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A new mouse model of Charcot-Marie-Tooth 2J neuropathy replicates human axonopathy and suggest alteration in axo-glia communication [1]

['Ghjuvan Ghjacumu Shackleford', 'Department Of Neurology', 'Institute For Myelin', 'Glia Exploration', 'Jacobs School Of Medicine', 'Biomedical Sciences', 'State University Of New York At Buffalo', 'Buffalo', 'New York', 'United States Of America']

Date: 2022-12 

Myelin is essential for rapid nerve impulse propagation and axon protection. Accordingly, defects in myelination or myelin maintenance lead to secondary axonal damage and subsequent degeneration. Studies utilizing genetic (CNPase-, MAG-, and PLP-null mice) and naturally occurring neuropathy models suggest that myelinating glia also support axons independently from myelin. Myelin protein zero (MPZ or P0), which is expressed only by Schwann cells, is critical for myelin formation and maintenance in the peripheral nervous system. Many mutations in MPZ are associated with demyelinating neuropathies (Charcot-Marie-Tooth disease type 1B [CMT1B]). Surprisingly, the substitution of threonine by methionine at position 124 of P0 (P0T124M) causes axonal neuropathy (CMT2J) with little to no myelin damage. This disease provides an excellent paradigm to understand how myelinating glia support axons independently from myelin. To study this, we generated targeted knock-in Mpz T124M mutant mice, a genetically authentic model of T124M-CMT2J neuropathy. Similar to patients, these mice develop axonopathy between 2 and 12 months of age, characterized by impaired motor performance, normal nerve conduction velocities but reduced compound motor action potential amplitudes, and axonal damage with only minor compact myelin modifications. Mechanistically, we detected metabolic changes that could lead to axonal degeneration, and prominent alterations in non-compact myelin domains such as paranodes, Schmidt-Lanterman incisures, and gap junctions, implicated in Schwann cell-axon communication and axonal metabolic support. Finally, we document perturbed mitochondrial size and distribution along Mpz T124M axons suggesting altered axonal transport. Our data suggest that Schwann cells in P0T124M mutant mice cannot provide axons with sufficient trophic support, leading to reduced ATP biosynthesis and axonopathy. In conclusion, the Mpz T124M mouse model faithfully reproduces the human neuropathy and represents a unique tool for identifying the molecular basis for glial support of axons.

Charcot-Marie-Tooth (CMT) neuropathies are a large family of incurable peripheral nerve disorders. Despite extensive clinical and genetic heterogeneity, axonal degeneration is the common end point of all the type of CMT. Thus, a major question is why axons degenerate and how to protect them. Over the years, it has become clear that myelinating glial cells, which are in close contact with axons, are essential for axonal support. However how glial cells support axons remains only partially understood. Here, we generated and characterized an animal model of Charcot-Marie-Tooth 2J (CMT2J), an axonal inherited neuropathy due to mutation in the Myelin Protein Zero (Mpz) gene. Mpz is expressed only in Schwann cells, the peripheral myelin-forming glia, but not in axons, making this the model unique to contribute to our understanding on how glia support axons. Our model reproduces very closely most aspects of the human neuropathy and reveals several alterations in domains crucial for axoglial communication. We also detected metabolic abnormalities in peripheral nerves of these mice that are known to be associated with axonal degeneration. Our work sheds light on the cellular and molecular mechanisms of axoglial communication and axonal degeneration, with implications for a plethora of neurodegenerative diseases.

Axonal degeneration is a common endpoint of peripheral neuropathies [ 16 ] and understanding the glial processes that contribute to axon protection and degeneration is considered the key to cure neuropathies. Yet, this topic is poorly understood. Axonal degeneration is uncoupled from demyelination in CMT2J, providing a unique opportunity to understand how myelinating glia support axons independently of myelin. We generated an authentic mouse model of CMT2J carrying the P0T124M mutation, representing, to the best of our knowledge, the first animal model for CMT2J disease and the first model of human axonal neuropathy caused by a mutation in a protein of compact myelin. Our findings show that the Mpz T124M mouse model closely recapitulates the axonopathy and clinical aspects observed in CMT2J patients. Alterations in non-compact myelin, metabolic changes, and mitochondria dysfunction in Mpz T124M mutants suggest that axonal loss is the result of a defect in SC-to-axon communication. Our results highlight potential mechanisms of how a mutation in P0 causes axonal degeneration without demyelination and underlie the fundamental role of SC in axonal support.

Charcot-Marie-Tooth (CMT) neuropathies are the most common inherited neurological disorders, affecting 1/2500 people [ 5 ]. CMT neuropathies are caused by alterations in over 80 genes encompassing approximately 1,000 independent mutations [ 6 ]. Based on nerve conduction velocity (NCV), CMT are mainly classified as demyelinating CMT type 1 (CMT1) (NCV<38 m/s) or axonal CMT type 2 (CMT2) (NCV>38 m/s) hereditary neuropathies. CMT1B, represents a special challenge because it is caused by more than 200 diverse mutations in MPZ, resulting in different toxic gain of function mechanisms and various related clinical phenotypes [ 7 , 8 ]. Initially, the majority of P0 mutations identified were associated with demyelinating neuropathies. CMT1B patients typically present with early onset disease with extremely slow NCV (<10–20 m/s). Surprisingly, since P0 is only expressed in SC but not in neurons, several mutations in P0 [ 9 – 12 ], such as the substitution of threonine by methionine at position 124 (P0T124M), were shown to cause an axonal neuropathy referenced as CMT2J. After around 40 years of age, patients with T124M-CMT2J begin experiencing symptoms, such as lancinating pain and fast-progressive weakness of the lower limbs. NCVs vary widely among T124M-CMT2J patients and can be normal, slightly reduced similar to that seen in CMT2, or as slow as in CMT1 but never as low as in CMT1B. However, the amplitudes of motor evoked potentials are strikingly reduced in patients carrying the P0T124M mutation. Sural nerve biopsy samples exhibit regenerative clusters and a marked reduction of myelinated axons, but with little to no myelin damage. Moreover, T124M-CMT2J patients exhibit early signs of sensory abnormalities, such as pupillary abnormalities and hearing loss [ 13 – 15 ]. Beside its myelin producing function, these observations suggest either that P0 in SC contributes to axonal support or that SC expressing P0T124M produce deleterious signals contributing to axonal degeneration.

In the peripheral nervous system (PNS), Schwann cells (SC) make myelin by wrapping their plasma membranes around axons. Myelin is a multilamellar lipid-rich structure containing a set of proteins (in the PNS: mostly myelin protein zero [MPZ, P0], peripheral myelin protein 22 [PMP22], and myelin basic protein [MBP]) essential for its compaction. Among these structural proteins, P0, encoded by the MPZ gene, is the most abundant, accounting for 45% of the total protein expressed in PNS myelin [ 1 ]. P0 is a single-pass transmembrane glycoprotein with an immunoglobulin-like fold in its extracellular domain that is expressed exclusively by SC. Crystallography of P0 and X-ray diffraction analysis of myelin suggest that P0 forms homotetramers interacting in trans to hold together adjacent wraps of myelin membrane [ 2 ]. As evidence of the crucial role of P0 in myelination and myelin compaction [ 3 ], Mpz deficient mice produce few myelin layers that are poorly compacted [ 4 ].

Results

MpzT124M mice display progressive motor defects Using homologous recombination, we engineered a genetically authentic T124M-CMT2J mouse model (S1 Fig). Like CMT2J patients, the mice harboring the P0T124M mutation (referred to as TM mice in figures) exhibit signs of late-onset, progressive peripheral neuropathy. Starting at 6 months of age, and more obviously at 12 months, we noticed limb clasping behavior, clawed hind paws, and Achilles’ tendon retraction (Fig 1A and 1B). These neurological abnormalities were not fully penetrant, reflecting the heterogeneity of symptoms observed in T124M-CMT2J patients, and were more prominent in MpzT124M homozygous (MpzT124M/T124M) mice than in heterozygous (MpzT124M/+) mice. PPT PowerPoint slide

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TIFF original image Download: Fig 1. Clinical impairments in MpzT124M mice. (A and B) External signs of neuropathy in 12-month-old MpzT124M/T124M (TM/TM) mice. (A) Representative picture of clasping behavior. (B) Clawed hind paws and Achilles’ tendon retraction are indicated by arrows. (C to G) Locomotion is impaired in MpzT124M mice. Accelerating rotarod test at 2 (C), 6 (D), and 12 (E) months of age. No differences were observed at 2 months of age. Compared to that of wild-type (WT) mice, the performance of MpzT124M mice was worse at 6 months of age and significantly altered at 12. n (animals) ≥ 4 per genotype. Two-way ANOVA [2 month old: Time: F (1.840, 38.64) = 30.54; P<0.0001, Genotype F (2, 21) = 2.177; P = 0.1382; 6 month old: Time: F (1.723, 43.06) = 19.55, P<0.0001; Genotype: F (2, 25) = 4.412; P = 0.0228; 12 month old: Time: F (1.291, 23.24) = 11.51; P = 0.0013; Genotype: F (2, 18) = 4.590; P = 0.0245] with Tuckey’s post hoc. *p < 0.05: significant difference between WT and MpzT124M/+ (TM/+); $p < 0.05: significant difference between WT and MpzT124M/T124M. (F to H) Beam walking test at 6 months of age. (F) Representative pictures of beam walking test for WT and MpzT124M/T124M mice. Arrow indicates a slip. Quantification of slips (G) and speed (H). n (animals) ≥ 5 per genotype. One-way ANOVA [Slip: F (2, 14) = 3.430; P = 0.0613; Speed: F (2, 14) = 7.839; P = 0.0052]. (I to K) Auditory neuropathy in MpzT124M mice. (I) ABR wave I latency measurements in WT and MpzT124M/T124M mice at 11 months of age. Cochleograms of WT (J) and MpzT124M/T124M (K) mice at 12 months of age. n (animals) ≥ 4 per genotype. (L) Plasmatic neurofilament light (Nfl) concentrations in WT, MpzT124M/+, and MpzT124M/T124M mice at 2 and 12 months of age. Plasma from 4-month-old P0 null mice was used as a positive control. n (animals) ≥ 5 per genotype. One-way ANOVA [2 month old: F (2, 23) = 4.857; P = 0.0174; 12 month old: F (2, 21) = 14.83; P<0.0001]. (M to R) Electrophysiological analysis. Amplitudes of compound muscle action potentials (CMAPs) and nerve conduction velocities (NCVs) were measured at 2 (M and N), 6 (O and P), and 12 (Q and R) months of age. n (nerves) ≥ 6 per genotype. One-way ANOVA [2 month old: amplitude: F (2, 19) = 2.53; P = 0.1058, NCV: F (2, 19) = 2.83; P = 0.0837; 6 month old: amplitude: F (2, 47) = 3.142; P = 0.05; NCV: F (2, 49) = 4.394; P = 0.0176; 12 month old: amplitude: F (2, 31) = 4.608; P = 0.0177; NCV: F (2, 31) = 10.95; P = 0.0003]. *p < 0.05, **p < 0.01, ***p < 0.001 by Tukey’s post hoc tests (G to R) after one-way ANOVA. Graphs indicate means ± SEMs. https://doi.org/10.1371/journal.pgen.1010477.g001 We tested the motor performance of MpzT124M mice in the rotarod and beam walking tests and found no differences from wild-type (WT) mice at 2 months of age (Fig 1C). However, at 6 months of age (Fig 1D), we noticed a trend towards a reduced motor capacity in MpzT124M mice. By 12 months (Fig 1E), MpzT124M mice remained on the accelerating rotarod half as long as WT mice. Results from the beam walking test further demonstrated motor impairments (Fig 1F and S1–S3 Movies). At 6 months of age, MpzT124M mice exhibited more foot slips (Fig 1F and 1G), and MpzT124M/T124M mice were 25% slower than the WT mice in crossing the rod (Fig 1H). Thus, MpzT124M mice develop a peripheral neuropathy and display locomotion impairment.

Hearing loss in MpzT124M mice Hearing loss is one of the most consistently observed symptoms in T124M-CMT2J patients [13,14]. To determine if hearing loss also occurs in our mouse model, we used click stimuli to obtain auditory brainstem evoked potential recordings (ABR). The ABR trace consists of five major peaks or waves occurring in the first 5–7 ms following stimulus onset. Wave I, which has a latency of approximately 1.8 ms at high intensities, reflects neural activity of the auditory nerve and subsequent peaks represent to a first approximation synchronized neural responses from successfully more proximal regions in the auditory brainstem (i.e. cochlear nuclei, superior olive, lateral lemniscus, and inferior colliculus). Because only the distal part of the auditory nerve is myelinated by SC, while more central structures in the CNS are myelinated by oligodendrocytes, we focused our attention on ABR wave I. By 11 months of age, the click-evoked latency of ABR wave I was markedly increased at all stimulus intensities in MpzT124M/T124M mice compared to WT mice (Fig 1I). The increase in latency is suggestive of a neural conduction delay in the auditory nerve similar to that observed in T124M-CMT2J patients. To determine if this latency prolongation was caused by loss of sensory hair cells, we performed a cochleogram assay to assess the percentage loss of outer hair cells (OHC) and inner hair cells (IHC) from the base to the apex of the cochlea. Mean cochleogram revealed losses of OHC and to a lesser extent IHC in the basal half of the cochlea of both MpzT124M/T124M and WT mice. Because there were no significant differences in the magnitude of OHC and IHC lesions between WT and MpzT124M/T124M mice, the longer wave I latencies in MpzT124M/T124M mice compared to WT mice are presumably caused by defect on peripheral auditory nerve fibers (Fig 1J and 1K). Abnormal ABRs are characteristic of the auditory neuropathy observed in late-onset CMT1B patients harboring the P0Y145S mutation [17], patients with PMP22 mutations [18,19], and those with GJB1 (gap junction beta 1) mutations [18,20], suggesting that hearing impairment in those with the P0T124M mutation may be due in part to degeneration of the distal part of the auditory nerve.

Level of plasmatic Neurofilament light is increased in MpzT124M mice Plasmatic neurofilament light (pNfl) is emerging as a biomarker for a variety of neurological diseases and positively correlates with the severity of peripheral neuropathies [21]. Remarkably, pNfl concentrations were increased 2-fold in MpzT124M/T124M mice at 2 months of age and 8-fold at 12 months of age. In MpzT124M/+ mice, pNfl concentrations were trending toward an increase (3-fold) at 12 months of age (Fig 1L). These data suggest that pNfl level could also serve as a biomarker in T124M-CMT2J patients and may correlate with disease progression.

Electrophysiological alterations in MpzT124M mice Electrophysiological analyses showed no differences between MpzT124M and WT mice at 2 months of age (Fig 1M and 1N). At 6 months of age, there was a trend toward reduced compound muscle action potential (CMAP) amplitude in MpzT124M/+ mice and a significantly reduced CMAP amplitude in MpzT124M/T124M mice (Fig 1O) but normal NCVs (Fig 1P). At 12 months, both MpzT124M/T124M and MpzT124M/+ mice exhibited significantly reduced CMAP amplitudes (Fig 1Q), consistent with the observed neuromuscular impairment, but only MpzT124M/T124M mice had significantly lower NCVs (Fig 1R). Altogether, these results show that the P0T124M mutation causes a peripheral neuropathy with negative functional impact on axons before myelin.

P0T124M mutation impedes N-glycosylation A denaturing Western blot revealed that P0T124M has a lower relative molecular weight than the WT protein (P0WT) (23 kDa and 28 kDa, respectively) (Figs S4, 4A and 4B). The methionine residue in place of threonine 124, which is within the acceptor sequence for N-glycosylation, may impede P0 N-glycosylation, resulting in the observed shift in migration. To test this, we removed sugars of N-glycosylation via enzymatic digestion with endoglycosidase H (EndoH) or peptide N-glycosidase F (PNGaseF), which decreased the molecular weight of P0WT to 25 kDa. By contrast, P0T124M was insensitive to digestion by both enzymes, suggesting that P0T124M is not N-glycosylated. However, the lack of glycosylation is not sufficient to explain the observed shift in the molecular weight of P0T124M, as N-glycosylation accounts for only 3 kDa (Fig 4A and 4B). To confirm this experimentally, we transfected an HA-tagged version of P0N122S, a non-glycosylatable mutant into COS-7 cells, and observed that it migrated at a higher relative molecular weight (29 kDa, comparable to deglycosylated P0WT-HA) than HA-tagged P0T124M (27 kDa) (Fig 4C). Therefore, additional modifications, likely independent of N-glycosylation and specific to the T124M mutation, may be responsible for the altered migration of P0T124M. PPT PowerPoint slide

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TIFF original image Download: Fig 4. T124M mutation is responsible for molecular P0 modifications but does not alter P0 trafficking or UPR activation. Western blot analysis of P0 in sciatic nerve lysate from wild-type (WT), MpzT124M/+ (TM/+), and MpzT124M/T124M (TM/TM) mice treated with (cut) or without (uncut) EndoH (A) and PNGase F (B). (C) COS-7 cells were transfected with P0-HA, P0T124M-HA, or P0N122S-HA. Samples were treated with (cut) or without (uncut) PNGase F and blotted with antibodies against HA. β-Tubulin (TUB) was used as a loading control. Asterisks indicate relative molecular weight. Bip, Chop, and Xbp1 spliced mRNAs were measured at postnatal day 10 (D) and postnatal day 30 (E, F, and G) by RT-qPCR TaqMan assay. GAPDH mRNA was used for normalization. ΔS63 sciatic nerve mRNA was used as a positive control. One-way Anova [Bip P10: F (2, 10) = 0.07536, p = 0.9279; Bip P30: F (3, 20) = 20.72, p<0.0001; Chop P30: F (2, 8) = 0.7230, p = 0.5145; Xbp1s P30: F (3, 16) = 9.800, p = 0.0007]. Representative immunofluorescence images of sciatic nerve teased fibers (H) and cross sections (I) at postnatal day 30 and 2 months of age, respectively. Fibers and cross sections were stained for P0 (green), endoplasmic reticulum (ER) (KDEL, red), and DAPI (blue). Note that P0T124M, like P0WT, is not retained in the ER and reaches the myelin sheath (arrows). P0ΔS63 is retained in the ER (arrowheads) and does not reach the myelin sheath. (J) Representative electron micrographs of WT and MpzT124M/T124M myelinating Schwann cell ER at 2 months of age. ER from MpzT124M/T124M SC is not enlarged in comparison to ER from WT SC. Arrowheads indicate ER. Scale bars: 2 μm. n (animals) ≥ 3 per genotype. *p < 0.05, **p < 0.01, ***p < 0.001 by multiple-comparisons Tukey’s post hoc tests after one-way ANOVA. Graphs indicate means ± SEMs. https://doi.org/10.1371/journal.pgen.1010477.g004

Pathogenesis of P0T124M does not involve the unfolded protein response N-Glycosylation occurs in the endoplasmic reticulum and Golgi apparatus, and it is important for proper protein folding and for protein quality control [41]. The lack of N-glycosylation could impede P0T124M trafficking and generate endoplasmic reticulum stress. For example, the unfolded protein response is activated in two CMT1B mouse models: P0ΔS63 [26,42] and P0R98C [43]. However, the levels of mRNA for unfolded protein response markers (binding immunoglobulin protein [Bip], C/EBP homologous protein [Chop], and X-box binding protein 1 spliced [Xbp1s]) were not increased in sciatic nerves from MpzT124M mice (Fig 4D–4G). Moreover, the trafficking of P0T124M was not altered, as the mutant protein was not retained in the endoplasmic reticulum and was able to reach the plasma membrane (Fig 4H and 4I). Consistently, EM analysis did not reveal ER morphology alteration in MpzT124M SC (Fig 4J). These results suggest that the unfolded protein response is not involved in the pathogenesis of T124M-CMT2J disease.

ATP and NAD+ but not glycolysis are decreased in MpzT124M mice Because of the alterations at axon-glia exchange areas, we reasoned that metabolite transport from SC to axons would be disturbed in MpzT124M mice. We studied the steady state levels of key metabolites for SC and axon functions by mass spectrometry in sciatic nerves from 12-month-old mice (Fig 8A–8H). ATP, the main energy source for neurons, decreased by 30% in MpzT124M/T124M mice (Fig 8A). In CNS, glycolytic activity in myelinating glia is fundamental for axonal energetics and survival [52–57]. We wondered if defect in glycolysis could explain ATP decrease in MpzT124M PNS. However, we did not notice significant differences between WT and MpzT124M/T124M mice in the amounts of glucose-6-phosphate (G6P) (Fig 8B), lactate (Fig 8C), or pyruvate (Fig 8D). Interestingly, there was striking and significant reduction (-55%) in the amount of oxidized nicotinamide adenine dinucleotide (NAD+) in sciatic nerves from MpzT124M/T124M mice (Fig 8E) along with a trend toward less reduced NAD (NADH) (Fig 8F). NAD+ and NADH are involved in many cellular and biological functions such as energy metabolism, mitochondrial function, and redox state [58], each of which is important for axonal physiology. Moreover, NAD+ balance is crucial for axonal survival. Cleavage of NAD+ by SARM1 (sterile alpha and toll/interleukin 1 receptor motif-containing 1) into nicotinamide (NAM) and adenosine phosphoribose (ADPR) leads irremediably to axonal degeneration [59–61]. Although ADPR expression appears to be normal in MpzT124M/ nerves (Fig 8G), it is rapidly converted to cyclic ADPR (cADPR), making it difficult to obtain a reliable measurement. However, NAM expression shows a trend toward increased levels in MpzT124M/T124M mice (Fig 8H). This combined with the reduced levels of NAD+ suggest an involvement of SARM1 in the axonal degeneration observed in MpzT124M mice. PPT PowerPoint slide

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TIFF original image Download: Fig 8. Metabolic and axonal mitochondrial impairments in MpzT124M mice. (A to H) Measurements of metabolites involved in axonal energy and degeneration programs. ATP (A), G6P (B), lactate (C), pyruvate (D), NAD+ (E), NADH (F), ADPR (G), and NAM (H) were quantified via liquid chromatography-tandem mass spectrometry of sciatic nerves from 12-month-old mice. n (animals) ≥ 5 per genotype. (I and J) Representative confocal pictures of sciatic nerve teased fibers stained with antibodies against mitochondrial protein HSP60 (red) and axonal marker neurofilament (NF) (green) from mice at 2 (I) and 12 (J) months of age. Arrows indicate swelled mitochondria. Scale bars: 10 μm. High magnification insets show mitochondria. (K) Relative frequency distributions of mitochondrial surface area (μm2) at 2 months of age. Nodal length Surface area was quantified from at least 1,096 mitochondria per animal. Proportions of axons with clustered and enlarged mitochondria at 2 (L) and 12 (M) months of age. One-way ANOVA [clustered mitochondria at 2 month old: F (2,6) = 2.558, p = 0.1573 and at 12MO month old: F (2,7) = 1.812, p = 0.2322; enlarged mitochondria at 12 month old F (2,7) = 14.43, p = 0.0033]. At least 263 axons per genotype were imaged. n (animals) ≥ 3 per genotype. *p < 0.05, **p < 0.01, ***p < 0.001 by two-tailed Student’s t test (A to H) and by multiple-comparisons Tukey’s post hoc tests after one-way ANOVA (K to M). Graphs indicate means ± SEMs. https://doi.org/10.1371/journal.pgen.1010477.g008

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