1–12 • The Journal of Neuroscience, February 28, 2024 • 44(9):e1324222024

Development/Plasticity/Repair

Static Magnetic Field Stimulation Enhances Shunting
Inhibition via a SLC26 Family Cl− Channel, Inducing
Intrinsic Plasticity
Adya Saran Sinha,1* Sumiya Shibata,2,3*
Tatsuya Mima6

Yasuyuki Takamatsu,4* Tenpei Akita,1,5

Atsuo Fukuda,1 and1Department of Neurophysiology, Hamamatsu University School of Medicine, Hamamatsu-shi 431-3192, Shizuoka, Japan, 2Department of Physical
Therapy, Niigata University of Health and Welfare, Niigata-shi 950-3198, Japan, 3Institute for Human Movement and Medical Sciences, Niigata University
of Health and Welfare, Niigata-shi 950-3198, Japan, 4Department of Rehabilitation Science, Faculty of Health Sciences, Hokkaido University, Sapporo-shi
060-0812, Hokkaido, Japan, 5Division of Health Science, Department of Basic Nursing, Hamamatsu University School of Medicine, Hamamatsu-shi
431-3192, Shizuoka, Japan, and 6The Graduate School of Core Ethics and Frontier Sciences, Ritsumeikan University, Kyoto-shi 603-8577, Kyoto, Japan

Magnetic ﬁelds are being used for detailed anatomical and functional examination of the human brain. In addition, evidence for their
efﬁcacy in treatment of brain dysfunctions is accumulating. Transcranial static magnetic ﬁeld stimulation (tSMS) is a recently developed technique for noninvasively modifying brain functions. In tSMS, a strong and small magnet when placed over the skull can
temporarily suppress brain functions. Its modulatory effects persist beyond the time of stimulation. However, the neurophysiologicalmechanisms underlying tSMS-induced plasticity remain unclear. Here, using acute motor cortical slice preparation obtained frommale C57BL/6N mice, we show that tSMS alters the intrinsic electrical properties of neurons by altering the activity of chloride (Cl−)
channels in neurons. Exposure of mouse pyramidal neurons to a static magnetic ﬁeld (SMF) at a strength similar to human tSMS
temporarily decreased their excitability and induced transient neuronal swelling. The effects of SMF were blocked by DIDS and
GlyH-101, but not by NPPB, consistent with the pharmacological proﬁle of SLC26A11, a transporter protein with Cl− channel activity.
Whole-cell voltage-clamp recordings of the GlyH-101-sensitive Cl− current component showed signiﬁcant enhancement of the component at both subthreshold and depolarized membrane potentials after SMF application, resulting in shunting inhibition and reduced
repetitive action potential (AP) ﬁring at the respective potentials. Thus, this study provides the ﬁrst neurophysiological evidence for the
inhibitory effect of tSMS on neuronal activity and advances our mechanistic understanding of noninvasive human neuromodulation.
Key words: Cl− channels; intrinsic excitability; intrinsic plasticity; shunting inhibition; SLC26A11; transcranial static magnetic ﬁeld
stimulation

Signiﬁcance Statement
Transcranial static magnetic ﬁeld stimulation (tSMS) is a recently developed noninvasive brain stimulation technique.
In tSMS, a strong, small magnet placed over the skull temporarily suppresses brain functions, and its modulatory effects persist beyond the stimulation time. To elucidate the neurophysiological mechanisms of tSMS, we evaluated the excitability ofmouse pyramidal neurons exposed to a static magnetic ﬁeld at a strength similar to that of human tSMS using whole-cellpatch-clamp experiments. We demonstrated that the static magnetic ﬁeld temporarily decreased neuronal excitability byincreasing the activity of a speciﬁc type of Cl− channel in the plasma membrane, and it also induced transient neuronal swelling. This study reveals for the ﬁrst time the neurophysiological mechanism of tSMS-induced suppression of brain functions.

Received July 4, 2022; revised Jan. 9, 2024; accepted Jan. 13, 2024.
Author contributions: S.S., T.A., A.F., and T.M. designed research; A.S.S. and Y.T. performed research; A.S.S.,
Y.T., T.A., and A.F. analyzed data; A.S.S., S.S., and T.M. wrote the paper.
This study was partly supported by Grants-in-Aid for Scientiﬁc Research [KAKENHI; grant number 15H05872(A.F.), 18K17739 (Y.T.), 19H01091 (A.F., T.M.), 21H05687 (A.F.), 21K17671 (S.S.), 21K19745 (T.M.), 22H04788(T.M.)] from the Japan Society for the Promotion of Science.

*A.S.S., S.S., and Y.T. contributed equally to this work and share ﬁrst authorship.
The authors declare no competing ﬁnancial interests.
Correspondence should be addressed to Tatsuya Mima at t-mima@fc.ritsumei.ac.jp.
https://doi.org/10.1523/JNEUROSCI.1324-22.2024
Copyright © 2024 the authors2 • J. Neurosci., February 28, 2024 • 44(9):e1324222024

Introduction
The clinical application of biomagnetism has been a powerfultool for recent breakthroughs in human neuroscience.
Magnetic resonance imaging is used to visualize brain structuresand cerebral metabolism. Magnetoencephalography detects
brain activity in the order of milliseconds. Repetitive transcranial
magnetic stimulation (rTMS) can alter human brain functions
noninvasively, the mechanism of which is probably via the longterm potentiation/depression (LTP/LTD) of synaptic transmission (Hallett, 2000; Kobayashi and Pascual-Leone, 2003;
Hoogendam et al., 2010; Freitas et al., 2013).
Transcranial static magnetic ﬁeld stimulation (tSMS) is thelatest type of biomagnetism technique (Oliviero et al., 2011;
Kirimoto et al., 2014; Gonzalez-Rosa et al., 2015; CarrascoLópez et al., 2017; Shibata et al., 2020, 2021). By holding a strong
and compact magnet over the skull, tSMS can temporarily suppress brain function just below it and cause behavioral eﬀects
on cognition, motor control, learning, and memory (GonzalezRosa et al., 2015; Nojima et al., 2019; Kirimoto et al., 2020;
Tsuru et al., 2020). Furthermore, the eﬀect of brain inhibition
persists beyond the time of stimulation (Oliviero et al., 2011;
Dileone et al., 2018), suggesting induction of brain plasticity inhumans. As a safe, easy, and low-cost therapeutic tool, tSMS
has recently been applied for neuromodulation in children(Hollis et al., 2020) and for self-administered daily treatment
for neurological disorder (Di Lazzaro et al., 2021).
The neurophysiological mechanisms underlying the prolonged eﬀects of tSMS remain to be elucidated. In contrast torTMS, in which neuronal ﬁring followed by synaptic transmission is repeated, tSMS does not directly produce neuronal action
potentials (APs). The eﬀects of tSMS may result from changes in
the function of membrane ion channels caused by magnetic torque forces on magnetically anisotropic membrane phospholipids(Rosen, 1996; Coots et al., 2004). Thus, tSMS may induce longlasting changes in nonsynaptic intrinsic neuronal excitability or
intrinsic plasticity. Intrinsic plasticity is usually represented by
the neuronal input–output (I–O) function (Zhang and Linden,
2003) and is mediated by changes in the properties of membrane
ion channels (Titley et al., 2017). Moreover, intrinsic plasticity
has been implicated as a cellular correlate of certain types oflearning and memory (Daoudal and Debanne, 2003; Jang et al.,2020) as well as a background for some neuronal disorders,
including epilepsy, Alzheimer’s disease, and addiction (Beckand Yaari, 2008; Kourrich et al., 2015). However, the involvement of intrinsic plasticity in the eﬀects of human noninvasive
brain stimulation has never been hypothesized.
Here we show that tSMS induces intrinsic plasticity by increasing membrane chloride (Cl−) conductance. This increased conductance is mediated by a speciﬁc type of Cl− channel whose
pharmacological proﬁle is consistent with SLC26A11 (Rungta
et al., 2015). Thus, this study provides the ﬁrst molecular andcellular evidence for the inhibitory eﬀects of tSMS on brain
function.

Materials and Methods
Slice preparation. All procedures were in accordance with guidelines
issued by the Hamamatsu University School of Medicine on the ethicaluse of animals for experimentation and were approved by theCommittee for Animal Care and Use (2019008). All experiments were
performed to minimize the number of animals used. Postnatal day (P)
21–27-old male C57BL/6N mice were anesthetized with 50–90 mg/kg
intraperitoneal injection of pentobarbital and decapitated. Brains werequickly removed and placed in ice-cold oxygenated modiﬁed artiﬁcial

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• Intrinsic Plasticity in tSMS

cerebrospinal ﬂuid (ACSF) consisting of the following (in mM): 220
sucrose, 2.5 KCl, 1.25 NaH2PO4, 2.0 MgSO4, 0.5 CaCl2, 26.0 NaHCO3,
and 30.0 glucose, pH 7.4. Coronal slices containing the motor cortex
with a thickness of 350 µm were prepared in modiﬁed ACSF by using
a vibratome (Campden Instruments). This region was identiﬁed using
the mouse brain atlas (Paxinos and Franklin, 2019). Slices were allowed
to recover for 60 min on nylon meshes (with 1 mm pores) placed ondishes and submerged in standard ACSF consisting of the following(in mM): 126 NaCl, 2.5 KCl, 1.25 NaH2PO4, 2.0 MgSO4, 2.0 CaCl2,
26.0 NaHCO3, and 20.0 glucose, saturated with 95% O2 and 5% CO2
at 25–26°C.
Experimental design. To investigate the eﬀects of a static magnetic
ﬁeld (SMF) on the brain, we evaluated the intrinsic excitability of mouselayer II/III pyramidal neurons in acute coronal brain slices containing
the motor cortex after SMF exposure. The setup for the SMF exposureis shown in Figure 1a. The intensity of the SMF at the cortical slices
was 300 mT, and the duration of exposure was 30 min. These conditions
are similar to those of tSMS in humans. As in humans, eﬀects of tSMS
produce a period of ∼10 min attenuated neural activity followed by waning of eﬀects within ∼11–20 min; a similar strategy to evaluate eﬀects of
SMF was devised. Coronal brain slices for the experiments were divided
into three groups: control with no SMF exposure (CTRL), and following
a 30 min period of SMF, groups of post-SMF 1, and post-SMF 2. AP
ﬁring induced by current injection (0.5 s) was recorded in the neurons
under current-clamp conditions within 0–10 and 10–20 min after SMF
exposure in the post-SMF 1 and post-SMF 2 groups, respectively(Fig. 1b). The recorded neurons were synaptically isolated by bath application of drugs that blocked both glutamatergic and GABAergic neurotransmissions, as described below.
Application of SMF (Fig. 1). Following a 1 h incubation of the slices
at 25–26°C for recovery, slices on the nylon meshes were submerged instandard ACSF in thin-walled (0.5 mm) borosilicate glass beaker withthe placement of a neodymium (NdFeB) magnet (10 cm × 10 cm ×3 cm) underneath. The distance between the slice tissue and the magnet
surface was maintained at 1 cm. The whole slices were separated into
hemispheres, and one half was positioned during incubation, such thata magnetic ﬁeld of 300 mT was maintained during stimulation. This
position was calibrated using a teslameter/gaussmeter (PCE Instruments).
The duration of the stimulation was 30 min. The control half of the slices
was not exposed to a magnetic ﬁeld and was maintained at 25–26°C in standard ACSF.
Whole-cell patch-clamp recordings. The slices were then transferred
to a recording chamber on the stage of an upright microscope(BX51WI; Olympus) and were continuously perfused with oxygenated
ACSF at a ﬂow rate of 2 ml/min at 25–26°C. Whole-cell patch-clamp
recordings were obtained from layer II/III pyramidal neurons of the
motor cortex. The electrode resistance ranged from 3 to 5 MΩ whenthe electrode was ﬁlled with a solution containing (in mM) 140.0 potassium methanesulfonate (CH3SO3K), 10.0 KCl, 2.0 MgCl2, 10.0 HEPES,
3.0 Na2 ATP, 0.2 Na GTP, 1.0 EGTA, pH adjusted to 7.3 with KOH,
and an osmolarity value of 310 mOsm. The liquid junction potential(LJP) was calculated to be 4.8 mV and corrected during the recording.
In another set of experiments, a high Cl− internal solution of the following composition (in mM) was used: 150.0 KCl, 2.0 MgCl2, 10.0 HEPES,
3.0 Na2 ATP, 0.2 Na GTP, 1.0 EGTA, with pH adjusted to 7.3 with KOH,
and osmolarity of 306 mOsm. The corresponding calculated LJP value of
6.8 mV was corrected during the recording. Voltage changes wererecorded using a MultiClamp 700B ampliﬁer (Axon Instruments) witha Bessel preﬁlter at 5 kHz, and digitized at 25 kHz using a Digidata
1440A data acquisition system (Axon Instruments).
Recordings were performed in the presence of bath-applied ionotropic glutamate receptor blockers: 10 µM 6-cyano-7-nitroquinoxaline-2,3dione (CNQX), 50 µM D-2-amino-5-phosphonopentanoic acid
(D-AP5), and 50 µM picrotoxin (PTX), a GABAA receptor
antagonist. Additionally, during rescue experiments, either 300 µM
4,4′ -diisothiocyano-2,2′ -stilbenedisulfonic acid (DIDS) or 200 µM

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J. Neurosci., February 28, 2024 • 44(9):e1324222024 • 3

Figure 1. Application of SMF and strategy for our experiments. a, Setup for SMF exposure using a NdFeB magnet. The ex vivo motor cortical slices were placed on a nylon mesh with a pore
diameter of 1 mm and submerged in ACSF continuously bubbled with an oxygen and carbon dioxide mixture (95% O2 + 5% CO2) at 25–26°C. The distance between the surface of the slice andthe magnet was maintained at 1 cm. b, Workﬂow for experiments. After recovery for 1 h at 25–26°C, the two hemispheres of the coronal brain slices were separated and assigned to threegroups: control without SMF exposure (CTRL), post-SMF 1, and post-SMF 2. Current-clamp recordings were performed from layer II/III pyramidal neurons—0–10 min or 11–20 min after 30 minof SMF exposure in the post-SMF 1 and post-SMF 2 groups, respectively.
5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) was applied in
the bath. Furthermore, for additional experiments, a more speciﬁc
blocker for SLC26A11-mediated currents, N-2-naphthalenyl-2-[(3,5dibromo-2,4-dihydroxyphenyl) methylene] glycine hydrazide (GlyH101), at 50 µM concentration was bath applied.
To understand the current–voltage (I–V) characteristics of theenhanced Cl− currents, we performed voltage-clamp recordings for
GlyH-101-sensitive currents. We used the internal solution and drug
cocktail combination together with the voltage step protocol reported
earlier (Rungta et al., 2015) for isolation of SLC26A11-mediated currents. The internal solution of the following composition (in mM) was
used: 108.0 CsCl, 8.0 TEA-Cl, 8.0 K-gluconate 2.0 MgCl2, 10.0 HEPES,
4.0 Na2 ATP, 0.2 Na GTP, 1.0 EGTA, with pH adjusted to 7.3 with
CsOH, and osmolarity of 290 mOsm. The corresponding calculated
LJP value of 4.8 mV was corrected during the recording. The bath solution during this experiment contained 10 µM CNQX, 50 µM PTX,
50 µM D-AP5, 50 µM nickel, 60 µM cadmium, and 100 µM 4-aminopyridine. GlyH-101-sensitive currents were obtained by subtracting the current response obtained by the voltage step protocol in the presence of
50 µM GlyH-101 from the current response obtained without GlyH-101.
Electrophysiological analysis. The pClamp suite was used, including
Clampex 10.7 software for data acquisition and Clampﬁt 10.4 foranalysis. Records obtained with series resistance <25 MΩ were selected
for analysis. To evaluate the input resistance (RIn) of the recorded neurons, hyperpolarizing currents with amplitudes starting at −60 pA anddecreasing by 20 pA were repeatedly injected for 500 ms. The voltage
responses obtained were plotted against the corresponding current injection amplitudes to create a current–voltage relationship. The data points
were then ﬁtted using linear regression analysis. The calculated slope
indicates the RIn of the recorded neuron. The rheobase current, that is,
the amount of current injection at which a neuron ﬁres one AP, was estimated by applying depolarizing current steps of 500 ms in 20 pA stepincrements. The repetitive AP ﬁrings elicited by the depolarizing current
steps were used to examine the I–O curve of the recorded neurons.
Furthermore, short (2 ms) depolarizing pulses were applied to induce
single APs, and waveform parameters of AP amplitude, half-width, maximum rise, and decay slope were analyzed. The resting membrane potential (RMP) was calculated from the trace of the zero current injectionlevel before application of the short-pulse protocol. The AP amplitude
was calculated from RMP to AP peak voltage. The AP threshold voltage
was determined at the inﬂection point where the AP was triggered.
Chemicals. PTX was from Tocris Bioscience. D-AP5, NPPB, and
GlyH-101 were from Cayman Chemical. DIDS and TTX were from
Wako, Fujiﬁlm. CNQX and 4-AP were from Sigma-Aldrich.

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Neuronal size analysis. Surface area and perimeter analyses were performed using Fiji (ImageJ) platform, version 2.1.0/1.53 C. Brieﬂy,
IR-DIC 8 bit images were obtained during electrophysiological recordings under a 40× objective. From the images, neurons with a well-deﬁned
pyramidal-shaped soma and sharp contrasts along the edges wereselected for analysis. The freehand selection tool was used to deﬁne theregion of interest (ROI). These ROIs were further reﬁned using the selection brush tool with a 2 pixels tip. Then, using the measurement dialog
box, parameters of surface area and perimeter were analyzed for theselected ROIs. Measurements from cells within a single slice were averaged to obtain a single data point. Therefore, the number of data points
in plot indicates the number of slices analyzed.
Statistical analysis. Statistical analysis of data obtained by patchclamp experiments was performed using IBM SPSS Ver.23 software. In
brief, the data were tested for normality and equality of variance using
the Kolmogorov–Smirnov (K–S) test and Levene’s test, respectively.
For independent samples, two-sided t test with conﬁdence interval set
at 95% was used for comparisons between the two groups. For multiple
group comparisons, one-way ANOVA was performed, followed by apost hoc Ryan–Einot–Gabriel–Welsch F test. However, if the K–S statistic conﬁrmed a non-normal distribution, the one-sided Mann–Whitney
U test was employed for the two groups, and for multiple groups, theK–W test followed by stepwise step-down comparisons was used. Data
are presented as mean ± standard error of the mean (SEM). Statisticalanalysis of the imaging data was also performed based on a similar
algorithm. Statistical signiﬁcance is presented as follows: *p < 0.05,**p < 0.01, ***p < 0.001, ns (not signiﬁcant).

Results
Temporary suppression of neuronal excitability after SMF
exposure
The relationship of injected currents to the number of AP spikes(I–O curve) showed that their numbers in the post-SMF 1 groupwere signiﬁcantly lower than those in the CTRL and post-SMF 2groups over the full range of injected currents [Fig. 2a,b; Kruskal–
Wallis test (K–W test); **p < 0.01]. Rheobase, the minimum current necessary for AP generation, was signiﬁcantly higher in the
post-SMF 1 group (313.75 ± 26.82 pA; n = 16 cells, ﬁve mice)than that in the CTRL group (183 ± 17.49 pA; n = 20 cells, ﬁve
mice; p = 0.000136; Fig. 2c). The rheobase in the post-SMF 2group (197.14 ± 19.87 pA; n = 21 cells, six mice) was similar tothat in the CTRL group. The higher rheobase suggested changes
in the passive membrane properties in the post-SMF 1 group.
Indeed, the RIn, the reciprocal of membrane ion conductanceunder resting conditions, in the post-SMF 1 group (69.35 ±6.34 MΩ; p = 0.003) was signiﬁcantly lower than that in the
CTRL (128.1 ± 16.51 MΩ) and post-SMF 2 groups (119.75 ±13.12 MΩ), whereas the RIn in the CTRL and post-SMF 2 groups
was statistically similar (Fig. 2d). Together, these ﬁndings indicate that 30 min of exposure to SMF temporarily suppressed
the excitability of pyramidal neurons by reducing RIn, that is,
by increasing the resting membrane conductance, and the excitability recovered within 20 min after the end of the exposure.
The increase in the resting membrane conductance can be
mediated by various ions. The RMP shifts toward a more negative or positive value on an increase in the K+ or Na+ conductance, according to the polarity of the equilibrium potential for
K+ or Na+, respectively. However, the RMP did not diﬀer
between the three groups (CTRL, −70.65 ± 1.80 mV; post-SMF
1, −70.13 ± 1.89 mV; and post-SMF 2, −70.91 ± 1.73 mV;
p = 0.931; Fig. 3a), suggesting that the increase in K+ or Na+ conductance was less likely. The higher rheobase may also haveresulted from an increase in the threshold membrane voltage
level for AP generation; however, it also did not diﬀer between

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them (Fig. 4a). Moreover, the parameters characterizing the
waveform of a single AP remained unchanged (Figs. 3b, 4b–e).
Thus, the voltage-gated Na+ and K+ channels responsible for AP
generation appeared to be unaﬀected by the SMF exposure. A previous study (Rosen, 2003a) reported a reduction in Na+ conductance in the GH3 cell line within 200 s after the end of the SMF
exposure. In our experimental setup, it took ∼4–8 min to complete
the AP recordings after the SMF exposure. Thus, the reduced Na+
conductance, if any, was restored to its initial level by the time ofdata acquisition in our experiments. Therefore, the increased resting membrane conductance underlying the temporary suppression of neuronal excitability within 10 min of SMF exposure was
likely mediated by ions other than Na+ and K+.
SMF exposure increases resting membrane conductancethrough Cl− channels
The equilibrium potential for Cl− under the experimental conditions described above was approximately −60 mV. Thus, theincrease in Cl− conductance, if any, would have had little impact
on the RMP. Therefore, to test whether the SMF increased Cl−
conductance, we next examined passive membrane properties
and AP ﬁring using a patch pipette solution containing a highconcentration of Cl− (154 mM) and compared them between
the conditions of the CTRL and post-SMF 1 group. If the SMF
increased the Cl− conductance under this condition, it would
shift the RMP toward a more positive value, because the equilibrium potential for Cl− was high (∼0 mV). Indeed, we found asigniﬁcant depolarization of the RMP in the post-SMF 1 group(−70.15 ± 1.05 mV; n = 18 cells, ﬁve mice; p = 0.016) in comparison with the CTRL group (−73.69 ± 1.33 mV; n = 18 cells, ﬁve
mice; Fig. 5a). Higher rheobase (Fig. 5b), lower RIn (Fig. 5c),and a smaller number of AP spikes (Fig. 5d) were also reproduced
under this condition. The parameters of the AP waveform were
unchanged (Figs. 5e, 6), except for a small but signiﬁcant reduction in the AP amplitude after SMF exposure (CTRL, 109.31 ±1.64 mV; post-SMF 1, 102.37 ± 1.56 mV; p = 0.004; Fig. 6b).
Thus, the SMF exposure temporarily suppressed neuronal excitability by enhancing resting Cl− conductance.
The ligand-gated Cl− channel γ-aminobutyric acid receptors(GABAAR) are important regulators of neuronal excitability.
However, the eﬀect of SMF on intrinsic excitability was independent of GABAergic action, as the experimental conditions
included PTX, a potent blocker of these channels.
Identiﬁcation of the other types of Cl− channels involved in
SMF action is inherently complex because of the lack of highly
speciﬁc and sensitive pharmacological tools at our disposal(Verkman and Galietta, 2009). However, a reasonable assessmentof the type of Cl− channel responsible for the SMF action can beobtained from their sensitivity to classical Cl− channel blockers
with divergent spectra. Therefore, we examined the eﬀects ofthese drugs on the shifts in RMP, RIn, and rheobase after SMF
exposure using a high Cl− pipette solution. Blockers were applied
during the SMF exposure. We found that a nonselective blocker,
DIDS (300 µM), signiﬁcantly inhibited the SMF-induced shifts ofall parameters (Fig. 7a–c; post-SMF 1 + DIDS), whereas anothernonselective blocker, 5-nitro-2-(3-phenylpropylamino) benzoic
acid (NPPB; 200 µM), had no eﬀect (Fig. 5a–c; post-SMF 1 +NPPB).
In addition, since increase in Cl− conductance is often associated with increase in cell volume (Akita and Okada, 2014), we
compared the perimeter and surface area of neuronal somata
before whole-cell recording between the CTRL and post-SMF 1
groups (Fig. 8a,b). We found that both perimeter (CTRL, 45.04

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J. Neurosci., February 28, 2024 • 44(9):e1324222024 • 5

Figure 2. Reduced intrinsic excitability of layer II/III pyramidal neurons of the motor cortex following SMF application. a, Representative subthreshold current injections (from −60 to +60 pA)
with consequent voltage responses and repetitive ﬁring by suprathreshold current injection (+480 pA) in the control (CTRL; blue), post-SMF 1 (orange), and post-SMF 2 (purple) groups. At thesame subthreshold current injection, the voltage responses decreased in the post-SMF 1 group. At +480 pA current injection, the number of spikes decreased in the same group. b, A signiﬁcantdecrease (K–W test; **p < 0.01) in the I–O relationship for the post-SMF 1 (orange) group in comparison with both CTRL (blue) and post-SMF 2 (purple) groups. Error bars represent SEM. c, d,
Individual data points are shown as open circles in blue (CTRL, n = 20), orange (post-SMF 1, n = 16), and purple (post-SMF 2, n = 21). The mean values are indicated by solid bars next to thedata points. Error bars represent SEM. Rheobase from layer II/III pyramidal neurons signiﬁcantly increased in the post-SMF 1 group compared with CTRL and post-SMF 2 (one-way ANOVA;
p = 0.000136; post hoc, Ryan–Einot–Gabriel–Welsch F test; post-SMF 1 vs post-SMF 2 with **p < 0.01; post-SMF 1 vs CTRL with *p < 0.05) groups (c). Input resistance of pyramidal neurons
signiﬁcantly decreased in the post-SMF 1 group (K–W test; **p < 0.01; post hoc, stepwise step-down; post-SMF 1 vs post-SMF 2 with **p < 0.01; post-SMF 1 vs CTRL with *p < 0.05; d).

± 1.27 µm, n = 17 slices, 6 cells/slice; post-SMF 1, 53.80 ±0.57 µm, n = 17 slices, 5.5 cells/slice, 8 mice, p = 1.0 × 10−5;
Fig. 8d) and surface area (CTRL, 121.41 ± 6.13 μm2; post-SMF
1, 167.62 ± 3.60 μm2; p = 3.0 × 10−6; Fig. 8e) were signiﬁcantly
larger in the post-SMF 1 group. DIDS treatment blocked thistransient increase in perimeter (CTRL, 45.04 ± 1.27 µm; post-SMF
1 + DIDS, 46.05 ± 0.59 µm; n = 13 slices, 6 cells/slice, 5 mice) andsurface area of neuronal somata (CTRL, 121.41 ± 6.13 μm2;
post-SMF 1 + DIDS, 128.50 ± 3.03 μm2; Fig. 8c–e). Thus, theconductance through a DIDS-sensitive, NPPB-insensitive type of6 • J. Neurosci., February 28, 2024 • 44(9):e1324222024

Cl− channel would have been predominantly increased by SMF,
thereby temporarily suppressing the excitability of pyramidal
neurons and also causing neuronal swelling.
Among neuronal Cl− channels, the one with a pharmacological proﬁle consistent with our ﬁndings of DIDS sensitivity and
NPPB insensitivity was the channel function of the transporter
protein SLC26A11 reported by Rungta et al. (2015). In thatreport, they showed that CFTR inhibitor GlyH-101 inhibited

Figure 3. RMP values and single AP waveform following SMF application. a, Individual
data points are shown as open circles in blue (CTRL, n = 20), orange (post-SMF 1,
n = 16), and purple (post-SMF 2, n = 21). The mean values are indicated by solid bars
next to the data points. Error bars represent SEM. The RMP values were not signiﬁcantly
diﬀerent among the three groups (one-way ANOVA; ns). b, Representative single AP waveforms elicited by a short-pulse (2 ms) protocol in the CTRL (blue), post-SMF 1 (orange), andpost-SMF 2 (purple) groups. SMF application had no apparent eﬀect on the AP waveform.

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Cl− transport via SLC26A11 by 80% at 10 µM concentration.
Therefore, we investigated whether GlyH-101 could rescue thedepression of intrinsic excitability of neurons after SMF using
the high Cl− (154 mM) internal solution. The comparisons of
I–O curves revealed a reversal of eﬀect of SMF following
GlyH-101 application. Moreover, the number of AP spikes
ﬁred by these neurons exceeded those of CTRL neurons signiﬁcantly (Fig. 9a,b; K–W test; ***p < 0.001). We also observed areversal of rheobase currents (Fig. 9c), RIn (Fig. 9d), and RMP levels (Fig. 9e) similar to CTRL group, respectively. Thus, the Cl−
channel aﬀected by SMF application was sensitive to DIDS and
GlyH-101 but not to NPPB, suggesting the SLC26A11 channel.
SMF increases voltage-dependent GlyH-101-sensitive currents
both at subthreshold and depolarized membrane potentials
Rungta et al. (2015) reported that the Cl− currents mediated by
SLC26A11 in layer 4 cortical neurons were dependent on themembrane voltage. We therefore next evaluated if Cl− currents
modulated by SMF exhibit any voltage dependence. Using thesame internal solution, blockers and voltage step protocol(from −70 to +20 mV, 300 ms duration) as in Rungta et al.
(2015), whole-cell Cl− currents were recorded before and after
application of 50 μM GlyH-101, and the GlyH-101-sensitive current component was isolated by subtraction and compared
between the CTRL and post-SMF 1 groups (Fig. 10a). The
peak current–voltage (I–V) plots (Fig. 10b) revealed that the
GlyH-101-sensitive current component was signiﬁcantly

Figure 4. AP parameters following SMF application. a–e, Individual data points are shown as open circles in blue (CTRL, n = 20), orange (post-SMF 1, n = 16), and purple (post-SMF 2,
n = 21). The mean values are indicated by solid bars next to the data points. Error bars represent SEM. AP thresholds (CTRL, −36.08 ± 2.16 mV; post-SMF 1, −38.09 ± 2.46 mV; and post-SMF 2,−31.48 ± 3.30 mV; p = 0.400; a), AP amplitudes (CTRL, 108.45 ± 1.75 mV; post-SMF 1, 107.77 ± 1.84 mV; and post-SMF 2, 108.38 ± 2.11 mV; p = 0.946; b), maximum rise slopes (CTRL,
210.50 ± 15.20 mV/ms; post-SMF 1, 224.26 ± 12.49 mV/ms; and post-SMF 2, 200.78 ± 16.09 mV/ms; p = 0.286; c), maximum decay slopes (CTRL, 52.09 ± 2.66 mV/ms; post-SMF 1, 47.26± 1.41 mV/ms; and post-SMF 2, 48.41 ± 2.75 mV/ms; p = 0.394; d), and AP half-maximal widths (CTRL, 1.91 ± 0.08 ms; post-SMF 1, 1.94 ± 0.06 ms; and post-SMF 2, 2.22 ± 0.14 ms;
p = 0.08; e) were not signiﬁcantly diﬀerent among the three groups (one-way ANOVA, ns).

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Figure 5. Intrinsic excitability of layer II/III pyramidal neurons of the motor cortex under high [Cl−]i internal solution. a–c, Individual data points are shown as open circles in blue (CTRL,n = 18) and orange (post-SMF 1, n = 18). The mean values are indicated by solid bars next to the data points. Error bars represent SEM. a, Quantitative analysis of current-clamp recordings using
154 mM Cl− internal solution revealed a signiﬁcantly depolarized RMP in the post-SMF 1 group (Mann–Whitney U test; *p < 0.05), indicating an enhancement of resting Cl− conductance(compare Fig. 3a). b, Rheobase was signiﬁcantly increased in the post-SMF 1 group (t test; **p < 0.01). c, Input resistance of neurons from the post-SMF 1 group was signiﬁcantly reduced (t test;**p < 0.01). d, A signiﬁcant decrement in the input–output relationship for the post-SMF 1 group (orange) in comparison with the CTRL group (blue; Mann–Whitney U test; *p < 0.05). Error
bars represent SEM. e, Representative single AP waveforms elicited by a short-pulse (2 ms) protocol in the CTRL (blue) and post-SMF 1 (orange) groups.

enhanced after SMF application in the whole voltage range from−70 to +20 mV (at −70 mV, post-SMF 1: 32.24 ± 7.20 pA, n = 14
cells, four mice; CTRL: 4.27 ± 7.55 pA, n = 14 cells, ﬁve mice, p =0.012548; at −60 mV, post-SMF 1: 48.96 ± 8.26 pA, n = 14 cells,
four mice; CTRL: 12.56 ± 6.35 pA, n = 14 cells, ﬁve mice, p =0.001727 and at +20 mV, post-SMF 1: 627.57 ± 72.59 pA, n =14 cells, four mice; CTRL: 233.61 ± 43.19 pA, n = 14 cells, ﬁve
mice, p = 0.000082). The component further consisted of the rapidly activating and inactivating current component generated at
the beginning of the voltage step and the slowly activating current
component throughout the voltage step. The rapid component
was already generated at the subthreshold levels of the voltage
steps. The slow component showed outward rectiﬁcation withincreasing voltage, similar to the current reported by Rungta
et al. (2015). Both the rapid and slow current components weresigniﬁcantly enhanced after SMF application. Thus, the rapid
Cl− current component induced by SMF is responsible for the
shunting inhibition that oﬀsets membrane excitability at subthreshold voltages and the slow Cl− current component is
responsible for the reduction of AP spikes during repetitive AP
ﬁring at depolarized voltages.
TTX-sensitive Na+ currents contribute to induction of SMF
actionConsidering that Cl− entry is secondarily driven by the electrochemical driving force for Na+, we investigated the role oftetrodotoxin (TTX)-sensitive Na+ channel activity in the
SMF-induced eﬀects. For this purpose, we applied 1 µM TTX
during 30 min of SMF application and evaluated RMP, RIn
and rheobase currents after TTX washout using the high Cl−
pipette solution. We found that the TTX treatment abolishedthe eﬀects of SMF. RMP (CTRL: −73.69 ± 1.33 mV, n = 18 cells,
ﬁve mice; post-SMF1 + TTX: −74.20 ± 1.71 mV, n = 15 cells,
four mice) remained similar to the CTRL group, indicatingthat the increase in basal conductance after SMF was prevented. Similarly, the changes in RIn and rheobase
currents became nonsigniﬁcant (data not shown). Thus, theNa+ entry through voltage-gated Na+ channels was found tobe necessary for the SMF-induced increase in membrane Cl−
conductance.

Discussion
Our electrophysiological experiments investigating the mechanisms underlying tSMS action show that mouse pyramidal cells
exhibited same prolonged suppression of intrinsic excitability(∼10 min) after exposure to the similar magnetic parameters as
in human tSMS studies (Oliviero et al., 2011). This intrinsic plasticity is mediated by a Cl− channel sensitive to both DIDS and
GlyH-101. The I–V characteristics of these GlyH-101-sensitive
Cl− currents indicate SMF action produces increase in conductance at both subthreshold and depolarized membrane potentials
essentially enhancing a shunting inhibition.

8 • J. Neurosci., February 28, 2024 • 44(9):e1324222024

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• Intrinsic Plasticity in tSMS

Figure 6. AP parameters under high [Cl−]i internal solution following SMF application. The maximum rise slopes (a), maximum decay slopes (c), AP half-maximal widths (d), and AP
threshold voltages (e) were not signiﬁcantly diﬀerent between the CTRL and post-SMF 1 groups (t test; ns). b, AP amplitudes were signiﬁcantly reduced in the post-SMF 1 group (t test;**p < 0.01).

Figure 7. Inhibition of SMF-induced shifts in RMP, rheobase, and input resistance by DIDS, not by NPPB. a–c, Individual data points are shown as open circles in blue (CTRL, n = 18), orange(post-SMF 1, n = 18), purple (post-SMF 1 + DIDS, n = 18), and red (post-SMF 1 + NPPB, n = 18). The mean values are indicated by solid bars next to the data points. Error bars represent SEM.
a, The signiﬁcant increase in RMP after SMF application was reversed by DIDS (300 µM) treatment; however, not by NPPB (200 µM) application (p = 0.003; CTRL vs post-SMF 1 + DIDS, ns; CTRLvs post-SMF 1, **p < 0.01; post-SMF 1 vs post-SMF 1+ NPPB, ns). b, The signiﬁcant increase in rheobase levels was also reversed following DIDS treatment. NPPB treatment showed no eﬀect(p = 0.000149; CTRL vs post-SMF 1+ DIDS, ns; CTRL vs post-SMF 1, *p < 0.05; post-SMF 1 + DIDS vs post-SMF 1, *p < 0.05; post-SMF 1 vs post-SMF 1 + NPPB, ns). c, Application of DIDS rescued
the reduction in input resistance after SMF application. However, NPPB treatment showed no eﬀect (**p < 0.01; CTRL vs post-SMF 1 + DIDS, ns; CTRL vs post-SMF 1, **p < 0.01; post-SMF 1 +DIDS vs post-SMF 1, **p < 0.01; post-SMF 1 vs post-SMF 1 + NPPB, ns).

The neuronal membrane expresses diﬀerent classes of Cl−
channels and transport proteins (Duran et al., 2010). In tandem,
they regulate pH, cell volume, and GABAergic and glycinergic
neurotransmission, enabling a multitude of physiological functions related to neuronal migration, brain development, andinformation processing. The identity of the Cl− channels
involved in the SMF action may be surmised based on their pharmacological sensitivities to two broad-spectrum Cl− channel

blockers, DIDS, and NPPB. Among the voltage-gated Cl− channels (ClC family), ClC-2 channels are expressed in neurons andother cell types, including astrocytes, and are active at voltages
close to the RMP (Jentsch and Pusch, 2018). As a candidate
mediator of the SMF action, ClC-2 channels are feasible; however, their pharmacological proﬁles indicate poor antagonism
by DIDS (Alexander et al., 2013). In addition, they are sensitive
to the NPPB blocks. This spectrum is in contrast to ourSinha et al.

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Figure 8. Transient neuronal swelling following SMF application. Examples of IR-DIC images from the motor cortical slices incubated with CTRL (a), post-SMF 1 (b), and post-SMF 1 + DIDS
(300 μM; c). Pyramidal neurons in the post-SMF 1 slice had an apparent swollen appearance unlike in the DIDS-incubated slices. Scale bar, 50 μm. d, Pyramidal neurons from slices in the
post-SMF 1 group (orange open circles, n = 17 slices, 8 mice) signiﬁcantly increased in the soma perimeter when compared with that in the CTRL group (blue open circles, n = 17 slices, 8 mice).
DIDS treatment (purple open circles, n = 13 slices, 6 mice) prevented this transient increase in the soma perimeter (**p < 0.01; CTRL vs post-SMF 1 + DIDS, ns; CTRL vs post-SMF 1, **p < 0.01;
post-SMF 1 + DIDS vs post-SMF 1, **p < 0.01). e, Quantitative analysis of the soma surface area of neurons showed a signiﬁcant increase in the post-SMF 1 group when compared with both CTRL
and DIDS application groups (*p < 0.01; CTRL vs post-SMF 1 + DIDS, ns; CTRL vs post-SMF 1, **p < 0.01; post-SMF 1 + DIDS vs post-SMF 1, **p < 0.01).

experimental ﬁndings. Another prospective target is volumesensitive outwardly rectifying (VSOR) anion channels (Akita
and Okada, 2014) important in cell volume regulation. These
VSOR channels are reportedly sensitive to both DIDS andNPPB (Inoue et al., 2005). Our pharmacological proﬁling experiments indicated that the SMF-activated Cl− conductance wasinsensitive to NPPB action, thereby deviating from classical
VSOR channel properties.
Among the other mediators of neuronal Cl− transport, afavorable pharmacological proﬁle consistent with our ﬁndingsof DIDS and GlyH-101 sensitivity and NPPB insensitivity is
the channel activity of the transporter protein SLC26A11(Rungta et al., 2015, Rahmati et al., 2016). These proteins arereported to express at moderate to high levels in the neocortex(Rahmati et al., 2013) and accumulate Cl− intracellularly accompanying sustained increase in [Na+]i during cytotoxic neuronalswelling (Rungta et al., 2015). In addition to the increase in Cl−
conductance, we also reported transient neuronal swelling following SMF application (Fig. 8) which could be reversed byapplication of DIDS. These evidences support the involvementof SLC26A11 channels in Cl− accumulation.
Our experiments using GlyH-101 as a potential blocker of
SMF eﬀects indicate that SMF action could be reversed bythem. In addition, our voltage-clamp data indicates similaritiesin I–V characteristics with SLC26A11-mediated currents(Rungta et al., 2015). Considering these similarities, we proposethat SMF modulates these same channels. In addition to thealready ascribed functions of these channels, we submit thatthey would act as a weak inhibitory shunt during normal neuronal activity mostly at depolarizing membrane potentials.
However, on application of SMF, the basal conductance of thesechannels is enhanced (Fig. 10), enhancing the shunting eﬀect at
subthreshold membrane voltages reducing RIn and increasingrheobase currents. Furthermore, during sustained depolarization
similar to periods of repetitive ﬁring, the voltage-dependent
strong currents produced after SMF application (Fig. 10) resultsin the reduction of the number of AP spikes ﬁred (Fig. 9) producing an overall decrease in intrinsic excitability of neurons following SMF.
Delving deeper into the plausible mechanism of tSMS action
on neurons, the neuronal membrane comprising the anisotropic
lipid bilayer (Rosen, 2003b) interspersed with transmembraneproteins may undergo a plethora of changes in response to the
SMF. An important clue to this eﬀect is our observation thatblockade of TTX-sensitive Na+ channels during SMF application
failed to depolarize the RMP, reduce RIn, and increase rheobase
currents in our current-clamp recording using a high Cl− internal
solution. TTX binds deep into the pore of voltage-gated Na+channels in close proximity to the selectivity ﬁlter and exhibits
use-dependent enhancement of block (Hille, 2001). In addition,binding of TTX impedes the return of voltage sensor in domain
IV to resting conformation (Capes et al., 2012). This aﬀects the10 • J. Neurosci., February 28, 2024 • 44(9):e1324222024

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• Intrinsic Plasticity in tSMS

Figure 9. Reduced intrinsic excitability of layer II/III pyramidal neurons of the motor cortex following SMF application rescued by GlyH-101 block of Cl− channels. a, Representative
subthreshold current injections (from −60 to +40 pA) with consequent voltage responses and repetitive ﬁring by suprathreshold current injection (+320 pA) in the CTRL (blue), post-SMF 1(orange), and post-SMF 1 + GlyH-101 (pink) groups. At the same subthreshold current injection, the voltage responses decreased in the post-SMF 1 group. At +320 pA current injection, thenumber of spikes for post-SMF 1 + GlyH-101 group exceed number of spikes elicited by CTRL group. Scale bar as shown in inset. b, A signiﬁcant decrease (K–W test; **p < 0.001) in the I–
O relationship for the post-SMF 1 (orange) group in comparison with both CTRL (blue) and post-SMF 1 + GlyH-101 (pink) groups. I–O relationship for post-SMF 1 + GlyH-101 signiﬁcantly increased
in comparison with CTRL group (K–W test; *p < 0.05; **p < 0.01). Error bars represent SEM. c–e, Individual data points are shown as open circles in blue (CTRL, n = 18), orange (post-SMF 1, n =18), and pink (post-SMF 1 + GlyH-101, n = 18). The mean values are indicated by solid bars next to the data points. Error bars represent SEM. Rheobase from layer II/III pyramidal neurons
signiﬁcantly increased in the post-SMF 1 group compared with CTRL and post-SMF 1 + GlyH-101 (one-way ANOVA; p = 0. 000004; post hoc, Ryan–Einot–Gabriel–Welsch F test; post-SMF 1
vs post-SMF + GlyH-101 with ***p < 0.001; post-SMF 1 vs CTRL with ***p < 0.001) groups. No signiﬁcant diﬀerences between post-SMF + GlyH-101 and CTRL groups (c). Input resistanceof pyramidal neurons signiﬁcantly lower in the post-SMF 1 group (K–W test; **p < 0.01; post hoc, stepwise step-down; post-SMF 1 vs post-SMF 1+ GlyH-101 with **p < 0.01; post-SMF 1
vs CTRL with **p < 0.01). No signiﬁcant diﬀerences between post-SMF + GlyH-101 and CTRL groups (d). RMP of pyramidal neurons signiﬁcantly lower in the post-SMF 1 group (K–W test;**p < 0.01; post hoc, stepwise step-down; post-SMF 1 vs post-SMF 1+ GlyH-101 with *p < 0.05; post-SMF 1 vs CTRL with *p < 0.05). No signiﬁcant diﬀerences between post-SMF +GlyH-101 and CTRL groups (e).

electromechanical coupling between the inner gating pore andvoltage sensor. A recent modelling study (Hernando et al.,2020) indicated that the application of SMF with inhomogeneous
gradients induced surface tension along the neuronal membrane,eﬀectively stretching them. They proposed that this mechanicalforce would be suﬃcient to interfere with the energy barriers governing the activation, inactivation, and deactivation of ion channels. Thus, we hypothesize that SMF action produces membrane
deformation facilitating the movement of voltage sensor of Na+
channels enhancing the Na+ entry into the neurons. Thereafter,
post-SMF the Cl− entry through open Cl− channels driven by
the electrochemical driving force for Na+ produce the prolonged
shunting inhibition. As application of TTX blocks Na+ entry andhinders transitions of the voltage sensor domain during SMF
application, it prevents alteration in intrinsic excitability.
This study showed the neurophysiological mechanism of
intrinsic plasticity in mice induced by moderate-intensity SMF
with the same magnetic parameters as those of tSMS in humans.
Intrinsic plasticity phenomenon is a nonsynaptic modulation ofneuronal excitability (Titley et al., 2017; Debanne et al., 2019).
Intrinsic plasticity has been reported to result from modiﬁcationsof voltage- or calcium-dependent potassium channels (Schreurs

et al., 1998; Frick et al., 2004; Belmeguenai et al., 2010) as wellas hyperpolarization-activated and cyclic nucleotide-gated(HCN) channels (Brager and Johnston, 2007). This is the ﬁrst
study to show that intrinsic plasticity may also be mediated by
Cl− channels. The relationship between intrinsic excitabilitychanges and synaptic plasticity has been proposed to be synergistic (Debanne et al., 2019). This is important because in vitro correlates of learning paradigms (LTP/LTD) strictly adhere to
protocols that essentially modify neuronal excitability. In addition, the observed temporary suppression of the intrinsic excitability of pyramidal neurons may lead to a consequential
reduction in the excitatory synaptic output. However, consideringthat the dynamic phenomena of homeostatic synaptic scaling andHebbian plasticity mechanisms are also at play, the intrinsic plasticity aﬀected by the SMF action may impact neural coding. The
enhanced shunting inhibition produced as a result of SMF (Figs.
2, 5, 9) may impact synaptic plasticity. Another interesting observation was that the SMF-induced GlyH-101-sensitive currents
also showed a rapidly activating and inactivating current component during the step voltage protocol (Fig. 10a). This componentof the Cl− current would aﬀect the summation of excitatory postsynaptic potentials (EPSPs). The regulation of [Cl−]i in neurons isSinha et al.

• Intrinsic Plasticity in tSMS

J. Neurosci., February 28, 2024 • 44(9):e1324222024 • 11

Figure 10. SMF application enhances GlyH-101-sensitive Cl− currents with similar voltage dependence to SLC26A11. a, Samples of GlyH-101-sensitive currents elicited by a voltage step
protocol (from −70 to +20 mV; 300 ms) in the CTRL (blue) and post-SMF 1 + GlyH-101 (magenta) groups. b, A signiﬁcant enhancement of GlyH-101-sensitive currents (independent samples ttest; **p < 0.001) as evidence by the current–voltage (I–V) relationship for the post-SMF 1 (magenta) group in comparison with CTRL (blue) group (CTRL, n = 14 cells; post-SMF 1, n = 14, 5
mice; t test, *p < 0.05; **p < 0.01; ***p < 0.001). Scale bar as shown in inset. Error bars represent SEM.

another fundamental factor for proper neural coding (Doyonet al., 2016) that aﬀects GABA action and the subsequent summation of synaptic inputs. GABA decrease in the primary motor area(M1) was positively correlated with the degree of motor learning(Stagg et al., 2011), and GABAA receptor agonist depressed theuse/practice-dependent plasticity of the M1 (Büteﬁsch et al.,
2000; Ziemann et al., 2001). SMF action-mediated temporarychanges in both intrinsic excitability and [Cl−]i would appear toaﬀect learning. Indeed, it was reported that tSMS alters sequential
implicit motor learning in humans (Nojima et al., 2019). Thiseﬀect of tSMS on motor learning, in addition to our ﬁndings,
would depend on the summation of network-level changes contributed by the SMF action on GABAergic interneurons, although
the eﬀect of SMF on motor learning remains to be investigated.
Taken together, our ﬁndings suggest that tSMS can induce motor
cortical plasticity in humans by regulating the activity of a certain
type of Cl− channel in neurons.

Data Availability
Data that support the ﬁndings will be made available upon
reasonable request to the corresponding author.

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