112: 2895-2901Crossref PubMed Google Scholar, 8Zammit P.S. Golding J.P. Nagata Y. Hudon V. Partridge T.A. Beauchamp J.R. J. Cell Biol. 2004; 166: 347-357Crossref PubMed Scopus (665) Google Scholar). Muscle regulatory factors form heterodimers with ubiquitous E proteins and activate myogenic differentiation through their subsequent binding to specific sequences, termed E boxes, in the promoter regulatory regions of muscle-restricted target genes (4Pownall M.E. Gustafsson M.K. Emerson Jr., C.P. Annu. Rev. Cell Dev. Biol. 2002; 18: 747-783Crossref PubMed Scopus (457) Google Scholar). The transcriptional activities of muscle regulatory factors are negatively regulated by a family of inhibitors of DNA-binding (Id) proteins. The four Id proteins (Id1, Id2, Id3, and Id4) are helix-loop-helix proteins that contain no basic region and thus do not bind DNA. However, they are able to dimerize with one another and with MyoD or E proteins, albeit with different affinities (9Edmondson D.G. Olson E.N. J. Biol. Chem. 1993; 268: 755-758Abstract Full Text PDF PubMed Google Scholar, 10Benezra R. Rafii S. Lyden D. Oncogene. 2001; 20: 8334-8341Crossref PubMed Google Scholar, 11Langlands K. Yin X. Anand G. Prochownik E.V. J. Biol. Chem. 1997; 272: 19785-19793Abstract Full Text Full Text PDF PubMed Scopus (196) Google Scholar). Id1 is most active in terms of MyoD binding. The binding affinity of Id1 for the E proteins is considerably higher than its affinity for MyoD. Sequestering the ubiquitous E proteins allows Id1 to control the transcriptional activity of muscle-specific MyoD. In cultured myoblasts, Id1 over-expression via a "dominant-negative" effect inhibits the transactivation by MyoD, thereby inhibiting the synthesis of proteins participating in muscle differentiation and consequently the fusion of myoblasts to myotubes (12Jen Y. Weintraub H. Benezra R. Genes Dev. 1992; 6: 1466-1479Crossref PubMed Scopus (393) Google Scholar). MyoD and Id1 have both been shown to be degraded by the ubiquitin-proteasome system (13Floyd Z.E. Trausch-Azar J.S. Reinstein E. Ciechanover A. Schwartz A.L. J. Biol. Chem. 2001; 276: 22468-22475Abstract Full Text Full Text PDF PubMed Scopus (63) Google Scholar, 14Lingbeck J.M. Trausch-Azar J.S. Ciechanover A. Schwartz A.L. J. Biol. Chem. 2003; 278: 1817-1823Abstract Full Text Full Text PDF PubMed Scopus (50) Google Scholar, 15Trausch-Azar J.S. Lingbeck J. Ciechanover A. Schwartz A.L. J. Biol. Chem. 2004; 279: 32614-32619Abstract Full Text Full Text PDF PubMed Scopus (47) Google Scholar, 16Ciechanover A. Breitschopf K. Hatoum O.A. Bengal E. Mol. Biol. Rep. 1999; 26: 59-64Crossref PubMed Google Scholar, 17Breitschopf K. Bengal E. Ziv T. Admon A. Ciechanover A. EMBO J. 1998; 17: 5964-5973Crossref PubMed Scopus (232) Google Scholar, 18Fajerman I. Schwartz A.L. Ciechanover A. Biochem. Biophys. Res. Commun. 2004; 314: 505-512Crossref PubMed Scopus (59) Google Scholar, 19Abu Hatoum O. Gross-Mesilaty S. Breitschopf K. Hoffman A. Gonen H. Ciechanover A. Bengal E. Mol. Cell. Biol. 1998; 18: 5670-5677Crossref PubMed Google Scholar). This pathway involves the activation of ubiquitin by the ubiquitin-activating enzyme, E1, followed by transfer of ubiquitin to E2, a ubiquitin-conjugating enzyme. E2 shuttles the ubiquitin to the substrate-specific ubiquitin ligase, E3, which then delivers the ubiquitin to the protein substrate to be degraded. The ubiquitin-proteasome proteolytic system is recognized as a versatile and efficient mechanism for the control of gene expression. The level of expression of MyoD and Id1 are vitally important during muscle differentiation. Both mRNA and protein levels of Id1 are down-regulated upon initiation of differentiation (20Benezra R. Davis R.L. Lockshon D. Turner D.L. Weintraub H. Cell. 1990; 61: 49-59Abstract Full Text PDF PubMed Scopus (1785) Google Scholar). MyoD mRNA levels change only slightly during differentiation (21Shimokawa T. Kato M. Ezaki O. Hashimoto S. Biochem. Biophys. Res. Commun. 1998; 246: 287-292Crossref PubMed Scopus (84) Google Scholar), yet the level of MyoD protein is decreased in reserve cells during muscle differentiation (22Yoshida N. Yoshida S. Koishi K. Masuda K. Nabeshima Y. J. Cell Sci. 1998; 111: 769-779Crossref PubMed Google Scholar). Reducing the level of MyoD protein by its destabilization has been shown to be associated with the inhibition of myogenic differentiation under abnormal or pathophysiological conditions. For example, accelerated MyoD degradation resulting from hypoxia blocked the accumulation of early myogenic differentiation markers such as myogenin, p21, and pRb and prevented both permanent cell cycle withdraw and terminal differentiation (23Di Carlo A. De Mori R. Martelli F. Pompilio G. Capogrossi M.C. Germani A. J. Biol. Chem. 2004; 279: 16332-16338Abstract Full Text Full Text PDF PubMed Scopus (105) Google Scholar). In addition, tumor necrosis factor α inhibits myogenic differentiation through destabilizing MyoD protein in a NF-κB-dependent manner, interferes with skeletal muscle regeneration, and may contribute to muscle wasting (24Acharyya S. Ladner K.J. Nelsen L.L. Damrauer J. Reiser P.J. Swoap S. Guttridge D.C. J. Clin. Investig. 2004; 114: 370-378Crossref PubMed Scopus (401) Google Scholar, 25Guttridge D.C. Mayo M.W. Madrid L.V. Wang C.Y. Baldwin Jr., A.S. Science. 2000; 289: 2363-2366Crossref PubMed Scopus (740) Google Scholar, 26Langen R.C. Van Der Velden J.L. Schols A.M. Kelders M.C. Wouters E.F. Janssen-Heininger Y.M. FASEB. J. 2004; 18: 227-237Crossref PubMed Scopus (247) Google Scholar). Studies in a cell culture model that has a phenotype similar to that observed in myoblast cultures derived from myotonic dystrophy 1 patient muscle suggest that C2C12 myogenic differentiation is disrupted by mutant myotonic dystrophy protein kinase 3′-untranslated region transcripts via posttranscriptional reduction of MyoD protein levels (27Amack J.D. Reagan S.R. Mahadevan M.S. J. Cell Biol. 2002; 159: 419-429Crossref PubMed Scopus (45) Google Scholar). Previous studies in vitro or in non-muscle cells also show that MyoD degradation is regulated by phosphorylation, DNA binding, and protein-protein interactions (19Abu Hatoum O. Gross-Mesilaty S. Breitschopf K. Hoffman A. Gonen H. Ciechanover A. Bengal E. Mol. Cell. Biol. 1998; 18: 5670-5677Crossref PubMed Google Scholar, 28Song A. Wang Q. Goebl M.G. Harrington M.A. Mol. Cell. Biol. 1998; 18: 4994-4999Crossref PubMed Scopus (138) Google Scholar). Phosphorylation of MyoD is required for its rapid degradation. The specific DNA sequence to which MyoD binds can inhibit MyoD degradation. Id1, which inhibits the binding of MyoD complexes to DNA, abrogates the effect of DNA on the stabilization of MyoD. Furthermore, protein degradation studies following co-transfection of MyoD and Id1 to HeLa cells have shown that MyoD is able to modulate both the localization and the degradation of Id1 (15Trausch-Azar J.S. Lingbeck J. Ciechanover A. Schwartz A.L. J. Biol. Chem. 2004; 279: 32614-32619Abstract Full Text Full Text PDF PubMed Scopus (47) Google Scholar). It remains unclear, however, how the morphological and biological changes involved in myogenic differentiation affect the ubiquitin-proteasome-mediated degradation of MyoD and Id1 in muscle cells and how their degradation and interaction contribute to their cellular abundance and thus regulate differentiation. Mouse C2C12 myoblast cells are a well characterized myogenic cell line. In the presence of mitogen-rich serum, they proliferate as an undifferentiated population expressing MyoD and Myf5. Terminal differentiation of C2C12 cells may be induced by serum deprivation, which initiates a series of chronologically ordered events with expression of myogenin as the earliest event detected. Thereafter, cells permanently withdraw from the cell cycle, contractile proteins such as myosin begin to accumulate, and cell fusion then takes place resulting in the formation of myotubes. Using C2C12 cells as our model, we show here that MyoD and Id1 co-localize within the nucleus in proliferating myoblasts. In mature myotubes, in contrast, they reside in distinctive subcellular compartments, with MyoD within the nucleus and Id1 exclusively in the cytoplasm. Both MyoD and Id1 are rapidly degraded by the ubiquitin-proteasome pathway during the differentiation of myoblast to myotube. Furthermore, the rate of their degradation appears to be unaffected by the differentiation state, whereas a reduction of MyoD and Id1 synthesis rate was observed during myogenic differentiation. Plasmids and Construction of Id1-HA—Wild type MyoD in pCIneo and Id1 in pcDNA3 have been described previously (15Trausch-Azar J.S. Lingbeck J. Ciechanover A. Schwartz A.L. J. Biol. Chem. 2004; 279: 32614-32619Abstract Full Text Full Text PDF PubMed Scopus (47) Google Scholar). Id1 with a 1× HA 1The abbreviations used are: HA, hemagglutinin; GM, growth medium; DM, differentiation medium; TRITC, tetramethylrhodamine isothiocyanate; CHX, cycloheximide; RT, reverse transcription; DAPI, 4′,6-diamidino-2-phenylindole. tag at the C terminus was constructed by the insertion of the HA tag into pcDNA3 harboring wild type Id1 encoding DNA. PCR primers were purchased from Integrated DNA Technologies. DNA sequencing using Big Dye version 3.1 (ABI Biosystems) was used to confirm all sequences. Cell Culture—The C2C12 mouse myoblast cell line was obtained from the American Type Culture Collection. The cells were routinely propagated in growth medium (GM) and Dulbecco's modified Eagle's medium (Sigma) supplemented with 10% fetal bovine serum, 100 units/ml penicillin G, and 100 μg/ml streptomycin (Invitrogen) and maintained in a humidified chamber at 37 °C with 5% CO2. Myogenic differentiation was induced by changing the growth medium to differentiation medium (DM) and Dulbecco's modified Eagle's medium supplemented with 2% horse serum (HyClone), 100 units/ml penicillin G, and 100 μg/ml streptomycin when cells reached confluence. Cells were then maintained in differentiation medium for 6 days with medium being changed every 24 h. Transient transfections of C2C12 myoblasts were performed using the FuGENE 6 reagent (Roche Applied Science) according to the manufacturer's instruction. Immunofluorescence—C2C12 cells on glass coverslips were first washed with PBSc, a phosphate-buffered saline solution (PBSa, Fisher) supplemented with 100 mm CaCl2 and 50 mm MgCl2, fixed in 4% paraformaldehyde, quenched in 0.1 m ethanolamine (pH 8.0), and permeabilized in 1% Triton X-100 (Sigma). Subcellular localization of MyoD, Id1, myogenin, and myosin heavy chain in C2C12 myoblasts or myotubes was then determined by indirect immunofluorescence using the rabbit polyclonal antibodies (anti-MyoD (C-20), anti-Id1 (C-20), anti-myogenin (M-225), and anti-myosin heavy chain (H-300), Santa Cruz Biotechnology) followed by incubation with Alexa Fluor 568 goat anti-rabbit IgG (heavy and light chains) (Molecular Probes) after blocking the cells in PBSc containing 1% bovine serum albumin and 0.01% TW-80. Cells were observed with a Zeiss Axioskop microscope, and images were taken with a Zeiss AxioCam digital camera. Blocking control experiments for the detection of MyoD and Id1 were performed by preincubating the probing primary antibody with the corresponding peptides (MyoD (C-20)P and Id1 (C-20)P, Santa Cruz Biotechnology) that were initially used to generate the antibodies. For double immunofluorescence of MyoD and Id1, a mouse monoclonal anti-MyoD (Novocastra) coupled with TRITC-conjugated donkey anti-mouse IgG (Jackson ImmunoResearch Laboratories) and the rabbit polyclonal anti-Id1 (C-20) coupled with fluorescein isothiocyanate-conjugated goat anti-rabbit IgG (Jackson ImmunoResearch Laboratories) are used as probes. Cells were viewed with a confocal laser scanning microscope (Fluoview 500, Olympus) with a 60× oil immersion objective lens. Protein Expression Level during Myogenic Differentiation—C2C12 cells were washed with PBSa twice on ice and harvested at different stages of myogenic differentiation (day -1, 70-80% confluent C2C12 myoblasts in GM; day 0, 100% confluent C2C12 cells and the time point for switching the medium to DM; day 1 to day 6, C2C12 cells cultured in DM for 1 to 6 days). All of the cells were lysed for at least 30 min in PBSa containing 5% Igepal, 1 mm dithiothreitol, 1 μm pepstatin, 2.5 μg/ml leupeptin, and 0.2 mm phenylmethylsulfonyl fluoride. The collected cells were then sonicated briefly and centrifuged at 14,000 rpm for 10 min at 4 °C to remove cellular debris. Protein concentration of each freshly prepared cell lysate was determined by Bradford assay (Bio-Rad). After mixing with an equal volume of 2× Laemmli sample buffer (Bio-Rad), cell lysates containing equal amounts of total protein were loaded in each lane, resolved by SDS-PAGE, and transferred onto nitrocellulose membrane (Osmonics). Membranes were probed with rabbit polyclonal antibodies (anti-MyoD (C-20), anti-Id1 (Z-8), anti-myogenin (M-225), and anti-myosin heavy chain (H-300), Santa Cruz Biotechnology) followed by incubation with a secondary horseradish peroxidase-conjugated antibody. Protein bands were detected by chemiluminescence (ECL, Amersham Biosciences). Blocking control experiments for MyoD Western blot were performed by preincubating the probing primary antibody with peptide (MyoD (C-20)P, Santa Cruz Biotechnology) before applying to the membrane. Determination of Protein Degradation Half-life—C2C12 cells were incubated with cycloheximide (CHX, 100 μg/ml, Sigma) to inhibit further protein synthesis. The proteasome inhibitor MG132 (N-benzyloxy-carbonyl-Leu-Leu-leucinal, 20 μm, International Peptides) was added along with CHX when necessary. Following incubation for 0, 0.5, 1, 2, and 3 h, cells were harvested and lysed, and cell lysates were collected as described above. After treatment with Laemmli sample buffer (Bio-Rad), equal volumes of each sample were loaded in each lane for gel electrophoresis. Western blotting was performed in the same fashion as described above. A rabbit polyclonal anti-HA antibody (Upstate) was used for the detection of Id1-HA protein. Desired protein bands from the Western blottings were quantitated using the EDAS system (Eastman Kodak Co.), and the data were graphed using the EXCEL graphing program (Microsoft). Protein degradation rate is expressed as half-life (t½), the time for degradation of 50% of the protein. Each of the half-life data reported was evaluated by three to six independent determinations and is expressed as mean ± S.D. Comparison of Protein Synthesis Rate—C2C12 cells were incubated with MG132 (20 μm). Following incubation for 0, 1, 2, and 4 h, cells were harvested and lysed, and cell lysates were collected as described above. After treatment with Laemmli sample buffer (Bio-Rad), equal volumes of each sample were loaded in each lane for gel electrophoresis. Western blottings were performed in the same fashion as described above. The pixels for each band were measured and normalized so that the number of pixels at t = 0 was 1. The pixels of each band were plotted versus time. Protein synthesis rates were compared based on the initial slope from plots of data from 0 to 4 h. RNA Isolation and RT-PCR—Total RNA samples from C2C12 myoblasts and differentiated myotubes were obtained using RNAzol reagents (Tel-Test, Inc.) following the manufacturer's instructions. RT-PCR was performed for Id1, MyoD, and glyceraldehyde 3-phosphate dehydrogenase using ProSTAR Ultra HF RT-PCR system (Stratagene). The Id1 primers are as follows: 5′-TGGACGAGCAGCAGGTGAACG-3′ and 5′-GCACTGATCTCGCCGTTCAGG-3′ (with a product of 243 bp). The MyoD primers are: 5′-GACAGGACAGGACAGGGAGG-3′ and 5′-GCACCGCAGTAGAGAAGTGT-3′ (with a product of 358 bp). To study MyoD and Id1 degradation during muscle differentiation, we chose the murine C2C12 myoblast cells as our model (12Jen Y. Weintraub H. Benezra R. Genes Dev. 1992; 6: 1466-1479Crossref PubMed Scopus (393) Google Scholar, 29Liu C.J. Ding B. Wang H. Lengyel P. Mol. Cell. Biol. 2002; 22: 2893-2905Crossref PubMed Scopus (75) Google Scholar). By shifting confluent C2C12 cells from mitogen-rich growth medium to 2% horse serum containing differentiation medium, we were able to observe the expected morphological changes of C2C12 myogenic differentiation, namely cell alignment and fusion into multinucleated myotubes (Fig. 1A). By day 1 in DM, C2C12 cells were compact and aligned, but there was no obvious cell fusion. By day 2 in DM, about 5% of cells were fused to form scattered, small multinucleated cells. During the next 2 days, the cell fusion rate was rapid. By day 4 in DM, about 50% of cells completed fusion, and a large number of tubular syncytial cells were observed with various numbers of nuclei mostly arranged in linear arrays. These myotubes further expanded by incorporating additional neighboring mononucleated cells and via fusion with nearby myotubes and increased in size. By day 6 in DM, fused cells accounted for up to 70% of the cells, whereas a subpopulation of cells remained undifferentiated as reserve cells (22Yoshida N. Yoshida S. Koishi K. Masuda K. Nabeshima Y. J. Cell Sci. 1998; 111: 769-779Crossref PubMed Google Scholar). The observed morphology transition from myoblasts to myotubes was also confirmed by immunofluorescent staining of myosin heavy chain, a major muscle contractile protein, coupled with DAPI nuclear staining. As a marker for mature muscle cells, myosin heavy chain was detected only in myotubes with multi-DAPI-positive nuclei inside single cells (Fig. 1A). Previous studies have shown that MyoD subcellular localization markedly influences its degradation rate and that the degradation of Id1 can be modulated by MyoD (14Lingbeck J.M. Trausch-Azar J.S. Ciechanover A. Schwartz A.L. J. Biol. Chem. 2003; 278: 1817-1823Abstract Full Text Full Text PDF PubMed Scopus (50) Google Scholar, 15Trausch-Azar J.S. Lingbeck J. Ciechanover A. Schwartz A.L. J. Biol. Chem. 2004; 279: 32614-32619Abstract Full Text Full Text PDF PubMed Scopus (47) Google Scholar). To determine the locus of MyoD and Id1 degradation during differentiation and the possible interaction between this pair, we first set out to determine the localization of MyoD and Id1 in C2C12 myoblasts and myotubes (Fig. 1B). By indirect immunofluorescent examination, MyoD was localized to the cell nucleus in mononucleated myoblasts as well as in multinucleated myotubes. In myoblasts, Id1 was observed to be predominantly co-localized to the nucleus with MyoD, with low level cytoplasmic staining, suggesting that they interact in vivo. In contrast, MyoD and Id1 appeared to reside in distinctive subcellular compartments in myotubes, with MyoD within the nucleus and Id1 exclusively in the cytoplasm. Preincubation of the probing antibody with its corresponding peptide completely abolished the observed immunofluorescent signals for MyoD and Id1 in our blocking control experiments (data not shown), demonstrating that the localization of both proteins in myoblasts and myotubes is not the result of nonspecific binding. We then determined MyoD and Id1 protein abundance during the time course of differentiation via immunoblot. Cellular abundance of Id1 is markedly diminished from the very onset of muscle differentiation, whereas MyoD abundance is reduced to a much lesser extent and only at the later stages of differentiation (Fig. 2). After only 2 days in differentiation medium, the Id1 protein level had already decreased to less than 20% of that found in proliferating myoblasts. During this same period of differentiation, the MyoD protein level remained about the same, if not somewhat increased. By day 6 in differentiation medium Id1 protein was reduced 10-fold, whereas MyoD protein was reduced less than 3-fold. Overall, the relative ratio between MyoD and Id1 increased as the cells committed to differentiation. The abundance of the myogenic regulatory factor myogenin and the contractile protein myosin heavy chain were also determined in the same cell lysates (Fig. 2). Myogenin, which promotes terminal differentiation, became detectable the first day in differentiation medium after cells reached confluence. We also observed progressive accumulation of myosin heavy chain from day 1 onward. These observations verified that our model exhibited the biological characteristics of muscle differentiation in a chronologically appropriate manner. To determine how ubiquitin-proteasome-mediated degradation contributes to the regulation of MyoD and Id1 abundance during muscle differentiation, we first compared their protein degradation half-lives in C2C12 myoblasts and myotubes. As shown in Fig. 3, MyoD was rapidly degraded in both cases with the same half-life (t½ ≃ 0.9 h). Incubation of cells with MG132, a potent and selective inhibitor of the proteasome, markedly slowed the rate of MyoD degradation (t½ ≥ 10 h). For Id1, rapid degradation was also seen both in myoblasts and in myotubes (t½ ≃ 0.8 h). The presence of MG132 greatly stabilized Id1 (t½ > 10 h). All of these results suggested that both MyoD and Id1 are degraded by the ubiquitin-proteasome pathway during muscle differentiation and that the degradation rate is unchanged in myoblasts and in myotubes regardless of the morphological and biological changes. Herein, notably only the hyperphosphorylated MyoD species (i.e. the top, slower migrating, bands) in Fig. 3A was used to calculate the half-life. This practice is somewhat different from that used in previous reports in which both of the MyoD bands, although not well separated in many cases, are used for the calculation of MyoD degradation. We believe that this method best serves our purpose because MyoD degradation requires its prior phosphorylation (28Song A. Wang Q. Goebl M.G. Harrington M.A. Mol. Cell. Biol. 1998; 18: 4994-4999Crossref PubMed Scopus (138) Google Scholar) and because the hypophosphorylated band appears to be much more stable. Although no change was observed in the degradation rate of MyoD or Id1 in myoblasts (day -1) and myotubes (day 6), the possibility remains that MyoD or Id1 could be degraded at a different rate at a specific stage between the onset of muscle differentiation and its completion. We thus determined MyoD and Id1 degradation rates at various time points throughout C2C12 differentiation. As seen in Fig. 4, MyoD, as well as Id1, was degraded at about the same rate during differentiation from proliferating, mononucleated myoblasts to mature syncytial muscle cells. These results suggest that ubiquitin-proteasome-mediated MyoD and Id1 degradation is independent of the muscle differentiation state. Studies in HeLa cells following co-transfection of MyoD and Id1 have suggested that MyoD can modulate the rate of Id1 degradation (15Trausch-Azar J.S. Lingbeck J. Ciechanover A. Schwartz A.L. J. Biol. Chem. 2004; 279: 32614-32619Abstract Full Text Full Text PDF PubMed Scopus (47) Google Scholar). We have shown that MyoD and Id1 co-localize in the nucleus of myoblasts (Fig. 1B), suggesting that there is possible interaction between the pair. To determine whether the relative abundance of MyoD or Id1 influences the degradation rate, we over-expressed MyoD or Id1 in myoblasts by transient transfection. Following the transient transfection of MyoD or Id1 to myoblasts, each was rapidly degraded (t½ ≃ 0.9 h for MyoD and t½ ≃ 0.8 h for Id1) at the same rate as was observed for endogenous MyoD or Id1 in C2C12 myoblasts (Fig. 5). Furthermore, incubation with MG132 greatly stabilized the degradation of the protein in each case. An HA tag was attached to the C terminus of the Id1 protein to allow discrimination from the endogenous protein. Western blots using both anti-Id1 and anti-HA antibodies yielded identical half-lives. Because no obvious alteration in protein stability was seen for MyoD or Id1 during myogenic differentiation, we compared their protein synthesis rates in myoblasts and myotubes. We took advantage of MG132, the proteasome inhibitor, and determined the rate of MyoD and Id1 accumulation. Under the experimental conditions in which MyoD and Id1 protein degradation was abolished, the relative rate of MyoD and Id1 protein accumulation correlated with the relative rates of protein synthesis in vivo. As seen in Fig. 6, within2hof incubation in MG132, the rate of accumulation of both Id1 and MyoD was substantially slower in myotubes than in myoblasts, suggesting that the rate of Id1 (∼4-fold) and MyoD (∼3.5-fold) protein synthesis is more rapid in myoblasts than in myotubes. To determine whether the down-regulation of the protein synthesis rates is a general phenomenon of myogenic differentiation, similar studies were also performed on a variety of other proteins. For several, we observed no decrease in protein synthes
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