1. Neuroscience

PRRT2 as an auxiliary regulator of Nav channel slow inactivation

  1. Bin Lu  Is a corresponding author
  2. Qi-Wu Xu
  3. Jing Zhang
  4. Xue-Mei Wu
  5. Jun-Yan He
  6. Jing-Qiu Peng
  7. Guang Yang
  8. Ke-Xian Li
  9. Ling Zhuang
  10. Yu-Xian Zhang
  11. Zhi-Ying Wu
  12. Zhi-Qi Xiong  Is a corresponding author
  1. Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, China
  2. University of Chinese Academy of Sciences, China
  3. School of Life Science and Technology, ShanghaiTech University, China
  4. Department of Neurology, Zhongshan Hospital, Fudan University, China
  5. Department of Medical Genetics and Center for Rare Diseases, Second Affiliated Hospital, Zhejiang University School of Medicine and Zhejiang Key Laboratory of Rare Diseases for Precision Medicine and Clinical Translation, China
  6. Shanghai Center for Brain Science and Brain-Inspired Technology, China
https://doi.org/10.7554/eLife.109327.4
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Cite this article
  1. Bin Lu
  2. Qi-Wu Xu
  3. Jing Zhang
  4. Xue-Mei Wu
  5. Jun-Yan He
  6. Jing-Qiu Peng
  7. Guang Yang
  8. Ke-Xian Li
  9. Ling Zhuang
  10. Yu-Xian Zhang
  11. Zhi-Ying Wu
  12. Zhi-Qi Xiong
(2026)
PRRT2 as an auxiliary regulator of Nav channel slow inactivation
eLife 14:RP109327.
https://doi.org/10.7554/eLife.109327.4
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eLife Assessment

This important study identifies PRRT2 as an auxiliary regulator of Nav channel slow inactivation in vitro and in vivo, demonstrating that PRRT2 facilitates entry into, and delays recovery from, the slow-inactivated state. The revised manuscript has been substantially strengthened, providing compelling evidence that PRRT2 is relevant to normal brain physiology and disease pathophysiology, providing a mechanistic link between PRRT2 mutations and episodic neurological phenotypes. Overall, this study will be of interest to ion channel biophysicists and neurophysiologists, particularly those studying channelopathies.

https://doi.org/10.7554/eLife.109327.4.sa0

Abstract

During sustained activity, voltage-gated sodium (Nav) channels enter a slow-inactivated state to limit cellular hyperexcitability. Disruption of this regulatory process has been implicated in skeletal, cardiac, and neurological disorders. While the kinetics of this process are well characterized, its endogenous modulators remain unclear. Here, we identify Proline-Rich Transmembrane Protein 2 (PRRT2) as a native regulator of Nav channel slow inactivation. We show that PRRT2 facilitates the entry of Nav channels into the slow-inactivated state and delays their recovery, a regulatory effect conserved from zebrafish to humans. PRRT2 forms molecular complexes with Nav channels both in vitro and in vivo. In the mouse cortex, PRRT2 deficiency impairs the slow inactivation of Nav channels in neuronal axons, leading to reduced cortical resilience in response to hyperexcitable challenges. Together, these findings establish PRRT2 as a physiological modulator of Nav channel slow inactivation and reveal a mechanism that supports cortical resilience to pathological perturbations.

Introduction

Voltage-gated sodium (Nav) channels are critical for the initiation and propagation of electrical signals in excitable cells, contributing to cellular excitability in various tissues (Hodgkin and Huxley, 1952; Ulbricht, 1977). In mammals, including humans, Nav channels consist of nine α-subunit isoforms (Nav1.1-Nav1.9), which are differentially expressed and distributed across different tissues to regulate specific physiological functions (Catterall et al., 2005). Mutations in Nav channels that lead to either loss- or gain-of-function of these channels are associated with a wide spectrum of diseases, including neurological disorders (Escayg and Goldin, 2010; Hedrich et al., 2019; Meisler et al., 2021), pain syndromes (Dib-Hajj et al., 2017; Waxman, 2013), muscle illnesses (Cannon, 2018; Mantegazza et al., 2021), and cardiac diseases (Remme, 2013). Consequently, the Nav channels are considered promising therapeutic targets for treating these conditions (Noreng et al., 2021; Wisedchaisri and Gamal El-Din, 2022).

Voltage-dependent conformational changes enable Nav channels to transition between resting, open, and inactivated states (Catterall et al., 2020). Eukaryotic Nav channels undergo two major forms of inactivation (fast and slow inactivation), which are distinguished by their kinetics of onset and recovery (Goldin, 2003). Fast inactivation is triggered within milliseconds by brief depolarization and is rapidly reversed upon hyperpolarization, typically within tens of milliseconds (Armstrong and Bezanilla, 1977; Bezanilla and Armstrong, 1977). In contrast, slow inactivation develops during prolonged depolarizations and the slow inactivated channels require substantially longer periods to recover, ranging from hundreds of milliseconds to minutes (Rudy, 1978). Previous studies have shown that slow inactivation regulates Nav channel availability during sustained depolarization or high-frequency repetitive activity, thereby modulating the cellular excitability (Goldin, 2003; Silva, 2014; Vilin and Ruben, 2001). Impairment of this regulatory process due to mutations in Nav channels has been associated with disorders such as periodic paralysis (Hayward et al., 1999), cardiac arrhythmias (Richmond et al., 1998; Vilin et al., 1999), and epilepsy (Ghovanloo et al., 2023).

Beyond the intrinsic hierarchical gating mechanisms, auxiliary regulators further modulate Nav channel inactivation dynamics (Goldin, 2003). For example, β subunits of sodium channels (SCN1B) regulate inactivation of Nav channels in the heterologous expression system, although their effects vary across different Nav isoforms and cell types (He and Soderlund, 2014; Vilin et al., 1999; Webb et al., 2009). Fibroblast growth factor homologous factors (FHFs) interact with the carboxy terminus of Nav channels (Gade et al., 2020; Goetz et al., 2009), promoting intermediate- or long-term inactivation, with recovery times spanning hundreds of milliseconds (Dover et al., 2010; Venkatesan et al., 2014). Although these auxiliary proteins have been identified as regulators of fast or intermediate inactivation, native modulators that specifically target slow inactivation of Nav channels remain largely unknown.

Recent studies on proline-rich transmembrane protein 2 (PRRT2) provide new insights into this issue. The PRRT2, a member of the Dispanins B subfamily (DspB) (Sällman Almén et al., 2012), has been identified as a causative gene for paroxysmal kinesigenic dyskinesia (Chen et al., 2011; Lee et al., 2012; Wang et al., 2011). In vitro studies have shown that PRRT2 regulates Nav channel surface expression and delays recovery of these channels from inactivated states induced by brief depolarization (Fruscione et al., 2018; Lu et al., 2021; Valente et al., 2023). However, these studies did not clearly distinguish between the effects of PRRT2 on fast versus slow inactivation, nor did they provide direct in vivo evidence for its role in regulating Nav channel inactivation. Our original observations that PRRT2 expression increased the accumulation of inactivated Nav channels during repetitive stimulation (Lu et al., 2021) suggest that PRRT2 may be a previously unrecognized auxiliary regulator of Nav channel slow inactivation.

In this study, we combined whole-cell voltage-clamp recordings, co-immunoprecipitation, and in vivo approaches to investigate the role of PRRT2 in regulating Nav channel slow inactivation, and to examine how deficits in PRRT2-mediated modulation of this process affect neuronal function in awake mice.

Results

PRRT2 regulates the slow inactivation of Nav1.2 channels

Fast and slow inactivation of Nav channels proceed through distinct gating pathways (Goldin, 2003). Although previous studies have implicated PRRT2 in the regulation of Nav channel inactivation (Fruscione et al., 2018; Lu et al., 2021), it remains unclear whether PRRT2 primarily influences fast inactivation, slow inactivation, or both. To address this, we expressed mouse PRRT2 in HEK293T cells stably expressing Nav1.2 (Figure 1A) and applied classic electrophysiological paradigms to distinguish between the two inactivation processes.

Figure 1 with 1 supplement see all
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PRRT2 regulates slow inactivation of Nav1.2 channels.

(A, B) Representative immunoblots showing PRRT2 protein expression (A) and schematic of whole-cell recording (B) in Nav1.2-stably expressing HEK293T cells transfected with constructs encoding either EGFP or mouse PRRT2 (mPRRT2). MW, molecular weight. (C) Fast-inactivation decay time constants of sodium currents evoked by 20 ms depolarizing steps to the indicated command voltages (EGFP: n=13 cells, mPRRT2: n=13 cells). Insert: representative sodium current traces elicited by a brief depolarization (left) and voltage-clamp protocol used to assess fast inactivation (right). The arrow indicates the decay of the sodium current attributable to fast inactivation. (D) Recovery from fast inactivation of Nav1.2 channels (EGFP: n=10 cells, mPRRT2: n=7 cells). Insert: Protocol for examining fast inactivation recovery. (E) Schematic diagram of entry into and recovery from slow inactivation of voltage-gated sodium channels. Slow inactivation modulates the availability of the sodium channels. (F) Entry into slow-inactivated state of Nav1.2 channels induced by progressively longer depolarization (EGFP: n=6 cells, mPRRT2: n=6 cells). (G) The effects of mPRRT2 on the recovery of Nav1.2 channels from slow-inactivated state induced by 5 s depolarization (EGFP: n=6 cells, mPRRT2: n=6 cells). Data are presented as mean ± s.e.m. Two-tailed, unpaired Student’s t-test was used in (C), and two-way ANOVAs were used in (D), (F), and (G). ****p<0.0001. n.s., not significant. Source data are provided in Figure 1—source data 1–3.

Figure 1—source data 1

Original western blot images for panel A.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig1-data1-v1.zip
Figure 1—source data 2

PDF file containing original western blot images with the relevant bands labeled for panel A.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig1-data2-v1.zip
Figure 1—source data 3

Raw data used for the quantifications shown in panels C, D, F, and G.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig1-data3-v1.xlsx

Fast inactivation of Nav channels is rapidly triggered by brief depolarization and manifests as a current decay following a transient pore opening (Figure 1B and C), consistent with previous reports (Misra et al., 2008; Thompson et al., 2023). To evaluate the kinetics of fast inactivation, we measured the time constant of current decay across a range of test potentials and found that PRRT2 expression did not significantly alter the rate at which Nav channels entered the fast-inactivated state (Figure 1C). Moreover, the majority of Nav1.2 channels recovered from fast inactivation within ~10 ms at a hyperpolarized potential of –120 mV, and PRRT2 expression had a minimal effect on this recovery (Figure 1D).

In contrast to fast inactivation, Nav channel slow inactivation develops and recovers over much longer timescales (Figure 1E), typically ranging from hundreds of milliseconds to seconds, or even to minutes (Silva, 2014). Because slow-inactivated channels do not recover during brief hyperpolarization, they can be functionally distinguished from fast-inactivated channels using a classic two-pulse protocol. In this approach, a brief hyperpolarization step (e.g., –‍120 mV, 10 ms) is inserted between a conditioning pulse and a test pulse to recover fast-inactivated channels. Channels that have entered the slow-inactivated state remain unavailable during the test pulse, resulting in a reduction in peak sodium current (Figure 1E; Featherstone et al., 1996).

To assess the development of slow inactivation, we applied depolarization pulses of variable duration, followed by a fixed 10 ms hyperpolarization before the test pulse (Figure 1F). We found that a 5 s depolarization at 0 mV was sufficient to drive most Nav1.2 channels into the slow-inactivated state (Figure 1F), consistent with previous studies (Ganguly et al., 2021). PRRT2 significantly promoted the entry of Nav1.2 channels into the slow-inactivated state during sustained depolarization (Figure 1F).

To examine the role of PRRT2 in channel recovery from slow inactivation, we used a protocol in which a fixed 5 s depolarization at 0 mV was followed by varying durations of hyperpolarization at –120 mV to allow channel repriming (Figure 1G). PRRT2 expression markedly slowed the recovery of Nav channels from the slow-inactivated state (Figure 1G).

To further investigate whether the regulation of PRRT2 in slow inactivation is voltage dependent, we performed a steady-state slow inactivation protocol. In this assay, 10 s depolarizing steps from –110 to –10 mV were applied to drive Nav channels into steady state, followed by a fixed hyperpolarization and a test pulse to assess channel availability (Figure 1—figure supplement 1A). We found that PRRT2 promotes the transition of Nav channels into the slow-inactivated state across a broad range of membrane potentials. PRRT2 expression induced a pronounced hyperpolarizing shift in the inactivation curve, with a V0.5 of –76.27 mV compared to –48.05 mV in controls (Figure 1—figure supplement 1B).

In addition to prolonged depolarization, high-frequency repetitive depolarization can also induce Nav channel slow inactivation (Mickus et al., 1999). To determine whether PRRT2 regulates slow inactivation under these conditions, we applied a pulse-train protocol with an inter-pulse recovery interval of 10 ms. The use-dependent attenuation of peak sodium current was quantified as the fraction of current remaining across successive depolarizations (Figure 1—figure supplement 1C). Compared with the EGFP control, PRRT2 expression produced a greater decline in peak current during the train, consistent with enhanced slow inactivation during repetitive depolarizations (Figure 1—figure supplement 1C). Together, these findings indicate that PRRT2 is a potent regulator of Nav1.2 channel slow inactivation in vitro.

Functional auxiliary factors in the regulation of Nav channel slow inactivation

Several auxiliary factors of Nav channels have been identified, including the SCN1B and FHF2A (also known as FGF13a), both of which are implicated in modulating Nav channel inactivation (Dover et al., 2010; Isom et al., 1992). In this study, we compared the effects of mouse SCN1B and FGF13a with those of PRRT2 on the development and recovery of Nav channel slow inactivation (Figure 2A).

Figure 2 with 1 supplement see all
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Functional auxiliary factors in the regulation of Nav channel slow inactivation.

(A) Diagram of Nav channel with potent auxiliary factors, encompassing SCN1B, FHF2A (also known as FGF13a), and PRRT2. (B) The effects of auxiliary factors on the entry of Nav1.2 channels into slow inactivation (EGFP: n=12 cells, mPRRT2: n=14 cells, SCN1B: n=9 cells, FGF13a: n=12 cells). (C) The effects of auxiliary factors on the recovery of Nav1.2 channels from slow-inactivated state induced by 5 s depolarization (EGFP: n=12 cells, mPRRT2: n=12 cells, SCN1B: n=12 cells, FGF13a: n=12 cells). (D) Schematic representation of the putative topology of mouse PRRT2 and three truncations. Numbers indicate the positions of the amino acid. REL, re-entrant loop. TM, transmembrane domain. N, amino terminus. C, carboxyl terminus. aa, amino acid. (E) Effects of PRRT2 truncations on the entry of Nav1.2 channels into slow inactivation (EGFP: n=5 cells, mPRRT2(1–266): n=14 cells, mPRRT2(222–346): n=12 cells, mPRRT2(256–346): n=12 cells). (F) Effects of PRRT2 truncations on the recovery of Nav1.2 channels from slow-inactivated state induced by 5 s depolarization (EGFP: n=5 cells, mPRRT2(1–266): n=10 cells, mPRRT2(222–346): n=10 cells, mPRRT2(256–346): n=12 cells). Data are presented as mean ± s.e.m. The main effect of group was assessed using two-way ANOVAs (B, C, E, F). **p<0.01, ****p<0.0001, n.s., not significant. Source data are provided in Figure 2—source data 1.

Figure 2—source data 1

Raw data used for the quantifications shown in panels B, C, E, and F.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig2-data1-v1.xlsx

We found that SCN1B had negligible effects on both the entry into and recovery from the slow-inactivated state of Nav1.2 compared to controls (Figure 2B and C). In contrast, FGF13a markedly accelerated the onset of inactivation (Figure 2B), consistent with previous reports (Dover et al., 2010; Venkatesan et al., 2014). Notably, FGF13a had minimal effect on the recovery of Nav channels from the slow-inactivated state induced by a 5 s depolarization (Figure 2C). These distinct effects of FGF13a on inactivation are consistent with the idea that FGF13a does not primarily modulate classical slow inactivation, but rather promotes a form of rapid-onset and long-term inactivation (Dover et al., 2010; Venkatesan et al., 2014). By comparison, PRRT2 uniquely regulated slow inactivation by both facilitating entry into and delaying recovery from the slow-inactivated state of Nav channels (Figure 2B and C).

The rodent Prrt2 gene encodes a 346-amino-acid protein containing a proline-rich domain in the N-terminus, a re-entrant loop, and a transmembrane domain in the C-terminus (Figure 2D). To identify the functional domain(s) responsible for regulating slow inactivation, we generated a series of protein-truncating variants (Figure 2D).

We first examined PRRT2(222–346), which lacks the N-terminal proline-rich domain (Figure 2D). This variant retained activity similar to the full-length PRRT2, enhancing slow inactivation development and delaying recovery (Figure 2E and F). A shorter construct, PRRT2(256–346), which retains the intact C-terminus, also preserved regulatory function in modulating slow inactivation compared to the EGFP control (Figure 2E and F). To further test whether the C-terminal region is necessary for the effect of PRRT2 on Nav channel slow inactivation, we examined PRRT2(1–266), a truncated variant lacking the membrane-associated C-terminal domains (Figure 2D). Unlike variants containing the C-terminal region, PRRT2(1–266) showed no detectable effect on either the development or recovery of Nav channel slow inactivation compared to the control (Figure 2E and F). Notably, this lack of effect was not attributable to absent expression of the truncated construct (Figure 2—figure supplement 1A and B). Together, these results indicate that the C-terminal region of PRRT2, including the re-entrant loop and transmembrane domain, is both necessary and sufficient to mediate its regulatory effect on slow inactivation of Nav1.2 channels in vitro.

Evolutionarily conserved effects of PRRT2 on Nav channel slow inactivation

The PRRT2 and PRRT2-like genes emerged in vertebrates and have been conserved throughout evolution (Sällman Almén et al., 2012). The alignment of amino acid sequences from zebrafish, mouse, and human PRRT2 revealed 77.46% identity between human and mouse PRRT2, and 43.60% identity between zebrafish and mouse PRRT2 (Figure 3A and B). Notably, the majority of conserved residues are located within the C-terminal region of the PRRT2 proteins (Figure 3A). Given the essential role of the C-terminus of mouse PRRT2 in regulating Nav channel slow inactivation, we examined whether PRRT2 proteins from different species exhibit similar effects.

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Evolutionarily conserved effects of PRRT2 on Nav channel slow inactivation.

(A) Sequence alignment of PRRT2 protein in human, mouse, and zebrafish. Conserved amino acids are highlighted and the membrane-associated domains in the carboxyl terminus of PRRT2 are underlined. REL, re-entrant loop. TM, transmembrane domain. (B) Sequence identity of PRRT2 protein in human (hPRRT2), mouse (mPRRT2), and zebrafish (zfPRRT2). Sequence identity is calculated by using the protein BLAST tool from NCBI. (C) The effects of PRRT2 from different species on the entry of Nav1.2 channels into slow inactivation (EGFP: n=16 cells, mPRRT2: n=19 cells, hPRRT2: n=20 cells, zfPRRT2: n=11 cells). (D) The effects of PRRT2 from different species on the recovery of Nav1.2 channels from the slow-inactivated state induced by 5 s depolarization (EGFP: n=16 cells, mPRRT2: n=19 cells, hPRRT2: n=19 cells, zfPRRT2: n=11 cells). (E) Diagram for the chimeric construct of PRRT2 originated from human and zebrafish. (F) The effect of chimeric PRRT2 on the entry of Nav1.2 channels into slow inactivation (EGFP: n=11 cells, hPRRT2: n=6 cells, zfPRRT2: n=10 cells, Chimera: n=11 cells). (G) The effect of chimeric PRRT2 on the recovery of Nav1.2 channels from the slow-inactivated state induced by 5 s depolarization (EGFP: n=11 cells, hPRRT2: n=6 cells, zfPRRT2: n=10 cells, Chimera: n=9 cells). Data are presented as mean ± s.e.m. The main effect of group was assessed using two-way ANOVAs (C, D, F, G). **p<0.01, ****p<0.0001, n.s., not significant. Source data are provided in Figure 3—source data 1.

Figure 3—source data 1

Raw data used for the quantifications shown in panels C, D, F, and G.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig3-data1-v1.xlsx

By comparing the effects of zebrafish, mouse, and human PRRT2 on Nav channel slow inactivation, we found that all three PRRT2 orthologs similarly promoted the entry of Nav1.2 channels into the slow-inactivated state (Figure 3C). Notably, in recovery assays, zebrafish PRRT2 was more effective at delaying the recovery process compared to its mammalian counterparts (Figure 3D). To identify the domain responsible for this enhanced effect, we generated a chimeric PRRT2 construct by fusing the C-terminal region of zebrafish PRRT2 to the N-terminal region of human PRRT2 (Figure 3E). The chimeric PRRT2 exhibited comparable effects on the rate of slow inactivation development to those observed with both human and zebrafish PRRT2 (Figure 3F). Notably, the chimeric variant displayed enhanced modulation of recovery, resembling zebrafish PRRT2 and significantly differing from human PRRT2 (Figure 3G). These findings indicate that the C-terminal region of zebrafish PRRT2 confers its greater ability to modulate recovery of Nav1.2 from the slow-inactivated state.

We next extended our analysis to other DspB family members, including Trafficking regulator of GLUT4-1 (Trarg1) and Transmembrane protein 233 (Tmem233), both of which encode proteins with C-terminal regions similar to that of PRRT2 (Figure 3—figure supplement 1A and B). Given the critical role of the PRRT2 C-terminus in modulating slow inactivation, we hypothesized that TRARG1 and TMEM233 might exert similar regulatory effects. Under heterologous expression conditions, both TRARG1 and TMEM233 facilitated the entry of Nav1.2 channels into the slow-inactivated state and delayed their recovery (Figure 3—figure supplement 1C and D). These effects were comparable to those observed with mouse PRRT2, although TMEM233 exhibited slightly weaker modulation than PRRT2 and TRARG1 (Figure 3—figure supplement 1C and D). Together, these results suggest that the role of PRRT2 in regulating Nav channel slow inactivation is evolutionarily conserved.

PRRT2 promotes slow inactivation across multiple human Nav channel isoforms

In humans, the nine Nav channel isoforms (Nav1.1–1.9) share high sequence homology and similar structural features (Goldin, 2001). To determine whether PRRT2 modulates slow inactivation across different Nav channel isoforms, we selected Nav1.1, Nav1.4, Nav1.5, and Nav1.6, which are expressed in distinct tissues and cell types, and assessed the effects of human PRRT2 on their slow inactivation properties (Figure 4A–J).

Figure 4
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PRRT2 promotes slow inactivation across multiple human Nav channel isoforms.

(A) Protocol for examining the entry of Nav channels into slow-inactivated state. (B–E) The effects of human PRRT2 (hPRRT2) on the entry of Nav1.1, Nav1.4, Nav1.5, and Nav1.6 channels into slow inactivation. In (B), analyses for Nav1.1 (EGFP: n=10 cells, hPRRT2: n=12 cells); in (C), analyses for Nav1.4 (EGFP: n=11 cells, hPRRT2: n=11 cells); in (D), analyses for Nav1.5 (EGFP: n=17 cells, hPRRT2: n=17 cells); in (E), analyses for Nav1.6 (EGFP: n=5 cells, hPRRT2: n=4 cells). (F) Protocol for examining the recovery of Nav channels from slow-inactivated state. (G–J) The effects of human PRRT2 on the recovery of Nav1.1, Nav1.4, Nav1.5, and Nav1.6 channels from slow-inactivated state induced by 5 s depolarization. In (G), analyses for Nav1.1 (EGFP: n=10 cells, hPRRT2: n=12 cells); in (H), analyses for Nav1.4 (EGFP: n=11 cells, hPRRT2: n=11 cells); in (I), analyses for Nav1.5 (EGFP: n=16 cells, hPRRT2: n=15 cells); in (J), analyses for Nav1.6 (EGFP: n=5 cells, hPRRT2: n=4 cells). Data are presented as mean ± s.e.m. The main effect of group was assessed using two-way ANOVAs (B–E and G–J). **p<0.01, ***p<0.001, ****p<0.0001. Source data are provided in Figure 4—source data 1.

Figure 4—source data 1

Raw data used for the quantifications shown in panels B, C, D, E, G, H, I, and J.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig4-data1-v1.xlsx

Nav1.1, Nav1.4, and Nav1.6 displayed similar kinetics in the development and recovery of slow inactivation (Figure 4B, C, E, G, H, and J). In contrast, Nav1.5 showed greater resistance to entering the slow-inactivated state (Figure 4D and I), consistent with previous findings (Richmond et al., 1998). Despite these isoform-specific differences, the heterologous expression of PRRT2 consistently promoted entry of Nav channels into the slow-inactivated state (Figure 4A–E) and delayed recovery from it (Figure 4F–J) across all four Nav isoforms tested. These results suggest that PRRT2 modulates slow inactivation through a common mechanism shared by multiple Nav channel isoforms, despite differences in their intrinsic slow-inactivation properties.

Association between PRRT2 and Nav channels in vitro

PRRT2 may regulate slow inactivation through association with Nav channels. To examine the potential physical association between human PRRT2 and Nav1.2, we co-expressed PRRT2-HA and Flag-Nav1.2 in HEK293T cells and performed co-immunoprecipitation (Co-IP) assays (Figure 5A and B). As controls, cells were transfected with one of the two interacting proteins (either PRRT2-HA or Flag-Nav1.2) together with the empty vectors carrying the corresponding tag of the other protein (Figure 5A and B). Immunoprecipitation using an anti-Flag antibody to capture Flag-Nav1.2 successfully co-precipitated PRRT2 in the co-transfection group, but not in the control groups where either protein was expressed (Figure 5A). Conversely, immunoprecipitation with anti-HA antibody showed that Nav1.2 was specifically co-precipitated with PRRT2-HA in the co-transfection group but not in controls (Figure 5B). These results confirm that PRRT2 and Nav1.2 formed a protein complex in vitro.

Figure 5
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Association between PRRT2 and Nav channels in vitro.

(A, B) Schematic diagram of the co-immunoprecipitation assay used to detect association between human PRRT2 and Nav1.2. Flag-tagged Nav1.2 (+), HA-tagged PRRT2 (+), and empty (-) vectors were transfected in HEK293T cells as indicated. The cell lysates were immunoprecipitated with anti-Flag (A) or anti-HA (B) magnetic beads, the captured proteins were analyzed by SDS-PAGE and immunoblotting. (C) Diagram for HA-tagged truncation of human PRRT2 (PRRT2(1–268)), in which the carboxyl terminus of PRRT2 was deleted. REL, re-entrant loop. TM, transmembrane domain. (D, E) Co-immunoprecipitation assay for potential association between Nav1.2 and PRRT2(1–268). The proteins immunoprecipitated by anti-Flag (D) or anti-HA (E) magnetic beads were analyzed by immunoblotting. Note that HA-tagged PRRT2(1–268) was detected by anti-HA antibody in (D, E). (F) Diagram for HA-tagged truncation of human PRRT2 (PRRT2(250–340)), in which the amino terminus of PRRT2 was deleted. (G, H) Co-immunoprecipitation assay for potential association between Nav1.2 and PRRT2(250–340). The proteins immunoprecipitated by anti-Flag (G) or anti-HA (H) were analyzed by immunoblotting. Note that HA-tagged PRRT2(250–340) was detected by anti-HA antibody in (G, H). (I, J) Co-immunoprecipitation assay for potential association between Nav1.1 and PRRT2. The proteins captured by anti-Flag (I) or anti-HA (J) magnetic beads were analyzed by immunoblotting. In (A, B, D, E, G, H, I, J), blots shown are representative of at least three independent co-immunoprecipitation experiments with similar results. Red rectangles indicate the immunoblotting of bait and potential prey proteins in co-immunoprecipitation. IB, immunoblot. IP, immunoprecipitation. MW, molecular weight. Source data are provided in Figure 5—source data 1–16.

Figure 5—source data 1

Original western blot images for panel A.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data1-v1.zip
Figure 5—source data 2

PDF file containing original western blot images with the relevant bands labeled for panel A.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data2-v1.zip
Figure 5—source data 3

Original western blot images for panel B.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data3-v1.zip
Figure 5—source data 4

PDF file containing original western blot images with the relevant bands labeled for panel B.

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Figure 5—source data 5

Original western blot images for panel D.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data5-v1.zip
Figure 5—source data 6

PDF file containing original western blot images with the relevant bands labeled for panel D.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data6-v1.zip
Figure 5—source data 7

Original western blot images for panel E.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data7-v1.zip
Figure 5—source data 8

PDF file containing original western blot images with the relevant bands labeled for panel E.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data8-v1.zip
Figure 5—source data 9

Original western blot images for panel G.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data9-v1.zip
Figure 5—source data 10

PDF file containing original western blot images with the relevant bands labeled for panel G.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data10-v1.zip
Figure 5—source data 11

Original western blot images for panel H.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data11-v1.zip
Figure 5—source data 12

PDF file containing original western blot images with the relevant bands labeled for panel H.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data12-v1.zip
Figure 5—source data 13

Original western blot images for panel I.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data13-v1.zip
Figure 5—source data 14

PDF file containing original western blot images with the relevant bands labeled for panel I.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data14-v1.zip
Figure 5—source data 15

Original western blot images for panel J.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data15-v1.zip
Figure 5—source data 16

PDF file containing original western blot images with the relevant bands labeled for panel J.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig5-data16-v1.zip

To identify the Nav1.2-associating region within human PRRT2, we generated truncated constructs encoding either the N-terminus (residues 1–268) or C-terminus (residues 250–340) of PRRT2. Using the same Co-IP approach, we found that PRRT2(1–268), which contains the intracellular proline-rich domain, failed to associate with Nav1.2 (Figure 5C–E). In contrast, the PRRT2(250–340), which includes the re-entrant loop and transmembrane domain, was able to interact with Nav1.2 (Figure 5F–H). These results indicate that the C-terminal region of PRRT2 is necessary and sufficient, in this assay, for association with Nav1.2 channels.

Given that PRRT2 modulates slow inactivation across multiple Nav isoforms, we hypothesized that it might also associate with other Nav isoforms, such as Nav1.1. To test this, we co-expressed PRRT2-HA and Flag-Nav1.1 in HEK293T cells and performed Co-IP assays. Under these conditions, PRRT2 was found to associate with Nav1.1 (Figure 5I and J), similar to the association observed with Nav1.2 (Figure 5A and B).

These findings contrast with previous reports, which suggested that PRRT2 preferentially interacted with Nav1.2 over Nav1.1 in vitro (Franchi et al., 2023; Fruscione et al., 2018; Valente et al., 2023). This discrepancy may stem from differences in the experimental conditions used to assess protein–protein interactions. For example, in our study, Nav1.1 and PRRT2 were co-expressed in HEK293T cells, and cells were solubilized using the detergent n-Dodecyl β-D-maltoside (DDM). Variations in experimental conditions could influence the stability of the PRRT2-Nav channel complex and the efficiency of the Co-IP assay.

Collectively, our findings suggest that PRRT2 associates with Nav channels through its membrane-associated C-terminal domains, forming molecular complexes in vitro.

PRRT2 forms a molecular complex with Nav1.2 in the mouse brain

To extend our findings from the heterologous expression system to an in vivo context, we first investigated whether PRRT2 forms a protein complex with Nav channels in brain tissue. Due to the limited efficacy of available antibodies for immunoprecipitating native PRRT2 or Nav1.2 from brain lysates, we employed a tag-based capture strategy, which relies on the appropriate insertion of a commonly used epitope tag into the target protein.

We generated Prrt2-V5 knock-in mice via CRISPR/Cas9 technology (Yang et al., 2014), in which a V5 epitope tag was inserted at the C-terminus of PRRT2 (Figure 6A and B). Notably, compared with wild-type mice, PRRT2 protein levels in tissue lysates were markedly reduced in Prrt2-V5 knock-in mice (Figure 6C and D). These mice displayed susceptibility to dystonia attacks, resembling the phenotype previously observed in Prrt2-mutant mice (Wu et al., 2025), but with lower penetrance (Figure 6—figure supplement 1A–C).

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PRRT2 forms a molecular complex with Nav1.2 in the mouse brain.

(A) Schematic showing the generation of Prrt2-V5 knock-in (V5-KI) mice using Cas9 technology. V5-tag sequence was inserted precisely at the end of Prrt2 gene. PAM, protospacer-adjacent motif (NGG). ssDNA, single-stranded DNA. (B) Sanger sequencing around sgRNA targeting site in Prrt2-V5 knock-in mouse. The V5-tag, stop codon, and homologous arms were denoted. (C) Co-immunoprecipitation assay for potential interaction between Nav1.2 and PRRT2-V5 in the brain tissue. The proteins immunoprecipitated by anti-V5 nanobody were analyzed by immunoblotting. Blots shown are representative of four independent co-immunoprecipitation experiments. IB, immunoblot. IP, immunoprecipitation. MW, molecular weight. WT, wild-type. KI, knock-in. (D–F) Quantification of the density of protein bands for PRRT2 (D) and Nav1.2 (E) in lysate, and for Nav1.2 and ATP1B2 in co-immunoprecipitation experiments (F) (n=4 mice for each group). Data are presented as mean ± s.e.m. In (D–F), two-tailed, paired Student’s t-test was used for determining statistical significance. **p<0.01, ****p<0.0001, n.s., not significant. Source data are provided in Figure 6—source data 1–3.

Figure 6—source data 1

Original western blot images for panel C.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig6-data1-v1.zip
Figure 6—source data 2

PDF file containing original western blot images with the relevant bands labeled for panel C.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig6-data2-v1.zip
Figure 6—source data 3

Raw data used for the quantifications shown in panels D–F.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig6-data3-v1.xlsx

Despite the reduced PRRT2 expression, Nav1.2 expression remained unchanged (Figure 6E), and co-immunoprecipitation assays remained effective in the Prrt2-V5 knock-in mice. Using anti-V5 nanobody-conjugated magnetic beads, PRRT2 was effectively immunoprecipitated from membrane lysates of brain tissue (Figure 6C). A notable amount of Nav1.2 was co-immunoprecipitated in the Prrt2-V5 knock-in group, whereas only trace levels were detected in wild-type controls (Figure 6C and F). In contrast, ATP1B2, another membrane protein, was not co-immunoprecipitated in the Prrt2-V5 knock-in group (Figure 6C and F), supporting the specificity of the association between PRRT2 and Nav1.2. Together, although the V5 knock-in allele is hypomorphic, these results provide in vivo evidence that PRRT2 forms a molecular complex with Nav1.2 channels in the mouse brain.

PRRT2 deficiency impairs the regulation of Nav channel slow inactivation in cortical neuronal axons

Given the dense distribution of both PRRT2 and Nav channels in the cerebral and cerebellar cortices (Yamano et al., 2022), we next investigated whether PRRT2 regulates Nav channel slow inactivation in cortical neurons. Cortical slices were prepared from wild-type and Prrt2-mutant mice, the latter lacking PRRT2 expression (Tan et al., 2018). Unlike the relatively simple morphology of HEK293T cells, pyramidal neurons have complex arborizations, which present space-clamp limitations during whole-cell voltage-clamp recordings (Bar-Yehuda and Korngreen, 2008). To mitigate these issues, we followed established protocols to isolate axonal blebs from cortical pyramidal neurons (Hu and Shu, 2012). These spherical, membrane-bound blebs form at the cutting surface of cortical slices and are readily visualized (Figure 7A). Sodium currents recorded from axon blebs were voltage-dependent and sensitive to tetrodotoxin (TTX) (Figure 7A), confirming that they were primarily mediated by Nav channels (Hu and Shu, 2012).

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PRRT2 deficiency impairs the regulation of Nav channel slow inactivation and neuronal resilience in mice.

(A) Schematic showing the isolated axonal bleb recording in cortical neuron (left) and the representative brightfield image for the axonal bleb in brain slice (middle). The arrow indicates an axonal bleb. Scale bar, 20 μm. The representative sodium current traces recorded in axonal blebs (right). Scale bar, 0.5 nA, 5 ms. TTX, tetrodotoxin. (B) Sodium current density recorded in isolated axonal blebs from wild-type (WT) and Prrt2-mutant mice (WT: n=18 blebs from 7 mice, Prrt2-mutant: n=20 blebs from 8 mice). Sodium currents were measured with a brief depolarization to 0 mV preceded by a 200 ms hyperpolarizing prepulse to –110 mV. (C) Protocol for assessing Nav channel slow-inactivation (upper) and the representative traces of the sodium currents evoked by conditioning depolarization pulse and test pulses (bottom). Scale bar, 0.5 nA, 5 ms. (D) Fraction of available Nav channels after 5 s depolarization (WT: n=18 blebs from 7 mice, Prrt2-mutant: n=19 blebs from 8 mice). (E) Schematic illustration of the cortical electrostimulation and EEG recording in mice. Ref, reference electrode. Gnd, ground electrode. Stim, Stimulation. (F) Illustration showing the experimental schedule and the stimulation parameters. Stimulation was delivered once daily with stepwise increases in current intensity. (G) Representative traces and power of EEG signals in wild-type and Prrt2-mutant mice before and after 2 s cortical stimulation. After-discharges are indicated by the arrow and red line. Scale bar: 0.2 mV. Electrical stimulation (120 µA) was applied during 60–62 s period, where the stimulus-induced artifacts were removed prior to analysis. (H) Threshold of electrical stimulation to induce after-discharges in WT and Prrt2-mutant mice (WT: n=16 mice, Prrt2-mutant: n=8 mice). (I) Percentage of after-discharge occurrence in WT and Prrt2-mutant mice after electrical stimulation (WT: n=16, Prrt2-mutant: n=8). Data are presented as mean ± s.e.m. Two-tailed, unpaired Student’s t-test was used in (B, D) and two-tailed, unpaired Mann–Whitney test was used in (H). In (I), the main effect of group was assessed using two-way ANOVAs. ****p<0.0001, n.s., not significant. Source data are provided in Figure 7—source data 1.

Figure 7—source data 1

Raw data used for quantifications in panels B, D, H, and I.

https://cdn.elifesciences.org/articles/109327/elife-109327-fig7-data1-v1.xlsx

PRRT2 deficiency did not affect Nav1.2 protein expression in forebrain tissues (Figure 7—figure supplement 1A and B), nor did it alter the sodium current density in cortical axonal blebs when measured with a brief depolarization to 0 mV preceded by a 200 ms hyperpolarizing prepulse to –110 mV, a protocol designed to relieve inactivation before current elicitation (Figure 7B). Because prolonged voltage-clamp recordings from isolated axonal blebs are technically challenging, we adopted a simplified protocol to assess Nav channel slow inactivation, using a holding potential of –70 mV to approximate the resting membrane potential of cortical neurons. In this protocol, a single 5 s depolarization to 0 mV was applied, and the fraction of available Nav channels was determined after a 10 ms hyperpolarization (Figure 7C).

Approximately 20% of available sodium current remained in wild-type blebs, compared to ~40% in Prrt2-mutant blebs (Figure 7C and D), indicating that during sustained depolarization, fewer Nav channels in the Prrt2-mutant group entered the slow-inactivated state. Notably, because the proportion of channels entering slow inactivation differed substantially between wild-type and Prrt2-mutant groups, direct comparison of recovery kinetics was not feasible and was therefore not performed. Nevertheless, these findings provide direct evidence that endogenous PRRT2 is required for effective regulation of Nav channel slow inactivation in cortical neurons.

PRRT2 deficiency compromises cortical resilience in awake mice

Excitatory neurons exhibit intrinsic adaptation, whereby their responses progressively decline during sustained or repetitive activation (Fleidervish et al., 1996; Kim et al., 2012). We therefore hypothesized that PRRT2 may contribute to this process by promoting Nav channel slow inactivation.

To test this idea, we applied a repetitive stimulation protocol in the corpus callosum, a white-matter region of the brain enriched in both PRRT2 and Nav channels. Compound action potentials, recorded with a glass electrode positioned 150 μm from the stimulation site, were used as a functional readout related to Nav channel availability during repetitive activity (Figure 7—figure supplement 2A). Compared with wild-type mice, Prrt2-mutant mice exhibited a smaller decline in compound action potential amplitude during high-frequency stimulation (20 Hz), but not during low-frequency stimulation (1 Hz), indicating reduced adaptation (Figure 7—figure supplement 2B and C). This pattern is consistent with impaired slow inactivation during repetitive activity.

Neuronal adaptation is important for limiting excitability during episodes of aberrant network hyperactivity (Fleidervish et al., 1996; Zang et al., 2023). To further determine whether PRRT2-dependent impairment of neuronal adaptation contributes to hyperexcitability in vivo, we next assessed cortical resilience in awake mice, a measure of the resistance of the cerebral cortex to pathological perturbations. We used a stimulation–response paradigm in which stimulating electrodes were implanted into the left cerebral cortex, targeting both the sensory and visual cortices, while electroencephalogram (EEG) recording electrodes were placed in the contralateral hemisphere, which receives projections from the stimulated region (Figure 7E). Pulsed electrical stimulation (1 ms pulse width at 60 Hz for 2 s) was delivered to the cortex of mice once daily with stepwise increases in current intensity, and the minimal current required to evoke after-discharges in the contralateral cortex was determined in each animal (Figure 7F). The threshold current required to induce after-discharges was used as a functional indicator of cortical resilience to aberrant excitatory inputs (Blume et al., 2004).

Prrt2-mutant mice exhibited significantly lower stimulation thresholds for evoking after-discharges compared to wild-type controls (Figure 7G and H). Specifically, stimulation at an average intensity of 85 µA reliably induced after-discharges in Prrt2-mutant mice, whereas a significantly higher intensity (190 μA) was required to elicit similar responses in wild-type mice (Figure 7H and I), indicating that PRRT2 contributes importantly to maintaining cortical resilience in response to excitatory perturbations.

Discussion

In this study, we identified PRRT2 as a physiological auxiliary factor of Nav channel slow inactivation. We demonstrate that PRRT2 exerts a dual action by both facilitating the transition of Nav channels into a slow-inactivated state and delaying their recovery. Importantly, this regulatory action is evolutionarily conserved and requires the integrity of the PRRT2 C-terminal region. Using both Prrt2-V5 knock-in and Prrt2-mutant mouse models, we provide compelling in vivo evidence that PRRT2 forms a molecular complex with Nav channels and modulates their slow inactivation, thereby contributing to neuronal adaptation and cortical resilience to perturbations.

In eukaryotic Nav channels, ion permeability is regulated through at least four distinct and coordinated gating mechanisms: voltage-gated activation and deactivation, fast inactivation, and slow inactivation (Catterall, 2023). Slow inactivation, an evolutionarily conserved intrinsic property of Nav channels, is mediated by conformational rearrangements and has distinct molecular determinants from those governing fast inactivation (Payandeh, 2018; Silva, 2014; Vilin and Ruben, 2001).

The role of PRRT2 in the regulation of Nav channel slow inactivation appears to be unique compared to known regulators such as SCN1B and FHFs. The modulatory effects of the SCN1B on Nav channel inactivation were initially observed in Xenopus oocytes (Wallner et al., 1993), where it was found to influence sodium currents. Subsequent studies proposed that the SCN1B impedes slow inactivation, a hypothesis later confirmed in mammalian cells co-expressing Nav1.4 and SCN1B (Webb et al., 2009). However, its regulatory effects on slow inactivation varied across Nav isoforms (He and Soderlund, 2014; Makita et al., 1996; Vilin et al., 1999; Xu et al., 2007). In this study, we observed little effect of SCN1B on the onset and recovery of slow inactivation in Nav1.2, consistent with previous findings (Xu et al., 2007). In contrast to SCN1B, PRRT2 exerts a more consistent effect across Nav isoforms, enhancing slow inactivation regardless of the specific Nav subtype.

FHF proteins (FHF1-4, also known as FGF11-14) constitute a subfamily of FGFs that lack a secretion signal and function distinctly from traditional FGFs (Olsen et al., 2003). Previous studies demonstrated that FHFs mediate a rapid-onset, long-term inactivation, which differs mechanically from traditional slow inactivation (Dover et al., 2010). Unlike the FHF2A-mediated long-term inactivation, which can be triggered by a brief 2 ms depolarization, PRRT2 requires a longer depolarization to produce a detectable effect on the onset of slow inactivation of Nav channels. Additionally, PRRT2 significantly delays recovery of Nav channels from the slow-inactivated state induced by 5 s depolarization, whereas FHF2A has little effect on this process. These findings suggest that PRRT2 and FHF2A modulate Nav channel inactivation through different mechanisms.

Previous studies in heterologous overexpression systems have shown that PRRT2 can influence Nav channel trafficking and surface expression (Fruscione et al., 2018; Valente et al., 2023), raising the possibility that the observed effects on slow inactivation are secondary to altered channel abundance or localization. However, slow inactivation develops on a timescale of hundreds of milliseconds to seconds, whereas detectable changes in Nav channel trafficking and surface abundance generally occur over much longer intervals (minutes to hours) (Fréal et al., 2023; Higerd-Rusli et al., 2023). These distinct temporal profiles argue against trafficking as the primary basis for the effects of PRRT2 on Nav channel slow inactivation described here, although direct quantification of dynamic changes in Nav channel surface expression will be required to fully exclude such a contribution (Liu et al., 2022; Tyagi et al., 2025).

Our truncation analysis showed that the C-terminal region of PRRT2, which contains the re-entrant loop and transmembrane domain, is sufficient to reproduce the enhancement of slow inactivation. This finding suggests that PRRT2 modulates slow inactivation through a membrane-embedded or membrane-proximal mechanism, rather than through its intracellular proline-rich N-terminus. Current models of Nav channel slow inactivation propose that entry into the slow-inactivated state involves coupled conformational changes in both the voltage-sensing domains and the pore region (Catterall et al., 2024; Silva, 2014), including contributions from the external pore mouth (Balser et al., 1996; Xiong et al., 2006), the selectivity filter (Chatterjee et al., 2018; Todt et al., 1999), the S6 segments (Chancey et al., 2007; Chen et al., 2006; Wang and Wang, 1997), and regions encompassing S4, S4-S5 linker, and S5 (Bendahhou et al., 2002; Cummins and Sigworth, 1996; Hayward et al., 1999; Mitrovic et al., 2000; Wang et al., 2000). Within this framework, the membrane-associated C-terminal region of PRRT2 may enhance slow inactivation by stabilizing one or both of these conformational rearrangements, although the precise structural basis of this effect remains to be determined.

In mammals, Nav channel isoforms (Nav1.1–1.9) are distributed across the nervous system, skeletal muscle, heart, and secretory glands (Cusdin et al., 2008). Although we have shown that PRRT2 exerts a consistent effect on slow inactivation across various Nav channel isoforms in vitro, the regulation of Nav channel slow inactivation is likely to be more complex under physiological conditions. PRRT2 is predominantly expressed in the nervous system (Chen et al., 2011), with enrichment in excitatory neurons (Tan et al., 2018), making it unlikely to directly modulate Nav1.4 and Nav1.5 channels, which are primarily expressed in skeletal muscle and cardiac tissue, respectively (Cusdin et al., 2008). A similar rationale applies to Nav1.1, which is largely confined to inhibitory interneurons (Yu et al., 2006), where PRRT2 expression is minimal or absent. These observations suggest that the functional consequence of PRRT2 depends on the Nav isoform composition and cellular context of each tissue.

The partial mismatch between PRRT2 and Nav channel expression in vivo raises an important question about how Nav channel slow inactivation is regulated in cells that do not express PRRT2. One possibility is the expression of alternative regulators, such as TRARG1 or TMEM233, both of which belong to the DspB family and are expressed in tissues distinct from PRRT2 (Ehrlich et al., 2020; Oort et al., 2007; Santana-Varela et al., 2021; Shibata et al., 2007). These proteins may serve analogous functions in modulating Nav channel slow inactivation. Another possibility is that certain PRRT2-negative cells have a reduced requirement for Nav channel slow inactivation modulation due to their intrinsic rhythmic firing properties. For instance, in the cerebellar cortex, PRRT2 is absent in Purkinje cells, which exhibit repetitive firing (Lu et al., 2021). In such rhythmic firing cells, strong enhancement of Nav channel slow inactivation may be undesirable, as excessive accumulation of inactivated Nav channels could disrupt their regular firing patterns. Thus, the broad isoform activity of PRRT2 should be considered in any future attempt to manipulate PRRT2 function therapeutically. Notably, Nav channel slow inactivation represents only one of several mechanisms that regulate neuronal excitability. In both PRRT2-positive and PRRT2-negative neurons, excitability may also be fine-tuned through alternative mechanisms, including regulation of potassium channels such as Kv7.2/Kv7.3 and additional ion channel- or signaling-dependent pathways (Debanne et al., 2019; Jones et al., 2021). These distinct mechanisms likely operate in a coordinated manner to maintain neuronal excitability, with PRRT2-dependent regulation of Nav channel slow inactivation contributing as one component of this broader homeostatic framework (Marom and Marder, 2023).

Given the critical role of PRRT2 in regulating the slow inactivation of Nav1.2 and Nav1.6 channels, both of which are highly expressed in cortical neurons, PRRT2 deficiency may alter neuronal excitability and neuronal responses to prolonged or repetitive activity, thereby contributing to central nervous system dysfunction. Evidence from cultured neurons, animal models, and human genetics is broadly consistent with this view. Studies using human induced pluripotent stem cells (iPSC)-derived excitatory neurons have shown that homozygous PRRT2 mutation increases sodium currents and neuronal excitability (Fruscione et al., 2018). In mouse models, loss-of-function mutation or knockout of PRRT2 in cerebellar granule cells increases intrinsic excitability and facilitates spreading depolarization in the cerebellar cortex (Binda et al., 2021; Lu et al., 2021), whereas in the forebrain, PRRT2 deficiency impairs Nav channel slow inactivation in cortical neurons and reduces resistance to pathological perturbation in awake mice. In addition, PRRT2 mutations in humans are associated with infantile convulsions (Lee et al., 2012), paroxysmal dyskinesia (Chen et al., 2011; Wang et al., 2011), and hemiplegic migraine (Riant et al., 2022), all disorders linked to abnormal neuronal network excitability.

Although the present study supports a functional link between PRRT2-dependent regulation of Nav channel slow inactivation and cortical resilience, PRRT2 may influence cortical excitability and stability through additional Nav-dependent mechanisms, including reduced current density and shifts in the voltage dependence of channel inactivation (Fruscione et al., 2018; Lu et al., 2021; Valente et al., 2023). Notably, because PRRT2 facilitates Nav channel entry into slow-inactivated states both from closed states and from open states during prolonged depolarization, some of these previously reported effects may partly reflect enhanced slow inactivation and the resulting reduction in Nav channel availability. In addition to these Nav-dependent mechanisms, previous studies have shown that PRRT2 also regulates synaptic vesicle cycling (Coleman et al., 2018; Tan et al., 2018; Valente et al., 2016) and presynaptic surface expression of Cav2.1 channels (Ferrante et al., 2021). These effects are also expected to influence neurotransmitter release and, consequently, neuronal and network excitability, and may therefore contribute to cortical resilience.

Together, the identification of PRRT2 as an auxiliary factor regulating Nav channel slow inactivation provides new insight into the fine-tuning of Nav channel function and offers a mechanistic framework for understanding how PRRT2 deficiency may contribute to episodic neurological disorders associated with excitability dysregulation.

Limitations of the study

While our study identifies the functional domain of PRRT2 and provides biological evidence that PRRT2 associates with Nav1.2, the precise binding site on the Nav channel that mediates PRRT2-dependent regulation of slow inactivation remains undefined. Direct structural and biochemical mapping of the PRRT2-Nav interface, including targeted mutagenesis, crosslinking, and structural determination, will be necessary to elucidate the molecular basis of this interaction and its effect on channel gating. Cryo-electron microscopy (cryo-EM) represents a promising approach for resolving the structural details of the Nav-PRRT2 complex, particularly in light of recent advancements in determining high-resolution structures of various Nav channel isoforms and their complexes with auxiliary factors (Jiang et al., 2020; Pan et al., 2019; Pan et al., 2021).

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Gene (Homo sapiens)Human PRRT2GenBank112476
Gene (Mus musculus)Mouse Prrt2GenBank69017
Gene (Danio rerio)Zebrafish Prrt2GenBank100329612
Gene (M. musculus)Tmem233OriGene TechnologiesCat#:MR214829Also known as DSPB2
Gene (M. musculus)Trarg1OriGene TechnologiesCat#:MR218173Also known as DSPB1, Tusc5
Gene (M. musculus)Scn1bOriGene TechnologiesCat#:MR202467
Gene (M. musculus)Fgf13OriGene TechnologiesCat#:MR 221854Also known as Fhf2
Genetic reagent (M. musculus, male and female)Prrt2-mutantTan et al., 2018; PMID:29056747Zhi-Qi Xiong’s Laboratory
Genetic reagent (M. musculus, male and female)Prrt2-V5 knock-inThis paperSee ‘Materials and methods,’ ‘Animals’
Cell line (H. sapiens, female)HEK293TCell Bank/Stem Cell Bank, CASSCSP-502
Cell line (H. sapiens, female)Nav1.2-stably expressing HEK293TLab of ICE Bioscience Inc
Cell line (H. sapiens, female)Nav1.6-stably expressing HEK293TLab of ICE Bioscience Inc
Cell line (Cricetulus griseus, female)Nav1.1-stably expressing CHOLab of ICE Bioscience Inc
Cell line (C. griseus, female)Nav1.4-stably expressing CHOLab of ICE Bioscience Inc
Cell line (C. griseus, female)Nav1.5-stably expressing CHOLab of ICE Bioscience Inc
AntibodyAnti-human PRRT2 (rabbit polyclonal)Atlas AntibodiesCat#:HPA014447; RRID:AB_1855786WB (1:10,000)
AntibodyAnti-mouse PRRT2 (rabbit polyclonal)Wiiget BiotechCat#: Rp3246WB (1:2000)
AntibodyAnti-Nav1.2 (rabbit polyclonal)Alomone LabsCat#:ASC-002; RRID:AB_2040005WB (1:2000)
AntibodyAnti-Nav1.1 (rabbit polyclonal)Alomone LabsCat#:ASC-001; RRID:AB_2040003WB (1:2000)
AntibodyAnti-HA-Tag (rabbit monoclonal)Cell Signaling TechnologyCat#:3724S; RRID:AB_1549585WB (1:1000)
AntibodyAnti-ATP1B2 (rabbit monoclonal)AbcamCat#:ab185210WB (1:10,000)
AntibodyAnti-GAPDH-HRP (mouse monoclonal)KangChen Bio-techCat#:KC-5G5; RRID:AB_2631280WB (1:5000)
AntibodyAnti-HA nanobody magnetic beads (Alpaca nanobody)AlfaLifeBioCat#:KTSM1335Co-IP (30 uL/test)
AntibodyAnti-Flag magnetic beads (mouse monoclonal)BimakeCat#:B26102Co-IP (30 uL/test)
AntibodyV5-Trap magnetic beads (Alpaca nanobody)ChromotekCat#:v5tma; RRID:AB_2868498Co-IP (30 uL/test)
Recombinant DNA reagentpCAGIG (plasmid)AddgeneCat#:11159; RRID:Addgene_11159
Recombinant DNA reagentpCAG-Flag-Nav1.2Pan et al., 2019; PMID:30765605Huai-Zong Shen’s Laboratory
Recombinant DNA reagentpCAG-Flag-Nav1.1Pan et al., 2021; PMID:33712547Huai-Zong Shen’s Laboratory
Sequence-based reagentsgRNAThis paper5’-TCTCCCACAGTGTATAAGTG-3’
Commercial assay or kitSeamless Cloning kitBeyotime,Cat#:D7010
Chemical compound, drugn-Dodecyl β-D-maltosideAnatraceCat#:D310
Software, algorithmQuantity One 1-D analysisBio-RadRRID:SCR_014280
Software, algorithmGraphpad Prism (v8.4.3)GraphPad SoftwareRRID:SCR_002798
Software, algorithmClampfit (v10.7.0.3)Molecular DevicesRRID:SCR_011323
Software, algorithmFitmaster (v2x92)HEKARRID:SCR_016233

Molecular cloning

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Mammalian expression constructs were generated using the pCAGIG backbone (Addgene, 11159), which contains an internal ribosome entry site (IRES) followed by EGFP (Matsuda and Cepko, 2004). Coding sequences for full-length or truncated proteins of interest (POIs) were inserted in the pCAGIG vector using a seamless cloning method (Beyotime, D7010). The POIs included human PRRT2 (hPRRT2), hPRRT2(1–268), hPRRT2(250–340), mouse PRRT2 (mPRRT2), mPRRT2(222–346), mPRRT2(256–346), mPRRT2(1–266), zebrafish PRRT2 (zfPRRT2), mouse SCN1B, mouse FHF2A, mouse TRARG1, and mouse TMEM233.

A PRRT2 chimeric construct was generated by fusing the N-terminal fragment of human PRRT2 (hPRRT2(1–222)) with the C-terminal region of zebrafish PRRT2 (zPRRT2(110–226)), and cloning the resulting sequence into pCAGIG vector. Plasmids encoding human PRRT2 (Chen et al., 2011) were kindly provided by Dr. Zhi-Ying Wu’s Laboratory (Zhejiang University). Plasmids of pCAG-Flag-Nav1.2 (Pan et al., 2019) and pCAG-Flag-Nav1.1 (Pan et al., 2021) were provided by Huai-Zong Shen’s Laboratory (Westlake University). All constructs generated in this study were verified by Sanger sequencing (BioSune, Boshang Biology Technology, Shanghai). All plasmids generated in this study are available from the lead contact with a completed materials transfer agreement.

Cell culture and heterologous expression

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HEK293T cell lines stably expressing human Nav1.2 or Nav1.6 and CHO cell lines stably expressing human Nav1.1, Nav1.4, or Nav1.5 were obtained from the laboratory of ICE Bioscience Inc Wild-type HEK293T cells (SCSP-502) used for transient transfection were obtained from Cell Bank/Stem Cell Bank of Chinese Academy of Sciences. All cells were cultured in Dulbecco’s modified Eagle medium (DMEM; Gibco, 11995065) supplemented with 10% fetal bovine serum (FBS; Gibco, 10099141) and penicillin (50 U/mL)-streptomycin (50 μg/mL) (Gibco, 15140163), and maintained at 37°C in a humidified incubator with 5% CO2. The HEK293T cell line used in this study is of female origin. Short tandem repeat (STR) analyses were performed to authenticate the identity of each cell line. These cell lines were tested negative for mycoplasma contamination.

For heterologous protein expression in whole-cell voltage-clamp recording, 2 μg of plasmid DNA was transfected into HEK293T or CHO cells using Lipofectamine 2000 (Thermo Fisher, 11668-019) according to the manufacturer’s instructions. For co-immunoprecipitation assays, 2–3 μg of total plasmid DNA (encoding two proteins of interest) was used for co-transfection.

4 h after the transfection, the medium was replaced with fresh growth medium, and cells were maintained for an additional 20 h before subsequent experiments.

Animals

Prrt2-mutant mice, carrying a premature stop codon in exon 2 of the Prrt2 gene (Tan et al., 2018), were used in electrophysiological recording experiments.

Prrt2-V5 knock-in mice, used for co-immunoprecipitation (Co-IP) experiments, were generated in this study via CRISPR/Cas9 technology (Yang et al., 2014). Briefly, single-guide RNA (sgRNA: 5’-TCTCCCACAGTGTATAAGTG-3’) targeting the location nearby the stop codon of Prrt2 gene allele, together with a single-stranded DNA donor template encoding a V5 tag flanked by 53 bp homology arms (ssDNA: 5’-CTTCCCTGTC TGTCTTTCCC TCTCCTCTCC CACAGTGTAT AAGGGCAAGC CCATCCCCAA CCCCCTGCTG GGCCTGGACA GCACCTGAGG GGCTCTGCCC TGCATTCCAA GACTTTTCTT CCTGTTGG-3’) and Cas9 mRNA, were microinjected into zygotes from fertilized C57BL/6J mice. The injected zygotes were implanted into pseudopregnant female mice. Offspring were screened for correct insertion of the V5 tag immediately upstream of the Prrt2 stop codon by PCR, and the insertion was subsequently verified by Sanger sequencing. Positive founder (F0) mice were crossed with C57BL/6J mice to establish the Prrt2-V5 knock-in line.

Both Prrt2-mutant and Prrt2-V5 lines were backcrossed onto the C57BL/6J background for more than 10 generations. C57BL/6J mice used for breeding were obtained from the Shanghai Laboratory Animal Center (SLAC), Chinese Academy of Science.

Mice were housed in a 12 h light-dark cycle (light on at 7:00 a.m.) with ad libitum access to food and water. All procedures involving animals were approved by the Animal Care and Use Committee of the Center for Excellence in Brain Science and Intelligence Technology, Chinese Academy of Sciences (approval number, NA-009-2022). Both male and female mice were used in experiments.

All mice generated in this study are available from the lead contact with a completed materials transfer agreement.

Whole-cell voltage clamp recording for sodium currents in cell lines

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Sodium currents were recorded from HEK293T or CHO cell lines stably expressing individual human Nav channel isoforms (ICE Bioscience Inc). Cells were transfected with plasmids as described in the main text. 24 h after transfection, cells were trypsinized and re-plated at lower density onto poly-D-lysine-coated coverslips. EGFP-positive cells were selected for recording 24 h later.

Whole-cell voltage-clamp recordings were conducted at room temperature (23–24°C). The bath solution contained (in mM): 50 NaCl, 90 choline chloride, 3.5 KCl, 1 MgCl2, 2 CaCl2, 10 D-Glucose, 1.25 NaH2PO4, and 10 HEPES (pH adjusted to 7.4 with NaOH). The internal pipette solution contained (in mM): 50 CsCl, 60 CsF, 10 NaCl, 20 EGTA, 10 HEPES (pH adjusted to 7.2 with CsOH). The osmolarity of the external and internal solutions was adjusted to 300 and 290 mOsmol/kg, respectively.

Pipette electrodes were pulled from borosilicate glass capillary (Sutter Instruments, BF150-86-10) using a horizontal puller (P-97, Sutter Instruments). The pipette resistance was approximately 2 MΩ when filled with internal pipette solution. After whole-cell configuration was achieved, cells were allowed to equilibrate for 5 min prior to recording. Series resistance was compensated by 80% to minimize voltage errors. Currents were recorded with an amplifier (HEKA, EPC-10), sampled at 10 kHz, and acquired using Patchmaster (HEKA, v2x92).

Fast inactivation assay for Nav channels

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To assess the fast-inactivation kinetics of Nav channels, sodium currents were evoked by 20 ms depolarizing steps to test potentials between –20 and +30 mV in 10 mV increments, from a holding potential of –120 mV. The decay time constant (tau) was obtained by mono-exponential fitting of the current trace.

Recovery from fast inactivation was measured using a two-pulse protocol, in which a 10 ms depolarizing pulse to 0 mV (P1) was followed by a variable-duration recovery period at –120 mV and a second 10 ms test pulse at 0 mV (P2). Recovery was quantified as the normalized peak current of P2 relative to P1.

Slow inactivation assay for Nav channels

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To evaluate the development of slow inactivation of Nav channels, a series of conditioning prepulses of increasing duration at 0 mV was applied, followed by a brief 10 ms recovery at –120 mV and a 10 ms test pulse at 0 mV. Inter-sweep intervals were set at 15 s. Available fractions of Nav channels were assessed by normalizing the peak currents in the test pulse to that of the conditioning pulse.

For slow inactivation recovery, a 5 s conditioning pulse at 0 mV was followed by variable recovery durations at –120 mV, then a 10 ms test pulse at 0 mV. The inter-sweep interval was 15 s.

In experiments assessing the effect of zebrafish PRRT2 on slow inactivation recovery, a modified protocol was used to avoid cumulative inactivation. Within a single sweep, a 5 s depolarization was followed by a series of 10 ms test pulses to 0 mV, each separated by progressively longer recovery periods at –120 mV (Webb et al., 2009).

In this study, we chose 0 mV as the primary conditioning potential because it is widely used in conventional slow inactivation protocols and induces slow inactivation more robustly than more negative conditioning voltages, such as –70 mV.

Steady-state slow inactivation assay for Nav channels

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To measure steady-state slow inactivation, 10 s conditioning pulses ranging from –110 to –10 mV (in 10 mV increments) were applied, followed by a 10 ms hyperpolarizing step to –120 mV and subsequently a 5 ms test pulse to 0 mV. The peak test current was normalized to the maximum sodium current recorded across all conditions in the protocol and reported as I/Imax.

Slice preparation

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Mice (postnatal days 14–20) were anesthetized with 4% isoflurane. The brain tissues were rapidly extracted and immersed in ice-cold sucrose-based cutting solution containing (in mM) 2.5 KCl, 1.25 NaH2PO4, 26 NaHCO3, 10 MgSO4, 0.5 CaCl2, 10 D-glucose, 205 sucrose, 1 sodium pyruvate, and 1.3 sodium L-ascorbate. Coronal cortical slices (250 μm) were prepared using a vibratome (Leica, VT1200S).

Slices were then transferred immediately to an incubation beaker containing aerated artificial cerebrospinal fluid (ACSF) composed of (in mM) 126 NaCl, 2.5 KCl, 2 MgSO4, 2 CaCl2, 26 NaHCO3, 1.25 NaH2PO4, and 25 dextrose (pH 7.4, 315 mOsm). Slices were incubated at 35°C for 30 min, then maintained in the same solution at room temperature until use. The oxygenated ACSF was equilibrated with 95% O2 and 5% CO2.

Electrical stimulation and field recording in the corpus callosum

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Coronal slices were transferred to a recording chamber. A stimulation electrode (125 µm in diameter, concentric bipolar electrode, FHC, USA) was placed on the corpus callosum. The Ag-AgCl recording electrode (2–3 MΩ) was located at a site 150 µm away from the stimulation electrode along the corpus callosum. Stimulating pulses (0.1 ms pulse width, 100 µA) were delivered at 1 Hz for 100 s or at 20 Hz for 5 s. Compound action potentials were recorded as a functional readout related to Nav channel availability.

Electrophysiological recording in isolated axonal blebs

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Slices were transferred to a recording chamber and stabilized with a platinum anchor grid during recording. Resealed axonal ends (‘blebs’) in the cerebral cortex were visualized using an upright infrared differential interference contrast (IR-DIC) microscope. Individual blebs were mechanically isolated by sweeping a cutting pipette beneath it, and subsequently patched with glass electrodes (resistance ~7 MΩ).

For sodium current recordings, pipettes were filled with a potassium gluconate-based internal solution containing (in mM) 130 K-gluconate, 1 MgCl2, 11 EGTA, 1 CaCl2, 1 KCl, 2 Mg-ATP, 0.3 Na-GTP, 10 HEPES (pH adjusted to 7.2, ~290 mOsm), supplemented with 2 mM TEA-Cl. The external ACSF solution was supplemented with 5 mM 4-Aminopyridine (4-AP) and 200 μM CdCl2 to block K+ and Ca2+ currents, respectively. 1 μM tetrodotoxin (TTX) was used to block TTX-sensitive Nav channels.

Whole-cell voltage-clamp recordings were performed using a MultiClamp amplifier (Molecular Devices, 700B). Currents were filtered at 4 kHz (low-pass) and digitized at 20 kHz via a Digidata interface (Molecular Devices, 1440A). Series resistance was compensated by 80% to minimize voltage errors. Leak currents were subtracted by using an online P/4 procedure.

The holding potential was set to –70 mV, approximating the resting membrane potential of cortical neurons. To assess sodium current density, a hyperpolarization prepulse to –110 mV (200 ms) was used to remove inactivation, followed by a 20 ms depolarization to 0 mV. Sodium current density (nA/pF) was calculated by normalizing peak current to bleb capacitance.

To evaluate Nav channel slow inactivation, a single 5 s depolarizing pulse (0 mV) was applied to induce slow inactivation followed by test pulses (0 mV, 20 ms) separated by 10 ms recovery intervals at –70 mV. The fraction of available Nav channels was calculated by normalizing the peak current amplitude of each test pulse to the peak current measured at the onset of the 5 s depolarization.

Co-immunoprecipitation in transfected HEK293T cells

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In co-immunoprecipitation experiments, one protein-of-interest was tagged with an HA epitope and the other with a Flag epitope. The pCAGIG vector bearing either a HA or Flag tag alone was used as a control in paired constructs. HEK293T cells transfected with a pair of constructs were harvested 24 h post-transfection for protein expression. Cells were lysed in ice-cold lysis buffer (20 mM Tris-HCl, 150 mM NaCl, 0.5% n-dodecyl β-D-maltoside [DDM, Anatrace, D310], and protease inhibitor cocktail [Selleckchem, B14002]) with gentle agitation at 4°C for 2 h. Lysates were centrifuged at 20,000 × g for 20 min at 4°C to remove insoluble debris. The supernatant was transferred to a fresh tube and kept on ice.

Protein concentration of supernatant was determined using a colorimetric bicinchoninic acid (BCA) assay (Beyotime, P0012S). For each immunoprecipitation (IP), 400 μg of proteins was incubated with 30 μL of anti-HA nanobody magnetic beads (AlfaLifeBio, KTSM1335) or anti-Flag magnetic beads (Bimake, B26102) at 4°C with rotation for 1 h. Beads were then separated with DynaMag spin and washed three times with wash buffer (20 mM Tris-HCl, 150 mM NaCl, 0.05% DDM). The captured proteins were eluted with 25 μL SDS-PAGE loading buffer containing 100 mM DTT, followed by heating at 45°C for 10 min. Samples were stored at –20°C prior to western blot analysis.

Co-immunoprecipitation in mouse brain tissues

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Brain tissues (~100 mg per sample) were freshly dissected from wild-type or Prrt2-V5 knock-in mice, and homogenized on ice using glass homogenizers in 1 mL of Tris-buffered saline (TBS, 20 mM Tris-HCl, 150 mM NaCl, protease inhibitor cocktail). The homogenates were centrifuged at 800 × g for 10 min at 4°C to remove nuclei and cell debris. Supernatants were subjected to centrifugation at 40,000 × g for 45 min at 4°C to isolate membrane fractions. The resulting membrane-enriched pellets were resuspended in TBS and solubilized in lysis buffer containing 0.3% DDM for 3 h at 4°C. Lysates were then cleared by centrifugation at 20,000 × g for 20 min at 4°C. Protein concentration was measured via BCA assay. For each IP, ~200 μg of proteins was incubated with 30 μL of V5-Trap magnetic beads (Chromotek, v5tma-20) at 4°C with rotation for 1.5 h. Beads were washed three times with wash buffer (20 mM Tris-HCl, 150 mM NaCl, 0.15% DDM), and bound proteins were eluted with 25 μL SDS-PAGE loading buffer containing 100 mM DTT, followed by heating at 45°C for 10 min. Eluates were kept at –20°C until use in western blotting.

Western blotting

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Protein samples were separated using 4–12% PAGE Gel (Nanjing ACE Biotechnology, ET15412L) in MES-SDS running buffer at 140 V for 60 min. Proteins were transferred onto polyvinylidene difluoride (PVDF) membranes at 300 mA for 2 h at 4°C. The PVDF membranes were blocked in 5% nonfat milk in TBST (TBS with 0.1% Tween-20) for 1 h at room temperature, followed by incubation with indicated primary antibodies overnight at 4°C with gentle agitation.

After three washes in TBST, membranes were incubated with HRP-conjugated secondary antibodies for 2 h at room temperature. Following another three washes, chemiluminescent substrate (TIANGEN Biotech, PA112) was applied, and signals were detected using a blot imaging system (Analytikjena, ChemStudio 815). Densitometric analysis of protein bands was performed using QuantityOne software (Bio-Rad, Quantity One 1-D analysis, v4.6.2). The primary antibodies we used in western blotting were as follows: anti-human PRRT2 (Atlas Antibodies, HPA014447), anti-Nav1.2 (Alomone Labs, ASC-002), anti-Nav1.1 (Alomone Labs, ASC-001), anti-HA-Tag (Cell Signaling Technology, 3724S), anti-GAPDH-HRP (KangChen Bio-tech, KC-5G5), anti-ATP1B2 (Abcam, ab185210), anti-mouse PRRT2 (Wiiget Biotech, Rp3246), and anti-β Actin-HRP (Cell Signaling Technology, 5125S).

Animal surgery

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Mice were anesthetized with isoflurane (RWD Life Science, R510-22-10), using 4% for induction and 2% for maintenance. Body temperature was maintained throughout the procedure with a heating pad. Meloxicam (5 mg/kg, subcutaneous) was administered preoperatively for analgesia.

After securing the mice in a stereotaxic frame (RWD Life Science, Model 68528), 0.25% bupivacaine was locally injected at the incision sites for perioperative analgesia, and ophthalmic ointment was applied to protect the cornea. A midline incision was made to expose the skull, which was cleaned with 2% hydrogen peroxide using sterile cotton swabs to remove connective tissue. The skull surface was leveled to a stereotaxic plane, and the bregma was identified and marked. Once dry, the exposed skull was coated with a light-curing self-etch adhesive (3M ESPE Single Bond Universal).

For cerebellar electrical stimulation experiments, two small holes separated by 1.5 mm were drilled in the skull above the fourth/fifth cerebellar lobules (anterior–posterior [AP], –6.25 mm; medial–lateral [ML], ±0.75 mm; dorsal–ventral [DV], –1.0 mm from the dura). Two metal electrodes (100 μm in diameter; California Fine Wire Company, M426550) were implanted and secured with dental cement.

For cortical electrical stimulation and EEG recording, two small craniotomies (~0.6 mm in diameter) were made over the left sensory cortex (AP –0.8 mm; ML 1.5 mm) and visual cortex (AP –3.5 mm, ML 1.5 mm) for the insertion of bipolar stimulation electrodes (180 μm in diameter), which were implanted to a depth of –0.6 mm from the dura. Stainless steel screws were used as EEG electrodes: one was implanted into the right visual cortex (AP –2.25 mm, ML –1.5 mm), and the second into the right cerebellar vermis to serve as the reference. An additional screw was placed in the left cerebellar hemisphere as the ground. The electrodes and a custom-made head plate were affixed to the skull using dental cement.

Postoperatively, meloxicam (5 mg/kg, subcutaneously) was administered once daily for three consecutive days. Mice were allowed to recover for 1 week prior to electrical stimulation.

Cerebellar electrical stimulation and behavior assessment

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Before cerebellar electrical stimulation, mice were habituated to the head-fixed platform for 30 min per day over three consecutive days. Electrical stimulation (300 μA, 60 Hz, 1 ms pulse width, 2 s duration) was delivered to the cerebellar cortex through a stimulus isolator (AMPI, ISO Flex) driven by a Master-8 stimulator. Mouse behavior was recorded with a video camera at a frame rate of 30 frames per second for 10 min.

Dystonia in mice were defined by the presence of one or more of the following abnormal postures or movements: prolonged forelimb elevation; persistent backward extension of the ears, tightly apposed to the head; an elongated trunk posture with the abdomen close to the ground and prolonged elevation or excessive extension of hindlimbs.

Electrostimulation and EEG recording in the cerebral cortex

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1 week after surgery, mice were habituated to a custom-built head plate holder for 30 min per day over three consecutive days prior to testing.

For stimulation experiments, the pre-implanted electrodes were connected to a stimulus isolator (AMPI, ISO Flex). Electrical pulses (1 ms pulse width, 60 Hz frequency, 2 s duration) were delivered through the isolator and controlled via TTL signals from a pulse generator (RWD Life Science, R820). EEG signals (bandpass: 1–300 Hz) were amplified 1000-fold using an AC/DC differential amplifier (A-M Systems, Model 3000), digitized at 1 kHz with a Digidata (Molecular Devices, 1322A), and acquired using Clampex software (Molecular Devices, v10.7).

To assess cortical resilience in Prrt2-mutant versus wild-type mice, a stimulus-response protocol was employed using progressively increasing current intensities (0, 20, 40, 80, 120, 160, 200, 240, 280, 320 µA). Each stimulation was separated by a 24 h interval. The stimulation threshold was defined as the lowest current intensity that induced after-discharges (lasting ≥2 s) in the EEG trace. Each EEG recording lasted for 5 min, with electrical stimulation delivered immediately following a 1 min baseline period.

Statistics

All grouped data are presented as mean ± standard error of the mean (s.e.m.) unless otherwise stated. Statistical analyses were performed using GraphPad Prism 8 (GraphPad Software Inc, La Jolla, USA). No statistical methods were used to predetermine the sample sizes. The D'Agostino–Pearson omnibus (K2) test was used to assess normality.

Two-tailed, unpaired Student’s t-tests were applied in Figure 7B and D. A two-tailed, paired Student’s t-test was used in Figure 6D–F, Figure 7—figure supplement 1B. Two-tailed, unpaired Mann–Whitney test was applied in Figure 7H. Two-way ANOVAs were conducted for comparisons in Figures 1C ,D, F ,G—4B–E, G–J, 7I, Figure 1—figure supplement 1B and C, Figure 3—figure supplement 1C and D, and Figure 7—figure supplement 2B and C. If necessary, the Geisser–Greenhouse correction was applied to adjust for violations of sphericity.

Statistical tests and sample sizes (including number of cells, slices, or animals) are stated in the figure legends. In all experiments, n denotes the number of independent biological replicates. No data were excluded from analyses. Asterisks in figures denote significance levels: *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001, n.s., not significant.

Co-immunoprecipitation assays were independently repeated at least three times in both in vitro and in vivo tests. All attempts at replicating experiments presented in the manuscript have obtained consistent conclusions.

Electrophysiological recordings were conducted with the experimenter blinded to group identity (transfected condition or animal genotype).

Data availability

All data generated or analyzed during this study are included in the manuscript and supporting files; source data files have been provided for all figures.

References

Article and author information

Author details

  1. Bin Lu

    1. Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    2. University of Chinese Academy of Sciences, Beijing, China
    Contribution
    Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Investigation, Visualization, Methodology, Writing – original draft, Project administration
    Contributed equally with
    Qi-Wu Xu, Jing Zhang, Xue-Mei Wu and Jun-Yan He
    For correspondence
    lubin@ion.ac.cn
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1456-2517

  2. Qi-Wu Xu

    Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    Contribution
    Formal analysis, Investigation, Writing – review and editing
    Contributed equally with
    Bin Lu, Jing Zhang, Xue-Mei Wu and Jun-Yan He
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2236-8415

  3. Jing Zhang

    Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    Contribution
    Resources, Formal analysis, Investigation, Writing – review and editing
    Contributed equally with
    Bin Lu, Qi-Wu Xu, Xue-Mei Wu and Jun-Yan He
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0009-4050-8486

  4. Xue-Mei Wu

    1. Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    2. School of Life Science and Technology, ShanghaiTech University, Shanghai, China
    Contribution
    Formal analysis, Investigation, Writing – review and editing
    Contributed equally with
    Bin Lu, Qi-Wu Xu, Jing Zhang and Jun-Yan He
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0000-3387-7579

  5. Jun-Yan He

    1. Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    2. University of Chinese Academy of Sciences, Beijing, China
    Contribution
    Formal analysis, Investigation, Writing – review and editing
    Contributed equally with
    Bin Lu, Qi-Wu Xu, Jing Zhang and Xue-Mei Wu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0002-3342-9877

  6. Jing-Qiu Peng

    1. Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    2. School of Life Science and Technology, ShanghaiTech University, Shanghai, China
    Contribution
    Formal analysis, Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0008-1124-7726

  7. Guang Yang

    1. Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    2. University of Chinese Academy of Sciences, Beijing, China
    Present address
    Interdisciplinary Research Center on Biology and Chemistry, Shanghai Institute of Organic Chemistry, Chinese Academy of Sciences, Shanghai, China
    Contribution
    Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0009-1840-8411

  8. Ke-Xian Li

    Department of Neurology, Zhongshan Hospital, Fudan University, Shanghai, China
    Contribution
    Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0004-5401-4923

  9. Ling Zhuang

    1. Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    2. University of Chinese Academy of Sciences, Beijing, China
    Present address
    Hainan Academy of Medical Sciences, Hainan Medical University, Hainan, China
    Contribution
    Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0007-6846-883X

  10. Yu-Xian Zhang

    Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    Contribution
    Formal analysis, Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0005-1796-1443

  11. Zhi-Ying Wu

    Department of Medical Genetics and Center for Rare Diseases, Second Affiliated Hospital, Zhejiang University School of Medicine and Zhejiang Key Laboratory of Rare Diseases for Precision Medicine and Clinical Translation, Hangzhou, China
    Contribution
    Resources, Writing – review and editing, Conceptualization
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2106-572X

  12. Zhi-Qi Xiong

    1. Center for Excellence in Brain Science and Intelligence Technology (Institute of Neuroscience), Chinese Academy of Sciences, Shanghai, China
    2. University of Chinese Academy of Sciences, Beijing, China
    3. School of Life Science and Technology, ShanghaiTech University, Shanghai, China
    4. Shanghai Center for Brain Science and Brain-Inspired Technology, Shanghai, China
    Contribution
    Conceptualization, Data curation, Supervision, Funding acquisition, Methodology, Project administration
    For correspondence
    xiongzhiqi@ion.ac.cn
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7727-8412

Funding

Innovation of Science and Technology 2030-Major Project "platform of nonhuman primate models" (2021ZD0200900)

  • Zhi-Qi Xiong

National Natural Science Foundation of China (82271269)

  • Bin Lu

National Natural Science Foundation of China (82021001)

  • Zhi-Qi Xiong

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

The authors thank Fei Dong, Ting-Bin Ma, Xi Zhou, Wen Zhang, Wei Ke, Yue-Jie Xiao, Xue-Qin Jin, and You-Sheng Shu for technical support for electrophysiological recording. We thank Huai-Zong Shen, Zhang-Qiang Li, Xue-Jing Huang, and Shu-Jia Zhu for experimental support for protein expression. We thank Tong-Zhou Li, Hong Zhang, Jun-Jun Liu, and Hai-Yan Zhen from ICE Bioscience for experimental support in sodium current recording. We appreciate the staff at the Core Facilities for their generous technical support, and members of Xiong Lab for technical support and insightful discussions.

Ethics

All procedures involving animals were approved by the Animal Care and Use Committee of the Center for Excellence in Brain Science and Intelligence Technology, Chinese Academy of Sciences (Approval number, NA-009-2022).

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© 2025, Lu, Xu, Zhang et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Bin Lu
  2. Qi-Wu Xu
  3. Jing Zhang
  4. Xue-Mei Wu
  5. Jun-Yan He
  6. Jing-Qiu Peng
  7. Guang Yang
  8. Ke-Xian Li
  9. Ling Zhuang
  10. Yu-Xian Zhang
  11. Zhi-Ying Wu
  12. Zhi-Qi Xiong
(2026)
PRRT2 as an auxiliary regulator of Nav channel slow inactivation
eLife 14:RP109327.
https://doi.org/10.7554/eLife.109327.4

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