| | Limb–girdle muscular dystrophy: Diagnostic evaluation, frequency and clues to pathogenesisReceived 23 July 2007; received in revised form 23 July 2007; accepted 17 August 2007. Abstract We characterized the frequency of limb–girdle muscular dystrophy (LGMD) subtypes in a cohort of 76 Australian muscular dystrophy patients using protein and DNA sequence analysis. Calpainopathies (8%) and dysferlinopathies (5%) are the most common causes of LGMD in Australia. In contrast to European populations, cases of LGMD2I (due to mutations in FKRP) are rare in Australasia (3%). We have identified a cohort of patients in whom all common disease candidates have been excluded, providing a valuable resource for identification of new disease genes. Cytoplasmic localization of dysferlin correlates with fiber regeneration in a subset of muscular dystrophy patients. In addition, we have identified a group of patients with unidentified forms of LGMD and with markedly abnormal dysferlin localization that does not correlate with fiber regeneration. This pattern is mimicked in primary caveolinopathy, suggesting a subset of these patients may also possess mutations within proteins required for membrane targeting of dysferlin. 1. Introduction  The limb–girdle muscular dystrophies (LGMD) are a heterogeneous group of disorders characterized by weakness and wasting of the pelvic and shoulder girdle muscles [1], [2]. The clinical course of LGMD ranges from severe forms with disease onset and rapid progression within the first decade of life, to milder forms with later onset and slower progression [3]. There are currently 19 identified forms of autosomal dominant and autosomal recessive LGMD [4], and it is often difficult to distinguish between the various LGMD subtypes [5], [6]. Therefore, diagnosis of LGMD increasingly relies on a combination of immunohistochemical and immunoblot analyses, followed by DNA sequencing to identify the primary mutation, which is essential for the provision of accurate genetic and prognostic counseling. Despite these comprehensive studies, the genetic etiology of many cases of LGMD is not yet known [7]. The study of various dystrophy-associated pathogenic mechanisms has led to the identification of additional disease candidates. Increased susceptibility to membrane damage due to a loss of dystrophin is a major feature of Duchenne muscular dystrophy [8]. Dystrophin is closely associated with a large transmembrane complex of proteins known as the dystrophin-associated protein complex (DAPC); the absence of one or more members of this complex results in increased membrane damage in autosomal recessive forms of LGMD [9]. However, several muscular dystrophy disease genes have been identified that do not encode integral components of the DAPC and in which mechanisms underlying the dystrophic process are less clear. This group includes calpain 3, a calcium-sensitive intracellular protease (LGMD2A) [10], caveolin-3, a component of membrane caveolae (LGMD 1C) [11], [12], and lamin A/C [13] and emerin [14], nuclear envelope proteins (LGMD1B and Emery-Dreifuss muscular dystrophy). Mutations in the gene encoding dysferlin (DYSF) result in LGMD2B [15], [16]. Dysferlin localizes to the sarcolemmal membrane, and has been shown to associate with membrane patches in injured mouse myofibers [17], [18]. The disease mechanism underlying LGMD2B is thought to be due to impaired calcium-mediated muscle membrane repair, rather than increased susceptibility to muscle fiber damage. Therefore, it is likely that other genes involved in the membrane repair pathway are also responsible for as yet unidentified forms of LGMD. In this study, we have determined the frequency of LGMD subtypes within a cohort of Australian muscular dystrophy patients, using a combination of protein studies (immunohistochemistry and immunoblot analysis) and mutation analysis, and have compared the efficacy of using protein studies to guide mutation analysis for the diagnosis of LGMD subtypes. This extensive characterization has led to the identification of a group of muscular dystrophy patients in whom all common gene candidates have been excluded, thereby providing a valuable resource for the identification of new muscular dystrophy disease genes. A significant proportion of our patient cohort exhibit abnormal dysferlin localization. Given the potential role for dysferlin in membrane repair, we further investigated the mechanisms underlying abnormal localization of dysferlin and the likelihood that defective membrane repair may be a common pathway affected in muscular dystrophy. 2. Materials and methods 2.1. Patients We ascertained patients retrospectively through archived muscle biopsy specimens at the University of Sydney, and prospectively through the Neuromuscular Clinic at the Children’s Hospital at Westmead, as well as patients referred from other Neurologists for diagnostic workup of limb–girdle muscular dystrophy (LGMD). Patient evaluation included age of onset, muscle involvement, ambulation status, presence of facial weakness, family history, evidence of muscle hypertrophy, cardiac and/or respiratory muscle involvement, intellectual impairment and serum creatine kinase (CK) levels. We included in our study patients who had progressive weakness in proximal +/− distal muscles, and whose biopsies displayed a dystrophic or myopathic muscle pathology. We excluded patients with delayed early motor milestones and/or age of onset less than two years to rule out congenital muscular dystrophy (CMD). We also excluded patients with Duchenne or Becker muscular dystrophy, facioscapulohumeral dystrophy and myotonic dystrophy. This study was approved by The Ethics Committee of the Children’s Hospital at Westmead and the University of Sydney. 2.2. Antibodies Antibodies to the following proteins were used: dystrophin (NCL-DYS1), dysferlin (NCL-Hamlet), lamin A/C (NCL-LAM-A/C), calpain 3 (NCL-CALP-12A2 and NCL-CALP2C4), emerin (NCL-EMERIN), neonatal myosin heavy chain (NCL-MHCn) and β-spectrin (NCL-SPEC1) from Novocastra Laboratories (Newcastle, UK); caveolin-3 and annexin A2 from Transduction Laboratories (Lexington, KY); α-dystroglycan (VIA4-1) from Upstate Biotechnology (Lake Placid, NY); calcium channel alpha 1C subunit (dihydropyridine receptor, C1603) and skeletal myosin fast (MY32 from Sigma-Aldrich (St. Louis, MO)). 2.3. Immunohistochemistry and immunoblotting All muscle biopsies were snap frozen in liquid nitrogen-cooled isopentane and stored in liquid nitrogen until required. For stretched skeletal muscle, freshly harvested muscle was lightly stretched to approximately 125% of its resting length, clamped and fixed in 3% paraformaldehyde for 10 min at room temperature. Indirect immunofluorescence was performed on 8μm-thick cryosections. For immunodetection of dysferlin (using NCL-Hamlet), cryosections were fixed with methanol:acetone (1:1) at room temperature for 4min. For α-dystroglycan (using VIA4-1), we treated the muscle with ice-cold acetic acid:ethanol (1:1) for 1min. Sections were blocked in 2% BSA (bovine serum albumin) in PBS (phosphate buffered saline) at room temperature for 15min. Sections were incubated in primary antibody (diluted in 2% BSA) at room temperature for 2h or 4°C overnight, followed by three washes in PBS. Anti-mouse or anti-rabbit, CY3-conjugated or Alexa488-conjugated secondary antibodies (Jackson Immunolabs) were applied and sections were incubated at room temperature for 1h. After an additional three washes in PBS, coverslips were applied over Immumount (Shandon, Pittsburgh, PA, USA). Slides were examined under an Olympus BX50 fluorescence microscope and photographed using a Spot II cooled CCD camera. Alternatively, images were captured using a Leica SP2 scanning confocal microscope. Immunoblot analysis was performed as described previously [19]. All immunohistochemical staining and immunoblot studies were performed using age-matched muscle biopsies with normal histology as controls. 2.4. Mutation analysis Mutation analysis was performed on genomic DNA extracted from blood or frozen muscle, or on cDNA synthesized using RT-PCR of mRNA extracted from frozen muscle. PCR products were analyzed by direct sequencing. FKRP restriction enzyme analysis was performed on genomic DNA as described previously [20]. 3. Results 3.1. Characterization of an Australian cohort of limb–girdle muscular dystrophy patients We defined a cohort of 76 Australian patients that satisfied our inclusion criteria for a diagnosis of limb–girdle muscular dystrophy (LGMD). We screened this cohort for known causes of LGMD using a combination of protein analysis and DNA sequencing. We have not analyzed proteins involved in LGMD subtypes predominantly reported in genetic isolates; LGMD1A (myotilin) [21], [22], LGMD2G (telethonin) [23], LGMD2H (TRIM32) [24] and LGMD2J (titin) [25]. 3.1.1. Immunohistochemical analysis Muscle biopsies were initially screened by immunohistochemistry for abnormalities in dysferlin, caveolin-3, α-dystroglycan and the sarcoglycans. Of 76 patients, four were negative for dysferlin (dysferlin-deficient; Fig. 1), although a significant proportion of patients also exhibited abnormal dysferlin localization (see below). Two patients displayed reduced α-dystroglycan immunostaining (Fig. 2), two patients had markedly reduced or negative staining for the sarcoglycans (not shown), one patient had markedly reduced caveolin-3 [26]. 3.1.2. Immunoblot analysis We have previously described a method of immunoblot analysis which requires only a single 8μm cryosection of frozen skeletal muscle [19]. We used this method of immunoblot analysis to examine the expression of dystrophin, dysferlin, calpain 3, lamin A/C, emerin and caveolin-3 in the entire patient cohort (Fig. 3). All patients had normal results for dystrophin, confirming the exclusion of primary dystrophinopathy patients. No patients showed negative or reduced expression levels of emerin. However, increased expression of nuclear envelope proteins, lamin A/C and emerin, was frequently observed in patients with dystrophic biopsies, likely due to an increase in the number of nuclei (not shown). Abnormalities in the expression of dysferlin, calpain 3, lamin A/C and caveolin-3 were observed through immunoblot analyses in a subset of patients (Table 1). 3.1.3. Sequencing analysis We performed sequence analysis of dysferlin (DYSF), lamin A/C (LMNA) and caveolin-3 (CAV3) genes in patients with abnormal results by immunohistochemistry and/or immunoblot analysis (Table 2). Sequence analysis of DYSF in our dysferlin-deficient patients (in whom dysferlin was negative by both immunohistochemistry and immunoblot analysis) identified disease-causing mutations in all four patients. We also identified disease-causing mutations in LMNA for one patient who had normal lamin A/C immunolocalization (but abnormal Western blot results). We identified disease-causing mutations in CAV3 for two patients who had demonstrated reduced levels and/or migration abnormalities for caveolin-3 by immunoblot analysis; immunohistochemical screening detected caveolin-3 abnormalities in only one of these patients. Thus, our results demonstrate the application of single-section Western blot analysis for diagnosis of LGMD subtypes and confirm that immunohistochemical-based screening may not detect all cases of autosomal dominant LGMD, such as caveolinopathies and laminopathies. Recent studies have shown that LGMD2I, caused by mutations in the fukutin-related protein (FKRP), is the most frequent form of autosomal recessive LGMD in Northern European populations [7], [20]. Mutations within FKRP, or other putative glycosyltransferases, may present with a LGMD clinical phenotype and are characterized by hypoglycosylation and abnormal localization of α-dystroglycan [27], [28]. As noted above, only 2/76 of our LGMD cohort displayed reduced α-dystroglycan immunohistochemistry (Fig. 2). We screened our entire patient cohort (n=76) by FKRP restriction enzyme analysis for the common missense mutation 826C>A (Leu276Ile), present in the heterozygous or homozygous state in all cases of LGMD2I reported to date [20], [29], [30], [31], [32], [33]. Two patients were homozygous for the 826C>A mutation (Table 2), corresponding to the two patients with abnormal α-dystroglycan localization, confirming the use of α-dystroglycan localization as an effective tool for the diagnosis of LGMD2I. All remaining patients were normal; no carriers for the common FKRP missense mutation were identified. The lack of abnormal α-dystroglycan localization in these remaining patients further ruled out the possibility of primary mutations other glycosyltransferases, POMT1 and fukutin, which have been shown to cause LGMD2K [34], [35] and LGMD2L [36], respectively. Mutations in CAPN3 (causing LGMD2A) are a common cause of LGMD, however protein-based diagnostic approaches are unreliable for a definitive diagnosis [37], [38]. In fact, we have observed reduced/absent calpain 3 expression in 14/18 primary dystrophinopathy patients by immunoblot analysis (unpublished results). In our LGMD cohort, we observed a reduction/absence in the expression of calpain 3 in 30/76 patients (Table 1). Six of these patients had primary mutations in FKRP, DYSF, LMNA or CAV3. We performed CAPN3 sequencing analysis in the remaining 24 patients in whom all other forms of LGMD had been excluded (11 patients with reduced calpain 3 and 13 patients with absent calpain 3 expression by immunoblot analysis), as well as 9 patients with normal calpain 3 expression in whom the clinical picture was suggestive of LGMD2A and in whom all other causes of LGMD had been excluded. We identified primary CAPN3 disease-causing mutations in 6 of these patients (Tay et al., manuscript in preparation), including 2 patients with normal calpain 3 (by immunoblot analysis). These results confirm that protein-based screening of calpain 3 is unreliable for the identification of primary calpainopathies and that the level of calpain 3 expression may be secondarily reduced in many dystrophic biopsies, perhaps as a consequence of its rapid half-life [39] and/or the biopsy processing conditions. Successful diagnosis is reliant on genetic analysis, in combination with a strong clinical suspicion for LGMD2A. 3.2. Dysferlin is mislocalized in a significant proportion of limb–girdle muscular dystrophy patients While we observed deficiency of dysferlin in only 4/76 biopsies, 21 biopsies showed abnormal dysferlin expression, with markedly reduced and patchy sarcolemmal staining (hereafter referred to dysferlin-defective). We grouped these into three categories based on the pattern of cytoplasmic dysferlin staining. In transverse sections of control muscle, cytoplasmic staining was low, relative to membrane labeling (Fig. 1): dysferlin-defective Group I (14 biopsies) had reduced membrane staining with no increase in cytoplasmic staining; dysferlin-defective Group II (four biopsies) had reduced membrane staining with a significant increase in cytoplasmic dysferlin staining in all fibers; and dysferlin-defective Group III (three biopsies) had reduced membrane staining and variable cytoplasmic staining, with a mixed population of fibers with either low or markedly elevated levels of cytoplasmic dysferlin staining. This abnormal staining pattern was confirmed using two different anti-dysferlin antibodies (NCL-Hamlet and NCL-Hamlet-2), and an array of fixation techniques (not shown). Whilst the four dysferlin-deficient patients were also negative for dysferlin on immunoblot analysis (Fig. 3, lanes 1–4), dysferlin was present at normal (Fig. 3, lanes 5–7), or elevated levels (Fig. 5) in all dysferlin-defective biopsies despite their abnormal dysferlin immunohistochemistry. We have identified the primary cause of LGMD in 6/21 dysferlin-defective patients (Table 2), however 15 dysferlin-defective patients remain with unknown genetic causes of LGMD. We have sequenced the coding region of DYSF in six of these dysferlin-defective patients to date and have not identified any primary disease-causing mutations, suggesting that the mislocalization of dysferlin in our dysferlin-defective patients is a secondary abnormality. To determine whether there were any distinct clinical patterns in these dysferlin-defective patients, we reviewed a number of clinical parameters in detail including age of onset, progression, pattern of muscle hypertrophy/wasting and weakness, cardiac or respiratory involvement, intellectual impairment and facial weakness. There was no obvious common pattern of clinical characteristics in the dysferlin-defective patients: cardiac and respiratory involvement was observed in a small proportion of patients, mild-moderate intellectual impairment was observed in 3 patients and mild facial weakness was observed in 9 patients. 3.3. Abnormal dysferlin localization is not a consequence of fiber regeneration in a subset of dysferlin-defective patients Dysferlin has previously been reported to be abnormally localized in primary dystrophinopathies, sarcoglycanopathies and calpainopathies [40], [41]. Our studies support these findings (Fig. 4), and further demonstrate that dysferlin may be abnormally localized in other forms of muscular dystrophy such as FKRP (Fig. 2) and lamin A/C (Fig. 4). Huang et al [42] demonstrated that fiber regeneration is one cause of cytoplasmic dysferlin localization. We stained sequential muscle sections from 13 DMD patients, and the 6 dysferlin-defective patients with elevated levels of cytoplasmic dysferlin staining in whom the genetic cause had not been identified (Fig. 1, Groups II and III). Eight of 13 DMD patients demonstrated abnormal dysferlin localization in a subset of fibers, with a very strong correlation between cytoplasmic dysferlin localization and neonatal myosin heavy chain (nMHC) expression, a marker of fiber regeneration (Fig. 5a). Interestingly, in dysferlin-defective patients only occasional muscle fibers labeled positively for nMHC in all six biopsies (Fig. 5b), suggesting that dysferlin is mislocalized in many patients independently of fiber regeneration. Given the established role for dysferlin in membrane repair [17], [18], abnormal cytoplasmic localization of dysferlin in these muscular dystrophy patients may nevertheless reflect a degree of membrane damage occurring in these patients’ muscle that does not activate a significant degree of satellite cell repair. Alternately, in a subset of patients, abnormal dysferlin localization may be a direct consequence of a defect in a protein required for efficient membrane targeting of dysferlin, as is the case when caveolin-3 is deficient or mutated [43], [44]. We also examined the levels of expression in our dysferlin-defective patients by immunoblot analysis (Fig. 5c). Dysferlin expression was upregulated in all biopsies with an obvious dystrophic pathology, in whom many fibers stained positively for nMHC (not shown). Upregulation of dysferlin strongly correlated with increased levels of annexin A2, consistent with the known interaction of dysferlin with annexin A2 [18], and consistent with previous studies demonstrating a link between clinical severity and upregulation of annexin A2 in muscular dystrophy [45]. In contrast, dysferlin expression was not upregulated in dysferlin-defective patients with only mildly dystrophic or myopathic biopsies, and similarly, dysferlin expression levels were normal in a primary caveolinopathy patient. 3.4. High resolution confocal microscopy reveals an intracellular pool of dysferlin in normal and dysferlin-defective skeletal muscle To better define the normal (and abnormal) localization of dysferlin, we performed high resolution confocal microscopy in normal human skeletal muscle (Fig. 6A–D). In unfixed biopsy sections, dysferlin localizes to the sarcolemma in both transverse and longitudinal sections. However, when muscle sections were pre-treated with methanol:acetone, a network of cytoplasmic staining was observed in transverse sections, with a striated staining pattern in longitudinal sections. The use of methanol:acetone extracts lipids and dehydrates the samples, thus the appearance of cytoplasmic dysferlin staining following methanol:acetone extraction suggests that the epitope to the dysferlin antibody lies within a membranous structure normally masked by lipid molecules. Double labeling of human stretched vastus medialis skeletal muscle with anti-dysferlin and anti-DHPR antibodies (Fig. 7a) suggests that dysferlin co-localizes with the T-tubule system, consistent with previous studies [42], [43]. In longitudinal sections from dysferlin-deficient muscle, this striated staining pattern was not observed, suggesting that this staining is specific and not an artefact of the methanol:acetone treatment (Fig. 7b). In longitudinal sections of dysferlin-defective muscle however, despite the absence of dysferlin staining at the plasma membrane, the striated staining pattern is preserved (Fig. 7c). 4. Discussion We have characterized a cohort of Australian LGMD patients and determined the frequency of LGMD subtypes in the Australian population (Fig. 8). The most common causes of LGMD in our Australian-based cohort are calpainopathy (LGMD 2A, 8%) and dysferlinopathy (LGMD2B, 5%). Previous studies suggest that dystroglycanopathy is one of the most common causes of LGMD in the United States and in Europe; LGMD2I associated with the 826C>A FKRP mutation is the most common cause of autosomal recessive LGMD in the European population [7], [20], [46], [47]. However, primary FKRP abnormalities underlie less than 3% (2/76) of Australian muscular dystrophy patients, with no evidence of involvement of other glycosyltransferases. This result is surprising since the majority of our cohort is of White European descent. However, we have also found that congenital muscular dystrophy due to laminin-α2 deficiency is much less common in Australia (∼10%) compared to Europe (46–65%) [48]. We evaluated the efficacy of protein analysis to aid the diagnosis of LGMD subtypes. Primary dysferlinopathies can be identified by the absence of dysferlin on immunohistochemistry and immunoblot analysis [49], [50]. However, several patients in our cohort showed extremely low immunoreactivity to dysferlin by immunohistochemistry, despite normals levels of dysferlin by immunoblot analysis. The reason for this discrepancy is unclear, although it suggests that antibody-binding epitopes within dysferlin are obscured in some patients. Therefore, immunoblot analysis is a more effective method for distinguishing between dysferlin-deficient and dysferlin-defective patients. Primary caveolinopathies are often reported to have an absence/severe reduction of caveolin-3 in skeletal muscle [12], [51], [52]; here, we show that autosomal dominant LGMD caused by mutations in both caveolin-3 and lamin A/C can display normal immunohistochemistry, therefore, immunohistochemistry alone is an ineffective method for diagnosis of these disorders. Finally, while protein-based methods remain unreliable for the detection of primary calpainopathies, our application of the single-section Western blot method of analysis [19] allows the exclusion of other common forms of LGMD before undertaking CAPN3 genetic analysis. We have characterized the distribution and expression of dysferlin in a range of LGMDs. The patchy dysferlin membrane localization we have observed in patients with primary calpain 3, lamin A/C and FKRP mutations is similar to that reported for dystrophinopathies and sarcoglycanopathies [40]. We have also demonstrated that the cytoplasmic expression of dysferlin can be associated with the expression of neonatal myosin (nMHC) in dystrophic muscle, suggesting that dysferlin abnormally localizes and/or is upregulated in regenerating muscle fibers as reported by Huang et al. [42], and implying that dysferlin has a role in muscle regeneration in addition to its documented role in membrane repair [17]. Dysferlin localizes to the T-tubules in human skeletal muscle [42], [43]. The T-tubule system plays an important role in excitation–contraction coupling in skeletal muscle by regulating the release of calcium ions [53], raising the possibility that dysferlin-containing vesicles are transported to the T-tubules and sarcolemmal membrane in response to changes in calcium concentration that occur as a result of membrane damage. In a subset of our dysferlin-defective patients in whom the genetic cause is unknown (Groups II–III, Fig. 1), cytoplasmic dysferlin staining did not correlate with the expression of nMHC and thus cannot be attributed to muscle fiber regeneration. The intracellular striated pattern of staining despite loss of dysferlin membrane staining, suggests redistribution of dysferlin to the intracellular pool and may reflect minor damage that has not activated satellite cell repair. Alternatively, the abnormal distribution of dysferlin may reflect a defect in a protein required for efficient membrane targeting of dysferlin; this has been observed in studies primary caveolinopathies, which have shown that caveolin-3 is required for the correct membrane targeting of dysferlin [43], [44]. In conclusion, we have extensively characterized a cohort of Australian LGMD patients for the most common forms of LGMD using a combination of protein- and molecular-based methods. We have demonstrated that mislocalization of dysferlin may occur secondarily to muscle fiber regeneration, and that elevated levels of dysferlin correlate with dystrophic pathology. Finally, we have a well-characterized cohort of patients in whom all common muscular dystrophy disease candidates are excluded, that may provide a useful resource to the international community for identification of novel disease genes. Acknowledgements We are grateful to Drs. 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a Institute for Neuromuscular Research, The Children’s Hospital at Westmead, Sydney, Australia b Discipline of Paediatrics and Child Health, Faculty of Medicine, University of Sydney, The Children’s Hospital at Westmead, Locked Bag 4001, Westmead NSW 2145, Sydney, Australia c Centre for Clinical Neurosciences and Neurological Research, St. Vincent’s Hospital, Melbourne, Australia d Concord Repatriation General Hospital, Sydney, Australia  Corresponding author. Address: Discipline of Paediatrics and Child Health, Faculty of Medicine, University of Sydney, The Children’s Hospital at Westmead, Locked Bag 4001, Westmead NSW 2145, Sydney, Australia. Tel.: +61 2 98451906; fax: +61 2 98453389.
PII: S0960-8966(07)00716-X doi:10.1016/j.nmd.2007.08.009 © 2007 Elsevier B.V. All rights reserved. | |
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