DUX4 Differentially Regulates Transcriptomes of Human Rhabdomyosarcoma and Mouse C2C12 Cells

<div><p>Facioscapulohumeral muscular dystrophy (FSHD) is linked to the deletion of the D4Z4 arrays at chromosome 4q35. Recent studies suggested that aberrant expression of double homeobox 4 (<i>DUX4</i>) from the last D4Z4 repeat causes FSHD. The aim of this study is to determine transcriptomic responses to ectopically expressed DUX4 in human and mouse cells of muscle lineage. We expression profiled human rhabdomyosarcoma (RD) cells and mouse C2C12 cells transfected with expression vectors of <i>DUX4</i> using the Affymetrix Human Genome U133 Plus 2.0 Arrays and Mouse Genome 430 2.0 Arrays, respectively. A total of 2267 and 150 transcripts were identified to be differentially expressed in the RD and C2C12 cells, respectively. Amongst the transcripts differentially expressed in the RD cells, <i>MYOD</i> and <i>MYOG</i> (2 fold, p<0.05), and six <i>MYOD</i> downstream targets were up-regulated in RD but not C2C12 cells. Furthermore, 13 transcripts involved in germline function were dramatically induced only in the RD cells expressing DUX4. The top 3 IPA canonical pathways affected by DUX4 were different between the RD (inflammation, BMP signaling and NRF-2 mediated oxidative stress) and the C2C12 cells (p53 signaling, cell cycle regulation and cellular energy metabolism). Amongst the 40 transcripts shared by the RD and C2C12 cells, <i>UTS2</i> was significantly induced by 76 fold and 224 fold in the RD and C2C12 cells, respectively. The differential expression of <i>MYOD, MYOG</i> and <i>UTS2</i> were validated using real-time quantitative RT-PCR. We further validated the differentially expressed genes in immortalized FSHD myoblasts and showed up-regulation of <i>MYOD</i>, <i>MYOG</i>, <i>ZSCAN4</i> and <i>UTS2</i>. The results suggest that DUX4 regulates overlapped and distinct groups of genes and pathways in human and mouse cells as evident by the selective up-regulation of genes involved in myogenesis and gametogenesis in human RD and immortalized cells as well as the different molecular pathways identified in the cells.</p></div

Comparison of quantified proteins in atrophic and disorganized FSHD myotubes.

<p>(<b>A</b>) Experimental strategy. The proteomes of predominantly atrophic (aFSHD3) or disorganized (dFSHD12) myotubes were compared in two pairs (arrows) with one matching healthy control (CTL12) using post-digest ICPL and 2DLC-MS/MS. The number of <i>D4Z4</i> units, sex and age of the patients are indicated, in addition to the biopsy site. F: female, Q: quadriceps. In addition, another matching healthy control (CTL7) was compared to CTL12. (<b>B</b>) Venn diagram presenting the number of quantified proteins in TE and NE of atrophic (aFSHD3) or disorganized (dFSHD12) FSHD myotubes and their degree of overlap. (<b>C</b>) Fold change distribution of quantified proteins in total extracts of disorganized (blue: dFSHD12_TE) or atrophic myotubes (purple: aFSHD3_TE). (<b>D</b>) Fold change distribution of quantified proteins in NE of disorganized (blue: dFSHD12_NE) or atrophic myotubes (purple: aFSHD3_NE). The most dysregulated proteins in each analysis are indicated by the corresponding Hugo Gene symbol. The proteins that are representative of the most dysregulated categories (DAVID database, Functional Annotation Charts, Gene Ontology: GOTERM_BP_FAT) are also shown.</p

L'Athlète : journal hebdomadaire de tous les sports

03 juillet 19291929/07/03 (N633)-1929/07/03.Appartient à l’ensemble documentaire : Aquit

Workflow of the global procedure for protein extraction, labeling and proteomics.

<p>FSHD and control primary myoblasts were grown as described (<a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0051865#s4" target="_blank">materials and methods</a>) and differentiated into myotubes. Total extracts (TE) and a fraction enriched in nuclear proteins (NE) were analyzed. FSHD and control proteins were submitted to ICPL (Isotope Coded Protein Labeling) before (regular) or after (post-digest) enzymatic digestion with trypsin. FSHD and control samples were labeled with the heavy (H in red) or light (L in grey) ICPL tag, respectively. Equal amounts of FSHD and control labeled peptides were mixed and separated using two-dimensional liquid chromatography (2DLC) using SCX (cation exchange) and RP (reverse phase) columns. Two SCX columns were tested to improve the resolution. The peptides were then analyzed online by MS/MS. The H/L relative peptide quantification ratios were calculated based on the intensities of the peaks in the MS spectra (graph on the right panel). The data were systematically controlled manually, and the quantification data for 3 dysregulated proteins was validated by Western blot. The localizations of these proteins were verified using immunofluorescence. Data interpretation and determination of biological significance in the FSHD context were evaluated using different bioinformatics tools (David database, IPA: Ingenuity Pathway Analysis, Kegg pathway, UniProtKB). The bottom panel represents the fold change distribution of the quantified proteins in preliminary analysis #P1, using regular ICPL and 2DLC-MS/MS on TE of FSHD myotubes from non-affected quadriceps (FSHD8) and a matching control (CTL7), 6 days after differentiation. These cells were previously described in <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0051865#pone.0051865-Barro1" target="_blank">[27]</a>, and information relative to these patients is described in <b><a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0051865#pone.0051865.s006" target="_blank">Table S2</a></b>. Upregulated and downregulated proteins are circled in red or green, respectively. The most-represented functional categories were determined from the David database (Functional Annotation Charts and Clustering) using Gene Ontology (GOTERM_CC_FAT: Cellular Component, GOTERM_BP_FAT: Biological Process, GOTERM_MF_FAT: Molecular Function). GO annotation and accession numbers are indicated. The p-value is equivalent to the EASE score, which uses a conservative adjustment of Fisher’s exact probability and was applied to identify significantly enriched gene categories.</p

Western blot immunodetection of PTRF and MURC in FSHD and control myotubes.

<p>25 µg of TE from primary myotubes were separated by 12% SDS-PAGE and transferred to a nitrocellulose membrane for Western blotting. PTRF (<b>A</b>) and MURC (<b>B</b>) were immunodetected with specific antibodies as described in the <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0051865#s4" target="_blank">Materials and Methods</a> section. GAPDH was used as a loading control. aFSHD3 and dFSHD12 are FSHD myotubes that are predominantly atrophic or disorganized, respectively, and were compared to matching control myotubes (CTL12). The bottom panels present the densitometric analysis of biological replicates (n = 2).</p

Caveolar and Gpi anchor proteins for which a change in abundance was observed in FSHD myotubes using post-digest ICPL coupled to LC-MS/MS.

<p>AC: UniProt accession number; Hugo Gene symbol; Protein name; H/L: fold change (✓: identified protein without quantification); SD: geometric standard deviation; #: number of peptides used for quantification; *: statistical significance (p<0.05) determined by Student’s <i>t</i>-test. Proteins with an H/L ratio greater than 1.5 are highlighted in red; those with a ratio greater than 1.3 are in pink and those with a ratio greater than 1.2 are in light pink. Proteins with an H/L ratio less than 0.7 are highlighted in green and those with an H/L ratio of 0.7 - 0.8 are highlighted in light green. TE: total extract; NE: nuclear extract.</p

Quantification of myosin isoforms in atrophic or disorganized FSHD myotubes.

<p>(<b>A</b>) Histogram depicting the relative quantification (H/L ratio) of myosin heavy chains (MYH), myosin light chains (MYL) and myosin regulatory light chains (MLRS) determined by post-digest ICPL coupled to 2DLC-MS/MS analysis of TE and NE derived from atrophic (aFSHD3) or disorganized (dFSHD12) FSHD myotubes. The bottom table indicates the average H/L ratio for each myosin isoform, which is reported on the Y axis. The significance was evaluated using an ANOVA test and a multiple comparison of means (Tukey contrasts) using the R Foundation for Statistical Computing version 2.14.0 software. p<0.001 (***); p<0.01 (**) and p<0.05 (*) were considered significant. (<b>B</b>) Principal component analysis with quantified MYH9 and MYH3 peptides in TE and NE. Peptides from atrophic myotubes are represented by blue dots and peptides from disorganized myotubes by black dots. The quantification of these myosin isoforms allows the discrimination of two groups in the scatter plot: one group corresponding to aFSHD3 (underlined in purple) myotubes and the other corresponding to dFSHD12 (underlined in blue) myotubes. The proteomic quantitative data related to the myosin isoforms are presented in <b><a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0051865#pone.0051865.s008" target="_blank">Table S4A</a>,</b> and bibliographic information about the role of each isoform is given in <b><a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0051865#pone.0051865.s008" target="_blank">Table S4B and S4C</a></b>. (<b>C</b>) Scatter plot graph representing the H/L ratio corresponding to quantified peptides of skeletal myosin isoforms (MYH3, MYH7, MYH8, MLRS) in analyses dFSHD12_TE, aFSHD3_TE and CTL7_TE. The analysis to the right was conducted as a control for the variability among myotubes from two healthy individuals.</p

MURC and PTRF detection by immunofluorescence in FSHD and control myotubes.

<p>MURC (right panel, red), PTRF (left panel, red) and caveolin-3 (CAV3, green) were detected by co-immunofluorescence in primary FSHD (dFSHD12 and aFSHD3) and control (CTL12) myotubes following 4 days of differentiation. aFSHD3 and dFSHD12 myotubes are predominantly atrophic or disorganized, respectively. The 3 proteins were detected with specific antibodies as described in the <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0051865#s4" target="_blank">Materials and Methods</a> section. DAPI was used to visualize nuclei. The merged signal of CAV3 and PTRF or MURC, representing co-localization, is in yellow. PTRF or MURC staining at the plasma membrane is indicated by arrows and nuclear PTRF staining is indicated by arrowheads. Non-fused myoblasts had weaker MURC staining than myotubes (dotted circles).</p

Aberrant Splicing in Transgenes Containing Introns, Exons, and V5 Epitopes: Lessons from Developing an FSHD Mouse Model Expressing a D4Z4 Repeat with Flanking Genomic Sequences

<div><p>The <i>DUX4</i> gene, encoded within D4Z4 repeats on human chromosome 4q35, has recently emerged as a key factor in the pathogenic mechanisms underlying Facioscapulohumeral muscular dystrophy (FSHD). This recognition prompted development of animal models expressing the <i>DUX4</i> open reading frame (ORF) alone or embedded within D4Z4 repeats. In the first published model, we used adeno-associated viral vectors (AAV) and strong viral control elements (CMV promoter, SV40 poly A) to demonstrate that the <i>DUX4</i> cDNA caused dose-dependent toxicity in mouse muscles. As a follow-up, we designed a second generation of <i>DUX4</i>-expressing AAV vectors to more faithfully genocopy the FSHD-permissive D4Z4 repeat region located at 4q35. This new vector (called AAV.D4Z4.V5.pLAM) contained the D4Z4/DUX4 promoter region, a V5 epitope-tagged <i>DUX4</i> ORF, and the natural 3’ untranslated region (pLAM) harboring two small introns, <i>DUX4</i> exons 2 and 3, and the non-canonical poly A signal required for stabilizing <i>DUX4</i> mRNA in FSHD. AAV.D4Z4.V5.pLAM failed to recapitulate the robust pathology of our first generation vectors following delivery to mouse muscle. We found that the DUX4.V5 junction sequence created an unexpected splice donor in the pre-mRNA that was preferentially utilized to remove the V5 coding sequence and <i>DUX4</i> stop codon, yielding non-functional DUX4 protein with 55 additional residues on its carboxyl-terminus. Importantly, we further found that aberrant splicing could occur in any expression construct containing a functional splice acceptor and sequences resembling minimal splice donors. Our findings represent an interesting case study with respect to AAV.D4Z4.V5.pLAM, but more broadly serve as a note of caution for designing constructs containing V5 epitope tags and/or transgenes with downstream introns and exons.</p></div

Schematic of chromosome 4, D4Z4, and DUX4-expressing AAV vectors.

<p>A: A representation of the telomeric region of the chromosome 4 long arm (4q35). Drawing is not to scale. The 4q35 subtelomere harbors polymorphic, 3.3 kb D4Z4 repeat arrays, as well as other genes, some of which are indicated. This region is normally embedded in repressive heterochromatin. Contraction of the D4Z4 repeat array (in FSHD1) or mutations in SMCHD1 (in FSHD2) leads to epigenetic changes in the 4q35 region, and subsequently permits transcription of the DUX4 gene. An “FSHD permissive” haplotype creates a polyA signal in the pLAM region located downstream of the array. DUX4 transcripts initiated in the last D4Z4 unit extend to this signal and are stabilized by a polyA tail, thereby allowing the mRNA to be translated into the toxic, pro-apoptotic DUX4 protein. B: Two different AAV vectors were engineered to express DUX4. The first generation vector utilized a CMV promoter and SV40 polyA signal. The DUX4 ORF was tagged at the 3’ end with sequences encoding a V5 tag, thereby producing a full-length DUX4 protein containing a carboxy-terminal V5 epitope fusion. ITR, AAV2 inverted terminal repeats. The second generation AAV.D4Z4.V5 vector essentially recapitulates the terminal D4Z4 repeat and pLAM sequences isolated from an FSHD patient, but engineered to express DUX4 with a carboxy-terminal V5 epitope fusion.</p