Expression Profiling of Dystrophin Deficiency and α-SGD
The goal of this study was to determine downstream gene expression changes resulting from known primary biochemical defects in muscle. However, other sources of gene expression changes include variability in cell-type content of patient muscle biopsies and genetic background differences between individuals. These variables can complicate interpretation. To minimize the effect of these variables, we used the following experimental strategy (). First, each patient muscle biopsy to be studied was split and processed in duplicate. The duplication of each biopsy sample would be expected to control for all sources of both tissue and experimental variability, including cell-type heterogeneity within the tissue, variables in RNA isolation and biotinylated cRNA production, and variability in hybridization to GeneChip® microarrays. Second, to minimize genetic polymorphic variation in expression patterns between different individuals, we studied four or five patient biopsies simultaneously, with equal amounts of cRNA mixed for each of the groups, and the resulting cRNA pools were then hybridized to a single GeneChip® array. Polymorphic variations in expression profiles should be normalized by this approach, whereas gene expression changes correlating with the primary biochemical defect should be retained. The mixing protocol also reduced the cost of the analysis, requiring substantially less microarrays to carry out the experiments. This protocol resulted in six datasets (normal1, normal2, DMD1, DMD2, α-SGD1, and α-SGD2) (). An example of raw image data showing hybridization of cRNA to 20 probe pairs of a single gene is shown in A.
Experimental protocol employed for expression array analyses.
Figure 2 Expression profile data for pooled muscular dystrophy patient biopsies. (A) Example of raw data of probe sets for ERK6 in normal pooled controls (Control 1), DMD (DMD1), and α-sarcoglycan–deficient (α-SGD1) patient biopsies. Shown (more ...)
Description of genes tested on the GeneChip® HuGeneFL array is listed on our site (http://www.cnmcresearch.org; link to microarray). Among the 7,095 probe sets (~280,000 oligonucleotide features) on the Affymetrix HuGeneFL microarray, we found a consistent number of “present” calls for each of the six cRNA pools tested (control 32 and 37%; DMD 36 and 32%; α-SGD 30 and 36%). Data from each experiment is posted on a web site for public access, and comparison to other HuGeneFL datasets (http://microarray.CNMCResearch.org/resources.htm; link to “muscle, human”). Included on the web site is the raw image files for each of the six microarrays, text files containing absolute analyses of each chip (“present” calls; GeneChip® software output), and comparison analyses between different chips (difference calls; GeneChip® software output).
There was high concordance (88%) of “present” calls between duplicated datasets. However, the level of RNA found for each gene showed some variability between datasets; consistent with most microarray data published to date, the highest variability in levels was in genes showing low levels of cRNA hybridization ( B). The variability in levels of specific genes is likely a combination of tissue variability and experimental variability.
Genes that are consistently increased or decreased in all four possible iterative comparisons were determined (e.g., control 1 versus DMD1, control 1 versus DMD2, control 2 versus DMD1, and control 2 versus DMD2) (). Only ~40–60% of difference calls from a single comparison typically survived all four iterative comparisons of datasets. For this analysis, “marginal” difference calls assigned by GeneChip® software analysis were retained in the data sets. The cutoff used for difference calls was a twofold change in expression (either increase or decrease). The four iterative comparisons gave four values for “fold change,” which were then averaged.
Figure 3 Iterative comparisons of duplicate data sets result in stringent determination of differentially expressed genes. (A) The effect of sequential iterative comparisons between DMD and control gene expression profiles is shown. A single data set comparison (more ...)
It is important to note that we focused on difference calls that satisfied all four iterative comparisons of data. This can be considered a very stringent selection of data, as genes that showed significant changes in expression in three data comparisons, but not the fourth, would be excluded from further study. We studied a series of genes that showed difference calls in three comparisons, but not the fourth; in each case the fold changes were close to the twofold cutoff used (data not shown). All data is presented on the web site, with selected data presented in and .
List of Shared Expression Changes of Genes in Dystrophin Deficiency and α-SGD
List of Disease Specific Changes in Dystrophin Deficiency and α-SGD
From the four pairwise comparisons for each disease to normal controls, we identified 275 differentially regulated genes for dystrophin deficiency, and 233 differentially regulated genes for α-SGD. Thus, ~30% of probe sets tested were expressed in muscle, and ~10% of these showed differential regulation in dystrophin deficiency and/or α-SG deficiency. Expression of 138 genes was upregulated, and 137 genes downregulated in dystrophin deficiency versus control, 90 were upregulated and 143 genes downregulated in α-SGD versus control (). These data are also presented as a log scale graph of fold changes, with and without “tilda” values (). As explained in the figure legend, “tildas” are assigned when the denominator approaches zero (e.g., “absent” call), leading to possible exaggeration of the resulting ratio ().
Figure 4 Comparison of differentially expressed genes in dystrophin deficiency and α-SGD expressed as fold changes compared with normal muscle. Both graphs show average fold changes >2 on a log scale. Those spots on the diagonal represent genes (more ...)
Gene Expression Changes Shared by Dystrophin Deficiency and α-SGD
We expected many pathological processes to be involved in muscular dystrophy patient muscle, including degeneration cascades, regeneration programs, and fibrotic proliferation genes. We expected these changes to be shared between the two closely related primary biochemical defects. Consistent with this, we found 144 genes to show greater than twofold up and downregulation in both dystrophin deficiency and α-SGD. We then clustered these genes by pathological processes (). The largest functional group of upregulated genes were genes of cell surface and extracellular proteins (42%). Additional functional groups included genes involved in immune responses (20%) and cell growth, differentiation, and signaling (15%).
Among 80 downregulated genes, 36% of them were involved in mitochondria function and energy metabolism (). Importantly, this data suggests that there is a widespread disorder of both aerobic and anaerobic energy metabolism in patient muscle. Approximately 12% of downregulated genes were involved in cell growth, differentiation, and signaling. Specific genes were selected for verification of expression changes by immunostaining of patient muscle biopsies. As described below, all tested expression changes were confirmed by immunostaining data.
Gene Expression Changes Specific for Dystrophin Deficiency and α-SGD
We then tested for gene expression changes that were specific for either dystrophin deficiency or α-SG deficiency. We expected that this analysis would provide valuable transcriptional information regarding well documented secondary protein deficiencies in each disorder, and would help determine whether these secondary biochemical abnormalities were due to reduced RNA levels or protein instability. In addition, we hypothesized that such disease-specific changes might point to genes or proteins that could have a functionally significant association with dystrophin or α-sarcoglycan at the protein level or gene transcription level. Such biochemical or genetic partners could provide insights into novel pathways or pathophysiological cascades.
For dystrophin deficiency, 131 genes showed significant expression changes relative to normal muscle, yet were assigned as “no change” in α-sarcoglycan deficient muscle. Similarly, α-sarcoglycan deficient muscle showed 89 genes with difference calls that were not seen in dystrophin deficiency. However, the large majority of these genes, in both dystrophin deficiency and α-SGD, showed relatively small changes near the twofold cutoff for statistical acceptance of a difference call. Thus, we felt it was likely that many, if not most, of these potential disease-specific expression changes represented experimental and biological “noise” and not biochemical or genetic “partners” for dystrophin or α-sarcoglycan.
To focus on those gene expression changes most likely to represent biochemical or genetic “partners,” we selected only those genes showing fivefold or greater changes specifically for either dystrophin deficiency or α-SGD (). Using these criteria, only a relatively few genes showed disease-specific changes in gene expression (12 upregulated and 4 downregulated genes specific for dystrophin deficiency, and 11 upregulated and 9 downregulated genes specific for α-SGD).
Dystrophin-deficient patient biopsies showed a specific decrease in dystrophin mRNA (fourfold) that was not seen in α-SGD, suggesting that primary genetic defects can potentially be identified by expression profiling. However, there were other genes showing similar or greater disease-specific differences. Two disease-specific changes were an extracellular signal regulated kinase (ERK6; 10-fold decrease) and a protein tyrosine phosphatase (8-fold decrease). For α-SGD, nine genes showed disease-specific decreases. For example, distinct sets of probes for the uncoupling protein3 gene detected nine- and sevenfold decreases in α-sarcoglycan-deficient, but not dystrophin-deficient patients.
Importantly, there were no significant gene expression changes of well documented secondary protein deficiencies, such as dystroglycan, sarcoglycans (β, δ, and γ), or nNOS. Also, there was not a change in utrophin RNA levels in either Duchenne or α-sarcoglycan dystrophies. On the other hand, another dystrophin-binding protein, α1-syntrophin, showed dramatic reductions in RNA levels in both dystrophin deficiency (14-fold) and α-SGD (6-fold), and α-sarcoglycan showed 3–5-fold reductions in both diseases as well.
A recent report has shown that ERK6 associates with α1-syntrophin, and it is also known that the PDZ domain of α1-syntrophin interacts directly the COOH terminus of dystrophin (Hasegawa et al. 1999
). Thus, some of the biochemical partners of dystrophin (α1-syntrophin and ERK6) are also seen to be part of a coordinately regulated transcriptional group, as all three genes show dramatically reduced levels of RNA. Our data suggests this coordinately regulated gene cluster may also include a protein tyrosine phosphatase as a similar disease-specific mRNA reduction is seen.
Confirmation of Gene Expression Changes, and Cellular Localization of Differentially Regulated Gene Products
To confirm our expression array findings, we chose a series of differentially regulated genes to study by immunostaining patient muscle biopsies. This approach also allowed us to identify the localization of the differentially regulated gene product.
Serial 4-μm-thick frozen muscle sections were processed for immunostaining with antibodies against 15 protein products of differentially expressed genes. All were tested on both unfixed and acetone-fixed sections of normal controls, DMD, and α-sarcoglycan–deficient patient muscles. A subset of antibodies were also tested on female manifesting carriers of DMD (somatic mosaic for dystrophin expression in muscle) and unrelated muscular dystrophies of known causes (calpain III deficiency and partial merosin deficiency). Of the 15 antibodies tested, 9 provided adequate signal/noise ratios for interpretation of protein localization and amount. Results of antibody studies are summarized in .
Gene Expression Changes Confirmed by Immunofluorescence
Factor XIIIa, HLA-DRα Heavy Chain.
Factor XIIIa is a protein known to be involved in blood coagulation as a fibrin cross-linker. We found upregulation of this gene by 11–26-fold in both muscular dystrophies, though the expression in normal muscle was undetectable by GeneChip® array studies, hence possibly exaggerating the extent of upregulation. Immunolocalization of this protein showed positive cells in both epimysial and endomysial connective tissue in dystrophic muscle (). Double immunostaining with a marker for endothelium (laminin α1) showed that the staining for factor XIIIa did not colocalize with this protein, though the factor XIIIa–positive cells were often in close proximity to blood vessels (data not shown).
Figure 5 Immunolocalization of factor XIIIa shows colocalization with HLA-DR in tissue dendritic cells. (A–C) Immunofluorescent visualization of factor XIIIa in frozen muscle biopsy sections from DMD (A), α-SGD (B), and normal control (C) are shown. (more ...)
HLA-DR is a histocompatibility antigen highly expressed in antigen presenting cells. HLA-DRα was upregulated in both dystrophin-deficient (threefold) and α-sarcoglycan–deficient (threefold) patient muscle. Immunostaining for HLA-DRα showed strong immunolocalization to a subset of cells that resembled those immunostained by factorXIIIa. Indeed, double immunostaining for both factor XIIIa and HLA-DRα showed that most positively stained cells coexpressed these two proteins ().
This data suggested that these cells represented tissue dendritic cells (Sueki et al. 1993
). To test this, immunostaining was carried out with markers for circulating dendritic cell subtypes (CD1a, CD1b, and CD1c). Many of the factorXIIIa/HLADR-positive cells also stained with CD1a, and less frequently with CD1b and CD1c (data not shown). The data suggests that the infiltrating cells responsible for expression of factorXIIIa and HLA-DRα are related to tissue (dermal) dendritic cells. This is the first report of factor XIIIa+ and HLA-DR+ dendritic cell infiltration in dystrophic muscles.
Thrombospondin 4, SPARC, and Versican.
Thrombospondin 4 is an extracellular matrix calcium-binding protein particularly abundant in tendon and early osteogenic tissues. It has been shown to be upregulated in denervated muscle, though its function is poorly understood. Thrombospondin 4 was 15-fold increased in dystrophin deficiency, and 23-fold increased in α-SGD. Immunostaining of dystrophic patient muscle biopsies showed thrombospondin 4 to be localized to areas of macrophage infiltration, though the areas showing very strong staining for thrombospondin 4 extended beyond frankly necrotic regions (, D–G). This data suggests that thrombospondin 4 is expressed by interstitial cells in response to macrophage infiltration, denervation, and/or cellular damage of neighboring myofibers (Arber and Caroni 1995
Figure 6 Versican and thrombospondin IV show upregulation in dystrophin- and α-sarcoglycan–deficient muscle. (A–C) Shown are sections from dystrophin-deficient (A), α-sarcoglycan–deficient (B), and normal control (C) muscle (more ...)
SPARC/osteonectin is an extracellular glycoprotein that is strongly expressed during development and tissue regeneration, where it functions to mediate connections between cells and the extracellular matrix (Lane and Sage 1994
). SPARC showed fivefold upregulation in dystrophin deficiency, and fourfold elevation in α-SGD. Immunolocalization showed punctate immunostaining that was dramatically increased in the endomysial and perimysial connective tissue (data not shown).
Versican is a chondroitin sulfate proteoglycan that, like SPARC and thrombospondin, is prevalent in myogenesis of muscle (Carrino et al. 1999
). This gene showed eightfold upregulation in both dystrophies. Immunolocalization identified diffusely increased amounts of the protein in endomysial, but not perimysial, connective tissue of dystrophic muscle (, A–C).
α-Cardiac Actin, Embryonic Myosin Heavy Chain.
Both α-cardiac actin and embryonic myosin heavy chain are specific isoforms of proteins that are transiently expressed during normal muscle development and regeneration (Whalen et al. 1979
; Toyofuko et al. 1992
). Both of these proteins showed upregulation in dystrophic muscle: α-cardiac actin was increased 7–9-fold and embryonic myosin 124–140-fold. Immunolocalization of these proteins in muscle biopsies, similar to those used for expression profiling, showed high-level expression of embryonic myosin in ~20% of dystrophic myofibers, and of α-cardiac actin in ~80% of fibers ().
Figure 7 Developmentally regulated myogenic proteins are persistently upregulated in dystrophin- and α-sarcoglycan–deficient muscle. (A–E) Shown are immunostainings for α-cardiac actin, with quantitation of the percent of α-cardiac–actin (more ...)
Overt degeneration/regeneration of myofibers is a relatively rare event by histological assays (), and the large proportion of myofibers positive for these developmentally specific isoforms did not seem to be justified by the limited amount of regeneration in the dystrophic muscle biopsies. To test the association of myofiber regeneration with the amount of α-cardiac–actin positive myofibers, we studied both female mosaics for dystrophin deficiency ( and ) and a series of patient muscle biopsies from DMD patients (fetal, neonate, 3-, 5-, and 8-yr old), normal controls, merosin deficiency, and calpain deficiency ( F). In the manifesting carrier of dystrophin deficiency ( and ), both dystrophin-positive and dystrophin-negative myofibers were strongly positive for α-cardiac actin, and most appeared to be fully developed myofibers, suggesting that α-cardiac–actin expression persisted beyond the point of complete myofiber regeneration.
Immunostaining of α-cardiac actin in both normal and dystrophin-deficient fetal muscle (~18–22-wk gestation) showed 100% of myofibers to be positive for α-cardiac actin. By birth, the proportion of α-cardiac–actin positive myofibers in normal muscle declined to 0%, whereas in dystrophin-deficient muscle, ~60% of fibers remained strongly positive ( F). This high level was maintained throughout the disease process. Two unrelated dystrophic controls, partial merosin deficiency and calpain III deficiency, showed lower levels of α-cardiac actin (~15–20% positive fibers). As dystrophin-deficient neonatal muscle shows relatively little evidence of degeneration/regeneration by histopathology, we conclude that the overexpression of α-cardiac actin shows persistent expression beyond the normal windows of development and regeneration.