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Charcot–Marie-Tooth disease type 1A is the most common inherited neuropathy and is caused by duplication of chromosome 17p11.2 containing the peripheral myelin protein-22 gene. This disease is characterized by uniform slowing of conduction velocities and secondary axonal loss, which are in contrast with non-uniform slowing of conduction velocities in acquired demyelinating disorders, such as chronic inflammatory demyelinating polyradiculoneuropathy. Mechanisms responsible for the slowed conduction velocities and axonal loss in Charcot–Marie-Tooth disease type 1A are poorly understood, in part because of the difficulty in obtaining nerve samples from patients, due to the invasive nature of nerve biopsies. We have utilized glabrous skin biopsies, a minimally invasive procedure, to evaluate these issues systematically in patients with Charcot–Marie-Tooth disease type 1A (n = 32), chronic inflammatory demyelinating polyradiculoneuropathy (n = 4) and healthy controls (n = 12). Morphology and molecular architecture of dermal myelinated nerve fibres were examined using immunohistochemistry and electron microscopy. Internodal length was uniformly shortened in patients with Charcot–Marie-Tooth disease type 1A, compared with those in normal controls (P < 0.0001). Segmental demyelination was absent in the Charcot–Marie-Tooth disease type 1A group, but identifiable in all patients with chronic inflammatory demyelinating polyradiculoneuropathy. Axonal loss was measurable using the density of Meissner corpuscles and associated with an accumulation of intra-axonal mitochondria. Our study demonstrates that skin biopsy can reveal pathological and molecular architectural changes that distinguish inherited from acquired demyelinating neuropathies. Uniformly shortened internodal length in Charcot–Marie-Tooth disease type 1A suggests a potential developmental defect of internodal lengthening. Intra-axonal accumulation of mitochondria provides new insights into the pathogenesis of axonal degeneration in Charcot–Marie-Tooth disease type 1A.
Charcot–Marie-Tooth disease (CMT) type 1A is the most common inherited neuropathy, representing almost 50% of all CMT patients (Skre, 1974; Nelis et al., 1996). CMT1A is associated with 1.4 Mb duplication in chromosome 17p11.2 (Lupski et al., 1991; Raeymaekers et al., 1991) which includes the peripheral myelin protein 22 (PMP22) gene. PMP22 over-expression is believed to play an important role in the pathogenesis of CMT1A since over-expression of PMP22 in rodents causes a similar neuropathy. Deletion of the PMP22 gene also causes neuropathy with a different clinical phenotype, named hereditary neuropathy with liability to pressure palsies. Typical phenotypes in patients with CMT1A consist of childhood onset, distal and symmetrical weakness, muscle atrophy, sensory loss, areflexia and foot deformities (Harding and Thomas, 1980; Thomas et al., 1997). Nerve conduction studies show uniform slowing in conduction velocities, which can be observed even in young children (Nicholson, 1991; Garcia et al., 1998; Yiu et al., 2008) and is distinct from the non-uniform slowing of conduction velocities in the acquired demyelinating neuropathies (Lewis and Sumner, 1982). However, the mechanism of the uniform slowing in CMT1A is still elusive. In sural nerve biopsies of patients with CMT1A, increased myelin thickness followed by active de-/remyelination has been observed during the first years of life (Gabreels-Festen et al., 1992, 1995), with the amount of segmental demyelination significantly decreasing after the teenage years (Fabrizi et al., 1998). At the same time, onion bulbs, formed by supernumerary Schwann cells that are not attached to axons, gradually appear in the majority of myelinated fibres (Robertson et al., 2002; Hattori et al., 2003). Variable secondary axonal degeneration has been shown to gradually occur during late childhood (Gabreels-Festen et al., 1995). Axonal degeneration, but not de-/re-myelination, correlates with neurological disability in patients with CMT1A (Dyck et al., 1989; Krajewski et al., 2000). The mechanisms responsible for the axonal loss in CMT1A are yet to be determined.
Acquired demyelinating neuropathies such as chronic demyelinating inflammatory polyradiculoneuropathy (CIDP) are often asymmetric, both in their clinical presentation and in their nerve conduction studies. In particular, slowing in nerve conduction velocities is non-uniform in distinction to the uniform slowing described in CMT1 (Lewis and Sumner, 1982). Pathologically, CIDP is also non-uniform in that segmental demyelination is scattered along myelinated nerves with some internodes shorter than others as a result of the de-/remyelination (Hahn, 2005). Pathological distinctions between inherited and acquired demyelinating neuropathies often require invasive procedures such as sural nerve biopsies.
Furthermore, DNA testing provides reliable diagnosis and eliminates the necessity of sural nerve biopsy on patients with CMT1A. Thus, pathophysiological studies in patients with CMT have been limited by the difficulty in obtaining nerve samples due to the invasive nature of this procedure. As an alternative approach, we and others have begun to utilize skin biopsies to obtain morphological and molecular information from dermal myelinated nerve fibres (Nolano et al., 2003; Li et al., 2005; Provitera et al., 2007). This approach has taken the use of skin biopsy beyond the established application of determining unmyelinated epidermal nerve fibre density in small fibre sensory neuropathies (Holland et al., 1998; Polydefkis et al., 2001; Wendelschafer-Crabb et al., 2006).
In the present study, we applied this technique to a group of patients with CMT1A and CIDP and systematically investigated the morphological and molecular changes in internodes, paranodes, nodes and axons both at light and ultrastructural levels. Our data suggest that this minimally invasive procedure provides insights into the pathogenesis of both disorders and also demonstrated the potential to unveil mechanisms of axonal degeneration.
Skin biopsies were taken from 13 patients with CMT1A, 5 with axonal form of CMT (CMT2), 4 with CIDP and 12 healthy subjects. For electron microscopy and semi-thin section studies, an additional 19 patients with CMT1A and 12 healthy controls were included. All subjects signed a written informed consent before enrolling into the study. All patients underwent a neurological evaluation and a standard nerve conduction study. Neurological disability was assessed using the Charcot–Marie-Tooth Neuropathy Score (Shy et al., 2005).
The skin biopsy was done using a 2 mm skin biopsy punch. The skin was anaesthetized with lidocaine and the biopsies were taken from the lateral aspect of the index finger as previously described (Li et al., 2005). Patients with conditions that may cause or contribute to a neuropathy were excluded. The study protocol was approved by the Wayne State University Human Investigation Committee.
Skin biopsies were washed briefly with PBS buffer and fixed in 4% paraformaldehyde for 30 min at room temperature. The tissue was then embedded in optimal cutting temperature (OCT) medium and cut into vertical sections of 60 µm thickness using a cryomicrotome. Sections were washed in 1× phosphate buffered saline (PBS) in a nine-well glass-plate (Pyrex) and incubated for 1 h in a blocking solution composed of 5% fish skin gelatin and 0.5% triton X-100 in 1× PBS. The sections were then incubated with the primary antibody overnight at 4°C on a rocker. The following day, the sections were washed in 0.1% triton X-100 in 1× PBS three times, 1 h each, followed by incubation with the secondary antibodies overnight at 4°C on a rocker. On the third day, sections were washed three times in 0.1% triton X-100 in 1× PBS and once in 1× PBS, and mounted onto the slides. The slides were left to dry for 2 h and cover-slipped with Vectashield (Vector Labs) mounting media. For each biopsy, we collected 12 sections for double staining with protein gene product 9.5 (PGP9.5; Biogenesis; 1:1000), a panaxonal marker and myelin basic protein, a compact myelin protein (MBP; Ultraclone, Isle of Wight, England; 1:800) for labelling internodal length. Another six sections were double stained with MBP and contactin-associated protein (Caspr, kindly provided by Dr. Peles; 1:1000), a paranodal marker, to assess the paranodes. To evaluate the asymmetry of two hemiparanodes quantitatively, we used an asymmetry index for each hemiparanodal pair (Index = the difference between the longer and the shorter hemiparanode/the sum of the two hemiparanodes). To study whether sodium channels were correctly localized at the nodes of Ranvier, monoclonal pan antibodies against sodium channel (Sigma-Aldrich, 1:500) were used in six sections from CMT1A patients and six sections from healthy controls.
The sections were examined using a Nikon laser confocal microscope (Nikon D-Eclipse C1 confocal system). All fields containing at least an internode were imaged with a 20× objective. Confocal images were obtained at 2-µm increment. Measurements were performed using Reconstruct software (http://www.bu.edu/neural/Reconstruct.html) to trace the course of myelinated nerve fibres through the Z-stack images. Internodal length was measured in nerve fibres located either in the dermal papillae or the subepidermal plexus. This measurement could only be done in nerve fibres that allowed single internode to be individualized from other nerve fibres or internodes. For the evaluation of Meissner corpuscle density, these dermal mechanical receptors were identified, imaged and reconstructed using confocal microscopy in the skin sections stained with antibodies against PGP9.5.
Skin biopsies were fixed in 2.5% glutaraldehyde overnight, osmicated for 2 h in 1% osmium tetroxide, dehydrated in a series of ethanol dilutions (50, 80, 90 and 100%) and propylene oxide and then embedded in epoxy resin. Tissue blocks were sectioned in 1 µm thickness and stained with methylene blue for light microscopic examination to locate the dermal nerve fascicles. These semi-thin sections were also used to measure the number of axons in the dermal nerve fascicles (axonal density). The transverse area of each fascicle was measured by ImagePro Plus. Myelinated nerve fibres within the fascicle were manually counted. The axonal density of myelinated nerve fibres was calculated using total nerve fibres divided by total area. Tissue blocks were then trimmed and sectioned into ultrathin sections for electron microscopy examination (Zeiss EM900). Nerve fibres sectioned at a perpendicular angle were included for diameter measurement. Intra-axonal mitochondria were manually counted under electron microscopy. The axonal area at the transverse section was measured. The mitochondrial density was calculated (which is equal to mitochondrial number/axonal area).
Statistical analysis was performed using GraphPad Prism version 5.02 for Windows (GraphPad Software, San Diego, California USA). The Kruskal−Wallis test was used to compare non-parametric variables from three different disease groups (CMT1A, CMT2, CIDP) and controls. For two-group comparison, Mann–Whitney U-test was performed for non-parametric variables and unpaired t-test for parametric variables. A P-value of <0.05 was considered significant.
Study sponsors had no involvement in the study design, the collection, analysis and interpretation of data, in the writing of the report or in the decision to submit the paper for publication.
All subjects were prospectively enrolled into this study, including 32 with CMT1A (13 for immunohistochemistry and 19 for semi-thin section/electron microscopy studies) and three different control groups (12 normal controls; 5 patients with CMT2; 4 patients with CIDP). The two disease groups (CMT1A versus CMT2) had comparable distributions of ages (P = 0.59) and disease burden, determined by the Charcot–Marie-Tooth Neuropathy Score (Shy et al., 2005) (P = 0.74), with the majority presenting with moderate impairment (defined as neuropathy scores between 11 and 20). A significant age difference was found between healthy controls and both CMT1A and CMT2 groups (Table 1; P < 0.01). However, we reasoned that this age difference should not alter our data since there have been no correlations between internodal length and age in previous investigations of dermal myelinated fibres (Nolano et al., 2003; Provitera et al., 2007). Moreover, we identified no correlation between internodal length and age within any of the groups discussed below. Other demographic details are listed in Tables 1 and and22.
Clinically, patients with CMT1A or CMT2 showed classical phenotypes with slowly progressive distal sensory loss, muscle weakness and atrophy (Harding and Thomas, 1980), but the two groups were separable by nerve conduction studies that revealed uniform slowing of conduction velocities in the former but normal or near-normal conduction velocities in the latter group. As described previously (Lewis and Sumner, 1982), nerve conduction velocities in patients with CIDP were non-uniform with conduction block or temporal dispersion. Taken together, our patient population appeared well within the typical spectrum of clinical presentation of these forms of neuropathy.
A single skin biopsy (2 mm in diameter and 4 mm in depth) was taken from each enrolled patient and studied by immunohistochemistry with antibodies against MBP. The internodes were identified by the staining of MBP and their length was measured by confocal imaging (Fig. 1). In total, 437 internodes were evaluated in all patients, including on average 14.6 in each healthy control, 9.0 in each CMT1A patient, 17.0 in each CMT2 patient and 18.8 in each CIDP patient. There was one patient with severe CMT2 and a Charcot–Marie-Tooth Neuropathy Score of 34, whose skin biopsy yielded only one internode.
The mean of internodal length in patients with CMT1A was 73.9 ± 27.0 µm, which was significantly shorter than that in healthy controls (94.5 ± 28.6 µm; P < 0.001). Axonal degeneration has been demonstrated morphologically in patients with CMT1 (Dyck, 1975). To ensure the shortened internodes in CMT1A were not related to axonal loss, we compared the internodal length in patients with CMT1A with that in patients with CMT2. Again, there was a significant difference between CMT1A and CMT2 (73.9 ± 27.0 µm versus 92.0 ± 29.1 µm; Table 2; P < 0.001). Non-parametric Kruskal–Wallis analysis of the internodal length distribution confirmed a statistical significant difference between the three subgroups (Kruskal–Wallis = 39.4, P < 0.0001), with the distribution in patients with CMT1A shifted towards shorter values of internodal lengths (Fig. 2A). However, the variability around the mean internodal length was comparable between the three groups, suggesting that the reduction in internodal length observed in CMT1A affected nerve fibres uniformly. There was some overlapping of individual internodal lengths between different groups, especially between the 70 and 80 µm range, but no healthy controls or CMT2 patients showed a mean internodal length below 75 µm and no CMT1A patients had values above 92 µm (Supplementary Table 1). Taken together, these results suggest that shortened internodal length is an intrinsic feature of CMT1A, but is not secondary to axonal loss. In addition, we found that there was no correlation between the mean internodal length and our measure of clinical impairment, the Charcot–Marie-Tooth Neuropathy Score or nerve conduction values.
Segmental demyelination with subsequent remyelination is a recognized pathological feature of CMT1A (Gabreels-Festen et al., 1995). In this process, proliferating Schwann cells compete to contact and remyelinate the demyelinated axons; since multiple Schwann cells may contact and remyelinate denuded internodes the resultant internodes are typically shorter in length than normal (Atanasoski et al., 2002). However, during the studies outlined above, we identified no segmental demyelination in any of 117 internodes from patients with CMT1A. There was also no segmental demyelination in 176 internodes from controls and 69 internodes from patients with CMT2 (Table 2).
With this unexpected finding, we questioned whether our skin biopsy technique was capable of detecting segmental demyelination in demyelinating neuropathies. We therefore performed immunohistochemistry studies on skin biopsies from four patients with CIDP, an acquired peripheral nerve disease known to present with demyelination and remyelination. Segmental demyelination was found in all samples from the patients with CIDP. Interestingly, the demyelinating segments were always adjacent to the nodal regions and therefore they probably involved paranodes (Fig. 1E and F). In contrast, none of these features were found in biopsies from patients with CMT1A, CMT2 or normal controls. Thus, absence of segmental demyelination in CMT1A dermal nerves is unlikely to be the result of technical issues related to skin biopsies or the dermal location of the nerves. Supporting this interpretation, segmental demyelination was also undetectable in semi-thin sections or electron microscopic studies of skin biopsies from 19 patients with CMT1A and 12 normal controls.
Alterations in the molecular architecture of myelin paranodes and internodes are increasingly being recognized as features of demyelination and remyelination. For example, voltage-gated sodium channels may migrate from nodes of Ranvier into paranodes or even juxtaparanodes in demyelinated nerve fibres. Caspr, a protein localized to paranodes, may be redistributed into the juxtaparanodes or internodes (Dupree et al., 1999, Li et al., 2005). We therefore investigated the molecular architecture of dermal myelinated nerve fibres in CMT1A with immunohistochemistry. Voltage-gated sodium channels were restricted in the nodes of Ranvier (Fig. 3). Caspr was found within the paranodes, bilaterally flanking the nodes of Ranvier (Fig. 4). We did not observe Caspr dislocated into the internodal or nodal regions. These features were identical to those in myelinated nerve fibres from normal controls.
To determine whether paranodes were otherwise normal in our CMT1A patients, we utilized Caspr staining to measure paranodal lengths by confocal microscopy. The paranodal lengths were significantly reduced in patients with CMT1A, compared with normal controls (4.01 ± 1.25 versus 5.02 ± 1.3 µm; P < 0.001). This finding was consistent with the shortened internodal length in CMT1A (Fig. 2B), and was not consistent with a spreading of Caspr outside the paranodes as has been reported in other genetic and acquired demyelinating neuropathies (Rasband et al., 1998; Arroyo et al., 2002; Bai et al., 2006).
Asymmetry of the two hemi-paranodes that comprise a paranode has also been reported in demyelinating neuropathies. Accordingly, we calculated the indices of paranodal asymmetry on five healthy controls, five CMT1A and three CIDP patients. A small, but statistically significant increase of the index was identified in patients with CMT1A and CIDP, compared with normal controls (P < 0.02) (Fig. 4). Taken together, these results demonstrate that the skin biopsy technique is able to detect pathological features distinguishing CMT1A from acquired demyelinating neuropathy.
Axonal loss in CMT1A is a well-documented pathological feature and is probably responsible for clinical impairment in these patients (Krajewski et al., 2000). To determine whether skin biopsy could also assess axonal loss, we quantified axonal density in dermal nerve fascicles using electron microscopy. No significant difference was found between patients with CMT1A and healthy controls (5.08 × 10–3 ± 0.7 versus 7.73 × 10–3 ± 1.9 axons per μm2 of nerve fascicle, P = 0.16).
We noticed that dermal nerve fascicles were unevenly afflicted by axonal degeneration. If there was a biased sampling towards the less affected nerve fascicles, we were concerned that we may have over-estimated the density of dermal myelinated nerve fibres. We therefore also evaluated the density of Meissner corpuscles in five controls and eight patients with CMT1A. These mechanical receptors could be readily visualized on the skin biopsy sections that were stained with MBP and PGP9.5 antibodies. These corpuscles are primarily innervated by myelinated nerve fibres (Nolano et al., 2003) and their density should reflect the degree of axonal loss of these fibres. Indeed, we found a significant reduction of corpuscle density in patients with CMT1A, compared with normal controls (7.4 ± 5.2 versus 14.6 ± 3.4 corpuscles per mm2; P = 0.04). These results suggest that evaluation of the corpuscle density may be a better approach to assess axonal loss in CMT1A.
To explore further the potential mechanisms responsible for the axonal loss in CMT1A, we quantitatively evaluated the mitochondria in dermal myelinated nerve fibres in 12 controls and 8 CMT1A patients. The morphology of mitochondria appeared comparable between the CMT1A patients and normal controls. However, there was a significant increase in mitochondrial density in the CMT1A group, compared with normal controls (P < 0.01) (Fig. 5). Note that the age distribution for the mitochondrial study was comparable between the CMT1A group and normal control.
Our study has demonstrated that 2 mm punch biopsies are capable of identifying specific changes of molecular architecture and axonal loss in dermal myelinated nerve fibres from patients with CMT1A. These alterations provide pathological signatures that distinguish the inherited demyelinating neuropathy, CMT1A, from acquired demyelinating neuropathies such as CIDP. Moreover, the skin biopsy technique also demonstrated accumulation of intra-axonal mitochondria, which may provide new insights into the pathogenesis of axonal degeneration in CMT1A. We have found myelinated dermal nerves to be increasingly useful in investigating pathogenic mechanisms of inherited neuropathies since our initial report of the technique in 2005 (Li et al., 2005). For example, we have used skin biopsies to demonstrate abnormal RNA splicing of myelin protein zero in a patient with CMT1B (Sabet et al., 2006), to show abnormal trafficking of intracellular organelles in CMT4J (Zhang et al., 2008), and to quantify and identify variable PMP22 levels in compact myelin of patients with CMT1A (Katona et al., 2009). The present study has further expanded the utilities of this technique, and shown its potential to be used even in acquired demyelinating neuropathies. Taken together, our results have demonstrated that skin biopsy can be used to investigate pathophysiologic mechanisms in inherited, as well as acquired demyelinating neuropathies. Comparisons between findings from skin biopsies and sural nerve biopsies may further enhance our understanding of the pathogenesis of CMT1A. Moreover, skin biopsies also possess certain advantages, including its minimally invasive nature and potential repetitive use in the same subject for longitudinal studies.
Pathological studies of sural biopsies in different age groups of patients with CMT1A have suggested that extensive segmental demyelination occurs during the first decade of life in patients with CMT1A and are significantly less frequent in adulthood (Gabreels-Festen et al., 1992, 1995; Fabrizi et al., 1998; Gabreels-Festen and Wetering, 1999). Although active segmental demyelination was not found in dermal myelinated fibres of CMT1A patients, an increased asymmetry in hemiparanodal length suggests that previous segmental demyelination may have occurred in these fibres. The increased asymmetry of paranodes in CMT1A could have resulted from insidiously accumulated de-/remyelination that occurred in the early life of the patients, as has been suggested by investigations of sural nerve biopsies in other patients with CMT1A (Gabreels-Festen et al., 1992, 1995).
Interestingly, distinct to what has been described in sural nerve studies (Dyck et al., 1968), the variability around the mean internodal length was not increased in dermal fibres in CMT1A. In developmentally abnormal myelination, or dysmyelination, internodal length may be uniformly shortened. For example, in periaxin null mice, internodal elongation is impaired resulting in uniformly shortened internodes as well as slow nerve conduction velocities (Court et al., 2004). We speculate that a developmental defect in internodal lengthening, limiting Schwann cell extension along axons, could explain our findings and be a contributing pathological mechanism in CMT1A. Limited internodal lengthening has been demonstrated in a cell culture study using the Tembler-J mouse model of PMP22 point mutations (Liu et al., 2005). The recent observations suggesting that PMP22 is a binding partner of α6β4-integrin (Amici et al., 2006, 2007), a Schwann cell protein that mediates interactions with the basal lamina complex could provide a molecular basis for this hypothesis, although further studies are necessary to confirm this. Longitudinal studies to document the changes of internodal length and segmental demyelination during development in CMT1A would be helpful to test this hypothesis further.
It is not yet clear why active segmental demyelination is not observed in dermal myelinated fibres in CMT1A since this has been reported in sural nerve biopsies in patients with CMT1 prior to the era of molecular diagnosis (Dyck et al., 1968) and in cases of CMT1A (Gabreels-Festen et al., 1995; Fabrizi et al., 1998). One possible explanation is that the shorter internodal lengths and smaller fibre diameters of myelinated fibres typically observed in this territory may allow for a more efficient remyelination early in life. This discrepancy stresses the importance of distinguishing changes observed in dermal biopsies from those of sural nerve biopsies in order to understand better the pathogenic mechanisms involved in CMT1A. If this was the case, our skin biopsy technique would be a better approach to show the pathological changes distinguishing the inherited from acquired demyelinating neuropathy.
The finding of shortened internodal length fits well with the uniform slowing of conduction velocities in CMT1A (Lewis and Sumner, 1982), observed even in the youngest patients (Nicholson, 1991; Garcia et al., 1998; Yiu et al., 2008). Uniform slowing in CMT1A would not be adequately explained by active segmental de-/remyelination, since non-uniform slowing, temporal dispersion and conduction block are usually seen in acquired demyelinating neuropathies, where active segmental demyelination is the main pathological feature (Hahn, 2005). Shortened internodal length without segmental demyelination is sufficient to reduce nerve conduction velocities in periaxin-null mice (Court et al., 2004). Thus, uniformly shortened internodal length in patients with CMT1A may serve, at least partially, as a pathogenic mechanism for the uniform slowing of conduction velocity in CMT1A.
In the present study, we were able to quantify the axonal loss of myelinated nerve fibres by measuring the density of Meissner corpuscles. Attempts to quantify axonal loss by counting axons within fascicles produced variable results between fascicles, such that axonal loss could not be ascertained with confidence. Moreover, no significant differences between controls and patients with CMT1A could be determined by this technique. It is possible that the length of nerves innervating the hands may be insufficient to document axonal loss in mild forms of length-dependent hereditary neuropathies. However, areas in lower extremities typically have more severe axonal loss with few if any myelinated nerve fibres making it impossible to measure internodes and segmental demyelination. Thus, for the purpose of our study, skin biopsies from hands were the more appropriate area to perform these measurements. Alternatively, the density of Meissner corpuscles in CMT1A patients was significantly reduced compared with controls. Whether sensory nerves going to Meissner corpuscles are preferentially affected in CMT1A compared with other myelinated sensory axons in skin is unknown. Whatever the underlying reason for this difference is, Meissner corpuscle density appears to be a more sensitive index for evaluating axonal loss in CMT1A, which is consistent with the observation of a previous study (Dyck et al., 1966).
Since axonal loss is closely related to the neurological impairment in patients with CMT1A (Dyck et al., 1989; Krajewski et al., 2000), mechanisms underlying the axonal degeneration are necessary to understand the pathogenesis of the neuropathy. PMP22 is expressed in myelinating Schwann cells, but not in the neurons they ensheath (Welcher et al., 1991). Thus, axonal loss must result from abnormal interactions between mutant Schwann cells, which overexpress PMP22, and the underlying axons. Our study identified an increase of mitochondrial density in myelinated axons of CMT1A, suggestive of an impairment of mitochondrial transport along the axons. Abnormal mitochondrial trafficking resulting from mutations of nuclear encoded mitochondrial genes, causes severe axonal loss and impaired transport of mitochondria in the most common dominant (CMT2A) (Zuchner et al., 2004) and recessive (CMT4A) (Baxter et al., 2002, Cuesta et al., 2002) axonal forms of CMT. Thus, the mitochondrial abnormalities we have identified in axons of patients with CMT1A may provide important insights into potential mechanisms for axonal degeneration in this demyelinating disorder.
The number of mitochondria is known to increase in the nodal and paranodal regions of axons (Berthold et al., 1993). We recognized that one potential explanation for the observed increased mitochondrial density in CMT1A axons could be the increased mitochondrial number in the nodes/paranodes of dermal myelinated nerves, consequent to the internodal length reduction in CMT1A. However, this is an unlikely explanation for several reasons. First, there were no nodal regions in any of the sections we analysed for mitochondrial density. Second, in several axonal sections that were not in nodal or paranodal regions, there was clearly an increased mitochondrial density in patients with CMT1A. Consistent with this notion, an illustrative case in Fig. 5 shows a remarkable accumulation of intra-axonal mitochondria, but no morphological features of node or paranode. Therefore, this increase of mitochondrial number is independent of any regional effect of nodes/paranodes.
In summary, this study identifies several important features of CMT1A, including differences between CMT1A and CIDP. First, internodal length in CMT1A is shortened in the absence of active segmental demyelination which may, at least partially, explain the uniform slowing of conduction velocity in this disease. This finding also suggests a potential developmental defect in CMT1A during internodal lengthening. While sural biopsies from previous studies showed segmental demyelination in CMT1A, this difference between the sural and dermal myelinated nerve fibres stresses the importance of distinguishing changes observed in dermal and sural biopsies in order to understand better the biological property of dermal nerve Schwann cells or axons that protects them from demyelination. Second, the density of Meissner corpuscles proved to be a useful index of axonal loss in CMT1A. It remains to be determined whether this can be used as a surrogate marker for the progression of axonal degeneration in clinical trials. Finally, we found an abnormal accumulation of intra-axonal mitochondria in dermal myelinated axons of CMT1A, which may shed light on the pathogenesis of axonal degeneration in this disease.
Muscular Dystrophy Association (MDA 4029) and National Institute of Neurological Disorders and Stroke (NINDS) [K08 NS048204].
Supplementary material is available at Brain online.
The authors would like to thank Dr Elior Peles for his generous gift of Caspr antibodies and also Dr Yunhong Bai for her technical assistance.