Charcot-Marie-Tooth (CMT) syndrome is a clinically and genetically heterogeneous group of neuropathies affecting both peripheral motor and sensory nerves. Progressive sensorineural hearing loss, vestibular abnormalities, and dysfunction of other cranial nerves have been described. This is the second case report of otopathology in a patient with CMT syndrome. Molecular genetic testing of DNA obtained at autopsy revealed a missense variant in the MPZ gene (p.Thr65Ala), pathogenic for an autosomal-dominant form of CMT1B. The temporal bones were also prepared for light microscopy by hematoxylin and eosin and Gömöri trichome stains, and immunostaining for anti-myelin protein zero. Pathology was consistent with a myelinopathy of the auditory, vestibular, and facial nerves bilaterally. The pathophysiology of cranial nerve dysfunction in CMT is unknown. Findings in the current case suggested, at least in cranial nerves 7 and 8, that a myelinopathy may be causative.

Charcot-Marie-Tooth (CMT) syndrome is a clinically and genetically heterogeneous group of neuropathies affecting both peripheral motor and sensory nerves, and is also known as hereditary motor and sensory neuropathy (HMSN) or peroneal muscular atrophy. These disorders are caused by genetically induced abnormalities of the myelin sheath or axon of the peripheral nerves. Thus, CMT type 1 (CMT1) is a group of disorders of the myelin sheath of peripheral nerves, the most common of which (CMT1A) is inherited as an autosomal-dominant disorder of peripheral myelin protein 22 (PMP22), whereas CMT1B is inherited as an autosomal-dominant disorder of myelin protein zero (MPZ), of which there are greater than 120 different mutations. In contrast, CMT2 represents a group of disorders causing peripheral neuropathy in which the neuron itself is affected and of which there are several subtypes due to various mutations. CMT3, also known as Dejerine-Sottas syndrome, is a demyelinating progressive neuropathy caused most commonly by a mutation in either the MPZ or PMP22 gene.

Progressive sensorineural hearing loss has been described as part of the clinical presentation in several genetic forms of CMT, including various mutations in the MPZ gene [Chapon et al., 1999; De Jonghe et al., 1999; Alcin et al., 2000; Misu et al., 2000; Starr et al., 2003; Kochanski et al., 2004; Seeman et al., 2004; Leal et al., 2014; Tokuda et al., 2015; Duan et al., 2016], in kinships demonstrating mutations in the peripheral myelin protein (PMP22) gene [Alcin et al., 2000; Boerkoel et al., 2002; Kovach et al., 2002; Joo et al., 2004], in X-linked phosphoribosyl pyrophosphate synthetase-1 (PRPS-1) mutations [Synofzik et al., 2014; Gandia et al., 2015], and in CMT4C [Sivera et al., 2017].

The hearing loss that has been described in the CMT syndrome is progressive, often with poor word recognition scores [Alcin et al., 2000; Starr et al., 2003], consistent with auditory neuropathy. To date, otopathology has been described in 1 other patient with CMT syndrome. That patient had a mutation (Tyr145 Ser) in the MPZ gene [Starr et al., 2003]. The findings included loss of spiral ganglion cells with beading of both peripheral and central auditory neurons; patchy atrophy of the stria vascularis; loss of large myelinated fibers and thinning of the myelin sheath in the proximal auditory nerve; and little change in the auditory and vestibular hair cell populations [Starr et al., 2003].

Vestibular test impairments have also been described in CMT syndrome [Poretti et al., 2013]. In the report by Starr et al. [2003], the sensory epithelium of the vestibular system and the number of cell bodies in Scarpa’s ganglion were reported as normal. In this report, we present the histopathology of the inner ear in a case of CMT syndrome caused by a missense variant (p.Thr65Ala) in the MPZ gene.

This 70-year-old man was diagnosed several years previously with CMT syndrome affecting his hands and feet. There was no family history of a similar disorder. Although audiology was not done, the patient claimed deterioration in hearing on a hospi tal questionnaire 1 year before death. The patient succumbed to chronic obstructive pulmonary disease. Autopsy revealed a peripheral demyelinating neuropathy with “onion bulb” formation involving both motor and sensory fibers consistent with CMT syndrome. Motor and sensory nerve roots showed severe demyelination with only scattered myelinated fibers. The anterior and posterior spinal cord root entry zones showed a transition from complete demyelination in the periphery to normal central myelination (Fig. 1). The white and gray matter of the cord and brain were unremarkable. Electron microscopy of peripheral nerves confirmed nearly complete loss of myelination, and the few remaining myelinated fibers demonstrated no other abnormalities of the myelin sheath.

Fig. 1.

Section through the spinal cord at the level of dorsal rootlets. Myelination of the spinal cord (SC) itself appears normal, whereas that of the dorsal rootlet (DR) shown here is nearly totally devoid of myelinated fibers (MF). Luxol fast blue, HE counter stain.

Fig. 1.

Section through the spinal cord at the level of dorsal rootlets. Myelination of the spinal cord (SC) itself appears normal, whereas that of the dorsal rootlet (DR) shown here is nearly totally devoid of myelinated fibers (MF). Luxol fast blue, HE counter stain.

Close modal

Molecular testing was carried out at the Translational Genomics Core, Partners Health Care Personalized Medicine (Cambridge, MA, USA). DNA was obtained from scalp tissue at autopsy. Analysis revealed a missense variant in the MPZ gene (p.Thr65Ala), identical to that previously reported by Kochanski et al. [2004]. This mutation was considered pathogenic for an autosomal-dominant form of CMT1B. The present case is only the second case of CMT syndrome in which peripheral otopathology is available.

Exome Library Preparation and Sequencing

Exome capture and library construction was performed at the Translational Genomics Core at Partners HealthCare Personalized Medicine using the SureSelect XT Exome V5 kit (Agilent Technologies, Santa Clara, CA, USA) following the manufacturer’s standard protocol. As input, 1 μg of DNA extracted from fresh frozen scalp tissue was used. The Exome V5+ kit uses biotinylated RNA baits to enrich for exons and splice regions of genes, with a total target size of approximately 50 Mb. The kit utilizes standard library construction and hybrid capture methodology (see http://www.agilent.com/cs/library/usermanuals/Public/G7530-90000.pdf). The final exome library was then run on the Illumina HiSeq 2500 instrument in Rapid mode to generate 100-bp paired end sequencing reads.

Postsequencing Processing and Variant Calling

The sequencing yielded roughly 70 million paired reads, which were then aligned to the GRCh37 (hg19) reference genome by using Burrows-Wheeler Aligner followed by SNV and Indel variant detection with the use of the Genome Analysis Toolkit Unified software (GATK v.1.0.4705, Broad Institute, Cambridge, MA, USA). The sequencing yielded an average read depth of 141.52× across the target bases, with 96.7% of target bases having 15× coverage or higher, and a total of 72,356 variants were called. Variants were annotated using an in-house annotation tool that curates variant and gene-specific data from several disease databases, including ClinVar, HGMD, and OMIM, general population frequencies from databases such as dbSNP, ExAC, and 1,000 Genomes, and conservation and computational prediction data (PolyPhen2, SIFT, AlignGVGD).

Histopathology

Both temporal bones were removed and immersed in 10% formalin solution approximately 12 h after death and subsequently decalcified in ethylene diamine tetra-acetic acid. Following decalcification, both specimens were dehydrated in graded alcohols and embedded in celloidin. Serial sections with a thickness of 20 μm were cut in the horizontal (axial) plane, and every tenth section was stained with hematoxylin and eosin and mounted on glass slides. The cochlear duct and Rosenthal’s canal were reconstructed in two dimensions by a method described by Merchant [2010]. Special stains included Gömöri trichrome stain, Luxol fast blue, and immunostaining for anti-MPZ.

Celloidin sections containing the modiolus and cochlea from the subject patient and from a normal-hearing adult human as a control were chosen for immunostaining. Sections were mounted on gelatin subbed glass slides. The celloidin was removed with changes of a solution of sodium hydroxide and methanol followed by 100% methanol rinses. The tissue was rehydrated from 100 to 70% methanol and then distilled water. The sections were rinsed in phosphate-buffered saline (PBS) and incubated for 1 h in 5% normal horse serum. A chicken polyclonal primary antibody against MPZ (AB39375, Abcam, Cambridge, MA, USA) at a dilution of 1: 1,000 was applied and incubated overnight in a humid chamber at room temperature.

The following morning, sections were rinsed in 3 washes of PBS. Secondary antibody (made against chicken) diluted 1: 200 was applied for 1 h, and sections were then rinsed 3 times with PBS. Avidin-Biotin-horseradish peroxidase (standard ABC kit, Vector Labs, Burlingame, CA, USA) was applied for 1 h and rinsed in PBS. Colorization was accomplished with 0.01% diaminobenzidine and 0.01% hydrogen peroxide (H2O2) for 5–10 min. The sections were then rinsed in PBS, dehydrated in graded alcohols, and cover slips were applied.

Exome Sequencing

To identify candidate variants, we employed analysis filtration steps that removed variants with a high minor allele frequency (MAF) in general population databases (MAF ≥0.01) and prioritized variants matching the following criteria: (1) predicted functional impact on coding (missense, nonsense, frameshift), or conserved splice regions (±1–2 splice site positions), (2) variants in 152 genes having an association with CMT disease in the OMIM database (https://www.omim.org). Variants passing these filters were then assessed for pathogenicity based on ACMG guidelines [Richards et al., 2015].

Our filtration and variant prioritization approach identified a heterozygous missense variant in exon 2 of the MPZ gene (c.193A>G; p.Thr65Ala; Fig. 2). The MPZ gene is associated with autosomal-dominant CMT1B disease (MIM No. 118200), and this particular variant was previously reported in a sporadic CMT1B case presenting with early-onset polyneuropathy [Kochanski et al., 2004]. Two different missense variants that impact the same amino acid position, p.Thr65Ile and p.Thr65Asn, have also been reported in CMT-related neuropathy cases [Numakura et al., 2002; Brozková et al., 2010]. All three missense variants at this position were absent in over 200,000 alleles from a broad multiethnic population in the Genome Aggregation Database (http://gnomad.broadinstitute.org/). This evidence suggests that the p.Thr65 position in the MPZ gene is intolerant to variation, and supports a causative role for the p.Thr65Ala variant.

Fig. 2.

Representative sequencing reads from exome sequencing showing a heterogenous T>C transition (red box) on the genomic reference sequence (hg19), which corresponds to a c.193A>G (p.Thr65Ala) missense variant in exon 2 of the MPZ gene.

Fig. 2.

Representative sequencing reads from exome sequencing showing a heterogenous T>C transition (red box) on the genomic reference sequence (hg19), which corresponds to a c.193A>G (p.Thr65Ala) missense variant in exon 2 of the MPZ gene.

Close modal

Histopathology

The findings were similar in both ears and hence will be described together. There was minor scattered loss of both inner and outer auditory hair cells with mild atrophy of the stria vascularis (Fig. 3). The spiral ligament was atrophied in the middle and apical turns, but less so in the basal turn. On the right side, the corrected spiral ganglion cell count for segment 1 was 2,747 cells; for segment 2 was 7,058 cells; for segment 3 was 4,032 cells, and for segment 4 was 2,822 cells. The total corrected spiral ganglion cell count was 16,659, which represented 73% of normal for the age. On the left side, the corrected spiral ganglion count in segment 1 was 2,400 cells; in segment 2 was 6,888 cells; in segment 3 was 3,114 cells, and in segment 4 was 3,869 cells. The total corrected spiral ganglion cell count was 16,271, representing 71% of normal for age-matched controls.

Fig. 3.

a Mid-modiolar section of the left cochlea. The organ of Corti (OC) is present in all three turns of the cochlea. The spiral ganglion cell density (SPG) is reduced in the basal turn. b Basal turn of the right cochlea. Both inner (IHC) and outer hair cells (OHC) are present. The cochlear neurons in the osseous spiral lamina (OSL) are intact. Note the mild loss of cellular elements within the spiral ligament (SL) and the normal stria vascularis (SV). c Cribrose area seen in the mid-modiolar section of the right cochlea. There is deposition of eosinophilic amorphous extracellular material (EM) between the cochlear nerve fibers.

Fig. 3.

a Mid-modiolar section of the left cochlea. The organ of Corti (OC) is present in all three turns of the cochlea. The spiral ganglion cell density (SPG) is reduced in the basal turn. b Basal turn of the right cochlea. Both inner (IHC) and outer hair cells (OHC) are present. The cochlear neurons in the osseous spiral lamina (OSL) are intact. Note the mild loss of cellular elements within the spiral ligament (SL) and the normal stria vascularis (SV). c Cribrose area seen in the mid-modiolar section of the right cochlea. There is deposition of eosinophilic amorphous extracellular material (EM) between the cochlear nerve fibers.

Close modal

There was moderate degeneration of the ampullae of the superior and posterior semicircular canals. The ampullae of the lateral semicircular canals were normal. The maculae sacculi and utriculi were normal. Special stains demonstrated loss of myelinated fibers and increased collagen deposition in the auditory (Figs. 4, 5), vestibular (Fig. 6), and facial (Figs. 7, 8) nerves.

Fig. 4.

Gömöri trichrome stain of auditory nerves in the internal auditory canal. a Distal internal auditory canal near cribrose area of the left ear. Myelin is stained red and connective tissue blue. Only a few remaining myelinated fibers are present within a fibrillar deposit of connective tissue between residual axons. Schwann cell nuclei are also depleted compared to normal. b Control from a normal-hearing adult human. The density of myelinated axons (red) is considerably greater than that seen in a.

Fig. 4.

Gömöri trichrome stain of auditory nerves in the internal auditory canal. a Distal internal auditory canal near cribrose area of the left ear. Myelin is stained red and connective tissue blue. Only a few remaining myelinated fibers are present within a fibrillar deposit of connective tissue between residual axons. Schwann cell nuclei are also depleted compared to normal. b Control from a normal-hearing adult human. The density of myelinated axons (red) is considerably greater than that seen in a.

Close modal
Fig. 5.

Anti-MPZ immunostaining of the proximal auditory nerve in the internal auditory canal. a The myelin is sheath-stained brown. The number of myelinated axons is decreased. The caliber of the residual myelinated neurons and the thickness of the myelin sheath are more varied than normal. b Control from a normal-hearing adult human. The caliber of the myelinated axons is much more uniform than those in a.

Fig. 5.

Anti-MPZ immunostaining of the proximal auditory nerve in the internal auditory canal. a The myelin is sheath-stained brown. The number of myelinated axons is decreased. The caliber of the residual myelinated neurons and the thickness of the myelin sheath are more varied than normal. b Control from a normal-hearing adult human. The caliber of the myelinated axons is much more uniform than those in a.

Close modal
Fig. 6.

Vestibular neurons in the internal auditory canal (Gömöri trichrome stain). In a, there are only a few remaining myelinated nerve fibers (red) as compared to the control ear (b). The presence of macrophages (arrows, a) suggest ongoing degeneration.

Fig. 6.

Vestibular neurons in the internal auditory canal (Gömöri trichrome stain). In a, there are only a few remaining myelinated nerve fibers (red) as compared to the control ear (b). The presence of macrophages (arrows, a) suggest ongoing degeneration.

Close modal
Fig. 7.

Facial nerve trunk in the left (a) and right (b) temporal bones in the proximal descending segment. There is axonal degeneration with onion bulb formation and interstitial fibrosis (ND) indicative of ongoing demyelination and remyelination bilaterally. HE stain.

Fig. 7.

Facial nerve trunk in the left (a) and right (b) temporal bones in the proximal descending segment. There is axonal degeneration with onion bulb formation and interstitial fibrosis (ND) indicative of ongoing demyelination and remyelination bilaterally. HE stain.

Close modal
Fig. 8.

Gömöri trichrome staining of the facial nerve. There are only a few remaining myelinated fibers staining red (arrows, a) as compared to a normal control from an adult human (b).

Fig. 8.

Gömöri trichrome staining of the facial nerve. There are only a few remaining myelinated fibers staining red (arrows, a) as compared to a normal control from an adult human (b).

Close modal

The MPZ gene encodes myelin protein zero, a major structural adhesion glycoprotein of peripheral myelin expressed in Schwann cells, but not in the central nervous system. Variants impacting its function result in peripheral myelin defects that cause several types of neurological disease with varying severity, including progressive late-onset CMT1B (OMIM 118200), and the more severe Dejerine-Sottas syndrome (OMIM 145900) and congenital hypomyelinating neuropathy (OMIM 605253). The genotype-phenotype correlation is unclear across the different MPZ-related neuropathies; however, the more severe forms appear to result from variants having a dominant-negative mechanism, while variants resulting in a loss-of-function of MPZ protein are associated with milder disease [Inoue et al., 2004; reviewed in Houlden and Reilly, 2006]. Missense variants can result in either type of disease mechanism depending on the position and function of the amino acid residue affected by the variant [Boerkoel et al., 2002].

The family history for our patient was negative, suggesting the variant occurred de novo, but parental testing was unavailable. However, roughly 30–40% of causative MPZ variants occur de novo in sporadic cases of CMT-related neuropathy [Boerkoel et al., 2002; Brozková et al., 2010]. In addition, the previously reported CMT case with the same variant as our patient was also sporadic [Kochanski et al., 2004], and the 2 cases with causative missense variants impacting the same codon as our patient (p.Thr65Ile and p.Thr65Asn) were also sporadic, with de novo occurrence confirmed for 1 of these cases [Numakura et al., 2002; Brozková et al., 2010]. These data suggest that the p.Thr65 position is a mutation hotspot susceptible to variation arising de novo.

This is the second case report of otopathology in a patient with CMT syndrome, here with a mutation in the MPZ gene. In the first case, reported by Starr et al. [2003], a missense mutation in the MPZ gene (p.Tyr45Ser) was inherited in an autosomal-dominant fashion. This case was consistent with CMT1B. Audiology performed 9 years before death showed moderate to severe sensorineural loss and a speech discrimination of 4 and 8% on the right and left sides, respectively. The principal histopathologic correlate of the hearing loss seemed to be a severe loss of spiral ganglion cells, whereas the sensory epithelium of the organ of Corti was nearly normal. This was interpreted as consistent with a neuropathy of the auditory nerve. Both the sural and proximal auditory nerves showed a loss of nerve fibers, which was interpreted as consistent with a primary axonal disorder, and thinning of the myelin sheath and remaining axons consistent with incomplete remyelination.

The current case with a missense mutation in the MPZ gene (Thr65Ala) is also consistent with a CMT1B disorder. Unfortunately, no audiogram was done, although by the patient’s own report a mild hearing loss may have been present at the time of death. Histopathology in this case was consistent with a myelinopathy of the auditory nerve as well as the vestibular and facial nerves bilaterally, similar to the neuropathological findings at autopsy demonstrating severe demyelination of motor and sensory roots and nearly complete demyelination in the periphery compared to near normal central myelination.

Hearing loss has been described in CMT1A (PMP22 gene) [Alcin et al., 2000; Boerkoel et al., 2002; Kovach et al., 2002; Joo et al., 2004], in CMT1B (MPZ gene) [Starr et al., 2003], in CMTX (X-linked PRPS-1 mutations) [Synofzik et al., 2014; Gandia et al., 2015], in CMT4C [Yger et al., 2012; Sivera et al., 2017], and in CMT4B3 [Manole et al., 2017]. Vestibular abnormalities detected by electrophysiology have been described in CMT [Poretti et al., 2013].

In addition to vestibular and auditory cranial nerve involvement, other cranial nerves have been described as being affected in various genetic subtypes of CMT, including electrophysiologic involvement of cranial nerves VII, IX, and XII in CMT1A [Kumagai-Eto et al., 2004], subclinical abnormalities in cranial nerves III and V and unilateral cranial nerve III palsy in CMT1A [Posa et al., 2017], abnormalities in cranial nerves V and VII as detected by enlargement of the cranial foramina using MRI and CT, but without clinical symptoms in CMT1A [Das et al., 2017], involvement of cranial nerves III, V, and VII as detected by radiographic analysis of cranial nerve foramina by CT and MRI in CMT1A [Aho et al., 2004], cranial nerve X involvement in CMT of undefined subtype causing bilateral abductor vocal cord paralysis [Hollinger et al., 1979], electrophysiologic abnormalities of cranial nerves II, VII, and VIII in CMT1A [Triantafyllou et al., 1989], and electrophysiologic abnormalities of cranial nerve VII in CMT1A and in CMT3 [Glocker et al., 1999]. In addition, involvement of cranial nerves III, VIII, and X has been described in CMT1D [Pareyson et al., 2000], vocal cord paresis and dysfunction of cranial nerve III in CMT2C [Chen et al 2010], involvement of cranial nerve VII, IX, and X, in CMT3 [Boerkoel et al., 2001], vocal cord paresis in 8 out of 9 patients, and phrenic nerve dysfunction in 8 out of 8 patients in CMT4A [Sevilla et al., 2008], facial weakness in CMT4B3 [Manole et al., 2017], and involvement of cranial nerves VII, VIII, IX, and X in CMT4C [Gooding et al., 2005; Colomer et al., 2006; Yger et al., 2012].

Despite electrophysiologic or radiologic evidence of cranial nerve involvement, in many cases there was no clinical evidence of dysfunction [Glocker et al., 1999; Kumagai-Eto et al., 2004; Das et al., 2017]. This is consistent with the findings in the current case in which there was no clinical evidence of vestibular or facial dysfunction, despite histologic evidence of a myelinopathy of both vestibular and facial nerves.

The pathophysiology of cranial nerve dysfunction in CMT syndrome as evidenced by either electrophysiologic or clinical abnormalities, or both, is unknown. Findings in the current case suggest that, in at least cranial nerves VII and VIII, a myelinopathy may have been causative.

This work was supported by grant No. 5R01DC000152-35, National Institute on Deafness and Other Communication Disorders, Bethesda, MD, USA.

All patients consented to the sharing of clinical information and pathologic findings. The study protocol was approved by the Human Studies Committee under exemption No. 4.

There are no conflicts of interest reported by the authors.

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