All experimental animals were housed in the Roswell Park Division of Laboratory Animal Medicine vivarium. Procedures were approved by University at Buffalo’s Animal Care and Use Committee and conducted in accordance with the National Institutes of Health’s Guide for the Care and Use of Laboratory Animals. The following transgenic mice were obtained from The Jackson Laboratory: hM3Dq/mCitrine (JAX stock # 026220) and Sox10-iCreER^T2 (JAX stock #027651). Experimental animals were generated by crossing the hemizygous hM3Dq/mCitrine line with hemizygous Sox10-iCreER^T2 transgenic mice. For simplicity, conditional hM3Dq mice (hM3Dq^+/−, Sox10-iCreER^T2+/−) will be referred to as Sox10-hM3Dq for the remainder of this manuscript. In all the experiments presented in this work, mice of both sexes were used. To induce the expression of hM3Dq receptors in Sox10 expressing cells, Cre activity was induced by tamoxifen starting at postnatal day 2 (P2). Mice were intraperitoneal (IP) injected once a day for 5 consecutive days with 25 mg/kg tamoxifen (Sigma-Aldrich) dissolved in corn oil (Sigma-Aldrich). To activate hM3Dq receptors, P6 Sox10-hM3Dq mice were given either 1 mg/kg clozapine-N-oxide (CNO) (Hellobio), or vehicle (0.9% saline solution, BD) via IP injections once a day for 10 days. Sciatic nerves were collected at P10 and P15 for analysis. For experiments with adult mice, P50 Sox10-hM3Dq mice were injected intraperitoneally once a day for 5 consecutive days with tamoxifen (100 mg/kg). Then, at P60 these animals were given IP injections of 1 mg/kg CNO or vehicle (0.9% saline solution) once a day for 10 consecutive days and sciatic nerves were collected at P70, P80 and P90.

The DRGs were harvested from Sox10-hM3Dq mouse embryos at embryonic day 13.5. For each embryo, 20 DRGs were dissected and placed into 1 mL of Leibovitz’s L-15 medium (Invitrogen). The tissues were centrifuged at 200 × g for 5 min and resuspended in 500 μL of 0.25% trypsin solution lacking EDTA (Invitrogen). Enzymatic digestion was carried out at 37 °C for 45 min. To halt trypsin activity, 500 μL of L-15 medium supplemented with 10% fetal bovine serum (FBS; Invitrogen) was added. The supernatant was discarded, and the cells were washed once more with 1 mL of L-15 medium containing 10% FBS, followed by another centrifugation at 200 × g for 5 min. The resulting cell pellet was gently triturated and resuspended in C-medium consisting of Minimum Essential Medium (MEM; Invitrogen) supplemented with 4 g/L D-glucose, 10% FBS, 2 mM L-glutamine, 50 ng/mL nerve growth factor (NGF), and gentamycin. A 150 μL aliquot of the cell suspension was seeded at the center of a 15 mm collagen and poly-L-lysine (PLL)-coated coverslip. Cultures were incubated overnight at 37 °C in 5% CO₂. The following day, the medium was replaced with Neurobasal medium (Invitrogen) containing B27 supplement (Invitrogen), 2 mM L-glutamine, 50 ng/mL NGF, and gentamycin. After 5 days in culture, the cells were transitioned back to C-medium, which was subsequently refreshed every other day. To induce Cre-mediated recombination, cultures were treated with 0.5 μM 4-hydroxytamoxifen (Sigma-Aldrich) for two consecutive days beginning on day 5 post-plating. To promote SC maturation and myelination, cultures were supplemented with 50 μg/mL ascorbic acid (Sigma-Aldrich) starting on day 7 post-plating. In parallel, cultures received either vehicle (0.9% saline solution) or 10 μM CNO for 14 days to activate the hM3Dq receptor.

The DRG co-cultures were prepared from Sox10-hM3Dq mouse embryos at embryonic day 13.5, as described above. To induce hM3Dq expression in SCs, cultures were treated with 0.5 μM 4-hydroxytamoxifen for two consecutive days beginning on day 5 post-plating. On day 7, cells were transduced with an adenoviral vector encoding the Ca²⁺ indicator GCaMP6m (Ad-GCaMP6m; Vector Biolabs) at a multiplicity of infection (MOI) of 25 for 24 h. Following transduction, cultures were treated for 3 days with either vehicle (0.9% saline solution) or 10 μM CNO to activate hM3Dq signaling. Prior to imaging, cells were gently washed with sterile PBS and incubated in phenol red-free Hanks’ Balanced Salt Solution (HBSS; Gibco) containing Ca²⁺ and Mg²⁺, or in HBSS devoid of Ca²⁺ (Gibco). Ca²⁺ transients and resting Ca²⁺ levels were quantified at the single-cell level within SC somas, with data pooled from three independent biological replicates per condition. For each replicate, more than 100 cells were manually selected for analysis. GCaMP fluorescence was excited at 480 nm using a high-speed wavelength-switching system (Lambda DG4; Sutter Instruments). Emission was captured via a spinning disk confocal inverted microscope (Olympus IX83-DSU) equipped with a CCD camera (Hamamatsu ORCA-R2). Image acquisition and analysis were performed using MetaFluor software (Molecular Devices). To minimize photobleaching, both the excitation light intensity and sampling frequency were optimized for low exposure. Fluorescence measurements were acquired every 2 s for a total duration of 480 s (8 min). At 200 s into the recording, cells were stimulated with one of the following agents: ATP (100 μM), glutamate (100 μM), potassium chloride (K⁺ 50 mM), or acetylcholine (100 μM) (Sigma-Aldrich).

Cells were rinsed with PBS and fixed with 4% paraformaldehyde (PFA) for 20 min at room temperature. Permeabilization was carried out using methanol for 10 min at room temperature. Immunostaining was performed following the procedure described by Cheli et al. (2023), with minor modifications. Briefly, fixed cells were incubated in blocking solution containing 0.1% Triton X-100, 1.5% bovine serum albumin (BSA), and 5% goat serum, followed by overnight incubation at 4 °C with the primary antibodies. Secondary antibody incubation was performed using fluorophore-conjugated antibodies (1:600; Jackson ImmunoResearch Laboratories). Nuclear staining was conducted using DAPI (Invitrogen), and samples were mounted with Aquamount (Lerner Laboratories). Fluorescent imaging was carried out using a spinning disc confocal microscope (Olympus IX83-DSU). Myelin internodes and marker-positive cells were quantified in at least three independent biological replicates per condition, with 10 randomly selected fields analyzed per replicate. Semi-automated cell quantification was performed using MetaMorph software (Molecular Devices). Quantification of myelin internodes was performed using ImageJ Fiji version 1.54p. Images were pre-processed using a Gaussian blur filter to reduce background noise, followed by thresholding to identify internodes. Objects smaller than 50 μm² or with circularity more than 0.3 were excluded to eliminate nonspecific staining. Overlapping internodes that could not be accurately segmented were also excluded from analysis. DAPI staining was used to ensure the total number of cells was equivalent across experimental conditions. Primary antibodies: Ki67 (rat; 1:500; BD Biosciences), MAG (mouse; 1:1000; Abcam), MBP (mouse; 1:1000; Covance), P0 (chicken; 1:3000; Aves), Sox2 (mouse; 1:500; R&D Systems), Sox9 (rabbit; 1:500; Cell Signaling Technology), and Sox10 (rabbit; 1:1500; Cell Signaling Technology).

Sciatic nerves were dissected from mice and placed in tubes containing 4% paraformaldehyde (PFA) in PBS for overnight fixation at 4 °C. Longitudinal cryosections (10 μm thickness) were prepared using a clinical cryostat (Leica Microsystems) and mounted onto Superfrost Plus microscope slides (Thermo Fisher Scientific). Tissue sections were incubated in blocking solution containing 2% Triton X-100, 1.5% bovine serum albumin (BSA), and 5% goat serum for 2 h at room temperature. Primary antibody incubation followed overnight at 4 °C. The next day, sections were rinsed with PBS and incubated with the corresponding secondary antibodies (1:400; Jackson ImmunoResearch Laboratories) for 2 h at room temperature. Nuclear staining was performed using DAPI (Invitrogen), and slides were mounted with Aquamount (Lerner Laboratories) following final PBS washes and gentle air drying. The primary antibodies used for immunohistochemistry included: caspase-3 (rabbit, 1:500; Cell Signaling), Ki67 (rat, 1:500; Invitrogen), Krox20 (rabbit, 1:500; kindly provided by Dr. Dies Meijer, University of Edinburgh), MAG (mouse, 1:500; Abcam), MBP (mouse, 1:1000; Covance), P0 (chicken, 1:3000; Aves), Sox2 (rabbit, 1:500; Millipore), and Sox9 (rabbit, 1:500; Cell Signaling Technology). Quantification of myelin protein immunoreactivity and marker-positive cells was performed using MetaMorph software (Molecular Devices). Integrated fluorescence intensity was calculated as the product of the mean pixel intensity and the stained area. To ensure consistent detection thresholds and avoid inclusion of low-intensity background pixels, the same fluorescence threshold was applied to all sections from each nerve. Background fluorescence was subtracted for all imaging channels prior to quantification. Analyses were conducted on pooled data from a minimum of four sciatic nerves per experimental group. For each nerve, at least 12 tissue sections were analyzed using a stereological sampling strategy. DAPI-positive nuclei were used to determine the total number of cells per field, which served as the denominator for calculating the percentage of marker-positive cells.

Coordinated motor performance was assessed using a rotarod apparatus following the standardized EMPReSS protocol (European Mouse Phenotyping Resource of Standardized Screens). Mice were acclimated to the testing room for at least 30 min prior to testing to minimize stress-related variability. Each animal was placed on a rotating rod under two testing protocols: (1) an accelerating protocol in which the rotation speed increased from 5 rpm to 40 rpm over a 5-min interval, and (2) a constant-speed protocol in which the rod rotated at 20 rpm for a maximum of 10 min. The latency to fall (time until the mouse dropped from the rod) was recorded for each trial using an automated timer integrated into the apparatus. To ensure reliability, each mouse completed three consecutive trials per protocol, spaced 20 min apart to allow recovery and reduce fatigue. Between trials, mice were returned to their home cages. A minimum of eight mice per experimental group were evaluated, and all testing was performed during the light phase under controlled environmental conditions (temperature, humidity, and noise). The apparatus was cleaned with 70% ethanol between trials to eliminate olfactory cues, and experimenters were blinded to group allocation to minimize bias.

Sciatic nerves were homogenized in lysis buffer containing 50 mM Tris–HCl (pH 8.0), 150 mM NaCl, 1% (w/v) Triton X-100, 0.5% (w/v) sodium deoxycholate, 0.1% (w/v) SDS, 1 mM PMSF, 1 mM NaF, 1 mM sodium orthovanadate, 1 mM AEBSF, and protease inhibitors including aprotinin (10 μg/mL), leupeptin (10 μg/mL), and pepstatin (10 μg/mL). Total protein concentrations were quantified using a bicinchoninic acid (BCA) assay. Equal amounts (20 μg) of protein were loaded onto NuPAGE Novex 4–12% Bis-Tris gels (Invitrogen) and transferred to polyvinylidene difluoride (PVDF) membranes. Membranes were blocked for 2 h at room temperature in PBS containing 5% non-fat milk and 0.2% Tween-20, followed by overnight incubation at 4 °C with primary antibodies. Detection was performed using horseradish peroxidase-conjugated secondary antibodies (GE Healthcare) and enhanced chemiluminescence (ECL; GE Healthcare). Fluorescent bands were imaged using a C-Digit Blot Scanner (LI-COR) and quantified using Image Studio Software (LI-COR). Primary antibodies: CNP (mouse, 1:3000; NeoMarkers), MBP (mouse, 1:1000; Covance), and α -tubulin (mouse; 1:10,000; Proteintech).

Sciatic nerves were dissected from mice and fixed in 2% glutaraldehyde overnight. Samples were then processed for resin embedding, sectioned into ultrathin slices, and stained with uranyl acetate and lead citrate. Imaging was performed using a Tecnai F20 transmission electron microscope (FEI). Quantitative ultrastructural analysis included measurements of the g-ratio and the percentage of myelinated axons. For each nerve, a minimum of 200 individual fibers and approximately 1,000 axons were evaluated. Image analysis was conducted using MetaMorph software (Molecular Devices) in a semi-automated manner. In addition, 100 randomly selected Remak bundles per nerve were classified based on established criteria (Feltri et al., 2016). At least six nerves per experimental group were included in the analysis.

For single between-group comparisons, nested unpaired Student’s t-tests were performed using a 95% confidence interval. Multiple group comparisons were analyzed using nested one-way ANOVA followed by Bonferroni’s post hoc test to identify pairwise differences. All statistical analyses were conducted using GraphPad Prism software. Statistical significance was defined as p < 0.05 (two-tailed). Data are presented as mean ± SEM. Morphological and biochemical endpoints were assessed using a minimum of four animals per condition, with at least 12 slices per animal. Behavioral assays included a minimum of eight animals per group.

To target hM3Dq receptor expression specifically in Schwann cells (SCs), transgenic mice were generated by crossing hemizygous hM3Dq mice with hemizygous Sox10-iCreER^T2 mice. The Sox10-iCreER^T2 line expresses a tamoxifen-inducible Cre recombinase under the control of the Sox10 promoter, restricting Cre activity to SCs within the peripheral nervous system (PNS) (McKenzie et al., 2014). For clarity, conditional hM3Dq mice (hM3Dq^+/−, Sox10-iCreER^{T2 +/−}) will hereafter be referred to as Sox10-hM3Dq. Initial in vitro experiments were conducted using dorsal root ganglion (DRG) co-cultures prepared from Sox10-hM3Dq embryos. To determine how hM3Dq activation influences SC Ca²⁺ dynamics, DRG co-cultures were transduced with an adenoviral vector encoding the Ca²⁺ indicator GCaMP6m. Changes in intracellular Ca²⁺ within SC somas were monitored by measuring GCaMP6m fluorescence intensity (dF/F₀). Acute CNO application triggered robust Ca²⁺ transients in approximately 98% of SCs, with response kinetics consistent with classical intracellular Ca²⁺ release patterns (Figures 1A,C). Immunostaining further revealed that ~97% of Sox10-expressing cells in DRG co-cultures were positive for hM3Dq and displayed the characteristic morphology of immature SCs (Figure 1B). These findings confirm high recombination efficiency and functional expression of the hM3Dq receptor in SCs.

To evaluate the effects of prolonged hM3Dq activation on SC Ca²⁺ dynamics, DRG cultures were treated with CNO for three consecutive days. Compared to vehicle-treated controls, CNO-treated SCs exhibited a significant elevation in resting Ca²⁺ levels, indicating persistent hM3Dq activation (Figure 1D). Spontaneous Ca²⁺ oscillatory activity was monitored over a 10-min recording period in the same cells. In CNO-treated SCs, the frequency of spontaneous Ca²⁺ transients was reduced, whereas their amplitude was markedly increased, suggesting enhanced mobilization of intracellular Ca²⁺ stores or altered channel dynamics (Figure 1D). To further investigate channel and receptor function, SCs were stimulated with high K⁺ (to induce membrane depolarization), glutamate, ATP, or acetylcholine. In the presence of extracellular Ca²⁺, these assays measured both Ca²⁺ influx through plasma membrane channels and Ca²⁺ release from internal stores. Under these conditions, CNO-treated SCs showed a significantly smaller Ca²⁺ increase compared to controls (Figures 1E,F). In contrast, when experiments were performed in Ca²⁺-free media—where only Ca²⁺ release from internal stores is assessed—no differences were observed between groups (Figure 1F). These results indicate that hM3Dq activation primarily impairs ionotropic Ca²⁺ channel activity rather than metabotropic receptor signaling. Collectively, these findings demonstrate that hM3Dq activation elevates basal Ca²⁺ levels and increases the amplitude of spontaneous Ca²⁺ oscillations, while concurrently suppressing Ca²⁺ influx mediated by voltage- and ligand-gated channels in SCs.

We next examined how sustained hM3Dq activation affects SC differentiation and myelination. To model this, DRG co-cultures were treated with ascorbic acid to induce myelination and simultaneously exposed to CNO for 2 weeks, ensuring continuous hM3Dq activation during the differentiation period. Myelination was assessed by quantifying the number and total area of internodes positive for MAG, MBP, and P0—key structural components of the myelin sheath. SC identity and maturation were evaluated using Sox2, Sox9, and Sox10 immunostaining. These transcription factors were chosen for their distinct roles in SC biology: Sox10 is essential for lineage specification and initiation of myelination; Sox9 is expressed throughout development and persists in mature SCs, serving as a marker of overall SC density; and Sox2 marks immature SCs and acts as a negative regulator of myelination when its expression is sustained (Monk et al., 2015). Two-week CNO treatment nearly abolished internode formation, markedly reducing both the number and coverage area of MAG-, MBP-, and P0-positive segments (Figures 2A,B). In parallel, the proportion of Sox9- and Sox10-expressing cells decreased significantly, while Sox2-positive SCs remained unchanged, indicating a failure to progress toward mature SC identity (Figures 2A,B). SC proliferation, assessed by Ki67 staining within Sox9-positive cells, was unaffected, and no evidence of cell death—such as pyknotic nuclei—was observed in either the Sox10- or Sox9-positive cell populations (Figures 2A,B). Nuclear density (DAPI) also remained consistent across conditions (Figure 2A). Collectively, these findings demonstrate that sustained hM3Dq activation strongly inhibits SC maturation and myelination during in vitro differentiation, without impacting cell proliferation or survival.

To investigate SC development in vivo, Sox10-hM3Dq mice received tamoxifen injections from postnatal day (P) 2–6 and were treated daily with CNO for 10 days starting at P6 (Figure 3A). CNO is an inert compound that activates only DREADDs receptors. Importantly, we have previously reported that CNO administration does not affect CNS myelination in the absence of hM3Dq expression (Cheli et al., 2026). At P10, immunohistochemical analysis of sciatic nerves revealed a significant reduction in myelination in CNO-treated animals compared to vehicle controls, as assessed by myelin protein expression (MAG, MBP, P0) and stereological measurements (Figures 3B,C). Both the signal area—defined as the spatial extent of pixels exceeding a specified threshold—and fluorescence intensity—calculated as the product of mean pixel intensity and stained area (reflecting integrated signal)—were quantified. On average, the signal area of myelin proteins was reduced by approximately 40%, while fluorescence intensity decreased by 25% relative to vehicle-treated nerves (Figures 3B,C). Further analysis revealed a reduction in Sox2-positive cells in CNO-treated nerves, whereas the numbers of Sox9-positive cells and Sox9/Ki67 double-positive cells remained unchanged (Figures 3B,C). Consistent with impaired differentiation, the percentage of Krox20-positive SCs—a transcription factor essential for myelinating SC maturation (Monk et al., 2015)—was markedly reduced. To determine whether these changes were accompanied by cell loss, we assessed apoptotic cell death. No differences were detected in the percentage of active caspase-3-positive cells in CNO-treated nerves (Figure 3C). Total DAPI-positive cell counts per field/nerve section were likewise unchanged (data not shown), indicating that overall SC numbers were not affected by hM3Dq activation. Together—and consistent with our in vitro findings—these results demonstrate that hM3Dq activation disrupts SC progression toward a myelinating phenotype without altering proliferation or inducing cell death.

To validate the immunohistochemical observations, we examined myelin ultrastructure using transmission electron microscopy, which revealed pronounced deficits in CNO-treated mice. At both P10 and P15 (Figure 4A), sciatic nerves exhibited significantly thinner myelin sheaths—reflected by an increased g-ratio—and a reduced number of myelinated axons compared to vehicle-treated controls (Figures 4B–G). Myelin thinning was observed across axons of all calibers, indicating a global impairment rather than a size-specific effect. The proportion of myelinated axons was approximately 10% lower in CNO-treated nerves at both time points (Figures 4E,F), and the mean diameter of myelinated axons was significantly smaller, suggesting delayed myelination (Figure 4G). At the molecular level, Western blot analysis of sciatic nerve extracts at P10 confirmed reduced expression of key myelin proteins, including MBP and 2′,3′-cyclic-nucleotide 3′-phosphodiesterase (CNP), in CNO-treated mice (Figure 4H). Together, these findings demonstrate that sustained hM3Dq activation in SCs profoundly impairs myelination in vivo, resulting in thinner myelin sheaths, fewer myelinated axons, and decreased expression of essential myelin components.

Remak bundles are specialized structures in peripheral nerves where non-myelinating SCs ensheath groups of small-caliber axons, providing trophic support and maintaining axonal integrity. These bundles are essential for proper nerve function, and their abundance and morphology can serve as indicators of developmental progression or pathology. An increase in unmyelinated axon bundles often reflects delayed or impaired myelination (Feltri et al., 2016). In our study, both the total number and the area occupied by unmyelinated axon bundles were significantly higher in CNO-treated mice at P10 and P15 compared to vehicle controls (Figures 5A,B). At both time points, bundle numbers increased by approximately 50%, and the area occupied by these axon groups rose from ~18% at P10 to ~24% at P15, whereas control nerves consistently showed ~10% (Figures 5A,B). Moreover, degenerated Remak bundles—characterized by disorganized axons, axonal swelling or fragmentation, and vacuolization—were markedly more prevalent in CNO-treated nerves. Quantitative analysis revealed that over 75% of bundles in CNO-treated mice exhibited abnormal morphology at both time points (Figures 5A,B).

To further characterize bundle organization, we applied the classification system described by Feltri et al. (2016) (Figure 5C). CNO-treated mice at P10 and P15 exhibited a marked increase in the proportion of early-stage bundles, accompanied by a pronounced reduction in late-stage and morphologically normal bundles (Figure 5C). Although polyaxonal bundles were not observed, we detected a substantial rise in bundles lacking associated SCs in CNO-treated nerves (Figure 5C). Together, these findings indicate that sustained hM3Dq activation delays SC maturation, leading to impaired myelination and abnormal Remak bundle morphology.

To investigate the effects of hM3Dq activation in the adult PNS, Sox10-hM3Dq mice were administered tamoxifen at postnatal day 50 (P50) and subsequently treated with CNO daily from P60 to P70 (Figure 6B). Electron microscopy analysis of sciatic nerves at P70, P80, and P90 revealed increased g-ratios and a reduced percentage of myelinated axons in CNO-treated animals (Figures 6A,C–E,H). G-ratio values plotted against axonal diameter indicated that axons of all calibers were affected (Figure 6H). While reductions in myelin thickness were consistent across time points, the decline in the proportion of myelinated axons progressively worsened from P70 to P90, falling below 80% at the latest time point (Figures 6D,E). In parallel, the diameter of myelinated axons was significantly reduced in CNO-treated sciatic nerves (Figure 6G). Signs of axonal degeneration—including axonal swelling, mitochondrial abnormalities, and the presence of autophagic vacuoles—were prominent in CNO-treated mice (Figure 6F). The percentage of abnormal axons increased significantly over time, exceeding 10% by P90 (Figure 6F). Furthermore, CNO-treated mice at P70 and P90 exhibited a significant increase in the number, size, and total area of unmyelinated axon bundles (Figures 7A,B). Notably, over 20% of these bundles displayed abnormal morphology, including axonal swelling, fragmentation, and vacuolization at both time points (Figures 7A,B). Western blot analysis of sciatic nerve lysates at P90 confirmed these findings, showing reduced expression of key myelin proteins including MBP and CNP (Figure 7C). These findings demonstrate that hM3Dq activation in adult SCs induces progressive myelin degradation and neurodegeneration in peripheral nerves.

Consistent with previous results, immunohistochemical analysis revealed a marked reduction in myelin content within the sciatic nerve following CNO treatment (Figures 8A–C). Quantification of myelin-associated proteins—MAG, MBP, and P0—at P90 confirmed significantly decreased myelination in CNO-treated mice compared to vehicle controls (Figures 8B,C). On average, the signal area of myelin proteins was reduced by ~40%, while fluorescence intensity declined by more than 45% relative to controls (Figures 8B,C). Although the proportions of Sox2-positive cells and Sox9/Ki67 double-positive cells were unchanged, we observed a notable reduction in Sox9-positive cells (Figures 8B,C). In addition, the number of Krox20-expressing SCs was markedly decreased, indicating a diminished population of myelinating SCs in mature peripheral nerves (Figures 8B,C). Despite these shifts in SC marker expression, the proportion of active caspase-3–positive cells remained unchanged in CNO-treated nerves (Figure 8C), and the density of DAPI-positive nuclei also remained constant (data not shown). Together, these findings indicate that hM3Dq activation does not increase SC death in adult sciatic nerves.

Because demyelination disrupts ion channel distribution and impairs action potential conduction, leading to motor deficits (Coggan et al., 2010; Devaux and Scherer, 2005), we assessed motor coordination using the rotarod test under constant and accelerating speed protocols (Figure 8D). Vehicle-treated mice maintained stable performance across all time points, whereas CNO-treated mice exhibited a significant reduction in latency to fall at P70, P80, and P90, indicating impaired motor coordination following hM3Dq activation in mature SCs (Figure 8D). Collectively, these results demonstrate that hM3Dq activation in adult peripheral nerves disrupts SC homeostasis, compromises myelin maintenance, and leads to functional motor deficits.