Loss of MEF2D expression inhibits differentiation and contributes to oncogenesis in rhabdomyosarcoma cells

To understand the deregulation of myogenesis in RMS cells, we first determined the level of myogenin, MyoD and associated co-factors in RMS cells in comparison to the normal expression levels present during skeletal muscle differentiation (Figure 1A). Four independently derived RMS cell lines were used for this analysis. The ERMS subtype was represented by RD and RD2 cells and the ARMS subtype was represented by RH30 and RH28 cells. Murine C2C12 cells, a commonly used myogenic cell line, were used as a comparative cell line for RMS cells. Myogenin was not detectable in proliferating myoblasts, but was strongly induced upon differentiation. MyoD was expressed in proliferating myoblasts and maintained expression during differentiation. We found that myogenin was expressed in all assayed RMS cell lines (Figure 1A). The levels of myogenin in most RMS lines were higher than the level observed in normal differentiating myoblasts. The level of myogenin observed in RD2 cells was not as robust as was observed in the other RMS lines, but the level was still similar or modestly higher than that observed in normal differentiating myoblasts. We also assayed for MyoD expression and found that the expression of MyoD was similar to the expression of MyoD observed in myoblasts (Figure 1A). The cell lines of the ARMS subtype, RH30 and RH28, expressed MyoD at levels comparable or slightly higher to that observed in normal myoblasts. While expressed at a lower level than that found in ARMS cells, MyoD expression was also detected in both cell lines of the ERMS subtype, RD and RD2.

Next, we assayed the expression profile of the co-factors required by myogenin in C2C12 and RMS cells. We looked for the E proteins by assaying for both the E2A variants and HEB. The E2A locus encodes the two slice variants, E12 and E47, which differ by differential use of a single exon [33]. E12/47 and HEB are known to be expressed in proliferating and differentiating myoblasts. We found that the RMS cell lines showed apparently normal levels of expression of HEB (Figure 1A). RD and RH30 cell lines were used to confirm expression of E12/47 and we again observed high levels of the E proteins (Additional file 1: Figure S1).

We next examined the expression of the MEF2 family in C2C12 cells and RMS cells and found that while MEF2A, MEF2B and MEF2C were expressed (Additional file 1: Figure S2), MEF2D was dramatically down regulated in RMS cells when compared to the levels found in C2C12 cells (Figure 1B). The down regulation of MEF2D was also observed in primary cells derived from a mouse model of ERMS, JW41 (Figure 1B). The expression of MEF2D at the protein level was determined from extracts from proliferating cells and cells that were induced to differentiate for two days. MEF2D was robustly expressed in C2C12 cells, but was greatly reduced in all RMS cell lines tested (Figure 1C). HEK293 cells expressing exogenous MEF2D were used to confirm specificity of the antibody. Extracts from HEK293 cells expressing MEF2D were not recognized by antibodies against MEF2C and extracts from HEK293 cells expressing MEF2C were not recognized by antibodies against MEF2D (Additional file 1: Figure S3).

To confirm that muscle specific genes were down regulated in RMS cells, we assayed for the expression of several differentiation specific genes in C2C12 cells and RMS cell lines. Genes chosen for analysis were leiomodin2 (LMOD2), troponin I type 2, skeletal, fast (TNNI2), creatine kinase, muscle (CKM) and actin (ACTA1). We found that, as anticipated, these genes were robustly up regulated in response to differentiation in C2C12 cells. However, expression of these genes was at baseline levels in RMS cells and expression was not significantly induced by exposure to differentiation conditions (Figure 1D).

To determine if the loss of MEF2D affects promoter occupancy in RMS cells, chromatin immunoprecipitation assays were performed. We first assayed for the presence of MEF2D at muscle specific promoters. While MEF2D was highly down regulated, it was possible that low levels of MEF2D present in RMS cells could be associated with DNA. However, we were unable to detect MEF2D at the promoter of any gene tested. Shown are data from the TNNI2 promoter (Figure 2A), but the promoters of LMOD2, desmin (DES) and CKM were also assayed with similar results (data not shown). To determine if the MRFs and associated co-factors were present at promoters in the absence of MEF2D, we assayed for the presence of myogenin, MyoD and HEB as we have previously shown that myogenin, MyoD and HEB bind these promoters during normal myogenesis [34]. Here, we found that myogenin (Figure 2B), MyoD (Figure 2C) and HEB (Figure 2D) were bound to muscle specific promoters in RD and RH30 cells. As the MRF and E-protein binding profiles were unaffected by the down regulation of MEF2D, these data suggest that the lack of MEF2D proteins in RMS cells does not affect the binding of the MRFs or associated co-factors to muscle specific promoters, but is likely significant to the inactivity of the MRFs in RMS cells.

To determine if the loss of MEF2D contributed to the inactivity of muscle specific genes RMS cells, we assayed for activity using muscle specific luciferase reporters. We used several muscle specific reporters that show differentiation specific expression and respond to both myogenin and MyoD [35, 36]. Data from all tested reporters were similar and data for the Lmod2-luciferase reporter are shown. We have previously characterized the expression of these reporters and shown that they are active in differentiated C2C12 cells, consistent with the expression pattern of myogenin, and inactive in non muscle cells such as NIH3T3 cells [35, 36]. The Lmod2 reporter construct was transfected into RD and RH30 cell lines and assayed for luciferase expression (Figure 3A). In the ERMS line, RD, the Lmod2 reporter had minimal activity that was modestly above baseline values. The Lmod2 reporter was completely inactive in the ARMS cell line, RH30. The modest activity of the reporter in RD cells is interesting as it suggests that the degree of block to MRF function correlates with the oncogenic potential of the tumor type.

We next co-transfected MEF2D with the muscle specific reporters and assayed for expression. The muscle specific MEF2Dα2 isoform [26] was chosen for our study. Shown are the results for the Lmod2 reporter. We found that transfection of MEF2D promoted expression of the Lmod2 reporter in RD and RH30 cells, with a more robust effect noted in RH30 cells (Figure 3B). Exogenous MyoD and myogenin were also tranfected with or without MEF2D but we found that this did not further stimulate the activation conferred by MEF2D alone (data not shown). As MEF2D requires the MRFs to function [16, 37], the data suggest that the endogenous levels of MyoD and myogenin in RD and RH30 cells are sufficient to stimulate the activation driven by MEF2D.

Our data suggested that the loss of MEF2D might be responsible for the failure of RMS cells to differentiate, so we next assayed if exogenous expression of MEF2D could restore muscle specific gene expression and promote differentiation in RMS cells. RD and RH30 cells were transfected with a vector only control and an expression construct for MEF2D and stable drug resistant clones were selected. However, stable cell lines overexpressing MEF2D were not recovered for RD cells despite multiple experimental attempts. TUNEL analysis revealed a high level of apoptosis in the transfected cells (data not shown). Thus, we transiently transfected RD cells with vector control or MEF2D and examined the effect on muscle specific genes. We also assayed for the expression of the cyclin-dependent kinase (Cdk) inhibitor p21^CIP1/WAF1 (CDKN1A) which is induced early in myoblast differentiation and functions to block cell cycle progression [38, 39]. Induction of p21 in RMS cells is correlated with growth arrest and differentiation of RMS cells [40‐42] and is required for ceramide-induced G2 arrest [43]. We confirmed the expression of exogenous MEF2D in RD cells at the RNA (Figure 4A) and protein level (Figure 4B). We found that MEF2D expression led to an upregulation of muscle specific genes (Figure 4C) and the differentiation specific gene CDKN1A (p21) at the level of RNA (Figure 4D) and protein (Figure 4E).

Stable RH30 cell lines overexpressing MEF2D were recovered and screened to confirm expression at the level of RNA (Figure 5A) and protein (Figure 5B). RH30 cells transfected with vector only control or MEF2D were induced to differentiate for 2 days and gene expression analysis revealed an induction of differentiation specific gene expression in the presence of MEF2D at each gene tested (Figure 5C). We also found that expression of CDKN1A (p21) was robustly stimulated upon differentiation in the presence of MEF2D at the level of RNA (Figure 5D) and protein (Figure 5E). We also examined myosin heavy chain (MHC) expression, a hallmark of differentiated cells. As anticipated, C2C12 cells expressed low levels of MHC while proliferating, but MHC expression was strongly induced in differentiated cells (Figure 5F). In RH30 cells, almost no induction of MHC could be detected upon differentiation. However, RH30 cells tranfected with MEF2D robustly restored MHC expression upon differentiation (Figure 5F). RH30 cells transfected with MEF2D or vector controls were also immunostained with myosin heavy chain antibodies following exposure to differentiation conditions for 2 days. While myosin heavy chain positive cells could not be identified in RH30 cells transfected with a vector control, myosin heavy chain positive cells, including multinucleated myofibers, were readily observed in RH30 cells expressing MEF2D (Figure 5G). We also assayed for up regulation of myogenin as a marker of differentiation and found that myogenin was up regulated in the presence of MEF2D upon differentiation (Figure 5H). Thus, these results are highly suggestive that the lack of MEF2D is implicated in the failure of RMS cells to differentiate.

To evaluate the effect of MEF2D expression on cell proliferation, we measured the growth rate of RH30 cells with vector control or with MEF2D. We found that the expression of MEF2D inhibited the proliferation rate of RH30 cells by approximately 2 fold (Figure 6A). To assay for cell migration, we used the scratch wound assay. After 8 hours the wounds were colonized to a much higher degree by RH30 cells with vector control than RH30 cells with MEF2D (Figure 6B). This difference was still obvious at 18 hours after wounding. The degree to which wound healing was delayed appears to be beyond what could be attributed to the modest growth defect observed in the cells. Next, we examined the effects of MEF2D expression on attachment independent clonal growth of cells in a soft agar assay, a hallmark of cell transformation. We found that RH30 cells showed a strong capacity for colony formation in this assay and that MEF2D expression almost entirely blocked the ability of RH30 cells to grow in an anchorage independent manner (Figure 6C). The modest growth delay in MEF2D expressing cells cannot account for the lack of clonal growth observed in this assay as cells were grown for 30 days in soft agar.

Finally, we tested whether MEF2D expression in ARMS cells could act as an endogenous antitumor factor in vivo. 2 × 10⁶ cells from vector control RH30 cells or RH30 cells expressing MEF2D were injected into the hind limb of nude mice and the tumor size was measured every five days. RH30 cells transfected with a vector control formed visible tumors within the first 2 weeks. In contrast, overexpression of MEF2D led to a complete block of tumor growth (Figure 7A and B). Mice were sacrificed at 4 weeks and tumors resulting from the vector control RH30 cells were dissected, measured and weighed. The overall tumor sizes in each case were comparable (Figure 7C).

RD and SJRH30 (RH30) cells (ATCC) were grown in Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (Hyclone) according to standard protocols. RD2 and RH28 were obtained from Denis Guttridge, Ohio State University, and grown as described above. All cell lines were authenticated by Bio-Synthesis (Lewisville, TX) using STR analysis on September 14, 2011. JW41 cells, isolated from an ERMS tumor from a p53^−/−/c-fos^−/− mouse [51], were the gift of Charlotte Peterson, University of Kentucky. Proliferating C2C12 myoblasts (ATCC) and HEK293 cells (ATCC) were grown in DMEM supplemented with 10% fetal bovine serum (Hyclone). To induce differentiation of C2C12 myoblasts into myotubes, cells were grown to 70% confluence and the media switched to DMEM supplemented with 2% horse serum (Hyclone). C2C12 cells were grown in differentiation medium for the number of days indicated in each experiment.

Cell extracts were made by lysing PBS washed cell pellets in radio-immunoprecipitation assay buffer (RIPA) supplemented with protease inhibitors (Complete protease inhibitor, Roche Diagnostics). Following incubation on ice, clear lysates were obtained by centrifugation. Protein concentrations were determined by Bradford’s assay (Bio-Rad). For each sample, 30 μg of protein was loaded on each gel. Proteins were transferred onto a PVDF membrane using a tank blotter (Bio-Rad). The membranes were then blocked with 5% milk and 1X Tris buffered saline plus tween 20 (TBST) and incubated with primary antibody overnight at 4°C. Membranes were then washed with 1X TBST and incubated with the corresponding secondary antibody. Membranes were again washed with 1X TBST, incubated with chemiluminescent substrate according to manufacturer’s protocol (SuperSignal, Pierce) and visualized by autoradiography. The antibodies used include anti-MEF2D (P-17, Santa Cruz Biotechnologies), anti-MEF2C (E-17, Santa Cruz Biotechnologies), anti-HEB (A-20, Santa Cruz Biotechnologies), anti-myogenin (F5D, Developmental Studies Hybridoma Bank), anti-MyoD (5.8A, Santa Cruz Biotechnologies), anti-MHC (MF-20, Developmental Studies Hybridoma Bank) and anti-GAPDH (Millipore).

RNA was isolated from cells by Trizol extractions (Invitrogen). Following treatment with DNase (Promega), two micrograms of total RNA was reversed transcribed with MultiScribe™ MuLV reverse transcriptase (Applied Biosystems). cDNA equivalent to 40 ng was used for quantitative polymerase chain reaction (qPCR) amplification (Applied Biosystems) with SYBR green PCR master mix (Applied Biosystems). Samples in which no reverse transcriptase was added (no RT) were included for each RNA sample. The relative levels of expression of genes were normalized according to those of hypoxanthine guanine phosphoribosyl transferase (HPRT). qPCR data were calculated using the comparative Ct method (Applied Biosystems). Standard deviations from the mean of the [Δ] Ct values were calculated from three independent RNA samples. Primers are described in Additional file 1: Table S1. Where possible, intron spanning primers were used. All quantitative PCR was performed in triplicate and three independent RNA samples were assayed for each time point. qPCR gene expression data are shown using two formats. For measurements of relative gene expression (fold stimulation), a fold change was calculated for each sample pair and then normalized to the fold change observed at HPRT. For relative measurements of mRNA expression levels (mRNA expression), gene expression levels were quantitated using a calibration curve based on known dilutions of concentrated cDNA. Each mRNA value was normalized to that of HPRT. Fold change was calculated by dividing the mRNA expression values of each sample pair.

ChIP assays were performed and quantified as described previously [34] with the following modifications: 1 × 10⁷ cells were used for each immunoprecipitation and protein A agarose beads (Invitrogen) were used to immunoprecipitate the antibody:antigen complexes. The following antibodies were used: anti-MEF2D (P-17, Santa Cruz Biotechnology), anti-MyoD (5.8A, Santa Cruz Biotechnology), anti-myogenin (F5D, Developmental Studies Hybridoma Bank), anti-HEB (A-20, Santa Cruz Biotechnology). Rabbit IgG (Santa Cruz Biotechnology) was used as a non-specific control. Primers are described in Additional file 1: Table S1. The real time PCR was performed in triplicate. Values of [Δ] [Δ] Ct were calculated using the following formula based on the comparative Ct method: Ct, template (antibody) - Ct, template (IgG) = [Δ] Ct. Fold enrichments were determined using the formula : 2 ^{- [Δ] Ct.} (experimental)/2 ^-[Δ]Ct (reference, CHR19). Standard error from the mean was calculated from replicate [Δ][Δ] Ct values obtained from at least three individual experiments.

RD or RH30 cells were transfected with calcium phosphate according to standard protocols. The plasmids EMSV-myogenin (gift of D. Edmondson, U.T. Medical School at Houston) and pEMCIIs (provided by Andrew Lassar, Harvard Medical School) were used for expressing myogenin and MyoD, respectively. The plasmids pcDNA-MEF2C and pcDNA-MEF2D (gift of Eric N. Olson, University of Texas Southwestern Medical Center) were used for expressing MEF2C and MEF2D, respectively. pcDNA-MEF2D contains the MEF2Dα2 isoform of MEF2D. Luciferase activity was determined using the Dual-Luciferase Reporter Assay System (Promega). RH30 or RD cells were seeded at a density of 5 × 10³ cell per well in 96 well plates and transfected with 0.4 ug of DNA. Transfections were normalized to Renilla luciferase. Transfections were performed in triplicate and all data sets were repeated at least twice.

Stable SJRH30 cell lines overexpressing exogenous MEF2D were made by transfecting SJRH30 cells with linearized pcDNA-MEF2D plasmid or the empty vector, linearized pcDNA3.1, and selecting for geneticin (400 ug/ml) resistant colonies. Individual clones were isolated and propagated.

Cells were grown on cover slips, fixed with paraformaldehyde, incubated with goat serum and 1.0% NP-40 for one hour and washed with PBS. Primary antibodies against myosin heavy chain (1:100, MF20, Developmental Studies Hybridoma Bank) were incubated overnight at 4°C, washed with PBS and detected by Alexa Fluor-488 goat anti-mouse antibody (1:500, Invitrogen). Cell nuclei were then stained by incubating with DAPI (1 μM, Invitrogen) for 5 min.

Cells were seeded in a six well plate at 6 × 10⁴ per well and harvested every two days for cell counts with a hemocytometer. All counts were performed in triplicate and individual experiments repeated three times.

Cells were grown to 100% confluency and the cell monolayer was scraped in a straight line to create a “scratch” with a p200 pipet tip. The debris was removed and the edge of the scratch smoothed by washing the cells once with 1 ml of growth medium. Markings were created near the scratch to obtain the same field during the image acquisition. The tissue culture dish was then placed in a tissue culture incubator at 37°C for 0–18 hours.

Soft agar assays were carried out in 60 mm dishes in which 2 ml of 0.7% Noble agar (USB) in 1X DMEM with 10% FBS was overlaid with 2 ml of 0.35% agar in 1X DMEM with 10% FBS containing the cells. RH30-pcDNA3.1 (vector) and RH30-MEF2D cells were grown to 100% confluence, trypsinized, and dispersed. Cells of each clone (3 × 10⁵) were plated in triplicate. 1 ml of culture medium was added to the top of each plate every 5 days and cells were grown at 37°C for 30 days. The plates were stained with 1 ml of 0.05% Crystal Violet (Fisher) for > 1 hour and colonies were counted using a dissecting microscope.

For in vivo tumor formation, cells were harvested by trypsin treatment and counted. Cells were washed with PBS and suspended at 10⁶ cells/100 μl in PBS. 2 × 10⁶ cells were subcutaneously injected into the hind flanks of 10 week old female athymic nude mice (Foxn1 ^nu /Foxn1 ^nu , Jackson Laboratory). Eight animals were used, and each animal was injected with RH30-pcDNA3.1 cells in the right flank and RH30-MEF2D cells in the left flank. Mice were monitored every other day and tumor dimensions were measured with electronic calipers. Tumor size was estimated by using the modified ellipsoid formula 1/2(length × width²). All animal experiments were conducted according to procedures approved by the Institutional Animal Care and Use Committee at Southern Illinois University.