Acute invasive direct current stimulation of the lumbar spinal cord does not alter spinal motoneurons' firing characteristics in SOD-1 G93A mouse model of ALS.

Acute invasive direct current stimulation of the lumbar spinal cord does not alter spinal motoneurons' firing characteristics in SOD-1 G93A mouse model of ALS. | Authorea try { document.documentElement.classList.add('js'); } catch (e) { } var _gaq = _gaq || []; _gaq.push(['_setAccount', 'G-8VDV14Y67G']); _gaq.push(['_trackPageview']); (function() { var ga = document.createElement('script'); ga.type = 'text/javascript'; ga.async = true; ga.src = ('https:' == document.location.protocol ? 'https://ssl' : 'http://www') + '.google-analytics.com/ga.js'; var s = document.getElementsByTagName('script')[0]; s.parentNode.insertBefore(ga, s); })(); Skip to main content Preprints Collections Wiley Open Research IET Open Research Ecological Society of Japan All Collections About About Authorea FAQs Contact Us Quick Search anywhere Search for preprint articles, keywords, etc. Search Search ADVANCED SEARCH SCROLL European Journal of Neuroscience This is a preprint and has not been peer reviewed. Data may be preliminary. 29 August 2025 V1 Latest version Share on Acute invasive direct current stimulation of the lumbar spinal cord does not alter spinal motoneurons' firing characteristics in SOD-1 G93A mouse model of ALS. Authors : Bartosz Wasicki 0009-0008-0521-6851 [email protected] , Piotr Zawistowski , Tomasz Jankowiak , Leonor de Oliveira Pires , Sofia Fernandes , and Marcin Bączyk 0000-0002-2656-9655 Authors Info & Affiliations https://doi.org/10.22541/au.175648066.61520496/v1 233 views 149 downloads Contents Abstract Information & Authors Metrics & Citations View Options References Figures Tables Media Share Abstract In Amyotrophic Lateral Sclerosis (ALS), alterations of spinal motoneurons’ excitability form the hallmark of their degeneration. Trans spinal direct current stimulation (tsDCS) is based on the delivery of low-intensity DCS to the spinal column in order to alter spinal circuit excitability. Recently, this technique was applied to the management of ALS in the SOD1 G93A mice and resulted in a reduction of disease biomarkers and extended mouse survival. While indirect evidence suggests that these effects can be linked to a decrease in MNs excitability following tsDCS, this has never been directly confirmed. Therefore, in this study, we have utilized in vivo sharp intracellular recordings of spinal MN to directly investigate the impact of DCS on MN intrinsic excitability in SOD1 G93A mice. Electrophysiological properties of MNs recorded before DCS were compared to the properties of MNs recorded one hour after DCS application using linear mixed-effect models. We have found that direct DCS application significantly increases MN peak and plateau input resistance (by 31 and 35% respectively); however, this was not linked to any significant change to MN threshold and firing properties. Both computational modelling and in vivo recordings of the EF field indicate that our results may be explained by the low density of the DC field at the MN recording site. While our results indicate that invasive DCS is not efficient in modifying MN excitability, it may be effective in altering the excitability of afferent fibres traversing the dorsal column close to the DCS electrode. 1. Introduction Amyotrophic lateral sclerosis (ALS) is a fatal neurodegenerative disease characterized by progressive degeneration of upper (cortical) and lower (spinal cord and brainstem) motoneurons (MNs). In the spinal cord, the degeneration depends on the cell subtypes, with MNs innervating fast (group II) muscle fibres and forming fast motor units (FR, FF) being the most affected and deteriorating the most rapidly, while MNs innervating slow muscle fibers and forming slow motor units (S) remain unaffected until the disease end-stage (Pun et al. , 2006; Hegedus et al. , 2008). Interestingly, disease-resistant S MNs exhibit increased excitability in the neonatal SOD1 G93A mouse model of ALS. In contrast, disease-vulnerable FR and FF MNs appear to remain unaffected at this stage (Leroy et al. , 2014) of the disease. Then, at the presymptomatic stage, FR and FF MNs display decreased excitability levels, whereas slow (S) MNs’ excitability is unchanged (Delestrée et al. , 2014; Martínez-Silva et al. , 2018). This excitability-related neurodegeneration pattern gave rise to a radical new hypothesis that increased excitability of S MNs has a protective effect, whereas hypoexcitability of FR and FF MNs contributes to their degeneration in later stages of the disease. Indeed, the works of Saxena et al. (2013) and Bączyk et al. (2020) show that when MN excitability is increased with a chemogenetic approach, the cells display reduced misfolded SOD1 protein accumulation, indicating a potential neuroprotective effect of the treatment. Unfortunately, chemogenetic interventions are not mature enough for translation, and available pharmaceutical modification of MN excitability is subjected to a strong homeostatic response, abolishing its long-term effects (Antonucci et al. , 2024) Recently, a clinical trial utilizing a new non-pharmacological neuromodulation method of trans-spinal direct current stimulation (tsDCS) has been applied to alter ALS progression. TsDCS is based on the external application of weak DC to the spinal cord to modify neuronal activity, and its potential to alter motoneuron and motor unit excitability has already been confirmed in human (Berry et al. , 2017; Yamaguchi et al. , 2020) and animal (Song & Martin, 2022) studies. In the clinical trial NCT04293484, cortico-spinal DCS applied to symptomatic ALS patients slowed down the muscle force decrease and increased the overall patient functional scores (Benussi et al. , 2023). These results may be partially explained by the restoration of intracortical connectivity (Benussi et al. , 2023); however, the spinal component of the observed effects was largely neglected. Indeed, a more recent morphological study showed that anodal tsDCS applied with Multi-Path electrodes reduces misfolded SOD1 protein levels and promotes survival in SOD1 G93A mouse model of the disease (Ahmed et al. , 2025). While indirect evidence suggests that this might be linked with reduced MN excitability following the therapy, this was never directly proven, as no intracellular recordings of MN electrophysiological profiles were performed. Indeed, our recent data show that in the SOD1 G93A mouse model of ALS, the acute application of trans-spinal DCS transiently recovers the deranged MN synaptic excitation levels (Jankowiak et al. , 2022), indicating an elevated MN excitability. This does not, however, neglect the possibility of overall reduced MN excitability following anodal tsDCS, as MN firing characteristics are shaped by both synaptic inputs and intrinsic excitability levels (Burke, 2013), and reduced intrinsic MN excitability can well counteract the increased synaptic excitation. Therefore, in this study, we have applied anodal trans-spinal DCS to SOD1 G93A animals in a similar arrangement as Jankowiak et al. (2022), and we have implemented in vivo intracellular recording technique to investigate the impact of our intervention on MNs’ passive membrane properties, intrinsic excitability levels, and firing pattern. An invasive trans-spinal DCS application approach was chosen in order to bypass the potential diffusion of the current caused by skin and fat tissue, and furthermore to eliminate the confounding effects of the activation of the skin afferents, which can occur with the classical tsDCS approach. This allowed us to precisely determine the DC dose applied to the spinal cord and limit the potential source of the DCS effects to the spinal circuits. We hypothesized that, similarly to the trans-spinal DCS effects observed in WT rats, anodal trans-spinal DCS applied in a dorso-ventral arrangement to SOD1 G93A mice will significantly affect MNs’ intrinsic excitability and firing characteristics. 2. Materials and methods 2.1. Ethical approval All procedures performed for this study were approved by the Poznań Local Ethical Committee (approval number 47/2020). All authors were appropriately trained in all experimental procedures and had valid permits for working with laboratory animals. 2.2. Animals To estimate the impact of DCS on MNs’ passive membrane and firing properties, 15 B6SJL-Tg(SOD1*G93A)1Gur/J male mice bred at the Wielkopolska Center of Advanced Technologies at the Adam Mickiewicz University (Poznań, Poland) were used. Animals were housed two per cage at the Poznań University of Physical Education Animal Facility (Poznań, Poland) with a 12h/12h reversed light-dark cycle in the room, and unlimited access to food and water. Humidity in the room was set at (55 ± 10) %, while the temperature at (22 ± 2) °C. In vivo intracellular recordings were performed on mice between postnatal day (P) P45 and P55, weighing (23.3 ± 2.1) g. Importantly, at this age, SOD1 animals are still in the presymptomatic stage of the disease, just at the onset of fast-twitch muscle fiber denervation with no major motor symptoms (Hegedus et al., 2008), and with the majority of spinal MNs still present. Therefore, this time point is particularly important for treatment implementation as corrective mechanisms still have time to prevent MN degeneration. 2.3. Surgery for the in vivo intracellular recordings All experiments were performed on SOD1 mice under general anesthesia, similar to those performed by Manuel et al. (2009). First, to prevent salivation and edema, a subcutaneous injection of methylprednisolone (0.05 mg/kg Solu-Medrol; Pfizer, Poland) and atropine (0.20 mg/kg; Polfa, Poland) mix was administered. After 15 minutes, an intraperitoneal injection of an anesthetic mixture containing fentanyl (6.25 µg/ml; Polfa, Poland), midazolam (2.5 mg/ml; Polfa, Poland), and medetomidine (0.125 mg/ml; Cp-Pharma, Poland) at a dose of 10 ml/kg body weight was made. The depth of anesthesia was assessed by the absence of the hind limb withdrawal reflex, which typically disappeared five minutes after the injection. The heart rate was monitored with two subcutaneously inserted electrocardiogram (ECG) needles. An infrared heating lamp and an electric blanket TCAT-2DF; Physiltemp, USA) were used to maintain the animal’s core-body temperature at 37 °C. A tracheotomy was performed to artificially ventilate the animal with pure oxygen (SAR-1000 ventilator; CWE, USA) and to sustain the end-tidal CO 2 level at around 4% (MicroCapstar; CWE, USA). Right and left external jugular veins were catheterized, and used to deliver bolts of additional doses of the anesthetic mix at 1.7 ml/kg body weight every 25 minutes and constant infusion of physiological buffer (4% glucose solution containing 1% NaHCO3 and 14% gelatine; Tetraspan; Braun, Poland, 60 µL/h) to maintain animal physiological balance. Next, the sciatic nerve was dissected from surrounding tissues in preparation for electrical stimulation. The spinal column was immobilized with two pairs of horizontal bars (Cunningham Spinal Adaptor; Stoelting, USA), with the first pair fixed at the Th13/Th12 vertebrae boundary and the second at the L2 vertebra under the transverse processes. Afterward, a laminectomy was performed between Th13 and L1 vertebrae to expose L2-L4 spinal cord segments (SS). Finally, exposed tissues were covered with mineral oil, and the dura mater was removed from exposed spinal segments to allow the microelectrode to be inserted into the spinal cord. 2.4. DCS electrode arrangement and stimulation For this study, a dorso-ventral electrode placement was utilized. The dorsal electrode, a silver ball (1 mm diameter), was positioned directly on the spinal cord (L2 SS), one spinal segment above the MNs recording site (Fig. 1B). A thin layer of saline-soaked gelatin sponge was placed under the electrode to minimize the risk of spinal cord damage from direct contact with the electrode. The second electrode, a crocodile clip, was attached to the skin on the abdomen, diagonally caudal to the exposed recording zone. To enhance electrode contact and ensure effective current conduction, an electroconductive gel (Signa Gel; Parker Laboratories, USA) diluted 4:1 in tap water was applied to the crocodile clip. This electrode arrangement is identical to that previously used by Jankowiak et al. 2022, to modulate Ia synaptic excitation of spinal MNs with DCS and should maximize voltage shift in the motoneuron recording zone. The polarity of the DCS was determined by the dorsal electrode: for anodal DCS, the dorsal electrode was positive, while the crocodile clip served as the negative electrode. During each experiment, recordings from at least three MNs were made before the DCS application. Then, DCS was applied for 15 minutes with a current intensity of 30 µA, and no recordings were made during this period. After that time, DCS was turned off, and MNs’ recordings were again performed during the next 60 minutes (Fig. 1A). 2.5. Recordings In vivo intracellular recordings of the lumbar MNs were performed with glass micropipettes (resistance of 15-25 MΩ with a tip diameter of 1.0-1.5 µm) filled with 2M potassium acetate. All electrodes were compensated for their resistance, capacitance, and electrical offset. The recordings of the MNs’ properties were made with an Axoclamp 900A amplifier (Molecular Devices, USA), connected to a Power1401 interface (CED, UK), operated by Spike2 software (CED, UK). The amplifier system was used in bridge (BRIDGE) or discontinuous current clamp (DCC) mode. In the DCC mode, the switch rate was set at 8 kHz (Manuel, 2021) to avoid under- or overestimations of the MNs’ passive membrane and firing properties. To locate the MN pools, a monopolar silver ball electrode was placed on the exposed spinal column with a reference electrode inserted into the back muscles. Then the Sci nerve was stimulated with constant current pulses of 50 µA for 0.1 ms delivered at 3 Hz (DS4, Digitimer, USA) with the intensity set at 2-fold the threshold value required to recruit the most excitable Ia afferents, and the resulting afferent volley was recorded by the ball electrode (Fig. 1B). The spinal cord mapping was then performed to localize the zone with the highest Ia afferent volley amplitude, corresponding to the entry zones of the afferents within the sciatic nerve. Thereafter, the microelectrode was positioned at the spinal location of the highest afferent signal and descended into the spinal cord gray matter to localize the Sci MNs in the ventral horn as guided by the antidromic field potentials. A successful MN penetration was verified by an antidromic action potential evoked by Sci nerve stimulation (Manuel et al. 2009). To exclude the results contamination by unstable recordings in which spike-generating mechanisms are impaired by current leaks, only MNs with a stable resting membrane potential more hyperpolarized than -50 mV and an overshooting action potential were included in the analysis. FIGURE 1 HERE 2.6. Evaluating MNs’ electrophysiological profile After successfully penetrating and identifying each cell as Sci MN, the following protocol was used to establish the cell’s electrophysiological profile. All protocols were executed in DCC recording mode, with a single exception of membrane time constant (τ m ) measurements performed in the BRIDGE mode. First, the cell’s input resistance was estimated, and for that, three series of hyperpolarizing and depolarizing current pulses (-1 to 1 nA; 500 ms; 0.25 nA step) were injected intracellularly (Fig. 3A-B in results). The evoked voltage deflections at the beginning and end of the pulse were plotted against the injected current, and the slope of each relationship provided the peak (Rin Peak ) and plateau input resistance (Rin Plateau ), respectively. The difference between these two values was used to calculate the sag ratio , reflecting the hyperpolarization-activated current (I h ) activity (Manuel & Zytnicki, 2011). The membrane time constant (τ m ) was determined by analyzing the relaxation of the membrane potential following the injection of short, hyperpolarizing current pulses (−5 nA, 1 ms). T m was calculated by fitting a linear regression to the linear region of the voltage’s logarithmic decay, with the absolute value of the slope representing the membrane time constant. The first equalizing time constant (T 1 ) was estimated by subtracting the membrane time constant from the response, and similarly fitting a linear regression over the linear part (Fig. 3C-D). Electrotonic length (L) was established from the equation: L=𝜋/√(τ m /T 1 - 1), while MN’s capacitance (C) was calculated from the equation C = (τ m x L)/(Rin Peak x tanh(L)) (Manuel et al., 2009). Next, the ramp protocol was applied, during which a depolarizing current with a triangular pattern was slowly (0.5 - 2 nA/s) injected into the MN (Fig. 4A). This approach allowed us to determine several key parameters: the Ion (the current at which the MN starts to generate action potentials); the voltage threshold (the membrane potential at which the motoneuron begins generating action potentials); the resting membrane potential (RMP). When the current was applied, an initial linear membrane depolarization was observed before the initiation of firing. For this phase, a linear regression was plotted, and the slope of this regression provided the ramp input resistance . As the depolarization accelerated, a second phase was observed, and another linear regression was plotted (Fig. 4A). The slope of this regression represented the voltage upswing , reflecting the cationic persistent inward current (PIC) activity. Extending the ramp input resistance slope to its intersection with the voltage threshold allowed us to determine the theoretical Ion , representing the current at which the cell would have just started generating an action potential without the voltage upswing. Then, a difference between theoretical Ion and Ion was calculated to investigate the contribution of the voltage upswing to MN activation. Ramp current injection evokes two firing patterns in motoneurons. At first, the cell generates action potentials with irregular intervals and rapidly increasing frequency. At this stage, mixed-mode oscillations are visible on the voltage trace, indicating action potentials that were not able to fully develop (Manuel et al. , 2009). This initial firing is called the subprimary firing range. As the current intensity increases, the cell switches to the firing regime of gradually increasing frequencies with full-blown spikes and linear firing acceleration. This firing regime is called the primary firing range of firing (Fig. 5B). For both subprimary and primary ranges, the firing-frequency/injected current (F/I) relationship can be plotted, and a linear regression line can be fitted to the linear part of the F/I curve, which reflects the primary firing range. The slope of this regression provides the value of the F/I gain. The first 3 spikes of the subprimary firing range generated just above the voltage threshold, where the activity of potassium currents is the highest, were averaged and used to measure the orthodromic action potential parameters: spike width , after-hyperpolarization (AHP) duration, AHP amplitude, AHP time to peak, and AHP half-decay time (Fig. 5C). Finally, the cell’s rheobase (the lowest intracellular stimulation current intensity evoking an action potential) was verified. For this, 5-second current pulses were injected into the cell, with a 5-second break between each pulse (Fig. 6A). The current intensity was manually increased in 0.1 nA steps until the MN generated at least one action potential within the first 50 ms of the current injection. This protocol also allowed us to examine the spike generation threshold (the voltage at which the MN generated the spike) (Fig. 6A 1 ). The difference between the spike generation threshold and MN’s RMP was also calculated. 2.7. Statistics The acute effects of the DCS in the dorso-ventral electrode arrangement were evaluated by comparing the electrophysiological properties of the MNs recorded before and after the treatment. Due to the high variability of MNs threshold and firing properties in a single animal (Powers & Binder, 2001; Manuel & Zytnicki, 2011), MNs recorded from individual animals were pooled to form the Control group (consisting of MNs recorded before DCS application) and Anodal group (consisting of MNs recorded after DCS application). All the data was analyzed in R-Studio 2024.04.02 (Posit Software, PBC) with adequate libraries. Linear mixed models were used to statistically compare the data. Data was first fitted to an appropriate linear model using the “glmmTMB” package (Brooks et al. , 2024) with the DCS condition (Control, Anodal) set as the fixed effect, while the mouse (Animal) was set as the random effect. To validate the fitted model, a Residual Diagnostics for HierArchical (Multi-level/Mixed) Regression Model was performed using the DHARMa package. A plot of the scaled residuals was created by simulating from the fitted model, and data deviation, dispersion, and variance were assessed. If a significant deviation from the model was found, then the dispersion correction or data family correction was introduced and the model was refitted. Only the models that met all assumptions were used for further analysis. The significance of the fixed effect was established using joint test from the “emmeans” package (Lenth et al., 2024) with Tukey correction. Importantly, this approach allows us to exclude the potential dependence of the DCS effects on the animal and correct for data inflation from recording multiple MNs from a single mouse. The Pearson correlation coefficient was calculated from built-in RStudio functions, to estimate the correlation between voltage deflection and distance from the anode in the rostro-caudal axis. All plots were created using the “ggplot2” package (Wickham, 2016). The significance levels for both fixed effect and correlations was set at p<0.05. 2.8. 3D modelling and computer simulations To confirm the electric field (EF) direction and spatial pattern on the spinal cord, a 3D mouse model of the thoracic-lumbar region was designed, based on MRI scans of a P30 C57/B6 mouse. The model was designed in accordance with the experimental surgical protocol: the Th13 and L1 vertebrae were removed, and the CSF layer in the corresponding region was excluded (Fig. 2). ITK-SNAP (Yushkevich et al. , 2006) was used to segment the relevant anatomical regions. These were converted into 3D surface meshes refined using Blender 4.0 (www.blender.org). Corrected meshes were imported to the 3-Matic module from Mimics (v16, www.materialise.com) to add the spheric anode, placed dorsally on the exposed WM over a 0.25 mm layer representing the saline-soaked gelatin sponge. The crocodile clip was modelled as two rectangular-shaped surfaces over one layer of gel with a thickness of 0.25 mm, positioned ventrally over a skin tissue flap. Surface corrections, such as re-meshing and smoothing, were applied to obtain a non-manifold assembly feasible for volume meshing. The obtained anatomical regions included skin, vertebrae (from Th9 to L6), intervertebral discs, cerebrospinal fluid (CSF), white matter (WM), and gray matter (GM) of the spinal cord (SC), and several visceral organs (stomach, liver, spleen, pancreas, and intestines). The dura mater layer was not included as it was removed during the surgical procedures. The final volume mesh consisted of 1.1x10 7 tetrahedral elements, adequate for applying in simulations using the finite-element method (FEM). A literature review on the electrical properties of biological tissues was performed to compile electrical conductivity values for DC currents, focusing on studies involving mice, rats, and human estimates derived from multispecies data. To ensure consistency, priority was given to measurements obtained at body temperature (37 °C) and within the low-frequency range (<10⁴ Hz). The values listed in Table 1 were previously used in a full-body in silico mouse model and subsequently refined following validation against experimental data. The electrical conductivity of the gel was calculated as the weighted average of four parts Signa Gel (σ = 4.000 S/m; www.parkerlabs.com/products/signagel-electrode-gel/) and one part water (σ = 5.500 × 10⁻⁶ S/m, from the COMSOL Materials Library), yielding an effective conductivity of σ = 3.200 S/m. For the saline-soaked gelatin sponge, a conductivity of 1.45 S/m was used, corresponding to a 0.9% saline solution (Sauerheber & Heinz, 2015). The EF induced in the SC and surrounding tissues was calculated using the AC/DC module of COMSOL Multiphysics software, considering an injected current of 30 μA. EF was calculated as the gradient of the electric potential, assuming tissues as purely resistive (relative permittivity ϵ r = 1), consistent with the quasi-static approximation for DC currents (Miranda et al. , 2013). Simulations used an NVIDIA RTX 2000 Ada GPU, a 13th Gen Intel® Core™ i7-13800H processor, 32 GB RAM, and a 64-bit OS, with 1.5x10 7 degrees of freedom and an approximate solution time of 228 minutes. FIGURE 2 HERE Table 1: Electrical conductivity of the biological tissues represented in the thoracic-lumbar mouse model. Biological Tissue Electrical Conductivity (S/m) Literature Source Skin 0.435 Human (Geddes & Baker, 1967) Fat* 0.040 Human (Gabriel et al. , 1996) Muscle 0.355 Human (Rush et al. , 1963) Bone (vertebrae) 0.012 Rat (Kosterich et al. , 1983) Spinal WM 0.143 Human (Haueisen et al. , 1997) Spinal GM 0.333 Human (Haueisen et al. , 1997) CSF 1.790 Human (Baumann et al. , 1997) Intervertebral discs 0.200 Human (Haueisen et al. , 1997) Liver 0.123 Human (Surowiec et al. , 1987) Stomach 0.200 Human (Haueisen et al. , 1997) Intestines (large and small) 0.200 Human (Haueisen et al. , 1997) Pancreas 0.220 Human (Gabriel et al. , 1996) Spleen 0.100 Human (Gabriel et al. , 1996) Kidneys 0.100 Human (Gabriel et al. , 1996) *Subcutaneous fat and fat surrounding the spine 3. Results In total, 107 motoneurons were intracellularly recorded from 10 animals. The motoneurons were divided into two groups: the Control (n=64) group consisting of MNs recorded before the DCS and the Anodal (n=43) group consisting of MNs recorded within the first hour after the end of DCS. The 1h recording window was selected based on our previous reports showing a rapid decline of the polarization effects starting in the 2nd hour after DCS (Jankowiak et al. 2022). 3.1. Passive membrane properties We first focused our analysis on the MN passive membrane properties as the principal determinants of MN excitability (Gustafsson & Pinter, 1984). Figure 3, shows that 15 min of anodal DCS significantly increased both RIN peak (31.09% increase from (2.856 ± 0.991 MΩ) in Control group to (3.744 ± 1.297 MΩ) in Anodal group, Joint test p<0.001, Fig. 3E) and RIN plateau (35.00% increase from (2.314 ± 0.803 MΩ) in Control to (3.124 ± 1.250 MΩ) in Anodal, Joint test p<0.001, Fig. 3F) input resistance, however it did not change the MN SAG ratio (Joint test p=0.130, Fig. 3G) indicating no change in the underlying Ih current activity (Manuel & Zytnicki, 2011). FIGURE 3 HERE We did not observe any impact of polarization on the MN membrane time constant (Joint test p=0.167, Fig. 3H), first equalizing constant time (Joint test p= 0.355, Fig. 3I), electrotonic length (Joint test p=0.359, Fig. 3J), and electrical capacitance (Joint test p=0.089 Fig. 3K). Finally, there was no significant change in the MN membrane potential at rest (Joint test p=0.860 Fig. 3L). This indicates that anodal DCS only modestly affects the MN passive membrane properties, with the strongest impact on input resistance. 3.2. Threshold properties Having identified a significant effect of DCS on MN input resistance, we then moved to investigate its threshold properties. As MN RIN significantly correlates with cell excitability (Gustafsson & Pinter, 1984), we expected to see an alteration in MN firing profile after anodal DCS. We first analysed the response of MNs to the triangular ramp of depolarizing current (Fig. 4A-G). As expected, the MN input resistance calculated on the ascending part of the ramp (RIN Ramp ) was significantly increased in MN recorded after DCS (23.19% increase from 2.859±1.175 MΩ in Control group to 3.522±1.386 MΩ in Anodal group, Joint test p=0.023, Fig. 4B). To our surprise, this increase in ramp input resistance did not transfer into alterations of MN firing profile. The voltage upswing (Fig. 4C), Ion current (Fig. 4D), as well as the Theoretical Ion-Ion (Fig. 4E), voltage threshold (Fig. 4F), and the difference between voltage threshold and resting membrane potential (Fig. 4G), were not affected by DCS (all Joint test p>0.05). FIGURE 4 HERE 3.3. Firing properties A MN response to a depolarizing ramp of current develops from an initial spike to subprimary and finally the primary firing range (Manuel et al., 2009, Fig. 5A-B). As the F/I relationship is impaired in SOD1 MNs (Delestrée et al. , 2014), we investigated whether DCS can impact this firing scheme. We were unable to find any significant change in the F/I Slope following DCS intervention (Joint test p=0.504, Fig. 5D). This was in line with the lack of changes in the action potential properties analysed from the averaged first three spikes of the subprimary range (Fig. 5C). Indeed, the AHP duration (Joint test p=0.204, Fig. 5E), AHP time to peak (Joint test p=0.393, Fig. 5F), AHP half decay time (Joint test p=0.607, Fig. 5G), AHP amplitude (Joint test p=0.781, Fig. 5H), and spike width (Joint test p=0.212, Fig. 5I), were all not affected by DCS indicating no change to the AP generation mechanism. FIGURE 5 HERE Finally, we analysed the MN response to long depolarizing pulses of current (Fig. 6A, A 1 ), as these were previously found to be more effective in activating the SOD1 MNs in comparison to depolarizing ramps of current (Delestrée et al. , 2014). In line with the lack of changes in Ion and voltage threshold during injection of triangular ramps of current (Fig. 4), we found no significant impact of anodal DCS on the rheobase current (Joint test p=0.650, Fig. 6B), spike generation threshold (Joint test p=0.451, Fig. 6C) or in the difference between spike generation threshold and resting membrane potential (Joint test p=0.362, Fig. 6D). FIGURE 6 HERE Altogether, this data shows that while DCS can alter MN input resistance, this does not transfer to the alterations of threshold and firing properties. 3.4. Electric field The lack of significant impact of anodal DCS on MN firing profile was not expected, in light of the previous findings of Bączyk et al. (2020a, 2020b) showing a strong impact of DCS on MN firing profile in WT rats. Therefore, in order to find the source of this discrepancy, in 5 mice we analysed the geometric constraints of the induced electric field (EF) responsible for the neuromodulatory DCS actions (Fernandes et al. , 2018). Figure 7 shows the change of voltage recorded in the extracellular space of dorsal horns, intermediate zone, and ventral horn of the spinal grey matter when anodal DCS of 30 µA is activated. We have found a strong depolarization of the voltage in the majority of MN recording spinal segments immediately when anodal DCS was switched ON. Importantly, the voltage deflection was significantly correlated with the distance from the dorsal active electrode and a negative slope of the voltage/distance relationship (-1.63 mV/mm in dorsal, -1.00 mV/mm in intermediate, and -0.88 mV/mm in the ventral recording zones, (r(13) = -0.84, p<0.001 for all depths, Fig. 7 A-C), was found in all recording segments indicating a decrease in the EF field density with a increasing distance from the dorsal electrode. Additionally, we identified a significant difference in voltage deflection when the recording zone was moved in the dorso-ventral axis. For the most depolarized spinal segment, the voltage deflection was 81.9% higher in the dorsal than at the ventral horn (6.33 ± 2.66 mV vs. 3.48 ± 1.32 mV, for dorsal and ventral horn respectively, Joint test p = 0.03, Fig. 7D). This indicates that, in our experimental approach, the DCS preferentially targeted neuronal structures located in the dorsal part of the spinal column close to the stimulating electrode, with only modest impact on the ventral horn structures. The electrophysiological measurements were further confirmed by a modelling approach. Simulations of DCS in the mouse spinal cord confirmed that, within the analysed segment (L2-L4 SS), the longitudinal component of the electric field (EF_long) was negative (rostral-caudal oriented) in almost all sampling locations (Fig. 8A). Furthermore, a consistent dorsal–ventral gradient in EF_long magnitude was observed, with absolute dorsal values exceeding ventral ones (Fig 8B). Closer to the anode, in the L2 SS, we observe generally higher magnitude for the negative EF_long values, varying from 6.51 V/m to 39.7 V/m. Further the anode, EF_long magnitude decreases. In L3 SS, the maximum absolute value was 9.99 V/m, and the minimum was 6.80 V/m. In the L4 SS, the maximum absolute EF_long was 6.91 V/m, and the minimum was 4.63 V/m. In all three cross sections, the strongest EF_long are located dorsally, with the highest variance located nearer the anode due to the direct contact with the spinal cord. This observation is in line with the experimental result of preferential targeting of dorsal neuronal structures. Simulated EF_long magnitudes were systematically higher than those recorded experimentally. This outcome was expected, as the mice used in the experimental procedures were P45–55 of age, whereas the MRI scans underlying the computational model were obtained from a P30 mouse. Other two main factors may also likely explain this divergence: the model used literature-based conductivity values that may not fully reflect in vivo mouse tissue properties, and the finite element model represents a generic anatomical geometry that does not capture subject-specific morphological differences. Despite overestimation in absolute magnitude, simulations replicated the directionality and spatial pattern observed experimentally: a negative EF_long component on the analysed segment, with higher dorsal magnitudes. FIGURE 7 HERE FIGURE 8 HERE 4. Discussion In this paper, we provide direct evidence of acute DCS impact on spinal MNs passive membrane and firing properties in pre-symptomatic SOD1 G93A mice. While DCS was found to strongly affect MNs’ input resistance, this was not transferred into alterations of MNs’ threshold or firing characteristics. Our results indicate that direct DCS applied with ball electrodes is not effective in modifying spinal MN firing behaviour. 4.1. Passive membrane properties Fifteen minutes of anodal DCS at 30 µA in a dorso-ventral electrode arrangement significantly increased both peak and plateau input resistance in spinal MNs up to one hour after treatment application. This is largely in line with previous reports of Jankowiak et al. (2022), who also reported a strong increase in peak and plateau RIN when the same MN was recorded before and up to 15 min after DCS. The background of these changes is difficult to unravel, as MN’s input resistance is a result of an interplay between the cell’s membrane area and the number and density of active ion channels (Gustafsson & Pinter, 1984; Powers & Binder, 2001). We can, however, exclude several factors. First, it is unlikely that the changes in RIN can be attributed to a decreased activation of the Ih current (McLarnon, 1995), as the MN SAG ratio (Manuel & Zytnicki, 2011) was not changed by DCS in our MNs sample. Second, the activity of subthreshold persistent sodium currents (PICs) also likely did not affect the RIN as the voltage upswing (reflecting PICs activation (Kuo et al. , 2006)), was unaltered. Thirdly, there were no changes to the passive electrical properties of the MN membrane, including resting membrane potential, membrane time constant (T m ), electrotonic length, and electrical capacitance (Rall, 1969). Therefore, a change in the passive membrane properties was not the source of RIN alterations. At this point, we can only speculate that the increased RIN observed in our investigations results from the inactivation of unspecified ion channels upon application of current pulses during the RIN protocol. 4.2. Firing properties In opposition to our expectations, 15 minutes of anodal DCS failed to alter MNs’ threshold and firing characteristics. This is particularly surprising given the fact that the input resistance strongly correlates with MN excitability (Gustafsson & Pinter, 1984) and was indeed increased by our intervention. Furthermore, previous works of Bączyk et al. (2020a), showed that in WT rats, 15 min of anodal DCS of 100 µA evokes an increase in MN firing frequency and a decrease in rheobase current. The discrepancies between our results and those of Bączyk et al. can be explained by several methodological variables between the two investigations. While using the same dorso-ventral electrode configuration, Bączyk et al. (2020a) used a circle-shaped electrode (5mm diameter) for DCS application, which provided a larger contact area and likely resulted in a differently distributed electric field. In contrast, our study utilized a ball-shaped electrode, which led to the formation of a high current density directly beneath the active electrode, rather than in the ventral horn where MNs are localized. This result was validated by the computational modelling approach, in terms of EF directionality and spatial pattern. Within the L2-L4 SS of the in silico mouse model, we also observe a dorsal–ventral variance of the EF in the longitudinal axis, with negative direction and stronger magnitude values on the dorsal region of the GM, especially on a rostral section, near the stimulating electrode (Figure 8.B). This reasoning aligns with the reports of Gigliotti and Pereira (2025), who emphasized that it is the DC field distribution rather than the delivered current that is the key factor for DCS effectiveness. Nevertheless, the lack of change in firing characteristics in the presence of increased RIN indicates the activation of compensatory mechanisms that prevent the expected increase in cell intrinsic excitability. These may include upregulation of potassium or downregulation of sodium channels’ conductances, impacting several types of membrane currents such as SK (Li & Bennett, 2007), M-type (Sharples et al. , 2023), or L-Ca (Carlin et al. , 2000) currents. 4.3. Electrode arrangement and target area Our original hypothesis of strong DCS impact on MN firing behaviour was largely based on the results of Jankowiak et al., indicating strong alterations of MN synaptic excitation levels following 15 min DCS (Jankowiak et al. , 2022). This result was attributed to changes in the effectiveness of Ia synapse activation on the MN soma by DC application. However, the modelling data presented in this paper indicate that the current density, directly proportional to the EF for DC stimulation, diminishes rapidly with distance from the active electrode. As a result, the current density at the ventral horns (i.e., at the motoneuron recording site) may not be optimal to produce neuromodulatory effects. Indeed, recent works from Jankowska and Bennett group et al. (Li et al. , 2020; Sławińska et al. , 2025) point to the fact that the increase in MN synaptic excitation following DCS may be a result of increased afferent activity related to the activation of extrasynaptic GABA receptors at the Ia afferent branching points in the dorsal column. Indeed, when DC is applied to the dorsal surface of the spinal cord using a thin tungsten electrode, a strong increase in the activity of Ia afferent fibres can be observed (Jankowska et al. , 2017; Kaczmarek & Jankowska, 2018). This effect is linked to depolarization of the nodal GABA receptors, located at the Ia afferent branching points within the dorsal column (Li et al. , 2020), and is likely related to the increased activity of GABA-ergic astrocytes (Sławińska et al. , 2025), similar to the case of transcranial DC application (Monai et al. , 2016). However, this approach appears ineffective in modulating the MN firing properties as studied by intracellular application of stimulating currents. Indeed, with this approach, the afferent activity is excluded from the DC effects and can not impact MN intrinsic excitability. The remaining actions of DC are therefore limited to MN soma and are not potent enough to affect the firing behaviour. 4.4. Practical considerations and limitations In human applications, tsDCS is primarily used in pain management to inhibit the transmission of nociceptive signals (Truini et al. , 2011; Thordstein et al. , 2020; Guidetti et al. , 2021). On the other hand, the studies of tsDCS on motor output have shown both positive (Berry et al. 2017), and negative outcomes (Fava de Lima et al. , 2022). Our results favor the concept of preferential involvement of dorsal horn afferents in the tsDCS observed effects. These results can be further utilized to facilitate therapeutic approaches depending on afferent stimulation, however may have limited potential when ventral horn structures are involved. Still, one should be aware of the limitations of our study. First, this study involved placing a metal ball electrode directly on the surface of the spinal cord. This approach is not orthodox for tsDCS therapy, where DC is applied with larger electrodes located on the skin (Gómez‐Soriano et al. , 2019; Paget-Blanc et al. , 2019; Yamaguchi et al. , 2020). The latter application method may produce less localized current that has a better potential of penetrating the spinal column and therefore affecting the MNs (Yamaguchi et al. , 2020). Indeed, strong polarity-dependent alterations of MN passive membrane and firing properties were found in WT rats when tsDCS was applied with a plate electrode located on the skin above the MN recording site (Bączyk, Drzymała-Celichowska, et al. , 2020b). On the other hand, our approach reflects the spinal cord stimulation (SCS) method, where stimulation electrodes are implanted on the dura to deliver an alternating current (de Vos & Meier, 2024). As our previous results have shown that DCS applied with a ball electrode increases afferent fibre activity (Jankowiak et al. , 2022), this method of DCS could be applied before SCS to reduce afferent activation threshold and reduce the current intensity needed to produce therapeutic effects. Second. In our investigations, we were unable to directly predict the distance of the recorded MN from the active electrode. Although all recorded MNs were located within one spinal segment caudal to the active electrode, it is possible that MNs located closer to the electrode were preferentially affected by DCS. Third. We were unable to predict the spatial orientation of the recorded MN with respect to the induced electric field. It has already been shown by both modelling (Rahman et al. , 2013) and in vitro works (Kabakov et al. , 2012) that DC effects depend on the spatial orientation of neuronal compartments to the electric field. Finally, we can not exclude the possibility that part of the observed DCS effects are related to changes in pH evoked by electrical current. By definition, an anode located within an ionotropic environment will attract negative ions and may result in a negative pH shift close to the electrode, which can affect neuronal activity (Ruffin et al. , 2014). 5. Conclusions 30 µA anodal DCS applied with a ball electrode located directly on the surface of the spinal cord does not modify spinal MN firing properties. The lack of effect is largely explained by a low DC field intensity within the MN recording site. These results indicate that these DCS application parameters are not suitable to provide neuromodulation to neurological conditions where changes of MN intrinsic excitability may be beneficial. Data availability statement All data supporting the results of this study are included in the manuscript. The datasets generated during the current study are available from the corresponding author upon reasonable request. All raw recordings are stored in the private data repository at https://box.pionier.net.pl/, provided by the Poznan Supercomputing and Networking Center affiliated with the Institute of Bioorganic Chemistry of the Polish Academy of Sciences. Access to the raw data can be provided upon request. Funding T. Jankowiak was supported by National Science Center grant no. 2017/26/D/NZ7/00728. P. Zawistowski was supported by National Science Center grants no. 2019/35/B/NZ4/02058 and B. Wasicki was supported by National Science Center grant no. 2022/04/Y/NZ4/00117 under the JPND 2022 funding scheme. M. Bączyk was supported by all three aforementioned grants. S. Fernandes and L. de Oliveira Pires were supported by Fundação para a Ciência e Tecnologia (FCT) under the IBEB Strategic Project UID/00645/2025 and the DC4MND project from JPND, JPND/0003/2022. L. de Oliveira Pires was supported by Fundação para a Ciência e Tecnologia (FCT) under the Bolsa de Investigação para Doutoramento 2024.00602.BD. Author contributions All electrophysiological experiments and data analysis were performed at the Neurobiology Department of Poznan University of Physical Education, and EF modelling was performed at Instituto de Biofísica e Engenharia Biomédica, Faculdade de Ciências da Universidade de Lisboa. Conceptualization: M. Bączyk; methodology: M. Bączyk, S. Fernandes; formal analysis: B. Wasicki, P. Zawistowski, T. Jankowiak, L. de Oliveira Pires; investigation: B. Wasicki, P. Zawistowski, T. Jankowiak, L. de Oliveira Pires, M. Bączyk.; writing the original draft: B. Wasicki, P. Zawistowski, L. de Oliveira Pires, M. Bączyk, S. Fernandes; visualisation: B. Wasicki, P. Zawistowski, L. de Oliveira Pires; supervision: M. Bączyk, S. Fernandes; funding acquisition: M. Bączyk, S. Fernandes. All authors approved the final version of the manuscript. 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Ia afferents and motor axons are activated through electrical stimulation of the peripheral (sciatic) nerve, allowing for MNs identification based on the “all-or-nothing” principle. Figure 2. (A) Lateral view of the thoracic–lumbar mouse in silico model with the cathode (crocodile clip) positioned at the designated site. (B) Lateral view of vertebrae Th9–Th12 and L2–L6, illustrating the anode (silver ball electrode) placed on the exposed white matter of the spinal cord. Figure 3. Acute effects of anodal DCS on MNs’ passive membrane properties. Intracellular in vivo recordings were performed before (Control) and after (Anodal) 15 minutes of anodal DCS in a dorso-ventral electrode setup with the current intensity of 30 µA. ( A, C ) Voltage and current traces recorded during the RIN and τ m recording protocol. The green trace represents the voltage changes, while the blue trace represents the intracellularly injected current. ( B ) To measure a cell’s peak and plateau RIN, the voltage deflection is plotted against the current, and the slope of these linear relationships represents RIN. ( D ) A semilogarithmic plot of the voltage change is created, and a regression line (red) over the linear part is fitted to calculate a cell’s τ m . The first equalizing time (τ 1 ) is estimated by removing the membrane time constant from the response. ( E-L ) data distribution for peak input resistance ( E ), plateau input resistance ( F ), sag ratio ( G ), membrane time constant ( H ), first equalizing time constant ( I ), electrotonic length ( J ), electrical capacitance ( K ), and resting membrane potential ( L ). Note a strong increase in both peak and plateau input resistance after acute anodal DCS. In the graphs, the upper and lower hinges of the boxplot refer to the 75th and 25th percentiles, respectively, and each data point represents a single MN. The significance level was set at p<0.05. *** - p<0.001, ns - not significant. Data from 10 mice. Figure 4. Acute effects of anodal DCS on MNs’ threshold properties. A) An example of an MN’s response to the injection of the triangular ramp of depolarizing current. The green trace represents MN’s membrane potential, while the blue trace represents the intracellularly injected current. The boxed area has been extended , showing the area for electrophysiological properties measurements (see “Materials and methods - Evaluating MNs’ electrophysiological profile”). ( B-G ) data distribution for ramp input resistance ( B ), voltage upswing ( C ), Ion ( D ), theoretical Ion - Ion ( E ), voltage threshold ( F ), and difference between voltage threshold and resting membrane potential ( G ). Boxplot description as in Fig. 3. The significance level was set at p<0.05. * - p<0.05, ns - not significant. Data from 10 mice. Figure 5. The firing frequency gain is not affected by DCS. ( A ) MN’s response to the intracellularly injected triangular ramp of current. ( B ) The MN firing frequency is plotted against the injected depolarizing current. Following a rapid acceleration in firing frequency (subprimary range), a linear primary firing range is reached. A linear regression is then plotted for the primary firing range, and the slope of the regression determines the F/I gain. (C) The first three action potentials from the subprimary range are averaged, and AHP amplitude (CH), time to peak (CF), half-decay time (CG), and duration (CE) are calculated. Boxed area ( C ) has been expanded to show the spike width (CI). ( D-I ) data distribution for the slope of the F/I gain ( D ), AHP duration ( E ), time to peak ( F ), half-decay time ( G ) , amplitude ( H ), and spike width ( I ). Boxplot description as in Fig. 3. The significance level was set at p<0.05. ns - not significant. Data from 10 mice. Figure 6. Dorso-ventral anodal DCS does not alter MNs’ response to long depolarizing current pulses. ( A ) MN’s rheobase was identified by injecting 5-second depolarizing current pulses, with a 5-second break between them, with a 0.1nA step, until MN generated an action potential within the first 50ms of the stimulation. ( A1 ) The boxed area has been expanded to show the time profile of the measurements. Rheobase ( B ), spike generation threshold ( C ), and the difference between action potential generation threshold and resting membrane potential ( D ) are not changed by 15 minutes of anodal DCS. Boxplot description as in Fig. 3. The significance level was set at p<0.05. ns - not significant. Data from 10 mice. Figure 7. The dorso-ventral DCS electrode arrangement evokes a strong depolarization wave just beneath the dorsal electrode. The voltage deflection was measured at 3 depths (dorso-ventral axis) and 3 zones along the length (rostro-caudal axis) of the exposed spinal cord. At every depth (A-C) , a strong depolarization was observed next to the dorsal electrode, while the most caudal parts were largely devoid of the depolarizing wave. (D) In the rostral segments (L2), a significantly stronger depolarization is seen in the dorsal than in the ventral horns of the spinal gray matter. This difference is less prominent in the middle (L3) (E) and caudal (L4) (F) recording segments. *** - significant correlation at p<0.001 (A-C), * - significant difference at p<0.05 (D-F), ns - not significant, SS - spinal segment. Data from 5 mice. Figure 8. Distribution of the longitudinal electric field (EF_long) in the grey matter during spinal stimulation. (A) Sagittal view showing EF_long magnitude along the GM volume. (B) Cross-sectional views at rostral (L2 spinal segment), middle (L3 spinal segment) and caudal (L4 spinal segment) positions from the stimulating electrode, illustrating local EF_long distributions. Colour maps indicate EF intensity (V/m), normalised to the maximum EF_long value observed at the L2 segment (39.7 V/m). EF: electric field; SS: spinal segment; Max: maximum; Min: minimum. Information & Authors Information Version history V1 Version 1 29 August 2025 Copyright This work is licensed under a Non Exclusive No Reuse License. Collection European Journal of Neuroscience Keywords computational modelling firing properties in vivo intracellular recordings neurodegenerative diseases neuromodulation Authors Affiliations Bartosz Wasicki 0009-0008-0521-6851 [email protected] Akademia Wychowania Fizycznego im Eugeniusza Piaseckiego w Poznaniu View all articles by this author Piotr Zawistowski Akademia Wychowania Fizycznego im Eugeniusza Piaseckiego w Poznaniu View all articles by this author Tomasz Jankowiak Akademia Wychowania Fizycznego im Eugeniusza Piaseckiego w Poznaniu View all articles by this author Leonor de Oliveira Pires Universidade de Lisboa View all articles by this author Sofia Fernandes Universidade de Lisboa View all articles by this author Marcin Bączyk 0000-0002-2656-9655 Akademia Wychowania Fizycznego im Eugeniusza Piaseckiego w Poznaniu View all articles by this author Metrics & Citations Metrics Article Usage 233 views 149 downloads .FvxKWukQNSOunydq8rnd { width: 100px; } Citations Download citation Bartosz Wasicki, Piotr Zawistowski, Tomasz Jankowiak, et al. Acute invasive direct current stimulation of the lumbar spinal cord does not alter spinal motoneurons' firing characteristics in SOD-1 G93A mouse model of ALS.. Authorea . 29 August 2025. DOI: https://doi.org/10.22541/au.175648066.61520496/v1 If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. Simply select your manager software from the list below and click Download. For more information or tips please see 'Downloading to a citation manager' in the Help menu . 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