Application of microarray-based gene expression technology to neuromuscular disorders Sterrenburg, P.J.E.

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Application of microarray-based gene expression

technology to neuromuscular disorders

Sterrenburg, P.J.E.

Citation

Sterrenburg, P. J. E. (2007, January 18). Application of microarray-based

gene expression technology to neuromuscular disorders. Retrieved from

https://hdl.handle.net/1887/8914

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License: Licence agreement concerning inclusion of doctoral

thesis in the Institutional Repository of the University

of Leiden

Downloaded from: https://hdl.handle.net/1887/8914

Note: To cite this publication please use the final published version (if

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Gene expression profi ling highlights defective

myogenesis in DMD patients and a possible role

for bone morphogenetic protein 4

Ellen Sterrenburg, Caroline G.C. van der Wees, Stefan J. White,

Rolf Turk, René X. de Menezes, Gert-Jan B. van Ommen,

Johan, T. den Dunnen and Peter A.C. ’t Hoen

Neurobiology of Disease, 2006, 23: 228

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Gene expression profiling highlights defective myogenesis in DMD

patients and a possible role for bone morphogenetic protein 4

Ellen Sterrenburg, Caroline G.C. van der Wees, Stefan J. White, Rolf Turk,

¹

Rene´e X. de Menezes, Gert-Jan B. van Ommen, Johan T. den Dunnen, and Peter A.C. ’t Hoen*

Center for Human and Clinical Genetics, Leiden University Medical Center, Einthovenweg 20, 2333 ZA Leiden, The Netherlands Received 5 September 2005; revised 14 February 2006; accepted 17 March 2006

Available online 6 May 2006

Duchenne Muscular Dystrophy (DMD) is characterized by progressive muscle weakness and wasting. Despite the sustained presence of satellite cells in their skeletal muscles, muscle regeneration in DMD patients seems inefficient and unable to compensate for the continuous muscle fiber loss. To find a molecular explanation, we compared the gene expression profiles of myoblasts from healthy individuals and DMD patients during activation and differentiation in culture. DMD cultures showed significant gene expression changes, even before dystrophin is expressed. We found a higher expression level of bone morphogenetic protein 4 (BMP4) in DMD cultures, which we demonstrate to inhibit differentiation into myotubes. In the later stages of differentiation, we observed a significant decline in expression of sarcomeric genes in the absence of dystrophin, probably contributing to sarcomeric instability.

These results support the hypothesis that inefficient muscle regeneration is caused by impaired myoblast differentiation and impaired maintenance of the myotubes.

Introduction

Duchenne Muscular Dystrophy (DMD) is an X-linked disease caused by a frameshift mutation in the DMD gene (Koenig et al., 1987; Monaco et al., 1988). This frameshift results in a loss of function of the dystrophin protein (Hoffman et al., 1987).

Dystrophin plays an important structural role in the muscle fiber, connecting the extracellular matrix and the cytoskeleton. The N- terminal region binds actin, whereas the C-terminal end is part of the dystrophin-associated glycoprotein complex (DGC), which spans the sarcolemma (Campbell and Kahl, 1989). In the absence of dystrophin, mechanical stress leads to sarcolemmal ruptures, causing an uncontrolled influx of calcium into the muscle fiber interior, thereby triggering calcium-activated proteases and fiber

necrosis (Straub and Campbell, 1997). There is growing evidence that the DGC plays a role in signal transduction through physical interactions with proteins like Calmodulin, Grb2 and nNOS (Rando, 2001). Furthermore, mutations in non-DGC encoding genes (e.g.

Caveolin-3 and Integrin, alpha 7) are also known to cause muscular dystrophies, indicating that disturbed signaling contributes to the DMD pathology (Minetti et al., 1998; Mayer et al., 1997).

Previous research has shown that at the myoblast level, prior to the expression of the muscle-specific DMD isoform, differences can already be observed between healthy and DMD cells. DMD myoblasts in culture have an abnormal morphology, decreased adhesiveness and a reduced number of population doublings (Blau et al., 1983b; Delaporte et al., 1990). Furthermore, DMD myoblasts appear to fuse more slowly, resulting in smaller myotubes that contain fewer nuclei in comparison with normal myotubes (Delaporte et al., 1984). Blau et al., using clonally derived myoblasts, could not detect any morphological differences at the myotube level, stressing the importance of a mixed population cell system (Blau et al., 1983a).

Previously, we have performed microarray experiments to elucidate pathways involved in normal myogenesis. Monitoring gene expression profiles of healthy human primary myoblasts during fusion revealed clusters of potentially co-regulated genes which are involved in different stages of muscle cell differentiation (Sterrenburg et al., 2004). In this study, we set out to determine differences in myogenic gene expression patterns between healthy and DMD human primary cell cultures. To our knowledge, this is the first study looking at large-scale gene expression differences between healthy and DMD muscle differentiation. We have found gene expression differences that may account for the reduced regeneration capacity of DMD myoblasts.

Materials and methods Cell culture

doi:10.1016/j.nbd.2006.03.004

www.elsevier.com/locate/ynbdi Neurobiology of Disease 23 (2006) 228 – 236

* Corresponding author.

E-mail address:p.a.c.hoen@lumc.nl(P.A.C. ’t Hoen).

1Current address: Department of Physiology and Biophysics, 400 EMRB, University of Iowa, Iowa City, IA 52242, USA.

Available online on ScienceDirect (www.sciencedirect.com).

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(KM109, KM108 and HPP4) and three DMD patients (DL589.2 [exon 51 – 55 deletion], DL470.2 [exon 46 – 50 deletion] and 50685.1 [exon 48 – 50 deletion]) (Rando and Blau, 1994; Aartsma- Rus et al., 2002). The age at time of biopsy varied from 2 to 14 years. The cultures consist of myoblasts and other cell types that were present in the original biopsy. The proportion of myoblasts was determined for each biopsy by desmin staining and cell counting as described (Aartsma-Rus et al., 2003). Healthy and DMD cultures did not differ significantly in the average percentage of myoblasts (57 T 20%). Cells were grown in proliferation medium in collagen-coated culture flasks. When cells were 80%

confluent, differentiation was initiated by replacing the high-serum medium with low-serum medium (Sterrenburg et al., 2004). All cell cultures used for the experiments had passage numbers between 4 and 10.

cDNA hybridization

cDNA microarrays containing 4417 muscle-related genes and ESTs (spotted in triplicate) from a human sequence-verified 40K I.M.AG.E. cDNA library (Research Genetics) were used, and these were PCR-amplified, printed and pre-hybridized as described (Sterrenburg et al., 2002; Sterrenburg et al., 2004). Total RNA from the six different cell cultures was isolated at days 0, 1, 2, 4, 6, 10 and 14, amplified, labeled and co-hybridized with a common reference as described (Sterrenburg et al., 2002; Sterrenburg et al., 2004). The quality and quantity of the total RNA and cRNA were checked with the Bioanalyzer Lab-on-a-Chip RNA nano assay (Agilent Technologies).

Data analysis

All slides were scanned with an Agilent scanner (Model 2565BA), and spot intensities were quantified with the GenePix Pro 3.0 program (Axon Instruments). Raw intensity files were imported into Rosetta ResolverR v4.0 (Rosetta Biosoftware) and normalized with the Axon/Genepix error model. Per condition (healthy or DMD), 9 measurements per gene were considered (3 biological replicates, each in 3 technical replicates), and a stringent data analysis procedure was performed. Only genes with a normalized intensity higher than the average +2 standard deviations (SDs) of the negative array controls were analyzed further. This had to be consistent in one condition (healthy or DMD) and for at least one time point. Error-weighted two-way ANOVA was performed with time, disease state and the interaction between time and disease state as variables. Genes were considered differentially expressed when the P value for disease state was <1 10 ⁵ (Bonferroni corrected). Genes differentially expressed in time ( P < 1 10 ⁵) were functionally divided into groups using Gene Ontology Tree Machine (Zhang et al., 2004). Raw GenePix data and the normalized log 10 ratio of the hybridizations were submitted to the GEO database, accession number GSE2693 (Edgar et al., 2002).

Quantitative RT-PCR

cDNA was prepared from total RNA of all 6 cell cultures by reverse transcription using random hexamers and 0.5Ag total RNA as template. PCR primer pairs were designed using Primer3 (http://www-genome.wi.mit.edu/cgi-bin/primer/primer3.

cgi/). The oligonucleotide primer pairs used for each of the genes in this study correspond to the following nucleotides:

glyceraldehyde-3-phosphate dehydrogenase, 510 – 529 and 625 – 644 (NM_002046); BMP4 394 – 413 and 484 – 503 (NM_

001202) and AQP1 1129 – 1148 and 1227 – 1246 (NM_

198098.1). Quantitative PCRs (Lightcycler, Roche) were performed as described with an annealing temperature of 58-C (for GAPDH and BMP4) or 62-C (for AQP1) (Sterrenburg et al., 2004). Optimal cDNA dilutions and relative concentrations were determined using a dilution series per gene. Each gene was normalized to the abundance of glyceraldehyde-3-phosphate dehydrogenase mRNA (shows constant expression over time on the arrays).

RT-MLPA

MLPA probes were designed following the criteria described in Schouten et al. (2002). The reaction was carried out on 125 ng total RNA as described in Eldering et al. (2003), except that all oligonucleotides used as half probes were chemically synthesized (Illumina Inc, San Diego) and two fluorophores were used during the PCR reaction (White et al., 2004). Two microliters of labeled PCR product was mixed with 10 Al formamide and 0.05 Al ROX500 size standard and separated on an ABI3700 capillary sequencer (Applied Biosystems). As there was a considerable range in peak heights, 1:10 dilutions were loaded where necessary to obtain non-saturated signals. Data were exported to Excel for further analysis.

For each probe, the relative peak height was used as a measure of intensity. Normalization was performed by dividing the relative peak height of each probe by the sum of the peak height of two control probes amplified with the same fluorophore. The control probes were targeted to genes that showed no significant change in expression level in the array analysis (Calnexin and Protein phosphatase 3 regulatory subunit B). Standard error of the mean was calculated on basis of 6 measurements (3 cultures measured in duplicate).

Differentiation assay and immunohistochemical analysis

Recombinant Human BMP4 (R&D Systems Inc.) was added to the cells at different concentrations (between 0.3 and 30 ng/ml) upon initiation of differentiation. During medium change, new BMP4 was added (every 4 days). Cells were fixed with 100%

methanol ( 20-C) at 7 or 11 days after serum deprivation.

Immunohistochemical staining was performed as described previously (Aartsma-Rus et al., 2003). Differentiation index was calculated by dividing the number of myosin positive cells by the number of desmin positive cells (=myogenic cells able to differentiate) 100%. For the analysis, a generalized linear regression model was fitted to explain the differences in the differentiation as a function of day (7 or 11), BMP4 concentration (0, 0.3, 1, 3, 10 and 30 ng/ml) and cell line (healthy and DMD). The computations were run using R 2.0.1 (Ihaka and Gentleman, 1996). The model included a term representing the interaction between cell line and concentration, which is the term used to identify individual concentration levels for which the DMD cell line displayed statistically significant differentiation proportions. The model was fitted assuming either a normal or a binomial distribution associated with the error.

E. Sterrenburg et al. / Neurobiology of Disease 23 (2006) 228 – 236 229

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Results

To assess putative connections between gene expression and the differentiating potential of human DMD myoblasts, we performed large-scale gene expression analysis. We analyzed human primary skeletal muscle cell cultures of 6 different individuals (3 healthy, 3 DMD) with a muscle-related cDNA array. RNA was isolated at the myoblast stage (day 0) and at different days of differentiation (day 1, 2, 4, 6, 10 and 14). To find genes that were differentially expressed, an error-weighted two-way ANOVA was performed with time, disease state and the interaction between time and disease state as variables. Of the 4010 unique genes present on the array, 2423 gave a significant signal on at least one time point in either all the healthy samples or all the DMD samples. Ninety five genes were differentially expressed between healthy and DMD cultures (52 down, 43 up in DMD, P (disease state) < 1 10⁵).

Surprisingly, already in undifferentiated cells (t = 0), 7 genes showed more than a 2-fold difference in RNA expression level (Table 1and Supplemental Table 1a/b). Two of these (Aqua- porin 1 (AQP1) and Bone morphogenetic protein 4 (BMP4)) are continuously higher expressed in DMD over the whole time course. We confirmed this difference by quantitative RT-PCR (Fig. 1). Our results suggest that both BMP4 and AQP1 are expressed in the myoblasts because the cultures with the highest myogenicity also have the highest expression of BMP4 and AQP1.

FGF2 is expressed in the fibroblasts because the lines with the lowest myogenicity have the highest expression of FGF2. In this way, BMP4 and FGF2 are inversely correlated.

Dahlqvist et al. previously demonstrated in immortalized mouse myoblasts (C2C12) that BMP4 has an inhibitory effect on muscle differentiation (Dahlqvist et al., 2003). To determine if this also holds true in primary human myoblast cultures and to study if systematic differences between healthy and DMD cells are present, we added recombinant BMP4 to the cell cultures. As expected, addition of recombinant BMP4 to the fusion medium at t = 0 resulted in a concentration-dependent reduction of cell fusion (less multinucleated, myosin-positive cells) in both healthy and DMD cell cultures (Figs. 2 and 3and

Table 1

Genes showing differential gene expression (>2-fold) between healthy and DMD myoblasts

Genbank Symbol Sequence description Fold change

H24316 AQP1 Aquaporin 1

(channel-forming integral protein, 28 kDa)

6.48

AA463225 BMP4 Bone morphogenetic

protein 4

3.08

AA704587 EST ESTs, Weakly similar to hypothetical protein FLJ20958

2.07

R97066 TAL1 T-cell acute lymphocytic leukemia 1

2.30

R38539 FGF2 Fibroblast growth

factor 2 (basic)

2.20

AA598601 IGFBP3 Insulin-like growth factor binding protein 3

2.34

AA664101 ALDH1A1 Aldehyde dehydrogenase 1 family, member A1

2.05

Fig. 1. Gene expression of AQP1 (A) and BMP4 (B) determined with oligonucleotide microarrays (upper panel) or quantitative RT-PCR (lower panel) (Lightcycler, Roche). On the x axis, time is displayed in days. On the y axis, either the log 2 expression ratio (normalized to healthy at t = 0) or the D(Ct) (normalized to healthy at t = 0) are displayed. The Ct value is proportional to the log 2 of the initial amount of mRNA and thus comparable to the log 2 expression ratio. Vertical bars represent standard deviations of the different cultures (n = 3).

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Supplemental Fig. 1). However, DMD cell cultures were significantly more sensitive to BMP4 ( P < 0.05) since a 3- fold lower concentration causes a similar degree of differenti-

ation inhibition (tested with a generalized linear regression model, Supplemental Tables 2 and 3).

Along with genes that were differentially expressed between healthy and DMD over the whole time course, there was a small group of genes whose expression patterns start to diverge after induction of differentiation ( P (disease state) < 1 10 ⁵).

Fig. 4shows that these genes can be divided into two main groups. Firstly, a group of genes is upregulated during the fusion in healthy cell cultures and remains low in DMD cultures (B-factor (BF ), matrix remodeling associated 5 (MXRA5) and membrane metallo-endopeptidase (MME)).

Secondly, two genes with a constant low expression level in healthy cell cultures are upregulated during fusion of DMD cells (Mitochondrial tumor suppressor 1 (MTUS1) and Endo- thelin receptor type A (EDNRA)). We confirmed these results with reverse transcription multiplex ligation-dependent probe amplification (RT-MLPA, Supplemental Fig. 2). The RT-MLPA is a technique that allows the rapid and simultaneous quantification of up to 40 transcripts in a single reaction. We chose this technique as it allows multiple samples and transcripts to be tested in a faster and cheaper assay than quantitative RT-PCR (Eldering et al., 2003).

Fig. 2. Immunohistochemical staining of healthy and DMD myotubes incubated with different concentrations of BMP4, evaluated on day 7 of differentiation.

Cells were stained with DAPI (blue) and antibodies to desmin (red) and myosin (green).

Fig. 3. Differentiation index of healthy and DMD myoblasts after addition of different concentrations of recombinant BMP4. Cells were fixed at day 7.

Regression model, **P < 0.01, *P < 0.05.

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The two-way ANOVA also reveals genes whose expression changes equally in the time course of both healthy and DMD cultures (n = 68, P (time) < 1 10⁵). The role of these genes in myogenesis has been discussed in a previous paper (Sterrenburg et al., 2004). These genes were functionally annotated using Gene Ontology Tree Machine (Zhang et al., 2004).Fig. 5shows the average expression patterns for healthy and DMD cells (functional groups containing6 genes). The sarcomeric proteins (n = 10) showed a significantly decreased expression from day 6 in DMD compared to healthy cells (paired t test, P < 0.01). Other functional categories showed similar expression pattern changes in healthy and DMD cultures.

Discussion

Patients with Duchenne Muscular Dystrophy (DMD) suffer from a severe muscular dystrophy, with phenotypic characteristics appearing early in life. The genetic defect, a mutation in the DMD gene, is known, and many studies used gene expression profiling to find pathways involved in the disease mechanism (Chen et al., 2000; Haslett et al., 2002; Noguchi et al., 2003). These studies focused on the differences in gene expression in the overall muscle.

However, it also would be interesting to look at the gene expression profiles of the myoblasts present in this muscle. These cells are responsible for regeneration of the muscle, and in DMD patients this process may be compromised.

We performed a large-scale gene expression time course study using primary human myoblast cultures to explore muscle differentiation in DMD cells and monitor the reaction of the differentiating cell to the absence of dystrophin. Our results show a clear phasing of the different stages in myogenesis.

Already at the myoblast stage differences appear, and we show that the DMD cells differentiate less efficiently, although differentiation seems to initiate at the same time in healthy and DMD cultures.

Changes are DMD-specific

In order to investigate if the changes we found are DMD- specific and not just initiated by age or a muscular dystrophy, we compared the changes we found in DMD cell cultures with other gene expression studies. Bortoli et al. compared gene expression profiles of myoblasts derived from young individuals (5 days) with gene expression profiles of myoblasts derived from old individuals (52 and 79 years) (Bortoli et al., 2003). The genes we find differentially expressed in DMD cultures are not among the genes they find differentially expressed due to the age of the myoblasts.

Also in FSHD myoblasts, these genes are not differentially expressed, showing the results we obtained are DMD-cell- culture-specific (Winokur et al., 2003).

Healthy and DMD myoblasts

We observed significant differences in gene expression between healthy and DMD myoblast cell cultures despite the fact that the

Fig. 4. Log 2 gene expression ratios of genes differentially expressed between healthy and DMD myoblasts after induction of differentiation: BF, MME, MXRA5, MTUS1, EDNRA. Vertical bars show standard deviations of the different lines used (n = 3).

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full-length dystrophin is not yet expressed in myoblasts and the expression of Dp71, a dystrophin isoform expressed in myoblasts, is not changed by the DMD mutations as these are located upstream of the translation initiation site (Rapaport et al., 1992).

These differences are not likely due to the passaging of the cells as all cells used are in a passage number below 10 and only a few genes change during aging in vitro (comparison of passage number 1 with 15 – 19) (Bortoli et al., 2003). One of the differentially expressed genes, Fibroblast growth factor 2 (FGF2), is significantly lower expressed in DMD cell cultures. In vitro and in vivo studies demonstrated an important role for FGF2 in the recruitment of satellite cells into proliferation. The addition of recombinant FGF2 enhanced the number of proliferating myoblasts twofold and did not suppress the initiation of differentiation (Yablonka-Reuveni et al., 1999; Lefaucheur and Sebille, 1995; Yablonka-Reuveni and Rivera, 1997). In addition, Doukas et al. demonstrated that targeted transgene delivery of FGF2 and FGF6 genes led to an enhancement of skeletal muscle repair, showing the importance of the FGF genes in regeneration (Doukas et al., 2002). These observations indicate that the lower DMD cell culture FGF2 expression observed in our study can explain the decreased myoblast proliferation in DMD cultures previously reported (Dollenmeier et al., 1981; Blau et al., 1983b; Delaporte et al., 1990; Blau et al., 1983b). Most studies agree on the reduced proliferative capacity of DMD myoblasts, which is supported by our gene expression data. From previous studies, it is not clear, however, if this results in a reduced number of satellite cells in the DMD muscle (Renault et al., 2000; Roth et al., 2000; Ishimoto et al., 1983; Blau et al., 1983b). It is likely that additional defective processes, such as those described below, contribute to the severe phenotype of DMD patients.

Fusion into myotubes

Bone morphogenetic proteins (BMPs) belong to the trans- forming growth factor-beta (TGFh) superfamily and have been implicated in a wide variety of cellular processes (Hogan, 1996).

BMPs are involved in the differentiation of certain cell types, including myogenic cells. Addition of BMP2 or BMP4 to the culture medium can inhibit myotube formation in the immortalized murine myogenic cell line C2C12 (Katagiri et al., 1994; Dahlqvist et al., 2003). Our data show that BMP4 is highly expressed in undifferentiated DMD myoblasts, and not in myoblasts from healthy individuals. There are several possible explanations for this. The activation state of the cells could be different. However, this is improbable since most of the gene expression levels are initiated at the same time and the timing of the differentiation is not changed (Fig. 5). Furthermore, PAX7, MYOD and MYOG did not show any change in expression between healthy and DMD cell cultures. Another cause could be that the sub-populations of cells isolated from DMD and normal muscle biopsies are different. If this is the case, then the difference in sub-populations is intrinsic to the disease state (because all DMD lines show the same

Fig. 5. Mean log 2 gene expression of genes in different functional categories during healthy and DMD myoblast fusion. Sarcomeric genes show a decrease in gene expression in late DMD myogenesis (day 6, 10 and 14). *Paired t test, P < 0.01. Genes involved in other functional categories (cell growth and maintenance, protein binding, metabolism up and metabolism down) do not show a significant difference between healthy and DMD at any time point (n 6).

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overexpression in comparison to all healthy lines). This is however also unlikely since the expression profiles at t = 0 are highly similar in all lines. In addition, the percentage of myogenic cells is comparable in the healthy and DMD cultures used in this study.

Irrespective of the cause of the higher BMP4 expression, the higher expression itself probably contributes to the increased sensitivity of the DMD cells to recombinant BMP4 addition, with impaired differentiation as a result. Inhibition of myotube formation by BMP4 has been shown to occur through interaction with the Notch signaling pathway (Dahlqvist et al., 2003). In keeping with this finding, two downstream genes in this pathway, HEY1 and HEY2, are also upregulated in DMD cell cultures. These genes encode a new group of basic helix – loop – helix transcription factors that are related to the hairy/Enhancer of split genes that act by binding to DNA target sequences (E box and N box) and thereby specifically inhibiting transcription of genes involved in muscle differentiation (Nakagawa et al., 2000; Maier and Gessler, 2000; Iso et al., 2003).

Although activation of satellite cells is dependent on Notch signaling, this pathway needs to be downregulated during differentiation (Conboy and Rando, 2002; Conboy et al., 2003).

It is possible that the higher endogenous BMP4 expression in the DMD cell cultures not only reflects the higher activation status of the DMD myoblasts, but also is responsible for a less effective differentiation into myotubes by sustaining activity of the Notch signaling pathway.

Remarkably, we detected a significant increase in Aquaporin 1 (AQP1) expression in DMD myoblasts while Aquaporin 4 (AQP4, not present on the cDNA array) is the main aquaporin expressed in skeletal muscle (Frigeri et al., 2004) and downregulated in muscle of DMD patients (Frigeri et al., 2002). To check the expression level of AQP4 in healthy and DMD myoblasts/myotubes, we performed quantitative RT-PCR. In the cell cultures, AQP4 transcripts could not be detected with RT-PCR, probably due to low or no levels of mRNA of AQP4. This indicates that in myoblasts AQP1 is probably the main aquaporin. Aquaporins function in the maintenance of cellular osmotic environment and the whole body fluid balance (Verkman and Mitra, 2000).

Recently, however, AQP1 was reported to be involved in cell migration, and the higher AQP1 expression in DMD cell cultures may refer to the more active state of the cells in regeneration of the muscle (Verkman, 2005).

While the proliferative capacity is probably reduced and differentiation is inhibited in DMD cell cultures, our results indicate that the timing of the different processes is similar. Genes involved in proliferation are simultaneously downregulated after fusion induction in both healthy and DMD cell cultures (MCM6, CCNB2, CDC28, CKS2 and RPA3,Fig. 5, groupFcell growth and maintenance_). During the actual fusion process of myoblasts into myotubes, however, gene expression differences appear between healthy and DMD cell cultures, again pointing at an impaired fusion potential of DMD cells. It was previously suggested that impaired differentiation is one of the main causes of disease progression in DMD (Oexle and Kohlschutter, 2001). Additionally, after an expression profiling study on muscle biopsies (healthy and dystrophic), Chen et al. suggested that there is altered development and regeneration of myofibers in dystrophic muscle (Chen et al., 2000). In this study, we found genes involved in fusion of healthy myoblasts that probably do not participate in DMD myoblast fusion. Of these, membrane metallo-endopeptidase (MME) and matrix remodeling associated 5 (MXRA5) are thought to be involved in cell adhesion and cell – cell signaling, which is

important for cell fusion (Walker and Volkmuth, 2002). Angoli et al. previously reported that the Laminin-alpha-2-dependent adhesion force is absent in DMD myotubes (Angoli et al., 1997). In addition, in our gene expression study, laminin alpha 2 (LAMA2) is continuously lower expressed in DMD cell cultures, making them less adhesion competent. On the contrary, mitochondrial tumor suppressor 1 (MTUS1) and endothelin receptor type A (EDNRA), both presumed to be involved in signaling, are only upregulated in the DMD cells upon initiation of differentiation.

These genes are possibly part of alternative signaling pathways due to the absence of dystrophin.

Maintenance after fusion

In our time course study, differentiation and fusion of primary human myoblasts into myotubes take approximately 4 days. After this, almost no expression changes are visible and genes are stably expressed (Sterrenburg et al., 2004). A striking phenomenon is a significant decline of sarcomeric gene expression, starting at day 6, detectable in DMD myotubes only. As the myogenicity in the lines studied was comparable, we postulate that the absence of dystrophin in the DMD cell cultures causes sarcomere remodeling, resulting in a secondary response and initiating downregulation of structural genes. Other functional classes of proteins that are simultaneously up- or downregulated during myoblast differentiation do not show a difference in the later time points, indicating that the negative feedback at this time point is a unique characteristic of the sarcomeric proteins in DMD cultures.

Consequences for therapy

Current promising gene therapy studies for DMD are based on the re-establishment of dystrophin in the DGC complex by either myoblast transplantation, viral introduction of a mini-dystrophin gene or by AON-induced exon skipping to restore the reading frame of the DMD gene (van Deutekom and van Ommen, 2003;

Wang et al., 2000; Lu et al., 2003; Gregorevic et al., 2004;

Goyenvalle et al., 2004; Skuk et al., 2004). Thus far, these studies focused primarily on the presence and localization of the dystrophin protein. Our results along with previous studies show that impaired formation of new muscle fibers may complicate treatment protocol, and either intervention in very early life or additional (pharmaceutical) intervention is required to regain normal muscle regeneration capacity (e.g. BMP4 inhibitors, myostatin blockers (McCroskery et al., 2003; Bogdanovich et al., 2002; Wagner et al., 2002; Benabdallah et al., 2005), deflazacort (Anderson et al., 1996; Anderson et al., 2000)).

Conclusion

In conclusion, we show inherent molecular differences between healthy and DMD cell cultures during myogenesis. Decreased FGF2 levels and elevated expression of BMP4 in DMD cell cultures reduce proliferation capacity and make them less differentiation-competent. In addition, we observe lower expression of sarcomeric proteins in DMD myotubes. This combination of reduced proliferation, impaired fusion and impaired maintenance of the DMD myotubes leads to inefficient muscle regeneration and can contribute to the severe phenotype of DMD patients. These results might have important implications for the development of an effective therapy for DMD.

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Acknowledgments

This work was supported by grants from the Nederlandse Organisatie voor Wetenschappelijk Onderzoek (NWO, NL) and the Muscular Dystrophy Campaign (MDC, UK).

Appendix A. Supplementary data

Supplementary data associated with this article can be found in the online version atdoi:10.1016/j.nbd.2006.03.004.

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