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Endocrinology Vol. 148, No. 3 1108-1117
Copyright © 2007 by The Endocrine Society

Inositide-Dependent Phospholipase C Signaling Mimics Insulin in Skeletal Muscle Differentiation by Affecting Specific Regions of the Cyclin D3 Promoter

Irene Faenza, Giulia Ramazzotti, Alberto Bavelloni, Roberta Fiume, Gian Carlo Gaboardi, Matilde Y. Follo, R. Stewart Gilmour, Alberto M. Martelli, Katya Ravid and Lucio Cocco

Cellular Signalling Laboratory (I.F., G.R., R.F., G.C.G., M.Y.F., A.M.M., L.C.), Department of Human Anatomical Science, University of Bologna, 40126 Bologna, Italy; Laboratory of Cell Biology and Electron Microscopy (A.B.), Istituti Ortopedici Rizzoli, 40136 Bologna, Italy; Department of Pathology (R.S.G.), University of Cambridge, Cambridge CB2 1QP, United Kingdom; and Departments of Biochemistry and Medicine and Whitaker Cardiovascular Institute (K.R.), Boston University School of Medicine, Boston, Massachusetts 02118

Address all correspondence and requests for reprints to: Dr. Lucio Cocco, Cellular Signalling Laboratory, Department of Human Anatomical Sciences, University of Bologna, Via Irnerio 48, 40126 Bologna, Italy. E-mail: lcocco{at}biocfarm.unibo.it.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Our main goal in this study was to investigate the role of phospholipase C (PLC) ß1 and PLC{gamma}1 in skeletal muscle differentiation and the existence of potential downstream targets of their signaling activity. To examine whether PLC signaling can modulate the expression of cyclin D3, a target of PLCß1 in erythroleukemia cells, we transfected C2C12 cells with expression vectors containing PLCß1 or PLC{gamma}1 cDNA and with small interfering RNAs from regions of the PLCß1 or PLC{gamma}1 gene and followed myogenic differentiation in this well-established cell system. Intriguingly, overexpressed PLCß1 and PLC{gamma}1 were able to mimic insulin induction of both cyclin D3 and muscle differentiation. By knocking down PLCß1 or PLC{gamma}1 expression, C2C12 cells almost completely lost the increase in cyclin D3, and the differentiation program was down-regulated. To explore the induction of the cyclin D3 gene promoter during this process, we used a series of 5'-deletions of the 1.68-kb promoter linked to a reporter gene and noted a 5-fold augmentation of promoter activity upon insulin stimulation. These constructs were also cotransfected with PLCß1 or PLC{gamma}1 cDNAs and small interfering RNAs, respectively. Our data indicate that PLCß1 or PLC{gamma}1 signaling is capable of acting like insulin in regard to both the myogenic differentiation program and cyclin D3 up-regulation. Taken together, this is the first study that hints at cyclin D3 as a target of PLCß1 and PLC{gamma}1 during myogenic differentiation in vitro and implies that up-regulation of these enzymes is sufficient to mimic the actions of insulin in this process.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
MOUSE C2C12 CELLS have been used extensively to study the process of myogenic differentiation. Skeletal muscle differentiation is characterized by terminal withdrawal from the cell cycle, the activation of muscle-specific gene expression, and morphological changes including myoblast alignment, elongation, and fusion of mononucleated myotubes (1). These events are coordinated by a family of four muscle-specific basic helix-loop-helix transcription factors: MyoD1, Myf5, myogenin, and Mrf4, termed the muscle regulatory factors (MRFs). MRFs 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 (2). This process is stimulated by growth factors such as insulin and IGF-I and -II (3, 4). According to the primary structures, phospholipase C (PLC) is divided into four types: ß, {gamma}, {delta}, and {eta} (5) each of which consists of several subtypes (6). Phosphatidylinositol (PI)-PLC catalyzes the hydrolysis of phosphatidylinositol 4,5-bisphosphate to generate the second messengers inositol-1,4,5-trisphosphate (IP3) and diacylglycerol (DAG). PLCß1 has been shown to reside within the nucleus in many cell lines (7). The nuclear localization of this enzyme is determined by a cluster of lysine residues between positions 1055 and 1072 (8). Nuclear PLCß1 is the key enzyme responsible for the initiation of the nuclear phosphoinositide cycle, a nuclear signaling pathway that is activated by IGF-I and involves the hydrolysis of PI lipids in a manner that is analogous to, but quite distinct from, that at the plasma membrane (9, 10). In a previous investigation from our laboratory, we demonstrated that differentiation of C2C12 mouse myoblasts in response to insulin stimulation is characterized by a marked increase in nuclear PLCß1. Moreover, we have found that an imbalance of nuclear and cytoplasmic PLCß1 down-regulates myogenesis as evidenced by the overexpression of a cytoplasmic PLCß1 mutant that, because of the lack of a nuclear localization sequence, acts in a dominant negative fashion and suppresses the differentiation of C2C12 myoblasts (11). A more recent study reveals that the C2C12 muscle cell line contains at least three PLCß isoforms: PLCß1, PLCß3, and PLCß4 (12). Studies on signaling through the insulin/IGF-I receptors in muscle differentiation have revealed that PLC{gamma}1 is also involved during this process and that PLC{gamma}1 mRNA and protein levels were both increased during myogenesis (13, 14). Our previous observation showed that a downstream target of nuclear PLCß1 signaling is the cyclin D3/cdk4 complex (15). Even though the expression of most cyclins is down-regulated during cell cycle arrest, a notable exception is cyclin D3. In fact, previous work has shown that terminal differentiation of muscle cells is accompanied by transcriptional induction of cyclin D3 (16). It is also interesting to note that the cyclin D3 5' noncoding region contains several E-boxes. These DNA sequences display binding sites for myogenic transcription factors that activate muscle-specific structural genes. Myogenic factors binding to E-boxes, present in the cyclin D3 gene (17) could be the mechanism by which cyclin D3 is up-regulated during muscle differentiation. It has been proposed that myogenic factors regulate not only tissue-specific gene expression but also the exit from the cell cycle. For example, at the onset of differentiation, MyoD activates cyclin D3, which then sequesters unphosphorylated retinoblastoma protein, leading to the irreversible withdrawal of differentiating myoblasts from the cell cycle (18). Here we show that in C2C12, PLCß1 and PLC{gamma}1 reside in distinctive subcellular compartments, PLCß1 being predominantly in the nucleus and PLC{gamma}1 in the cytoplasm. Both PLCß1 and PLC{gamma}1 activate cyclin D3 promoter during the differentiation of myoblasts to myotubes, indicating that both PLCs are crucial regulators of the mouse cyclin D3 gene and that PLC signaling induced by insulin activates at least two distinct lipid-dependent signaling pathways.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell culture and differentiation induction
The mouse C2C12 myoblasts were grown in DMEM supplemented with 10% fetal bovine serum [growth medium (GM)] at 37 C and 5% CO2. For differentiation studies, cells at 80% of confluence (preconfluent cells) were cultured for the time indicated in the figures in a serum-free medium supplemented with 100 nM insulin [differentiation medium (DM)]. Cells were cultured for 48 h in a serum-free medium in the absence or presence of LY294002 (10 µM) and U-73122 (5 µM). Insulin signaling pathways were determined after pretreatment with the inhibitors for 20 min. Insulin and BSA were from Sigma Chemical Co. (St. Louis, MO). LY294002 and U-73122 were purchased from Calbiochem-Novabiochem International (La Jolla, CA).

Preparation of whole-cell extract
Whole-cell lysates were prepared by lysing cells in RIPA buffer [50 mM Tris (pH 7.5), 1% Nonidet P-40, 0.1% SDS, 100 mM NaCl, 50 mM NaF, 1 mM EDTA] supplemented with 0.5 mM phenylmethylsulfonyl fluoride, 10 mM ß-mercaptoethanol, and 1 mM EGTA, and a set of protease inhibitors (10 µg/ml leupeptin, 0.3 mM aprotinin, 15 µg/ml calpain I inhibitor, and 7.5 µg/ml calpain II inhibitor) was also added to the buffer.

Preparation of nuclei
Nuclei were purified as previously described (9). Briefly, 5 x 106 cells were lysed in 400 µl nuclear isolation buffer [10 mM Tris-HCl (pH 7.8), 1% Nonidet P-40, 10 mM ß-mercaptoethanol, 0.5 mM phenylmethylsulfonyl fluoride, 1 µg aprotinin and leupeptin/ml, 10 µg soybean trypsin inhibitor/ml, 15 µg calpain inhibitor I and II/ml, and 5 mM NaF] for 3 min on ice. MilliQ water (400 µl) was then added to swell cells for 3 min. The cells were sheared by eight passages through a 23-gauge hypodermic needle. Nuclei were recovered by centrifugation at 400 x g and 4 C for 6 min and washed once in 400 µl washing buffer [10 mM Tris-HCl (pH 7.4) and 2 mM MgCl2, plus protease inhibitors as describe above]. The purity of the isolated nuclei was analyzed by detection of ß-tubulin.

Construction of expression vectors and transient transfection
The full-length cDNA for rat PLCß1b was cloned into pRc/CMV (Invitrogen, Valencia, CA) expression vector plasmid as described elsewhere (19). The full-length cDNA for rat PLC{gamma}1 was kindly provided by P. G. Suh (Phoang University of Science and Technology, Korea). Transient transfection of these vectors into C2C12 cells was performed using an Amaxa Nucleofector apparatus (Amaxa, Koln, Germany), according to the manufacturer’s instructions. Briefly, 1 x 107 cells were resuspended in 100 µl of the specified electroporation buffer plus 10 µg of the plasmid. Two days later, cells were lysed and the amount of protein present in cell extracts was determined using the Bradford protein assay (Bio-Rad, Richmond, CA).

Stable transfection and reporter gene assay
pD3GH, the human GH (hGH) reporter gene plasmid containing 1680 bp of mouse cyclin D3 promoter sequences (pD3GH) and the deletion constructs pD3–957 (with +1 being the transcription start site), pD3–446, pD3–190, and pD3–37 were as described (20).

C2C12 cells were stably transfected by electroporation as described above (Amaxa). Each of the hGH gene reporter plasmids (10 µg) along with CMVß-galactosidase expression plasmid used for normalization of efficiency of transfection (pCMVß-gal; 10 µg) and the pcDNA3 plasmid (10 µg) containing a neomycin-resistant gene (Invitrogen) were added to 1 x 107 cells in 100 µl electroporation buffer. At 48 h after electroporation, the cells were cultured for 3 wk in medium containing 1 mg/ml G418 (GIBCO BRL, Carlsbad, CA), and the single resistant cell clones were selected and expanded separately. The hGH expression assay was performed in medium of cells cultured in the presence or absence of insulin. One tenth of cell lysates was used to determine ß-galactosidase activity as described (21) to normalize efficiency of transfection of different plasmids containing the cyclin D3 gene fragment. The level of hGH produced by cells was determined by using an hGH assay kit (hGH ELISA; Roche, Indianapolis, IN).

Immunochemical analysis
Fifty micrograms of proteins from the whole-cell extracts were separated on 6 or 10% polyacrylamide-0.1% SDS gels, as specified in the figure legends. Proteins were transferred to nitrocellulose membranes for subsequent immunodetection with the specific antibodies and detected by using the enhanced chemiluminescence method (Amersham Biosciences, Arlington Heights, IL) and visualized in a Kodak digital image station 2000R. The expression of specific proteins was analyzed by using the following antibodies: anti-ß-actin mouse monoclonal from Sigma; a monoclonal antibody specific for myogenin (sc-12732), polyclonal rabbit antibody specific for cyclin D3 (sc-182), PLCß1 (sc-9050), and histone H3 (sc-8653) from Santa Cruz Biotechnology (Santa Cruz, CA); monoclonal antibody specific for PLC{gamma}1 from Upstate Biotechnology (Lake Placid, NY); and a monoclonal antibody for ß-tubulin from Sigma. Analysis of the blots was performed by using a Kodak image station 2000R.

Construction of antisense RNA of the PLCß1 and PLC{gamma}1 gene
Knockdown of PLCß1 and PLC{gamma}1 gene expression in C2C12 cells was achieved by the small interfering RNA (siRNA) gene-silencing technique. For transient inhibition of PLCß1 and PLC{gamma}1 mRNA production, C2C12 cells were transfected with double-stranded 21-nucleotide-long siRNA targeting, respectively, PLCß1 and PLC{gamma}1. For efficient siRNA production, the Dicer siRNA Generation Kit from GTS (Gene Therapy System, Inc., San Diego, CA) was used. A mixture of siRNAs was obtained from a template of 1000 bp in length. The PLCß1 region selected for gene silencing is located between the sense sequence 5'-AGGATGCCAGGTGTGGGAAG-3' and the antisense sequence 5'-TCCAGCTCCACACAGCGACA-3'. The PLC{gamma}1 region selected for gene silencing is located between the sense sequence 5'-AAGTTGGAGACGCGCCAGAT-3' and the antisense sequence 5'-ATGGTAAATGACTGGCATCCC-3'. C2C12 cells were also transfected with siRNA for the green fluorescent protein as a negative control (supplied with Dicer siRNA Generation Kit from GTS). Transfection of the siRNA was carried out by electroporation using the Amaxa Nucleofector apparatus according to the manufacturer’s instructions.

Immunofluorescence microscopy
Myoblasts and myotubes grown on glass coverslips were fixed in 4% paraformaldehyde at 4 C and permeabilized with 0.15% Triton X-100 in PBS. All preparations were treated with PBS containing 4% BSA to saturate nonspecific binding. Incubation with monoclonal anti-PLCß1 and PLC{gamma}1 (Upstate Biotechnology) (1:100) was performed overnight at 4 C, whereas FITC-conjugated antimouse IgG antibody (1:200) was applied for 1 h at room temperature. Incubation with antimyogenin monoclonal antibody sc-12732 (Santa Cruz Biotechnology) (1:10) was carried out as for anti-PLCß1 antibody. Slides were washed and mounted with an antifade reagent in glycerol and observed with a Nikon E600 fluorescence microscope equipped with a digital camera.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Expression pattern of PLCß1, PLC{gamma}1, cyclin D3, and myogenin during myogenic differentiation of C2C12 cells
Our main goal in this study was to investigate the role of PLCß1 and PLC{gamma}1 in skeletal muscle differentiation. Therefore, we performed a simultaneous analysis of PLCß1 and PLC{gamma}1 expression in proliferative and differentiative conditions (Fig. 1Go). For this analysis, we used the C2C12 murine myoblast cell line. The expression of PLCß1 (Fig. 1AGo), which is quite low in GM cells, was induced during the onset of differentiation, increased at early time points after switching the cells to DM (24 h), and reached higher levels at later stages of differentiation (96 h of DM). The expression of PLC{gamma}1 (Fig. 1Go B) was already detectable in GM and reached higher levels at 72 h of DM. The presence of the myogenic regulatory factor myogenin was also determined in the same cell lysates (Fig. 1DGo). Myogenin, which promotes terminal differentiation, became detectable on the first day of differentiation after cells reached confluence. These observations verified that our model exhibited the biological characteristics of muscle differentiation in a chronologically appropriate manner.


Figure 1
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  		FIG. 1. Level of PLCß1, PLC{gamma}1, cyclin D3, myogenin, and ß-actin in C2C12 cells during muscle differentiation. Cultures of C2C12 cells were exposed to GM and to DM (medium with 100 nM insulin) for increasing periods of time. Whole-cell lysates from the same experiment were resolved by SDS-PAGE and then analyzed with specific antibodies. One of the membranes was reprobed with the ß-actin antibody to normalize the amount of loaded proteins.

 
It is known that cyclin D3 critically contributes to the irreversible exit of differentiating myoblasts from the cell cycle (18) and that the overexpression of PLCß1 in the nucleus of Friend erythroleukemia cells increases the expression of cyclin D3 (15). Here we show (Fig. 1CGo) that terminal differentiation of C2C12 cells was associated with an increase of cyclin D3 levels, which follows temporally the increase of both PLCß1 and PLC{gamma}1.

Immunolocalization of PLCß1 and PLC{gamma}1 during differentiation of C2C12 cells
To find out whether PLCß1 and PLC{gamma}1 could play a role in the control of cyclin D3 expression, first we analyzed the subcellular distribution of the two enzymes (Fig. 2Go). The presence and the subcellular localization of PLCß1 and PLC{gamma}1 was determined by immunofluorescence staining and by Western blot analysis in myoblasts and in differentiated multinucleated C2C12 myotubes. As shown in Fig. 2Go, PLCß1 is localized to the cell nucleus in mononucleated myoblasts as well as in multinucleated myotubes with a low level of cytoplasmic staining. In contrast, PLC{gamma}1 appears to be excluded from the nucleus. The presence and the subcellular localization of PLCß1 and PLC{gamma}1 were also confirmed by Western blot analysis (Fig. 2BGo). Cytoplasmic and nuclear fractions did not show, respectively, histone H3 and ß-tubulin, confirming the absence of cross-contamination of the two subcellular preparations.


Figure 2
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  		FIG. 2. Immunolocalization of PLCß1 and PLC{gamma}1 during differentiation of C2C12 cells. The subcellular localization of the two enzymes was examined by immunofluorescence in C2C12 cells induced to differentiate by switching the GM to DM (i.e. medium with 100 nM insulin) for 48 h. The cells were immunostained with anti-PLCß1 and anti-PLC{gamma}1 antibody (Upstate Biotechnology) followed by fluorescein-isothiocyanate-conjugated secondary antibody. Nuclear lysates from the same experiments were separated by SDS-PAGE and analyzed by immunoblot with the same antibodies. The membrane was reprobed with antihistone H3 antibody. Cytoplasmic lysates were tested for the presence of PLCß1 or PLC{gamma}1, and the membrane was reprobed with anti-ß-tubulin antibody.

 
Effect of the knockdown of PLCß1 and PLC{gamma}1 on the efficiency of myogenin and cyclin D3 expression in differentiating C2C12 cells
To investigate the involvement of PLCß1 and PLC{gamma}1 in the regulation of myogenic differentiation, the effect of loss of function of these enzymes was examined using siRNA specific to PLCß1 and PLC{gamma}1, respectively (Fig. 3Go). To knock down PLCß1 or PLC{gamma}1 protein expression, siRNA-PLCß1 or siRNA-PLC{gamma}1 oligos were transfected into C2C12 cells. When the expression of the two enzymes was examined by Western blot analysis, there was significant loss of protein expression in cells transfected with siRNA constructs compared with control cells (Fig. 3Go, A and B). The transfection of C2C12 cells with specific siRNA for PLCß1 (Fig. 3AGo) as well as for PLC{gamma}1 (Fig. 3BGo) resulted in a decrease in myogenin and cyclin D3 expression. In C2C12 cells, fusion into myotubes is an important methodological parameter of myogenic differentiation. The data reported in Fig. 3CGo indicated that C2C12 cell fusion into myotubes is significantly inhibited by the knockdown of both PLCß1 and PLC{gamma}1 expression. These results indicated that both PLCß1 and PLC{gamma}1 are necessary for efficient muscle differentiation.


Figure 3
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  		FIG. 3. The siRNA-mediated knockdown of PLCß1 and PLC{gamma}1 gene leads to reduced expression of cyclin D3 and myogenin. C2C12 cells were transfected with PLCß1, PLC{gamma}1-specific siRNA, and green fluorescent protein (si-gfp) as negative control. After 48 h of insulin stimulation, whole homogenates were separated by SDS-PAGE and analyzed by immunoblot. A, PLCß1, myogenin and cyclin D3 protein levels of wild-type and siRNA (PLCß1) transfected C2C12 cells; B, PLC{gamma}1, myogenin, and cyclin D3 protein levels of wild-type and siRNA (PLC{gamma}1) transfected C2C12 cells; C, light microscopy of C2C12 myoblasts (GM), myotubes (DM), and C2C12 cells cultured in DM and transfected with siRNA of PLCß1 and PLC{gamma}1. All transfection analyses were done in duplicate, and the results shown are representative of three independent experiments.

 
Activation of the cyclin D3 promoter by PLCß1 and PLC{gamma}1
To determine whether the activation of cyclin D3 expression by PLCß1 and PLC{gamma}1 was mediated via its promoter, C2C12 cells were stably transfected with a construct encoding the mouse cyclin D3 promoter sequence ligated to a GH reporter gene along with an expression plasmid. When pD3GH, which includes the 5' noncoding sequence from +85 to –1680, was stably transfected into C2C12 cells, a 4-fold induction of hGH activity was observed upon insulin stimulation. The stable pD3GH clone of C2C12 cells gave on its own a similar profile to the one found with transient transfection (17). This indicated the presence of an insulin ligand-inducible element within the cyclin D3 promoter. Notably, Fig. 4AGo shows that transient overexpression of PLCß1 or PLC{gamma}1 also increased cyclin D3 promoter activity without stimulation by insulin, consistent with increased levels of the expression of both enzymes detected in transient transfectants (Fig. 4BGo). To further assess the role of the PLC pathway in regulation of the cyclin D3 promoter, we tested cyclin D3 promoter activity in C2C12 cells in which the expression of PLCß1 or PLC{gamma}1 was knocked down by siRNA (Fig. 4BGo), and as shown in Fig. 4AGo, silencing of PLCß1 or PLC{gamma}1 gave rise to a reduction of activation of the cyclin D3 promoter even in the presence of insulin. The lack of activation was not complete because one of the two PLCs was still expressed. To verify the importance of the PLCß1 nuclear localization in activating the cyclin D3 promoter during differentiation of C2C12 cells, we simultaneously performed the same experiment by overexpressing the PLCß1-M2b mutant. Previous reports have shown that the M2b mutant induces a forced expression of PLCß1 in the cytoplasmic compartment and constitutes a good tool to discriminate the different role of PLCß1 as a function of its subcellular localization (11). As we expected, the cyclin D3 promoter activity was absent in cells in which we overexpressed the PLCß1-M2b mutant. Taken together, these experiments suggest a PLC-dependent activation of cyclin D3 promoter given that the nuclear PLCß1 and cytoplasmic PLC{gamma}1 expression is required for the induction of cyclin D3 promoter activity.


Figure 4
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  		FIG. 4. Cyclin D3 promoter activity is regulated by PLCß1 and PLC{gamma}1 expression. C2C12 cells stably transfected with pD3GH were transiently transfected with PLCß1 (ov-PLCß1), M2b-PLCß1 (ov-PLCß1 M2ß) mutant, and PLC{gamma}1 (ov-PLC{gamma}1) cDNA for the overexpression or with PLCß1 (si-PLCß1) or PLC{gamma}1 (si-PLC{gamma}1) siRNA for the knockdown. A, Cyclin D3 promoter activity; B, protein extracts from a representative experiment were analyzed by Western blotting to monitor the transient overexpression or the knock-down of PLCß1 and PLC{gamma}1 expression. ov, C2C12 transiently overexpressed; M2b, C2C12 transiently overexpressed with PLCß-M2b mutant; si-wt, C2C12 transiently silenced; wt, C2C12 wild type. To measure the level of expression of hGH in myotubes, cells were maintained in differentiation medium (100 nM insulin) after transfection for 2 d. The Western blot of transient transfectants reported here are representative of four other identical experiments. The data are the means of four determinations, with SD indicated by the error bars. *, Statistical significance (P < 0.001 compared with each non-insulin-stimulated control GM).

 
Induction of the cyclin D3 promoter by PLCß1 and PLC{gamma}1 occurs with identical kinetics
To assess whether PLCß1 and PLC{gamma}1 signaling pathways enhance the transcription of the cyclin D3 gene at different time points, we explored the inducibility of the cyclin D3 promoter at earlier time points. We assessed the cyclin D3 promoter activity at 6 and 12 h after insulin stimulation and after overexpression of PLCß1 or PLC{gamma}1. As shown in Fig. 5AGo, we demonstrated that the kinetics of the cyclin D3 promoter activity was similarly increased in C2C12 overexpressing PLCß1 or PLC{gamma}1, indicating no differences in the time point of action between the two signaling PLC.


Figure 5
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  		FIG. 5. PLCß1 and PLC{gamma}1 overexpression activates cyclin D3 promoter at the same time points. The cells were treated as described in Fig. 4Go. Cyclin D3 promoter activity was measured at different time points as indicated in the figure. The Western blot of transient transfectants reported here are representative of three other identical experiments. The data are the means of four determinations, with SD indicated by the error bars. *, Statistical significance (P < 0.001 compared with each non-insulin-stimulated control C).

 
PLCß1 and PLC{gamma}1 act on different transcriptional regulatory region of cyclin D3 promoter
To further the understanding of which cyclin D3 promoter regulatory sequences are responsive to PLCß1 or PLC{gamma}1, we used a series of 5'-deletions of the promoter linked to the hGH reporter gene (Fig. 6Go). Promoter deletion constructs were stably transfected into C2C12 cells, and the relative activities of these constructs in differentiating C2C12 myoblasts were measured after insulin stimulation and after overexpression of PLCß1 or PLC{gamma}1. Figure 7AGo reveals that insulin treatment resulted in the activation of all the cyclin D3 promoter constructs. It is further seen that overexpression of PLCß1 in the absence of insulin resulted in the activation of the cyclin D3 promoter and that this capacity was lost on deletion of sequences from –446 to –190 of the promoter, indicating that a responsive element is located in this region. The overexpression of PLC{gamma}1 in the absence of insulin activates the cyclin D3 promoter as well, and this capacity was lost on deletion of sequences from –1680 to –957. This capacity was reacquired, however, when the sequence –957 to –446 was deleted, suggesting the presence of a negative regulatory element in the cyclin D3 promoter, from –957 to –446, which controls promoter activity induced by PLC{gamma}1 overexpression. The nature of this element will require further exploration in the future. To demonstrate furthermore the effect of PLCß1 and PLC{gamma}1 on different regions of the cyclin D3 promoter, C2C12 cells stably transfected with deleted constructs of the cyclin D3 promoter were transiently transfected with PLCß1 or PLC{gamma}1 siRNA. PLCß1 or PLC{gamma}1 siRNA down-regulated the activity of the sequence deleted constructs of cyclin D3 promoter targeted by PLCß1 and PLC{gamma}1 overexpression, respectively (Fig. 7BGo). These data indicate that transcriptional activity of the cyclin D3 promoter can be induced by both PLCß1 and PLC{gamma}1 expression and that positive regions of regulatory DNA associated with their actions can be identified.


Figure 6
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  		FIG. 6. Schematic diagram representing the full cyclin D3 promoter with putative binding sites and each deleting sequence linked to the hGH reporter gene. Full promoter and each promoter deletion construct were transfected into C2C12 cells to obtain stable transfectants.

 

Figure 7
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  		FIG. 7. PLCß1 and PLC{gamma}1 act on different transcriptional regulatory regions of cyclin D3 promoter. A series of 5'-deletions of the cyclin D3 promoter linked to the hGH reporter gene was stably transfected into C2C12 cells, which were subsequently transiently transfected with PLCß1 or PLC{gamma}1 cDNA for the overexpression (A) and with PLCß1 or PLC{gamma}1 siRNA (B) for the knockdown. The constructs are designated by numbers corresponding to the length of cyclin D3 sequences upstream of the transcriptional start site. Protein extracts (30 µg) from a representative experiment were analyzed by Western blotting to monitor the transient overexpression or the knockdown of PLCß1 and PLC{gamma}1. c, C2C12; insulin, C2C12 treated with insulin; ov PLCß1-ins, C2C12 overexpressing PLCß1 without insulin stimulation; ov PLC{gamma}1-ins, C2C12 overexpressing PLC{gamma}1 without insulin stimulation; si PLCß1+ ins, C2C12 knocked down for PLCß1 expression treated with insulin; si PLC{gamma}1+ ins, C2C12 knocked down for PLC{gamma}1 expression treated with insulin. The Western blot of transient transfectants reported here are representative of four other identical experiments. The data are the means of four determinations, with SD indicated by the error bars. *, Statistical significance (P < 0.001 compared with each non-insulin-stimulated control C).

 
Effect of PI 3-kinase and PLC inhibitors on insulin stimulation of cyclin D3 promoter activity
To extend these results further, C2C12 myoblasts were incubated in DM in the absence or presence of the PI 3-kinase inhibitor LY294002 to inhibit signaling of the PI 3-kinase cascade. PLC activity was inhibited with the PLC inhibitor U-73122 (not shown). To investigate the signaling pathways involved in insulin-induced C2C12 myoblast differentiation to myotubes, the inhibitors LY294002 or U-73122 were added concomitantly with insulin in serum-free medium, and 72 h later, cells were analyzed for PLCß1, PLC{gamma}1, cyclin D3, and myogenin expression. Cells, when cultured in the presence of LY294002 or U-73122, upon insulin stimulation showed a low level of cyclin D3, myogenin, and PLC{gamma}1 expression comparable to that of undifferentiated C2C12 cells, whereas treatment with LY294002 did not appreciably affect PLCß1 expression. U-73122 is able to inhibit the induction of PLC expression after insulin stimulation (Fig. 8AGo). Figure 8BGo highlights the morphological phenotypes of C2C12 cells treated with LY294002 or U-73122 compounds. Both the inhibitors are capable of impairing the formation of myotubes, and the cell shape remains that of undifferentiated myoblasts. These results suggest that PLCß1 and PLC{gamma}1 regulate the expression of cyclin D3 by two distinct signaling pathways that flow into a unique nuclear target. To explore the inducibility of the cyclin D3 promoter by insulin and to delineate its important regulatory sequences, differentiated C2C12 cells stably transfected with the cyclin D3 deletion constructs were cultured in presence of the PLC inhibitor U-73122 and with the PI 3-kinase inhibitor LY294002 (Fig. 8CGo). In our experiment, pretreatment of C2C12 cells with 10 µM LY294002 greatly impaired insulin-induced cyclin D3 promoter activity in the sequences from nucleotide –190 to the transcription initiation site, which correspond to the same responsive regulatory sequences activated by PLC{gamma}1 overexpression. On the other hand, pretreatment of C2C12 cells with the PLC inhibitor U-73122 inhibited the activity of the full-length as well as the deleted constructs of cyclin D3 promoter, indicating that PLC signaling is necessary in insulin-dependent cyclin D3 transcription.


Figure 8
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  		FIG. 8. Effect of inhibitors on insulin stimulation of cyclin D3 promoter activity. A, Effect of inhibitors on PLCß1 and PLC{gamma}1, myogenin, and cyclin D3 expression after insulin stimulation. Whole-cell lysates were prepared and 50 µg of proteins were electrophoresed on SDS-polyacrylamide gels and immunoblotted with anti-PLCß1 and PLC{gamma}1, antimyogenin, and anti-cyclin D3 antibodies. B, Light microscopy of C2C12 myoblasts (GM), myotubes (DM), and C2C12 cells in the presence of 10 µM LY294002 (LY) or 10 µM U-73122 (U-73) for 48 h in DM. C, C2C12 cells stably expressing cycling D3 promoter deletion constructs, as described in Fig. 6Go, were assayed for hGH production. C2C12 overexpressing cells were cultured to 80% confluence in GM and then switched to DM for 48 h in the absence or presence of 10 µM LY294002 (LY) or 10 µM U-73122 (U-73). The data are the means of four determinations, with SD indicated by the error bars. *, Statistical significance (P < 0.001 compared with each non-insulin-stimulated control C).

 

   Discussion
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
PLCß1 and PLC{gamma}1 expression during C2C12 differentiation
In the present study, we have presented evidence that PLC signaling is required for the activation of cyclin D3 promoter in C2C12 cells. Differentiation of C2C12 cells leads to the up-regulation of cyclin D3 (18), which is also a target of nuclear PLCß1 signaling (15). We have previously demonstrated that differentiation of C2C12 myoblasts in response to insulin stimulation is characterized by a marked increase of nuclear PLCß1 expression (11). It has also been demonstrated that PLC{gamma}1 mRNA and protein levels were increased during myogenesis (13). In the current study, we attempted to find out the nuclear targets through which PLCß1 and PLC{gamma}1 regulate the skeletal muscle differentiation program. Because it has been demonstrated that overexpression of PLCß1 in the nucleus increases the expression of cyclin D3 (15), we wished to demonstrate a correlation between PLCß1 and cyclin D3 during myoblast differentiation and to determine whether this nuclear signaling could involve the one elicited by PLC{gamma}1. Expression of both PLCß1 and PLC{gamma}1 is low in undifferentiated cells and then increases as myoblasts elongate and fuse. Immunoblotting data and immunofluorescence analysis showed PLCß1 predominantly located in the nucleus and PLC{gamma}1 exclusively represented in the cytoplasmic fraction. The diverse location of the two enzymes hints at the likelihood that up-regulation of cyclin D3 during differentiation could be regulated by both enzymes but through different signaling pathways. Moreover siRNA-mediated knockdown of PLCß1 and PLC{gamma}1 genes showed that both PLCs were indeed required for the regulation of cyclin D3 and myogenin expression. It is worthwhile mentioning that siRNA for unrelated gene does not affect either PLCß1 or PLC{gamma}1 expression (Fig. 3Go, A and B) as well as that the knockdown or overexpression of one of the two PLCs does not affect the other one (not shown), confirming previously published data (19).

PLCß1 and PLC{gamma}1 expression affects cyclin D3 promoter
Our data show that cyclin D3 promoter activation is under the control of molecular events that depend not only on PLCß1 and PLC{gamma}1 expression levels but also on the nuclear localization of PLCß1, because the overexpression of the PLCß1 mutant (PLCß1-M2b), which localizes only in the cytoplasm, inhibits the promoter activity and differentiation as well. It should be noted that after knockdown of PLCß1 or PLC{gamma}1 expression, insulin signaling can still operate through any residual, unsilenced PLC mRNA and subsequently activate the cyclin D3 promoter. Indeed, complete inhibition of PLC activity, by means of the well known PLC inhibitor U-73122, abolishes the insulin-dependent differentiation (Fig. 8BGo). Given that the knockdown of either PLCß1 or -{gamma}1 is capable of inhibiting the differentiation of C2C12 myoblasts and the expression of cyclin D3 (Fig. 3Go), there is the likelihood that the transcription of cyclin D3 is dependent on the activation of each single region of the promoter, targeted by either PLCß1 or -{gamma}1. Therefore, only the activation of the full-length promoter is capable of inducing cyclin D3 transcription and in turn the differentiation of C2C12 myoblasts. The response to PLCß1 was dependent on the region from –446 to –190, which contains putative binding sites for E2-F, Ap1, STAT, and ATF/CREB (Fig. 6Go). Other studies have shown that the level of PLC expression changes during development (22) and that nuclear PLCß1 is involved in the activation of transcription factors like E2F and NF-E2 (15, 23). Modulation of PLC{gamma}1 expression targets and activates the cyclin D3 promoter in the region from –1680 to –957, which contains putative binding site for the transcription factor GATA and contains an E-box sequence (Fig. 6Go). This DNA sequence termed E-box displays binding sites to myogenic transcription factors that activate muscle-specific structural genes (24). Myogenic factors binding to E-boxes, present in the cyclin D3 gene (Fig. 6Go), could be the mechanism by which cyclin D3 is up-regulated during muscle differentiation (17). Moreover, also the regions –446 to –190 and –37 to +1 are activated by PLC{gamma}1. These regions contain putative binding sites for NF-{kappa}B, Ap2, Sp1, and NF-Y (Fig. 6Go). It is known that insulin produces a dramatic induction of NF-{kappa}B activity in myoblasts cultured for 72 h, correlating with the formation of myotubes in a PI 3-kinase-dependent manner (25).

The effect of PLC and PI 3-kinase inhibitors on cyclin D3 promoter
We found that inhibitors of PI 3-kinase reduced cyclin D3 promoter activity of the region targeted by PLC{gamma}1. Many studies have demonstrated that PI 3-kinase inhibitors can attenuate differentiation of different muscle cell lines. Wortmannin and LY294002 have been shown previously to curtail the normal induction of expression of muscle-specific mRNAs and proteins, including myogenin, and to reduce formation of multinucleated myotubes (26). Indeed, the inhibition of C2C12 differentiation is also linked to the loss of the induction of PLC{gamma}1 expression upon PI 3-kinase inhibition (Fig. 8AGo). Concerning the PLC inhibitor U-73122, it is worthwhile mentioning that it inhibits C2C12 differentiation as well and also gives rise to the loss of the induction of the expression of PLCß1 and PLC{gamma}1 when cells are treated with insulin. These findings suggest that PLC{gamma}1 signaling is downstream PI 3-kinase activation, whereas nuclear PLCß1 signaling is not regulated by PI 3-kinase. In rat skeletal muscle, a signaling pathway has been proposed by which gliclazide could stimulate insulin receptor substrate-1 that would allow its association with PI 3-kinase, promoting its activation. PI 3-kinase products could induce PLC activation whose hydrolytic activity could activate the DAG-dependent isoform protein kinase C (PKC)-{alpha}, -{theta}, and -{epsilon} (27). It has been demonstrated that in H9c2 cardiac myoblast IGF-I regulation of muscle differentiation is dependent on the activation of PLC{gamma}1 and Akt/PKB, both of which are downstream mediators of PI 3-kinase (13). Therefore, both IGF-I and insulin have a significant signaling activity. Insulin and IGF-I signaling involves the rapid phosphorylation of the receptor on tyrosine residues and the activation of PI 3-kinase (25). In skeletal muscle, inhibition of PI 3-kinase by pharmacological blockade using LY294002 or Wortmannin has shown that PI 3-kinase is an essential molecule for insulin-stimulated GLUT4 translocation and glucose transport. In this study, we show that PLC{gamma}1 plays an important role in muscle differentiation of C2C12 cells and that it is activated for the proper muscle differentiation. Moreover, recent evidence has suggested that PLC is involved in glucose transport in skeletal muscle (28, 29). Because PLC activation results in the production of DAG, it is plausible that DAG-sensitive members of the PKC family may be involved in insulin-stimulated glucose transport through a similar pathway. Another study demonstrates that {alpha}1-adrenoceptors mediate increases in glucose uptake in L6 muscle cells. This effect appears to be related to activation of PLC, PI 3-kinase, p38 kinase, and atypical PKC (30). It seems rather clear that PLC{gamma}1 action on cyclin D3 promoter activation is dependent on the PI 3-kinase signaling pathway taking place in the cytoplasm. Because PLC{gamma}1 does not enter the nucleus, it is reasonable that its action is mediated by the cytoskeletal filaments, which could affect both structure and function of the nuclear matrix (14). By means of a PLC inhibitor, the insulin-induced differentiation is abolished, indicating that PLC activity is necessary for this purpose.

PLCß1 location and C2C12 differentiation
The location of PLCß1 appears to be a key issue in the differentiation process. Indeed, the forced expression of the cytosolic mutant M2b, which has a dominant negative effect on endogenous PLCß1, is capable of abolishing myogenic differentiation. This accounts for the fact that the effect of PLCß1 and -{gamma}1 is dependent on their physiological location, i.e. in the nucleus and at the plasma membrane, respectively. The relationship between function and location seems to be supported by data showing the importance of the nuclear organization to achieve muscle differentiation. Nuclear PLCß1 could be implicated in lamina reorganization by acting on cyclin D3 function. Indeed, it has been demonstrated that changes in internal A-type lamins (A and C) organization in muscle cells are brought about by cyclin D3 with the involvement of pRb (31).

An inositol polyphosphate multikinase, unaffected by Wortmannin, has been shown to localize in the nucleus and to regulate transcription (32). The generation of IP3 by means of nuclear PLCß1 activity could allow IP3 sequential phosphorylation driven by inositol polyphosphate multikinase and could lead to molecular signals that could affect cyclin D3 promoter in nuclei of differentiating myoblasts. In summary, we have shown that both PLCß1 and -{gamma}1 are required for C2C12 differentiation as judged by both molecular and morphological analysis. Both PLCs are capable of inducing, once overexpressed in their physiological compartment, the differentiation of the C2C12 myoblast by targeting specific regions of the cyclin D3 promoter even in the absence of insulin, whereas the knockdown of PLCß1 and -{gamma}1 even in the presence of insulin blocks the differentiative program. This suggests that their expression is by its own sufficient for the increase of cyclin D3 during C2C12 myoblast differentiation.


   Footnotes
 
This work was supported by Italian Ministero dell’Università e della Ricerca-Fondo Investimenti Ricerca di Base (MIUR-FIRB), Ministero dell’Università e della Ricerca-Cofinanciamiento (MIUR-COFIN), Associazione Italiana Ricerca sul Cancro (AIRC), and the CARISBO Foundation.

Disclosure Statement: The authors have nothing to disclose.

First Published Online November 22, 2006

Abbreviations: DAG, Diacylglycerol; DM, differentiation medium; GM, growth medium; hGH, human GH; IP3, inositol-1,4,5-trisphosphate; MRF, muscle regulatory factor; PI, phosphatidylinositol; PKC, protein kinase C; PLC, phospholipase C; siRNA, small interfering RNA.

Received July 26, 2006.

Accepted for publication November 13, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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