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September 27, 2005; 65 (6) Articles

Early onset of inflammation and later involvement of TGFβ in Duchenne muscular dystrophy

Y. -W. Chen, K. Nagaraju, M. Bakay, O. McIntyre, R. Rawat, R. Shi, E. P. Hoffman
First published August 10, 2005, DOI: https://doi.org/10.1212/01.wnl.0000173836.09176.c4
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Abstract

Objective: To identify stage-specific induction of molecular pathology pathways in Duchenne muscular dystrophy (DMD).

Methods: We performed mRNA profiling using muscles from fetopsies, infants (aged 8 to 10 months), and symptomatic patients (aged 5 to 12 years) with DMD, and age- and sex-matched controls. We performed immunohistochemistry to determine changes at the protein level and protein localization.

Results: Activated tissue dendritic cells, expression of toll-like receptor 7, and strong induction of nuclear factor-κB pathways occurred soon after birth in DMD muscle. Two muscle wasting pathways, atrogin-1 and myostatin, were not induced at any stage of the disease. Normal muscle showed accumulation of glycolytic and oxidative metabolism capacity with increased age, but this accumulation failed in DMD. The transforming growth factor (TGF)-β pathway was strongly induced in symptomatic patients, with expression of TGFβ type II receptor and apoptosis signal-regulating kinase 1 proteins on subsets of mature DMD myofibers

Conclusions: Our data show stage-specific remodeling of human dystrophin-deficient muscle, with inflammatory pathways predominating in the presymptomatic stages and acute activation of TGFβ and failure of metabolic pathways later in the disease.

Duchenne muscular dystrophy (DMD) is caused by mutations in dystrophin gene, and leads to loss of the dystrophin-glycoprotein complex,1,2 with resulting membrane instability of myofibers, as well as abnormal signaling through nitric oxide synthase and other proteins.3 Patients show evidence of myofiber membrane instability (high serum creatine kinase levels) from birth, but weakness is not apparent until age 2 to 2.5 years and diagnosis is typically made at about age 4 years. The disease is progressive, with patients gradually losing most of their skeletal muscle (wasting) with increasing fibrosis and fatty tissue infiltration. Patients are typically wheelchair-bound around age 9.5 years and usually require ventilatory assistance by age 16 years. The progressive nature of dystrophin-deficiency in humans and animal models implies that there are important secondary, downstream effectors of muscle wasting and weakness.4–11

Previous work on the secondary changes in DMD has revealed possible involvement of mast cell degranulation,12–14 growth factors,15 phospholipase,16 nitric oxide synthase,17–19 metabolism crisis,20,21 oxidative stress,22 calcium homeostasis,23–25 and inflammation.26–30 All of these studies suggested that the progressive pathophysiology is undoubtedly a highly complex process, involving many different pathophysiological and biochemical pathways. However, the developmental timing of these pathways through the progression of DMD has not been investigated.

In this study, we studied skeletal muscle from patients with DMD from three different age groups (fetal, infant, and 5 to 12 years), representing relatively distinct stages of the cellular and clinical pathophysiology, and compared these to age- and sex-matched normal controls. We used genomewide mRNA profiling to characterize the stage in the disease at which normal developmental pathways were disrupted (e.g., induction of metabolic machinery) or pathologic pathways were induced. Our data define pathway targets for therapeutics in DMD and are consistent with recent findings of age-specific action of pharmacological interventions in DMD.31

Methods.

Patients and materials.

We used diagnostic muscle samples from patients with DMD and controls from three different age ranges. Fetopsies were from therapeutic terminations of pregnancies and all postnatal muscles were diagnostic biopsies referred for protein diagnosis. All biopsies were evaluated by the same person (E.P.H.). Fetal samples from DMD fetuses showed histology that was similar to age-matched controls with only rare hypercontracted fibers, consistent with previous reports.32 Infants with DMD were detected via a neonatal screening program using creatine kinase assays on blood spots, and showed an active dystrophic process with myofiber degeneration/regeneration, loose endomysial fibrosis, fiber size variation, and inflammatory infiltrate.33 Diagnostic muscle biopsies were taken at disease presentation from symptomatic patients (aged 5 to 12 years) and all biopsies showed dystrophic muscle, with extensive fiber size variation, degeneration/regeneration, and dense endomysial fibrosis. All samples were covered by Institutional Review Board protocols for preexisting pathologic specimens. Normal controls were age- and sex-matched individuals who showed no evidence of degeneration/regeneration or other pathologic abnormalities on muscle biopsy, and who tested normal for a muscular dystrophy protein panel including immunoblot and immunofluorescence analysis (dystrophin, dysferlin, sarcoglycans, merosin). All DMD and control subjects muscle biopsies were from the vastus lateralis or the quadriceps.

All patients with DMD showed complete dystrophin deficiency at the protein level by Western blot and immunostaining. Immunofluorescence was done using 60 kDa and d10 polyclonal antibodies1, dys III carboxyl-terminal antibody monoclonal antibodies, or both (Novocastra, Newcastle, UK).34 Immunoblot analyses were done using 30 kDa polyclonal antibodies1, dys II rod domain monoclonals (Novocastra),34 or both. All symptomatic patients showed clinical symptoms consistent with the diagnosis of DMD.

For expression profiling, we used frozen muscle biopsies from two male fetuses with DMD (13 and 24 weeks gestational age) and two age- and sex-matched fetal muscle controls (13 and 23 weeks gestational age), four male infant DMD patients (age 8 to 10 months), four age-matched infant controls, 10 symptomatic 5- to 12-year-old DMD patients, and five male patients aged 5 to 12 years for mixed controls. These samples were used individually for the MuscleChip. For the U95Av2 microarrays, the same hybridization cocktails of fetal, infant, and 5- to 12-year-old control samples were rehybridized to U95Av2 microarrays. Data for symptomatic DMD patients on MuscleChip and U95Av2 were taken from our previous publications,35,36 whereas the fetal and infant samples are new to this report. All profiles are available online via our web Oracle database resource (http://pepr.cnmcresearch.org).

For validation studies at the mRNA (reverse transcriptase PCR), U133A/B microarrays, and protein levels, we used a series of biopsies from patients that were distinct from the mRNA profiling studies, including both patients with DMD and controls fulfilling the criteria outlined above. One of the four infant DMD samples was shared in both test data (above) and validation data. The U133A/B data were done for nine distinct symptomatic patients with DMD, and these were part of a large data set of 130 patients and volunteers that will be published elsewhere. The controls for the U133A/B data were age- and sex-matched samples without pathologic findings as described above.

For some immunostaining validations, muscle biopsies from infant “disease controls” (congenital muscular dystrophy) were used (figures 2–4). These subjects all showed dystrophic histopathology on muscle biopsy and clinical and laboratory data consistent with a congenital dystrophy, but tested normal for merosin, dystrophin, dysferlin, and sarcoglycans. A subset of these patients showed reductions in alpha-dystroglycan glycosylation patterns, consistent with a congenital muscular dystrophy due to glycosylation enzyme defects. Mutation studies are currently under way.

Figure

Figure 2. Dendritic lineage cells are highly enriched in DMD muscle biopsies. Frozen muscle sections were stained with antihuman CD86 (A–C) and anti-DC-LAMP (D–F) antibodies, developed with DAB, and counterstained with hematoxylin. CD86 and DC-LAMP stained mainly infiltrating mononuclear cells. No significant staining was observed in the control muscle. Dystropglycan deficient muscle showed occasional CD86 or DC-LAMP positive cells.

Figure

Figure 3. TLR7 is highly upregulated in the muscle fibers of DMD but not normal control biopsies. Frozen muscle sections were stained with antihuman TLR-7 antibodies, developed with DAB, and counterstained with hematoxylin. (A) Some muscle fibers of normal control biopsies showed faint TLR-7 staining. (B) TLR7 staining was mostly restricted to subsarcolemmal regions of muscle fibers and infiltrating mononuclear cells in DMD biopsies. (C) No staining was observed with isotype control antibody on DMD biopsies. (D) Faint TLR-7 staining was observed in endothelial cells and infiltrating mononuclear cells in CMD patients.

Figure

Figure 4. Activated NF-κB and classic NF-κB target genes, HLA-A, HLA-B, and HLA-C, are highly upregulated in DMD biopsies. Frozen muscle biopsies from patients with DMD were stained with rabbit anti-NF-κB p65 antibody (A) and or competed with blocking peptide (B). To visualize nuclear localization, immunofluorescence was performed with rabbit anti-NF-κB p65 antibody and developed with antirabbit texas red and counterstained with DAPI (molecular probes) to visualize the nuclei (blue). (A) Activated muscle fibers showed subsarcolemmal and perinuclear NF-κB accumulation. Diffuse cytoplasmic staining was seen in some normal muscle fibers (C) as well in regenerating/degenerating muscle fibers of dystroglycan deficient muscle (D). (E) Infiltrating mononuclear cells and endothelial capillaries also intensely stain for MHC class I. (F) HLA-ABC staining was seen only in endothelial capillaries of normal muscle biopsies.

Expression profiling.

For the initial test data set, RNA was isolated from two distinct fragments of each biopsy, as we have previously described.20,35,36 Each RNA was processed for production of biotinylated cRNA and hybridization to microarrays, as we have previously described.20 Standard operating procedure and quality control was done as previously described.37 Each sample was then hybridized to both a custom MuscleChip and Affymetrix U95A v2 microarrays. The total number of samples was 24 with two fragments of each, leading to 47 MuscleChip profiles and 33 U95A profiles (some samples were not hybridized to U95A arrays). One infant sample did not pass quality control and was eliminated from further study. The Affymetrix U95Av2 contains approximately 12,000 full-length genes and expression sequence tags (ESTs), and our custom made Affymetrix MuscleChip contains approximately 1,000 full-length genes and 2,000 muscle ESTs.35,38

Generation of hybridization signals (probe-set algorithms) of the microarrays was done using MAS Version 5.0 (Affymetrix, CA), and dCHIP.40 All microarrays were required to fulfill quality control metrics, as we have recently described.35,37 After the absolute analysis using the MAS 5.0, the gene expression levels were imported into GeneSpring software. The DMD samples were normalized to the mean of all the profiles from control samples. Data filtering was done by retaining only those probe sets that showed at least one MAS 5.0 “present call” across all 47 (MuscleChip) or 33 (U95A) profiles. This resulted in retention of 50% to 54% of probe sets for the MuscleChip and 57% to 61% of probe sets for the U95A microarrays.

We focused data analyses on specific pathways or gene ontologies: inflammatory, nuclear factor (NF)-|RrB, transforming growth factor (TGF)-β, toll-like receptor (TLR), and metabolic pathways. Thus, correction for multiple testing was not as important because we limited our studies to specific transcripts. Welch t test was used to calculate the probabilities of significant gene expression changes. To visualize transcripts showing coordinate regulation as a function of disease progression, genes sharing temporal patterns were identified by hierarchical clustering using GeneSpring software. Clustering algorithm was based on standard correlation (r = 0.95). For hierarchical clustering, we included all genes with p < 0.05 in at least one group. A functional clustering tool, GenMAPP,41 was used to identify and present gene changes in the same pathway in DMD vs controls.

All profiles are publicly accessible via National Center for Biotechnology Information (NCBI) Gene Expression Omnibus (GEO) (http://www.ncbi.nlm.nih.gov/geo/), and the Children's National Medical Center Public Expression Profiling Resource (PEPR) (http://pepr.cnmcresearch.org). Online data queries are also publicly accessible, both through a single gene query via PEPR42,43 and via NCBI GEO.

Immunohistochemistry.

The study was conducted as previously described.44 Briefly, serial 4-μm thick frozen muscle sections were cut with an IEC Minotome cryostat, mounted to Superfrost Plus Slides (Fisher Scientific), and fixed in cold anhydrous acetone. Sections were then blocked for 30 minutes in 10% horse serum and 1× PBS, and incubated with primary antibody for 3 hours at room temperature. After three washes with 1× PBS, the sections were incubated in secondary antibodies conjugated with either Cy3 for immunoflorescence staining or biotin for immunohistochemistry. Polyclonal antibodies against TGFβ type II receptor (TGFβRII), apoptosis signal-regulating kinase (ASK1; Cell Signaling Technology, Beverly, MA), embryonic myosin heavy chain (Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA), dendritic cell-specific lysosome-associated membrane glycoprotein (DC-LAMP; Immunotech, Cedex, France); CD86, human leukocyte antigen (HLA)-DR (BD PharMingen, San Diego, CA), TLR7 (Imgenex, San Diego, CA), NF-|RrB and NF-|RrB blocking peptide (Santa Cruz Biotech Inc, Santa Cruz, CA), and major histocompatibility complex (MHC) class I ABC (Harlan Sera-labs, Loughborough, England) were used. Antibody dilutions were as follows: TGFβRII (1:50), p-ASK1 (1:50), embryonic myosin heavy chain (1:10), DC-LAMP (1:10), CD86 (1:20), HLA-DR (1:20), TLR7 (1:20), NF-|RrB (1:30) and MHC class I ABC (1:50). Secondary antibodies for immunoflorescence staining were purchased from Jackson ImmunoResearch Laboratories (West Grove, PA), including Cy3 conjugated donkey antirabbit and Cy3 conjugated donkey antimouse immunoglobulin (Ig) G. VECTASTAIN ABC Kit (Vector Laboratories. Burlingame, CA) was used for immunohistochemistry. The Cy3-conjugated secondary antibodies as well as biotin-conjugated secondary antibodies were used at 1:200 to 1:500 dilution.

TaqMan quantitative PCR analysis.

Quantitative reverse transcriptase (RT)-PCR was performed as previously described.44 Briefly, total RNA was reverse transcribed to cDNA using oligo dT primer (0.5 μg/ul) and reagents from Invitrogen, CA. cDNA was amplified in triplicate in SYBR Green PCR Master Mix (Applied Biosystems, CA). The thermal cycling conditions include 94 °C for 5 minutes, followed by 40 cycles of amplification at 94 °C for 30 seconds, followed by 60 °C for 1 minute. TaqMan PCR primers were designed using a Primer Express program version 1.01 (Applied Biosystems, CA). Primer sequences used for human atrogin-1 were (forward) 5|o:-TTTCCTGGAAGGGCACTGAC-3|o:, (reverse) 5|o:-ACGACTGACCTCTCGACCCTTAT-3. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as internal control. The primers of GAPDH were purchased from Applied Biosystem, CA. All primers were tested for nonspecific amplicons and primer dimers by visualizing PCR products on 2% agarose gels prior to performing qRT-PCR, as well as by dissociation curve analysis following the RT-PCR assays.

Results.

Hierarchical clustering of DMD vs normal controls as a function of age.

To understand the molecular mechanisms involved in the progression of DMD, we used mRNA profiling of skeletal muscle samples from patients with DMD in three different age groups (fetal, infant, and 5 to 12 years), and compared these to age- and sex-matched normal control muscles. Dystrophin mRNA was found to be significantly reduced in the DMD samples at all ages. This is expected given nonsense-mediated mRNA decay that occurs with premature stop codon mutations seen in the large majority of DMD patients.

For an initial overview of the profiling data, we used hierarchical clustering to visualize clusters of genes associated with specific stages of both normal and DMD muscle development (figure 1A). As expected, the proportion of expression changes increased as a function of disease progression (figure 1A). We then queried those branches of the dendrogram showing the most significant alterations in presymptomatic DMD infants, detected by neonatal serum creatine kinase screening, relative to age-matched controls (figure 1A, cluster 1 [bracket underneath dendrogram]). Query of this cluster showed that most cluster members (12 of 13) were found to be transcripts associated with inflammation (table 1). These included inductions of HLA class II and complement components. A subset of this early-onset inflammatory cluster showed continued increased expression as patients became symptomatic (e.g., HLA-DQ-beta [DR7 DQw2], HLA-DR beta [DR2.3], and complement factor D). However, most of the genes in this cluster showed induction in infants and did not change dramatically between 8 months and 5 to 12 years in DMD (table 1). This data shows that many inflammatory pathways are induced soon after birth in muscle from presymptomatic DMD patients.

Figure

Figure 1. Temporal clustering of transcriptome through clinical progression of DMD. (A) Gene tree generated by unsupervised hierarchical clustering using genes with p < 0.05 in any of the three groups: 1, early onset (including HLA-DR) cluster; 2, persistence of fetal cluster; 3, metabolic failure cluster. (B) Early onset (including HLA) gene cluster shows early activation of inflammatory cascades by neonatal period in DMD.

View this table:

Table 1 Immune response genes

A large group of transcripts was seen to be very strongly expressed in normal fetal muscle, with expression lost as a function of age; this same cluster showed abnormally high-level persistent expression in both muscle from DMD infants and symptomatic children (figure 1, cluster 2). Query of this large group of transcripts showed most to represent genes associated with muscle development and differentiation, and were hypothesized to represent mRNAs associated with myofiber regeneration. To confirm this conclusion, the components of this cluster were tested against a 27-timepoint murine muscle regeneration series45 (see figure E-1 on the Neurology Web site at www.neurology.org). Indeed, the large majority of genes in this cluster were strongly positively regulated during muscle regeneration.

The third large cluster showed strong induction as a function of age in normal muscle, but this induction failed in symptomatic children with DMD (figure 1, cluster 3). Query of this cluster showed a high proportion of transcripts associated with both oxidative and glycolytic energy metabolism. This analysis showed that normal muscle attains increasing capacity for substrate metabolism, consistent with the increased need for energy sources as a function of age. This accumulation of metabolic pathways clearly fails in DMD, with symptomatic children showing large-scale loss of metabolic capacity. These data are consistent with our previous suggestion of a metabolic crisis in DMD muscle.20 To determine the extent of the relative loss of both glycolytic and oxidative metabolism, we used the GenMapp resource (http://www.genmapp.org) (figure E-2). This analysis showed widespread downregulation of both glycolytic and oxidative metabolism pathways in symptomatic children with DMD. This expanded temporal series dataset also allows us to see that these metabolic changes become progressively worse as the disease progresses.

Progression of inflammatory responses.

Both the regeneration and metabolic changes were somewhat anticipated; however, the very early induction soon after birth of inflammatory cascades was surprising. We then focused on the inflammatory cluster, including protein and histologic validations. Further study of the 13 genes in cluster 1 (figure 1B) showed that 12/13 were immune-related transcripts (the last was a cDNA clone with unknown function) (table 1). We previously showed HLA-DQ-beta (DR7 DQw2) to be associated with activated dendritic cells and macrophages in the muscles of 5- to 12-year-old patients with DMD,20 but this new temporal series shows that this same transcript was elevated as early as the infant stage of DMD. We further characterized these cells and demonstrated that these infiltrating cells are positive for several markers of mature dendritic cells (CD86, HLA-DR, and DC-LAMP positive) in both infant (figure 2) and 5- to 12-year-old patients (data not shown).

TLRs have become increasingly recognized as some of the earliest detectable markers of an activated inflammatory response.46–49 Given our finding of very early inflammatory changes in DMD muscle, we specifically focused on the mRNA and protein status of TLRs in DMD muscle and the downstream NF-κB pathway components.

The TLR genes are poorly represented on both the U95A and MuscleChips used for clustering above. Therefore, we queried these genes in a set of U133A/B arrays done on symptomatic DMD patients (n = 9) relative to age- and sex-matched control samples (n = 4). We found elevations of TLR-7 mRNA (+6.6-fold, p < 0.0005) in 5- to 12-year-old patients with DMD. We studied TLR-7 by immunohistochemistry and found both infant and symptomatic DMD biopsies showed intense TLR-7 staining in the subsarcolemmal regions of most muscle fibers (figure 3), infiltrating mononuclear cells, and blood vessels at both ages. The immunostaining pattern was more intense than that seen in either age-matched congenital muscular dystrophy disease controls or age-matched biopsies showing no histologic abnormalities (figure 3).

We then extended our analysis of early inflammatory cascades to downstream targets of TLR pathways, the NF-κB pathway. NF-κB molecules exist as homo- or heterodimeric complexes found in the cytosol bound to IκB proteins. In response to various stimuli including physical and chemical stress, viral and microbial products, and inflammatory cytokines, IκB proteins are rapidly phosphorylated, ubiquitinated, and degraded. This frees NF-κB to translocate rapidly into the nucleus to regulate gene expression.50,51 We examined the DMD biopsies for NF-κB activation by immunostaining of a key marker protein, p65. Consistent with early activation of this pathway, we found NF-kBp65 localized to the nuclear/subsarcolemmal regions of skeletal muscle fibers of DMD patient muscle in both infants and symptomatic patients, and this staining was abolished in the presence of specific NF-kB peptide (figure 4, A and B). Diffuse cytoplasmic staining was also observed in some muscle fiber of control muscle (figure 4C). In a congenital muscular dystrophy (CMD) disease control muscle, there was an intense cytoplasmic staining in small regenerating fibers but much less nuclear/subsarcolemmal expression (figure 4D). Nuclear NF-κB was also seen on some of the infiltrating monocytes in DMD and CMD samples (data not shown).

It is known that NF-κB promotes the expression of many genes that participate in a variety of biologic processes, including induction of different cytokines, chemokines, MHC proteins, and adhesion molecules.52 We evaluated these biopsies for HLA-A, HLA-B, HLA-C, which are classic NF-kB target genes. We found a significant increase in HLA class I expression in DMD muscle fibers, infiltrating mononuclear cells, and endothelial cells (figure 4E). In age-matched biopsies showing no pathology, HLA-class I staining was restricted mainly to endothelial cells of the capillaries. When we checked the profiling data, we found upregulation of HLA-A (+4.4-fold, p < 0.05), HLA-B, and HLA-C (2.2-fold, p < 0.001) in 5- to 12-year-old patients with DMD. The genes were also found upregulated in infant samples (HLA-A: +1.6 fold, HLA-B and HLA-C, +1.2 fold), although not significantly. Finally, we examined one well-characterized NF-κB target gene in the mRNA profiling data, vimentin. Both infant and symptomatic DMD muscle biopsies showed upregulation of vimentin, although again, this was significant only in the symptomatic older children.

TGFβ pathways and muscle atrophy pathways.

We found that the TGFβ pathway was highly induced in symptomatic patients with DMD (aged 5-12 years) but not infants (table 2). When localizing the protein products of TGFβRII and ASK1 using immunohistochemistry, we identified a high proportion of positive fibers in the 5- to 12-year-old DMD samples, whereas only few fibers in the infant DMD samples were positive. These fibers were not regenerating, and TGFβRII and ASK1 were colocalized in some fibers but not all (figure 5).

View this table:

Table 2 Genes of TGFβ pathways were upregulated in 5- to 12-year-old DMD patients

Figure

Figure 5. Immunostaining of TGFβRII, ASK1, and embryonic myosin. Frozen muscle biopsies from DMD patients were stained with rabbit anti-TGFβRII, ASK1 antibody, and monoclonal antiembryonic myosin heavy chain antibody. A high proportion of fibers were positive for TGFβRII and ASK1 in the 5- to 12-year-old DMD samples, whereas only few fibers in the 8- to 10-month DMD samples were positive. TGFβRII and ASK1 were colocalized in some fibers but not all. Most TGFβRII and ASK1 positive fibers were not colocalized with fiber stained positively for embryonic myosin heavy chain, suggesting the fibers were not regenerating.

Members of the TGFβ family that are well-known to function in inhibiting muscle growth are myostatin (TGFβ8) and its inhibitor follistatin, the latter of which is growth-promoting. In DMD muscle, we found myostatin to be downregulated in both infants and symptomatic children, and follistatin to be upregulated. The degree of dysregulation was linear with age, with symptomatic children showing the most dramatic changes. These data indicate that the myostatin pathway is strongly shut down in DMD muscle, indicating that this pathway is not contributing to muscle wasting.

We then queried a key transcript associated with ubiquitin-mediated muscle atrophy, atrogin-1, by both microarray (U133A/B chips) and real-time quantitative RT-PCR. By both analyses, atrogin-1 did not show significant change in symptomatic DMD patient muscle compared to normal muscle.

Discussion.

In our original DMD profiling studies in symptomatic patients (aged 5 to 12 years), we showed extensive remodeling of dystrophic muscle, including activation of immune responses and dendritic cells, loss of metabolic capacity (both glycolytic and oxidative), and connective tissue remodeling.20 Here, we extended this data set to three age ranges (fetal, presymptomatic infants, children) to determine if a subset of these tissue remodeling pathways were present very early in the disease process, and if others were associated with later stages. We found little or no evidence of remodeling of fetal muscle; one of the very few significantly altered transcripts was dystrophin itself. The dystrophin mRNA would be expected to be significantly decreased in all DMD samples because patients harbor frame-shifting mutations that lead to nonsense-mediated mRNA decay at about 20% normal levels. Thus, this served as an internal control, demonstrating sensitivity and specificity of our approach. It is important to note that we studied only two DMD fetuses and two normal controls, and further sampling may need to be done to increase sensitivity for the earliest changes in dystrophin-deficient muscle.

Although fetal dystrophin-deficient muscle showed very few perturbations of the transcriptome, we found considerable evidence of molecular pathology relatively soon after birth (8 to 10 months), which typically does not appear until 2 to 2.5 years (figure 1). In infants with DMD, the most dramatic cluster of transcriptionally induced transcripts involved inflammatory genes (figure 1, A and B; table 1). Importantly, most of these transcripts showed high activation in infants, but then stayed at a relatively constant level through the disease progression (figure 1B). Thus, the inflammatory component is among the earliest pathways induced in dystrophin-deficient muscle, but induction of inflammation is not commensurate with symptoms.

The toll-like receptors (TLRs) have become an increasing focus for dissecting the cause/effect steps in complex tissue damage and remodeling inflammatory pathways.47–49 There is increasing evidence that endogenous ligands can stimulate TLRs and trigger an immune/inflammatory response. Signals from damaged or stressed cells may initiate an immune response even in the absence of infection, the so-called danger model of the immune response.53 The natural ligand for TLR7 is currently not known. For the studies, we used congenital muscular dystrophy muscle as a disease control; these patients have abnormalities of glycosylation of membrane proteins and defective regeneration due to inappropriate interactions with the myofiber basal lamina. Presymptomatic 8- to 10-month DMD muscle showed strong activation of TLR7 in most myofibers, with activation of the NFkB pathway (p65) (figures 4 and 5). Congenital muscular dystrophy patient muscle was used as a disease control and these patients also showed activation of both proteins, although expression was restricted to smaller regenerating fibers and did not show as much nuclear localization of activated p65 as DMD muscle. Further evidence of strong activation of the inflammatory pathways in DMD muscle was provided by detection of activated dendritic cells positive for CD86, DC-LAMP, and HLA-DR (figure 2). These same activated dendritic cells were largely absent from congenital muscular dystrophy disease controls. Thus, we conclude that early presymptomatic activation of TLR7 in most myofibers and infiltration of activated dendritic cells into the muscle are early triggers of the strong inflammatory response that we observed in infant muscle.

Given these data, we propose a model where myofiber membrane damage in DMD patient muscle stimulates the TLR pathways, and these in turn cause activation of NF-κB, infiltration/proliferation of dendritic cells, and widespread induction of inflammation pathways. This process is initiated soon after birth. The chronic activation of TLR, NF-κB, and inflammatory pathways results in a later crosstalk and activation of the TGFβ1 pathways, and HLA class I expression. The extent of activation of the early inflammatory pathways does not significantly change with age, despite the marked deterioration of the muscle that occurs much later. Our model is that NF-κB, TLRs, and inflammatory pathways are the keys to the early muscle damage seen in DMD, but that the muscle wasting is a later consequence of failed regeneration via induction of TGFβ1 pathways, and cross-talk between TGFβ1, IGF-1, and the constitutively active NF-κB, TLRs, and inflammatory pathways.

One of the only drugs used routinely in DMD is chronic administration of corticosteroids (prednisone). Most physicians begin prescribing daily prednisone when patients begin to show progression of symptoms. Thus, steroid use is typically delayed for many years after the patient is first diagnosed. Prednisone is a strong anti-inflammatory drug and also is known to inhibit the NF-κB pathways.54,55 Although the beneficial effect of steroids in DMD is not entirely understood, the anti-inflammatory activities of prednisone are believed to play an important role in its efficacy in DMD. Given that we find very early activation of inflammatory cascades in DMD muscle, our data suggests that very early, presymptomatic use of steroids may be indicated. This is particularly important given that recent clinical trial data suggests that DMD is a multistage disease.31 Indeed, these authors showed that creatine was beneficial to older patients but not younger ones; this is entirely consistent with our profiling data showing that the metabolic pathways targeted by this agent become important only in the later stages of the disease.

We also queried the three well-characterized muscle-wasting biochemical pathways to determine if these might contribute to muscle wasting in DMD: atrogin-1-mediated atrophy involving the AKT1/atrogin ubiquitin ligase pathway,56–58 myostatin-induced growth repression,59–61 and TGFβ/MAPK-related atrophy recently shown in the highly pathologic critical care myopathy setting.62 Atrogin-related atrophy is shared in most normal physiologic and some pathophysiological responses such as denervation, starvation, immobilization, corticosteroid myopathy, and cachexia. This pathway can be considered a “canonical” atrophy and muscle-wasting pathway. However, our quantitative RT-PCR in DMD muscle showed that atrogin-1 mRNA was not induced at any stage of the disease. Thus, we conclude that the AKT1/atrogin-1/FOXO pathway is not a major player in the muscle wasting seen in DMD.

The myostatin pathway has received considerable attention, where loss-of-function mutations or inhibition of myostatin by neutralizing antibodies leads to marked muscle hypertrophy. We queried myostatin mRNA in DMD muscle and found that myostatin was significantly decreased in muscle from symptomatic patients with DMD. A key inhibitor of myostatin is follistatin, and we found follistatin mRNA significantly increased at all stages of DMD (fetal, infant, and symptomatic children) by U95A results. Thus, we found evidence that the myostatin pathway is actively repressed in DMD muscle, suggesting that it may contribute to the presymptomatic muscle hypertrophy, but not the symptomatic muscle wasting.

The last muscle-wasting pathway involves TGFβ/MAPK networks. This network is less well described in muscle in human patients, with very recent papers suggesting that constitutive activation of this pathway may lead to muscle apoptosis in critical care myopathy.62,63 However, the TGFβ1 pathway is well studied as a regulator of numerous cellular responses such as cell proliferation, differentiation, migration, and apoptosis.64–66 It is also known to be a powerful modulator of inflammation, fibrosis formation, and myogenesis.67–73 TGFβ1 mRNA and protein has previously been shown to be increased in muscles of symptomatic DMD, congenital muscular dystrophy, and inflammatory myositis patients.74,75 However, here we show that the TGFβ1 pathway is not induced in infant DMD muscle, despite the strong inflammatory responses evident at this stage. Thus, activation of the TGFβ1 pathway appears to be associated with symptoms and muscle wasting in DMD, and should be targeted for therapeutic interventions in DMD.

Footnotes

  • Additional material related to this article can be found on the Neurology Web site. Go to www.neurology.org and scroll down the Table of Contents for the September 27 issue to find the link for this article.

    This article was previously published in electronic format as an Expedited E-Pub on August 10, 2005, at www.neurology.org.

    Supported by National Institutes of Health Grants 5R21AR048318 (Y.W.C.) and 5R01NS029525-09 (E.P.H), Vernon Lynch Memorial Fellowship in Arthritis Research (K.N.), Arthritis Investigator Award from National Arthritis Foundation (K.N.), Maryland Arthritis Research Centre grant (K.N.), Myositis Association grant (K.N.), Muscular Dystrophy Association Grant MDA3455 (M.B.), and donations from the Crystal Ball, Richmond, VA (M.B.).

    Received February 2, 2005. Accepted in final form June 2, 2005.

    Please

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