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PIP2-dependent coupling of voltage sensor and pore domains in Kv7.2 channel

Abstract

Phosphatidylinositol-4,5-bisphosphate (PIP2) is a signaling lipid which regulates voltage-gated Kv7/KCNQ potassium channels. Altered PIP2 sensitivity of neuronal Kv7.2 channel is involved in KCNQ2 epileptic encephalopathy. However, the molecular action of PIP2 on Kv7.2 gating remains largely elusive. Here, we use molecular dynamics simulations and electrophysiology to characterize PIP2 binding sites in a human Kv7.2 channel. In the closed state, PIP2 localizes to the periphery of the voltage-sensing domain (VSD). In the open state, PIP2 binds to 4 distinct interfaces formed by the cytoplasmic ends of the VSD, the gate, intracellular helices A and B and their linkers. PIP2 binding induces bilayer-interacting conformation of helices A and B and the correlated motion of the VSD and the pore domain, whereas charge-neutralizing mutations block this coupling and reduce PIP2 sensitivity of Kv7.2 channels by disrupting PIP2 binding. These findings reveal the allosteric role of PIP2 in Kv7.2 channel activation.

Introduction

Phosphoinositides are major constituents of biological membranes and key regulators of fundamental biological processes including signal transduction, membrane trafficking, and cytoskeletal dynamics1. Among the phosphoinositides, phosphatidylinositol-4,5-bisphosphate (PIP2) in the plasma membrane serves as a critical cofactor for many ion channels despite its low abundance (~1% of total acidic lipids)2,3. The affected channels include inward rectifier potassium (K+) channels4, voltage-gated calcium channels, transient receptor potential channels, hyperpolarization-activated cyclic nucleotide-gated channels, and voltage-gated potassium (Kv) channels2,5.

PIP2 activates all five members of the Kv channel subfamily Q (Kv7.1-Kv7.5) which control excitability of neuronal, sensory, and muscle cells6,7. Encoded by KCNQ1-KCNQ5 genes7, each Kv7 subunit has six transmembrane segments8,9. The first four segments (S1-S4) comprise a voltage-sensing domain (VSD) with the S4 being the main voltage-sensor8,9. The pore domain consists of the last two segments (S5-S6) flanking the pore loop which contains a highly conserved sequence and structure for K+ selectivity and permeability8,9. The C-terminal intersection of four S6 segments constitutes the main gate9,10. Each subunit also has a long intracellular C-terminal tail that harbors four α-helices (A-D)11. Helix-A and Helix-B interact with calmodulin (CaM)11, Helix-C mediates inter-subunit interaction, while Helix-D specifies the subunit assembly as a homotetramer or a heterotetramer11.

Despite the common core structure, each Kv7 subunit follows a distinct, cell-specific distribution that dictates its physiological roles in different tissues7,8. In the heart, Kv7.1 assembles with an auxiliary β subunit KCNE1 to produce the slow K+ current critical for repolarizing cardiac action potentials (APs)7,8. Kv7.4 is primarily found in cochlear hair cells of the inner ear8. In the central nervous system, Kv7 channels are mostly heterotetramers of Kv7.2 and Kv7.3, and to a lesser extent heterotetramers of Kv7.3 and Kv7.5 and homomeric Kv7.2 channels12,13. Neuronal Kv7 channels produce slowly activating and non-inactivating K+ current (IM) that suppresses repetitive firing of APs12,13, and dominant mutations in their subunits cause neonatal epilepsies including benign familial neonatal epilepsy (BFNE) and epileptic encephalopathy (EE) (rikee.org)14,15,16,17. EE is a collection of epileptic syndromes accompanied by profound neurodevelopmental delay and psychomotor retardation18,19.

Kv7 channels are inhibited by membrane PIP2 depletion upon activation of Gq-coupled receptors2,6,13,20. The underlying mechanism has been extensively investigated in Kv7.13. Voltage-clamp fluorometry studies have demonstrated that depolarization can activate the VSD of Kv7.1 but fails to open the pore upon PIP2 depletion21, suggesting that PIP2 is crucial for coupling the VSD to the pore domain. In the cryo-EM structure of Kv7.1 channel in complex with KCNE3 and CaM, PIP2 interacts with the S2-S3 and S4-S5 linkers, and this interaction may facilitate the channel opening by converting the unstructured loop between the S6 to Helix-A (pre-Helix-A) to a helix22. In addition to the S2-S3 and S4-S5 linkers and S6 as potential PIP2 binding sites in Kv7.121, in vitro binding studies with helices A-D of Kv7.1 have also identified basic residues in distal Helix-B that interact with PIP223.

Despite the accumulating mechanistic insights into PIP2-dependent modulation of Kv7.13, it remains unclear whether neuronal Kv7 channels are regulated by the same PIP2 binding residues and mechanism as Kv7.1. There are several differences in PIP2-dependent modulation between Kv7.1 and neuronal Kv7 channels. First, PIP2 sensitivity of Kv7.1 is regulated by KCNE124, whereas PIP2 directly modulates neuronal Kv7 channels without auxiliary β subunits13. Second, previous electrophysiology studies with site-directed mutagenesis have suggested potential PIP2 binding sites unique to Kv7.2 and Kv7.3 including the regions between Helix-A and Helix-B (AB linker) and between Helix-B and Helix-C (BC linker)14,25,26,27. Third, the AB linker of Kv7.2 is much longer than that of Kv7.1. Importantly, some epilepsy variants in Kv7.2 and Kv7.3 disrupt the channel sensitivity to the changes in cellular PIP2 level14,25,28,29. Therefore, detailed investigation of how PIP2 regulates neuronal Kv7 channels can increase our understanding of their physiological function in neurons and facilitate the development of new therapeutic strategies against epilepsy.

The first step toward understanding the molecular action of PIP2 on neuronal Kv7 channels is to identify PIP2 binding sites. To achieve this, we employed all-atom molecular dynamics (MD) simulations. This technique has been successfully employed to provide atomic-level structural insights on lipid-protein interactions in membrane proteins with high spatiotemporal resolution30,31,32,33,34, in close agreement with the experimental data30,31,32,33,34,35. We chose to identify PIP2 binding sites in homomeric Kv7.2 channels for several reasons. First, they produce robust K+ currents upon depolarization whereas homomeric Kv7.3 channels are nonfunctional36,37,38,39. Second, conditional deletion of Kv7.2 but not Kv7.3 results in cortical hyperexcitability, spontaneous seizures, and high mortality in mice40. Third, there are significantly more epilepsy mutations found in KCNQ2 than KCNQ3 (ClinVar Database, NCBI)41,42, and current suppression of homomeric Kv7.2 channels is a common feature of EE variants of KCNQ243.

Here our MD simulations reveal multiple PIP2 binding sites in homomeric Kv7.2 channels with more sites in the open state than the closed state. These sites include the S2-S3 linker, S4, S4-S5 linker, S6, pre-Helix-A, AB linker, Helix-B, and BC linker. Charge-neutralizing mutations of four PIP2-binding residues (R214Q in the S4, K219N in the S4-S5 linker, R325Q in pre-Helix-A, and R353Q in the AB linker) disrupt PIP2 binding to the mutated residues and decrease current densities of Kv7.2 channels. Importantly, R214Q, K219N, and R325Q mutations reduce channel sensitivity to PIP2 depletion, with the triple R214Q/K219N/R353Q mutation inducing the largest effect. Our simulations further show that R214Q, K219N and R325Q mutations decouple the VSD activation from the pore domain of Kv7.2 channels, while the R353Q mutation blocks the PIP2-induced increase in the propensity of helices A and B to interact with the inner leaf of the bilayer. These findings offer detailed mechanistic insights into PIP2-dependent modulation of Kv7.2 channels.

Results

Differential PIP2 binding in closed and open Kv7.2 channels

To identify PIP2 binding sites, we performed all-atom MD simulations on human Kv7.2 channels in explicit lipid bilayers composed of phosphatidylcholine and PIP2. By adopting an integrative structural modeling approach using X-ray and cryo-EM data, we first constructed the open and closed states of Kv7.2 channels within explicit lipid bilayers (Fig. 1a). The stability of the resulting models was investigated and reported in our previous publication14. At the beginning of the lipid-binding simulations, 8 PIP2 lipid molecules (2.2% of the total lipid in each leaflet) were distributed around the channel with their starting positions randomized in each of the three independent, 500ns-long simulation replicates (Fig. 1b, Table 1). Differential binding of PIP2 lipids to open and closed channels was captured and presented as the PIP2 headgroup occupancy maps extracted from the entire simulation trajectory set for each state (Fig. 1c–d). Upon binding, PIP2 remained stably bound throughout the rest of the simulation time in all identified binding sites in both states (Fig. 1e–f).

Fig. 1: Simulation design for identifying PIP2 localization in open and closed Kv7.2 channels.
figure 1

a Left: A tetrameric Kv7.2 channel used in our simulations, where each subunit is highlighted with a different color in surface representation. Right: Detailed view of the important structural features. A tetrameric human Kv7.2 channel in a closed state was modeled from the cryo-EM structure of human Kv7.1 channel with a closed pore (PDB ID: 5VMS). For modeling the Kv7.2 channel in an open state, TMD (targeted MD) was used to drive the transition of the closed Kv7.2 state model towards the template based on the open state of Kv1.2/ Kv2.1 channel (PDB: 2R9R). b Top view of the simulation replicates (Runs) showing the initial placement of 8 PIP2 molecules in both upper and lower leaflets of the lipid bilayer surrounding Kv7.2 channel in the absence of calmodulin. For each state/conformation (open or closed), 3 independent simulation replicates were performed in a tetrameric Kv7.2 channel by randomly shuffling the initial positions of PIP2 lipids, as shown by different colors (Runs 1-3). cd Volumetric maps of PIP2 headgroup occupancy extracted collectively from the simulation trajectories are shown as wireframes overlaid on the Kv7.2 channel structure for the closed state (c) or the open state (d). We label the PIP2 binding sites according to their proximity to the pore region. In the closed state, PIP2 headgroups bind to Site-C1 (N-terminal tail), Site-C2 (the S2-S3 linker), and Site-C3 (at the interface formed by Helix-A, the S2-S3 linker and the AB linker). In the open conformation, PIP2 headgroups are observed to bind Site-O1 (the interface formed by the N-terminal tail, the S2-S3 linker, distal Helix-B, and the BC linker), Site-O2 (distal Helix-B), Site-O3 (the interface formed by the S2-S3 linker, distal Helix-B, and the BC linker), and Site-O4 (the interface formed by the distal ends of the S4 and S6, the S4-S5 linker, and pre-Helix-A). ef Distance between the center of mass (COM) of PIP2 headgroup and the COM of the binding site is plotted against simulation time. Initially all PIP2 molecules were placed at least 15 Å away from any of the binding sites captured in Kv7.2 channel structure in the closed state (e) and the open state (f). Once PIP2 lipids bind to any of the binding sites, they remained stably bound throughout the simulation time.

Table 1 Details of the systems and the simulations performed.

We discover that the PIP2 headgroup interacts with 3 distinct sites in the closed state (Fig. 1c): Site-C1 (intracellular N-terminal tail), Site-C2 (the S2-S3 linker), and Site-C3 (the interface formed by Helix-A, the S2-S3 and AB linkers) (Fig. 1c). Notably, repeating the lipid-binding simulations on the recently solved cryo-EM structure of a closed Kv7.2 channel44 verified the formation of the same PIP2 binding sites (Supplementary Fig. S1). In the open state, we have identified four distinct PIP2-binding sites, all enriched with basic residues: Site-O1 (the interface formed by the S2-S3 and AB linkers), Site-O2 (distal Helix-B), Site-O3 (the interface formed by the N-terminal tail, the S2-S3 linker, distal Helix-B, and the BC linker), and Site-O4 (the interface formed by the ends of S4 and S6, the S4-S5 linker, and pre-Helix-A) (Fig. 1d).

Although PIP2 interact with the N-terminal tail and the S2-S3 and AB linkers in both closed and open states, there are key differences in the PIP2 binding sites of these two states. PIP2 headgroup occupancies at the four sites in the open channel are more focused than those at the three sites in the closed channel (Fig. 1c, d). Importantly, PIP2 localizes to the periphery of the VSD in the closed channel, whereas PIP2 binding spreads to a larger area in the open channel including the cytoplasmic ends of the VSD and S6, pre-Helix-A, distal Helix-B, and the BC linker (Fig. 1c, d), suggesting that the opening of Kv7.2 channels involves PIP2 interaction with VSD, the gate, and intracellular helices.

Characterizing specific lipid–protein interactions in Kv7.2 channels

To map PIP2-binding sites, we analyzed lipid–protein interactions during the last 200 ns of MD trajectories by quantifying the contact probability between each moiety of the PIP2 headgroup and the PIP2-binding residues. In the closed channel, Site-C1 is formed by a cluster of basic residues (K76, R87, and R89) in the intracellular N-terminal tail. R87 and R89 exclusively interact with P5-phosphate, whereas K76 establishes contacts with all the hydroxyl and phosphate groups on the inositol ring in PIP2 (Supplementary Fig. S2a, d). In Site-C2, a cluster of basic residues in the S2-S3 linker (R153, R158, and K162) coordinate PIP2. R153 and K162 show high contact probabilities to P4-phosphate and P5-phosphate, whereas R158 interacts with all groups in PIP2 (Supplementary Fig. S2b, e). Site-C3 is formed by R155 in the S2-S3 linker, Y347 in Helix-A, and R353 in the AB linker. R155 has high contact probabilities for P5-phosphate and the hydroxyl group at position 6 of the inositol ring, whereas R353 interacts preferentially with P4- and P5-phosphates (Supplementary Fig. S2c, f).

In the open channel, Site-O1 is formed by residues in the S2-S3 linker (K162 and R165) and the AB linker (F346, Y347, and R353) (Fig. 2a, b). All the three basic residues in this site show high contact probabilities for P5-phosphate on PIP2 headgroup (Fig. 2a). In Site-O2, a cluster of basic residues in distal Helix-B (K552, R553, and K554) coordinate PIP2 (Fig. 2b, f). K552 and R553 primarily bind to P5-phosphate, while K554 interacts with P4- and P5-phosphates and hydroxyl group at position 3 of the inositol ring (Fig. 2b). Site-O3 is formed by residues in the intracellular N-terminal tail (R87), the S2-S3 linker (R153, Y154, and K166), distal Helix-B (R553), and the BC linker (R560) (Fig. 2c, g). Y154 and K166 predominantly interacts with P4-phosphate, whereas the other basic residues bind to P5-phosphate. In Site-O4, PIP2 interacts with R214 at the end of S4, K219 in the S4-S5 linker, K319 and Q323 in distal end of S6, and R325 in pre-Helix-A (Fig. 2d, h). Most of these residues, except K319, show high contact probabilities for P5-phosphate in PIP2.

Fig. 2: PIP2 coordination in 4 binding sites captured in the modeled structure of the open Kv7.2 channel.
figure 2

ad PIP2 coordination in Site-O1 (a), Site-O2 (b), Site-O3 (c), and Site-O4 (d). Kv7.2 protein is shown in ribbon representation with S4 in red, helices A and B in brown, and the rest of the protein in ice blue. A PIP2 lipid (carbon atoms in yellow, oxygen in red, and phosphorus atoms in tan) and the residues of the binding pocket at each site at the end of simulations are shown in sticks (basic residues in blue, polar in green, acidic in red, and hydrophobic in white). eh The contact probability for each chemical moiety of PIP2 headgroup with the key residues in the binding Site-O1 (e), Site-O2 (f), Site-O3 (g), and Site-O4 (h). The chemical moieties of PIP2 headgroup include P1-phosphate (black), P4-phosphate (red), P5-phosphate (orange), 2-hydroxyl (cyan), 3-hydroxyl (brown), and 6-hydroxyl (pink) group of the inositol ring. A heavy-atom distance cutoff of 4 Å was used to define a contact between a protein residue and a phosphate group of PIP2, whereas a 3.5 Å cutoff was used to define a contact between a residue and a hydroxyl group of PIP2. Analysis of the contact probabilities was performed over the last 200 ns of the simulation trajectories. The number of PIP2 interacting events: Site-O1 (n = 6), Site-O2 (n = 5), Site-O3 (n = 2), and Site-O4 (n = 2). Data represent mean ± SEM for analysis of each monomer in 3 independent trajectories.

Overall, PIP2 binds to many more basic residues in the open state than the closed state. However, both states share 4 common PIP2 binding residues including R87 at the N-terminal tail, R153 and K162 in the S2-S3 linker, and R353 in the AB linker. Interestingly, the simulation trajectories (Supplementary Movies 13) show that R353 comes in contact with PIP2 first after which the lipid forms contacts with other residues in Site-O1, suggesting that R353 might act as an initial anchor point of PIP2 in Site-O1.

Charge-neutralizing mutations of potential PIP2 binding residues disrupt voltage-dependent activation of Kv7.2 channels

PIP2 is required for activation of all Kv7 channels3. To test the functional impact of PIP2 binding sites identified by our MD simulations, we introduced charge-neutralizing mutations in select residues that had high contact probability to PIP2 headgroups. In Site-O4, we introduced R214Q in the distal S4, K219N in the S4-S5 linker, and R325Q in pre-Helix-A (Fig. 3a–i). Since MD simulations identified R353 as an initial anchoring point for PIP2 in Site-O1, we also made R353Q in the AB linker (Fig. 3j–l). To test if charge-neutralizing mutations disrupt PIP2 binding to mutated basic residues, we performed 500-ns MD simulations after introducing each mutation in the PIP2-bound conformation of wild-type (WT) channels and monitored the distance between the PIP2 headgroup and the mutated residues.

Fig. 3: Charge-neutralizing mutations disrupt PIP2 binding to the mutated residues in the open Kv7.2 channel.
figure 3

a, d, g, j Time evolution of the distance between the COM of PIP2 headgroup and the COM of the mutated residue upon introducing mutations including R214Q (a), K219N (d), R325Q (g), and R353Q (j) in PIP2-bound Kv7.2 channel in an open state from three independent simulations (different colors). Introduction of these mutations resulted in the dissociation of bound PIP2 lipid, as indicated by the distance increase. b, e, h, k The contact probability for each chemical moiety of PIP2 headgroup with the key residues in open Kv7.2 channels containing R214Q (b), K219N (e), R325Q (h), and R353Q (k). The chemical moieties of PIP2 headgroup include P1-phosphate (black), P4-phosphate (red), P5-phosphate (orange), 2-hydroxyl (cyan), 3-hydroxyl (brown), and 6-hydroxyl (pink) group of the inositol ring. A heavy-atom distance cutoff of 4 Å was used to define a contact between a residue and a phosphate group of PIP2, whereas a 3.5 Å cutoff was used to define a contact between a residue and a hydroxyl group of PIP2. Analysis of the contact probabilities was performed over the last 200 ns of the simulation trajectories. The number of PIP2 interacting events: R214Q (n = 3), K219N (n = 3), R325Q (n = 3), and R353Q (n = 3). Data represent mean ± SEM for 3 independent trajectories. c, f, i, l PIP2 coordination in Site-O4 of Kv7.2-R214Q (c), Kv7.2-K219N (f), and Kv7.2-R325Q (i) and in Site-O1 of Kv7.2-R353Q (l) channel at the end of the respective simulations. The protein is shown in ribbon representation with the S4 in red, helices A and B in brown, and the rest of the protein in ice blue. A PIP2 lipid (carbon atoms in yellow, oxygen in red, and phosphorus in tan) and the residues of the binding pocket at each site are shown in sticks (basic residues in blue, polar in green, acidic in red, and hydrophobic in white).

Upon introducing R214Q, we observed that the bound PIP2 dissociated from the mutated residue and Site-O4 and diffused to basic residues in distal Helix-B in 2 out of 3 simulations (Fig. 3a–c). The introduction of K219N or R325Q resulted in dissociation of PIP2 from the mutated residue in all simulations (Fig. 3d–i). Compared to the WT channel (Fig. 2d, h), these mutations also decreased PIP2 binding to K319 and Q323 but not to other basic residues in Site-O4 (Fig. 3e–f, h–i). Introduction of R353Q resulted in dissociation of PIP2 from the mutated residue in all simulations, although PIP2 remained bound to K162 and R165 in Site-O1 (Fig. 3j–l).

To test if these mutations affect voltage-gated activation of Kv7.2 channels, we performed whole cell patch clamp recording in CHO hm1 cells14,25,45, which display depolarized resting membrane potential (Vm) due to a low level of endogenous K+ channels45,46. Application of depolarizing voltage steps from −100 to +20 mV in cells transfected with GFP and WT Kv7.2 produced a slowly activating outward K+ current with peak current density of 26.5 ± 1.4 pA/pF at +20 mV (Fig. 4a–c, Supplementary Fig. S3). Current activation was sigmoidal, and full activation was reached from 0 mV step with half-maximal current activation potential (V1/2) of −30.5 ± 0.5 mV (Fig. 4a–d, Table 2). Due to this increase in outward K+ currents, these cells also displayed hyperpolarized Vm compared to cells transfected with GFP alone (Supplementary Table S1).

Fig. 4: Charge-neutralizing mutations of potential PIP2 binding residues decrease voltage- and PIP2-dependent activation of homomeric Kv7.2 channels.
figure 4

Whole cell voltage clamp recordings of macroscopic K+ currents through Kv7.2 WT or mutant channels were performed in CHO hm1 cells cotransfected with GFP and PIP5K. The DNA plasmid ratio was 0.2:0.45:0.45 (GFP: Kv7.2 WT or mutant: PIP5K). Cells were held at −80 mV. K+ currents were evoked by depolarizing voltage steps for 1.5 s from −100 mV to +20 mV in 10-mV increments, followed by a step to 0 mV for 300 ms. a Representative traces after subtraction of leak currents. Leak current was defined as non-voltage-dependent current from GFP-transfected cells. Note different Y-axis scales for systems with and without PIP5K. b Average peak current densities (pA/pF) of Kv7.2 WT or mutant channels with or without PIP5K coexpression at all voltage steps. c Average peak current densities of WT or mutant Kv7.2 channels at +20 mV. p values are computed from one-way ANOVA post-hoc Fisher’s test. d Normalized conductance (G/Gmax) at all voltage steps. e Average activation constant (τ) αt +20 mV. The number of GFP-positive cells that were recorded without PIP5K coexpression: Kv7.2 WT (n = 31), R214Q (n = 16), K219N (n = 12), R325Q (n = 11), R353Q (n = 13), R214Q/ K219N (n = 15), R214Q/K219N/R353Q (n = 14). The number of GFP-cotransfected cells that were recorded with PIP5K: Kv7.2 WT (n = 33), R214Q (n = 15), K219N (n = 12), R325Q (n = 11), R353Q (n = 16), R214Q/ K219N (n = 19) or R214Q/K219N/R353Q (n = 14). The source data for Fig. 4b and Fig. 4d are available in Figshare93. Data represent the mean ± SEM. One-way ANOVA with post-hoc Fisher’s multiple comparison test was used. GFP + selected variant: *p < 0.05 for Kv7.2 WT vs. mutant (**p < 0.01, ***p < 0.005); p < 0.05 for Kv7.2 WT + PIP5K vs. mutants + PIP5K (††p < 0.01, †††p < 0.005); ^p < 0.05 for the difference between -PIP5K and +PIP5K within the same transfection (^^p < 0.01, ^^^p < 0.005).

Table 2 Biophysical properties of Kv7.2 homomers in CHO hm1 cells co-transfected with GFP and PIP5K.

Compared to WT channels, Kv7.2-K219N channels produced K+ currents with significantly smaller peak current density (8.2 ± 1.6 pA/pF at +20 mV), a large depolarizing shift in V1/2 (−13.7 ± 1.7 mV), and a slower activation kinetic (Fig. 4a–e, Table 2). The R325Q mutation significantly reduced peak current density by ~90% (3.5 ± 0.4 pA/pF at +20 mV) (Fig. 4a–c). At −20 to +20 mV, the R325Q-transfected cells produced currents that were larger than those in cells transfected with GFP alone (Supplementary Fig. S3g), suggesting that Kv7.2-R325Q channels are functional. The peak current densities of Kv7.2-R214Q and Kv7.2-R353Q channels were also decreased by ~30%, but their voltage dependence and activation time constant were unaffected (Fig. 4a–e, Supplementary Figs. S3S4). Similar surface expressions were observed for WT and all tested mutants except for Kv7.2-R214Q, which displayed lower surface expression than the WT (61.8 ± 11.0% of WT, Supplementary Fig. S5). The surface/total protein ratio of all tested mutants did not differ from WT Kv7.2 (Supplementary Fig. S5e), suggesting that none of the mutations affected the proportion of Kv7.2 to express on the plasma membrane.

Charge-neutralizing mutations of potential PIP2 binding residues alter PIP2 sensitivity of Kv7.2 channels

To test if charge-neutralizing mutations alter gating modulation of Kv7 channels by PIP2, we increased cellular PIP2 level by co-transfecting phosphatidylinositol-4-phosphate 5-kinase (PIP5K) which converts phosphatidylinositol 4-phosphate to PIP247. Since the endogenous membrane level of PIP2 is not enough to saturate Kv7 channel activation24, enhancing cellular PIP2 level by PIP5K expression is shown to increase single-channel open probability48 and whole-cell current densities of Kv7.2 channels14,28. Consistent with previous reports14,28,48, PIP5K expression significantly increased peak Kv7.2 current density by 4.4 ± 0.2-fold (118.7 ± 4.7 pA/pF) with a hyperpolarizing shift in their voltage dependence and a faster activation kinetic (Fig. 4a–e, Supplementary Figs. S3–S4).

In contrast, the R325Q mutation abolished the PIP5K-induced current potentiation (Fig. 4a–c, Supplementary Figs. S3S4). In the presence of PIP5K, Kv7.2 channels containing R214Q, K219N or R353Q mutations produced significantly less outward K+ currents compared to WT channels (Kv7.2-R214Q: 79.3 ± 6.8 pA/pF; Kv7.2-K219N: 51.1 ± 3.9 pA/pF; Kv7.2-R353Q: 70.3 ± 3.0 pA/pF) (Fig. 4a-e, Supplementary Figs. S3–S4, Table 2). However, the fold increases in their PIP5K-induced current potentiation were comparable to the WT channels (4.7 ± 0.4 fold for R214Q, 6.2 ± 0.5 fold for K219N, 4.0 ± 0.2 fold for R353Q) (Fig. 4a–e, Supplementary Figs. S3–S4, Table 2), suggesting that the charge-neutralizing mutation of a single residue may not be sufficient to fully dissociate PIP2 from Site-O4 and Site-O1 (Fig. 3b,e,h,k).

Therefore, we next generated the R214Q/K219N double mutant (DM) and the R214Q/K219N/R353Q triple mutant (TM). Compared to Kv7.2-WT channels, both DM and TM channels displayed smaller peak current densities (DM = 12.5 ± 0.6 pA/pF, TM = 12.9 ± 1.3 pA/pF) and activated at more depolarized voltages with slower activation kinetics (Fig. 4a–e, Supplementary Figs. S3–S4, Table 2). PIP5K co-expression increased the peak current density of DM channels by 4.3 ± 0.3-fold (53.5 ± 4.1 pA/pF) but that of TM channels only by 2.1 ± 0.2-fold (27.5 ± 2.7 pA/pF) (Fig. 4a–e, Supplementary Figs. S3–S4, Table 2), indicating that the triple mutation decreased the channel sensitivity to PIP2 enhancement.

To further investigate PIP2 sensitivity of mutant channels, we examined current decay upon membrane PIP2 depletion. To achieve this, we coexpressed Danio rerio voltage-sensitive phosphatase (Dr-VSP)14. Upon activation of Dr-VSP, K+ currents through WT channels reached a maximal decay of at +100 mV (0.52 ± 0.02, Fig. 5a–c). Kv7.2-R353Q channels displayed larger current decays at +40 and +60 mV, but showed similar decays to WT channels at +100 mV (Fig. 5a–c). In contrast, the VSP-induced current decays of R214Q and K219N channels were smaller than WT channels and were minimal in Kv7.2-R325Q, DM, and TM channels (Fig. 5a–c), indicating these mutants have decreased sensitivity to PIP2 depletion.

Fig. 5: Charge-neutralizing mutations of potential PIP2 binding residues alter Kv7.2 current response to Dr-VSP activation.
figure 5

a Representative current traces showing Dr-VSP-mediated Kv7.2 current decay in CHO hm1 cells co-expressing Dr-VSP and Kv7.2 WT or mutants from −20 mV to +100 mV. CHO cells were held at –70 mV, and a brief voltage step to –60 mV was applied to calculate the linear leak. 10 s step depolarizations were applied in 20-mV steps from –20 to +100 mV with 2 min inter-step intervals to allow PIP2 regeneration. Red trace shows current decay curve when cells were held at +40 mV. b Ratio of current decay in Kv7.2 WT or mutant channels at each voltage step. c Kv7.2 current decay ratio at +40 mV, +60 mV, +80 mV and +100 mV. The number of cotransfected cells that were recorded with EGFP-tagged Dr-VSP: Kv7.2 WT (n = 27), R214Q (n = 14), K219N (n = 13), R325Q (n = 12), R353Q (n = 15), R214Q/ K219N (n = 13) or R214Q/K219N/R353Q (n = 11). The source data for Fig. 5b is available in Figshare93. Data represent the mean ± SEM. One-way ANOVA Fisher’s test results are shown (*p < 0.05, **p < 0.01 and ***p < 0.005).

PIP2-mediated correlated motions of the VSD and the pore domain

Voltage-dependent conformational changes of the VSD and the pore domain are critical for the gating of Kv channels49. To examine PIP2-mediated allosteric interactions, we investigated the communities formed in the WT and mutant channels by employing dynamic network analysis and Pearson correlation. These communities correspond to sets of residues that move in a correlated manner during the MD simulations and thus represent strongly connected regions. We detected the presence of a pronounced community connecting the end of S4, the S4-S5 linker, S6, and pre-Helix-A in WT channels (Fig. 6a). The thickness of the edges connecting the amino acids corresponds to the strength of the correlation between them. All mutant channels show thinner edges within this community (Fig. 6b–d), indicating weaker correlated motions in this area compared to WT channels. Interestingly, we also detected the emergence of additional (sub)communities in all the studied mutants (Fig. 6b–d), in line with the described weaker interactions in this region. The presence of disconnected communities suggests uncorrelated dynamics of this region (Site-O4) after decreased PIP2 binding (Fig. 3). These data suggest that R214Q, K219N, and R325Q mutations disrupt the correlated motions of the VSD and the S6 gate in the pore domain of Kv7.2.

Fig. 6: Charge-neutralizing mutations of potential PIP2 binding residues in Site-O4 disrupt the correlated motions of the VSD and the S6 gate of Kv7.2 channels.
figure 6

Network-based community analysis in WT (a) and mutant Kv7.2 channels containing R214Q (b), K219N (c), and R325Q (d) mutations in Site-O4. A community represents a set of residues that move in a correlated manner during the MD simulations. The thickness of the edges between the amino acids within each community corresponds to the strength of correlation between them. The network analysis was performed after combining all 3 independent simulation trajectories. a Binding of PIP2 at Site-O4 in the WT channel results in correlated motions of the VSD (the S4 and the S4-S5 linker) and the gate S6, highlighted by a single community in green. b Introduction of the R214Q mutation leads to uncorrelated motions of the VSD and the gate, highlighted by 3 separate communities colored in pink, green, and gray. c Introduction of the K219N mutation leads to uncorrelated motions of the VSD and the gate, highlighted by the smaller community in green and the emergence of an additional community colored in pink. d Kv7.2-R325Q channels displayed weaker correlated motions of the S4 and the S4-S5 linker, and the decoupling of the S6 of the pore domain from the VSD, highlighted by the much smaller community in green and the emergence of 2 additional communities colored in pink and gray.

PIP2-dependent conformational change of helices A and B

The cryo-EM structure of Kv7.1/KCNE3 in complex with CaM suggests that PIP2 induces a conformational change in the cytoplasmic domain that may facilitate the opening of Kv7.1 channels22. To test the conformational impact of PIP2 on Kv7.2 channels, we compared the dynamics of the open channel in PIP2-containing lipid bilayers with a control simulation performed in the absence of PIP2 lipids. The most significant PIP2-induced conformational change was observed in the cytoplasmic helices A and B (Fig. 7a–b). The effect was quantified by calculating the orientation of the helical pair with respect to the membrane normal (Fig. 7c). In a PIP2-free lipid bilayer, these helices fluctuate around their initial position (θ = 83.8 ± 13.2°) and remain in a largely solvent-exposed conformation (Fig. 7a,c). In the presence of PIP2, the helices adopt a conformation where they interact directly with the lipid bilayer (θ = 95.5 ± 3.7°)(Fig. 7b–c). This large-scale conformational change also forms a pathway for PIP2 to move along Helix-B and ultimately bind to Site-O4 (Fig. 7d, Supplementary Movie 4). Since R353 in the AB linker acts as an initial anchor point for PIP2 binding (Supplementary Movies 13), we next tested if a charge-neutralizing mutation at this residue (R353Q) affects the PIP2-induced conformational change. We found that the R353Q mutation moves the helices back toward a more solvent-exposed conformation (θ = 86.7 ± 1.9°) (Fig. 7c).

Fig. 7: PIP2-mediated conformational changes of helices A and B in open Kv7.2 channel.
figure 7

a Snapshot of a representative Kv7.2 monomer in PIP2-free membranes. b Snapshot of a representative Kv7.2 monomer in PIP2-containing membranes. c Orientation angle of helices A and B with respect to the membrane normal. The distribution of the orientation angle of helices A and B in WT Kv7.2 channel over the last 200 ns of all the simulations in membranes with (w/) or without (w/o) PIP2 as well as that of mutant Kv7.2-R353Q channel in membranes with PIP2. In PIP2-free membranes, the helical pair of the WT channel adopts a largely solvent-exposed conformation. Addition of PIP2 induces a drastic conformational change in these helices from a solvent-exposed conformation to a bilayer-interacting conformation. However, the R353Q mutation in the AB linker moves the helices closer to a solvent-exposed conformation even in the presence of PIP2. d Large-scale conformational change in the cytoplasmic helices provides a pathway for PIP2 movement along Helix-B to its ultimate localization at Site-O4. All the phosphorus atoms of the bilayer are shown in vdW (van der Waals) representation. Kv7.2 protein is shown in ribbon representation with the S4 in red, helices A and B in brown, and the rest of the protein in ice blue.

Discussion

Neuronal Kv7 channels are known as the “M-channels” due to their inhibition by the activation of M1 and M3 muscarinic acetylcholine receptors13. PIP2 hydrolysis underlies current inhibition of M-channels, bringing attention to Kv7-PIP2 interaction6,20. Increasing the PIP2 level enhances current density and the open probability of Kv7.2 channels and induces a hyperpolarized shift in their voltage-dependence6,14,25,28,48. However, the detailed mechanism underlying PIP2-dependent regulation of neuronal Kv7 channel remains unclear. In this study, we address this knowledge gap by identifying PIP2 interaction sites in both open and closed Kv7.2 channels.

Our all-atom MD simulations have revealed PIP2 localization to 3 sites at the periphery of the VSD in the closed Kv7.2 channel, whereas PIP2 binds to 4 distinct sites at the VSD, the pore domain, and intracellular helices in the open channel (Fig. 1, Supplementary Figs. S1S2). The common PIP2 binding domains in both open and closed channels are the intracellular N-terminal tail, the S2-S3 and AB linkers. Importantly, PIP2 binding in the open channel is coordinated by multiple basic residues from different functional domains of Kv7.2.

Our identification of the S2-S3 and S4-S5 linkers as PIP2 binding domains is consistent with a previous simulation study reported by Zhang et al.50. However, our study has identified many more PIP2 binding regions in Kv7.2 channel compared to Zhang et al. These include the intracellular N-terminal tail, the distal ends of the S4 and S6 as well as pre-Helix-A, Helix-A, the AB linker, Helix-B, and the BC linker in the intracellular C-terminal tail. The difference could be attributed to structural templates used in each study. Our homology model of Kv7.2 was based on the cryo-EM structure of Kv7.1 containing cytoplasmic helices A-C and a part of intracellular N-terminal tail10. In contrast, Zhang et al modeled the Kv7.2 transmembrane domains only (residues 95–337) based on the crystal structures of Kv1.2 and an activated bacterial K+ channel KcsA50. In the following sections, we discuss the functional implications of the additional PIP2 binding sites, which we have identified in this study.

To the best of our knowledge, our MD simulations have identified a novel PIP2-interacting interface in the open Kv7.2 channel, Site-O4, which is comprised of the distal ends of S4 and S6, the S4-S5 linker, and pre-Helix-A. We show that R325 in pre-Helix-A interacts with PIP2 (Fig. 2), and its charge-neutralizing mutation, R325Q, abolishes this interaction (Fig. 3), basal current, and the sensitivity to PIP2 (Figs. 45). Severe reduction in current expression and PIP2 sensitivity has also been reported in Kv7.2 channels containing recurrent EE variant R325G28 which causes drug-resistant seizures, neurodevelopmental delay, and intellectual disability18,51. The R325 is the first residue in a “RQKH” motif conserved in all Kv7 subunits3,22,28. This motif in Kv7.1 undergoes a PIP2-induced conformational change from an unstructured loop to a helix22, further supporting that PIP2 binding to R325 is involved in activation of Kv7.2 channels.

In Site-O4, PIP2 also interacts with R214 in distal S4 and K219 in the S4-S5 linker in the open channel (Figs. 1 and 2). Although the S4-S5 linker was previously reported to mediate PIP2 modulation of Kv7.1-Kv7.33,21,50,52, to the best of our knowledge, the identification of R214 as a PIP2 binding site in Kv7.2 is a novel finding. R214 is also the target of two epilepsy mutations53,54,55, supporting its functional importance. Although the charge-neutralizing mutations R214Q and K219N reduce PIP2 interactions with the mutated residues (Fig. 3), their reduced sensitivity to PIP2 depletion (Fig. 5) suggests increased PIP2 affinity to other residues of the channel. K219N, but not R214Q, induces a depolarizing shift in voltage dependence (Fig. 4) similar to R243A and K248A in the corresponding residue in Kv7.327,52. However, the double R214Q/K219N mutation further shifts the activation curve to a depolarized potential and induces less sensitivity to PIP2 depletion compared to single mutants (Figs. 45), demonstrating the synergetic effects. These findings suggest that PIP2 interaction with both the distal S4 and the S4-S5 linker is needed for robust voltage-dependent activation of Kv7.2 channels.

Previous studies proposed the role of PIP2 in coupling of the VSD activation to pore opening in Kv7.1 and Kv7.310,21,22,52,56. In Kv7.2 channels, we observe that PIP2 binding to Site-O4 forms an allosteric network of interactions, leading to a correlated motion of the main voltage sensor S4, the S4-S5 linker, the gate S6, and pre-Helix-A (Fig. 6), suggestive of the coupling of the VSD to the pore domain. Charge neutralizing mutations (R214Q, K219N, or R325Q) induce PIP2 dissociation from the mutated residues (Fig. 3) and disrupt the coordinated movements of the VSD and the pore domain (Fig. 6), indicative of their decoupling. To the best of our knowledge, this is the first study to provide an atomic-level structural basis for the allosteric role of PIP2 in voltage-dependent activation of neuronal Kv7.2 channels.

Our MD simulations have identified Site-O2 comprised of the K552-R553-K554 motif in Helix-B and with R560 in the BC linker as a PIP2 binding site in the open state (Fig. 2). We have previously shown that current potentiation induced by increasing PIP2 level is abolished by EE mutations K552T, R553L, and R560W14,25 but enhanced by K554N25, highlighting the role of these basic residues in PIP2 modulation of Kv7.2 channels14,25. The corresponding basic residues in Helix-B of Kv7.1 bind to PIP2 in vitro23 but those in Kv7.3 do not affect its sensitivity to PIP2 depletion27, suggesting the subunit-specific difference of Helix-B in mediating PIP2 binding.

Our simulations also demonstrate that PIP2 interacts with Site-O1 formed by F346, Y347, and R353 at the beginning of the AB linker and with K162 and R165 in the S2-S3 linker (Fig. 2). Due to the presence of a lysine residue (K162) and two aromatic residues (F346 and Y347), this site fulfills the requirement for a canonical PIP2 binding site57. Consistent with the conserved sequence of the S2-S3 linker among Kv7 subunits3, PIP2 binding to this linker has also been reported in Kv7.13,21,22,58 and Kv7.327,52, suggesting the role of this linker in PIP2 modulation of Kv7 channels.

Although a “cationic cluster” (K452/R459/K461) in the distal region of the AB linker is implicated in PIP2 modulation of Kv7.2 current26, the charge-neutralizing mutation of R353 in the proximal region of this linker reduces basal current expression of Kv7.2 channels and increases current inhibition upon PIP2 depletion, suggestive of decreased PIP2 affinity (Figs. 45). Combination of R353Q with R214Q and K219N in Site-O4 further reduces basal current and the PIP5K-induced current enhancement (Fig. 4), suggesting that R353 modulates Kv7.2 channels by coordinating PIP2 binding with other residues. Indeed, R353 serves as an initial anchor point for PIP2 binding to Site-O1 and Site-O4 (Supplementary Movies 14). Furthermore, PIP2 induces the conformational change of helices A and B from the largely solvent-exposed conformation to a bilayer-interacting conformation, whereas the R353Q mutation attenuates this effect (Fig. 7), suggesting that PIP2 binding to R353 contributes to this conformational change.

The conformational change in Kv7.2 is different from that observed in the PIP2-bound Kv7.122,59 which lacks the analogous arginine residue and shows low sequence homology to the AB linker of Kv7.2. Kv7.3, on the other hand, has the analogous arginine residue in the AB linker with fairly well-conserved sequence, suggesting that Kv7.3 may adopt a similar conformation as Kv7.2 in the presence of PIP2. We propose that PIP2 binding to R353 affects the opening of Kv7.2 channels by inducing a bilayer-interacting conformation of the helical pair and facilitating the movement of PIP2 along Helix-B and its binding to distal Helix-B (Site-O2). Subsequent interaction with other basic residues in Site-O1 and Site-O4 will induce the transition from the closed to the open state by coordinating allosteric movement of the VSD and the pore domain. R353 is also the target of three epilepsy mutations (ClinVar Database, NCBI)60,61,62, further supporting the role of R353 in PIP2 modulation of Kv7.2 channels.

Neuronal Kv7 channels are highly enriched in the axonal surface where they regulate the AP firing threshold, frequency and shape13,63, whereas dendritic Kv7 currents increase the threshold of Ca2+ spike initiation64. Heterozygous deletion of KCNQ2 gene in mice leads to hippocampal hyperexcitability and increased seizure propensity65,66. Consistent with their critical role in inhibiting neuronal excitability, > 400 BFNE and EE mutations are found in KCNQ217. Computational algorithms have identified the S4, the pore loop, the S6, pre-Helix-A, Helix B, and the BC linker as hotspots for EE mutations14,67, whereas BFNE mutations are enriched in the S2-S3 linker67. The overlap between epilepsy mutation hotspots and the PIP2 binding domains identified in our study underscores the critical role of PIP2 in the pathophysiological mechanism underlying KCNQ2-related epilepsy.

Current anti-epileptic drugs are ineffective in treating many epilepsy patients with Kv7.2 EE variants18,19,68. M-current inhibition upon PIP2 depletion results in neuronal hyperexcitability69, and impaired PIP2 sensitivity of Kv7.2 channels is associated with EE variants14,25,28. Retigabine (INN; USAN ezogabine) is a selective agonist of Kv7.2-Kv7.5 channels, but not Kv7.1 channel70,71. Retigabine suppresses seizures in animal models and humans70,71, however, it has been discontinued as an anti-epileptic drug due to adverse side effects72. However, it may be effective in opening EE mutant channels with impaired PIP2 binding because it stabilizes the open state of KV7.2 and KV7.3 channels by binding to a hydrophobic pocket near the gate73. Alternatively, strengthening PIP2-Kv7.2 interaction may increase Kv7 current. For example, zinc pyrithione can rescue M-current in hippocampal neurons following PIP2 depletion by competing with PIP2 for Kv7.2 activation74. Recently, a compound, CP1, is shown to substitute PIP2 for the VSD-pore coupling in Kv7.1 channel, and to a less extent, Kv7.2 and Kv7.2/Kv7.3 channels75, suggesting its potential to inhibit neuronal hyperexcitability. Our in-depth investigation of PIP2-Kv7.2 interaction may provide the foundation to explore a new class of therapeutics for epilepsy that can control PIP2 modulation of neuronal Kv7 channels.

Methods

Structural models of open and closed states of Kv7.2

The open and closed conformations of Kv7.2 channel used in the molecular dynamics (MD) simulations were modeled following the procedure described in a recent study14. Briefly, the closed conformation of the channel was modeled based on the cryo-EM structure of Kv7.1 (PDB ID: 5VMS)10. Multiple sequence alignment of the target template and Kv7.2 was performed using TCoffee web server (https://www.ebi.ac.uk/Tools/msa/tcoffee/). After the alignment, the homology model of the closed state was built with MODELLER76. Our final model contains residues from 74–363 and 537–594. The open conformation of Kv7.2 was then constructed by performing non-equilibrium, driven MD simulations. We performed a 20-ns targeted MD (TMD)77 simulation during which the closed structure was driven towards an open form, while embedded in a lipid bilayer containing 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine (POPC) and 2.2% 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylinositol 4,5-bisphosphate (PIP2) lipids. The target of the TMD simulation was selected to be the highly homologous Kv1.2/Kv2.1 channel in an open-state conformation (PDB ID: 2R9R)78. As major structural changes occur in the pore region of the channel, we applied a restraint (k = 250 kcal/mol/Å2; only Cα atoms were driven) on the S4-S5 and S6 helices of each monomer to drive it towards the target open state. The structural stability of the open and closed conformations of Kv7.2 was evaluated by monitoring the degree of opening of the states, calculated by the number of water molecules in the pore helix and the selectivity filter during 400-ns MD runs in explicit lipid bilayers14. During these simulations we consistently observed stable and a higher number of water molecules in the open state of the channel, as compared to the closed state14.

PIP2-binding simulations

The open and closed conformations of Kv7.2 were embedded in multiple independently generated POPC membranes with or without 2.2% PIP2 lipids (Table 1). All the membranes were constructed using the Membrane Builder module in CHARMM-GUI79, and the initial placement of PIP2 lipids was intentionally varied in each membrane. All PIP2 lipid molecules were initially at least 15 Å away from the protein. The systems were solvated using TIP3P water and buffered at 150 mM KCl to neutralize. The final simulation systems consisted of ~300,000 atoms. All the simulations were performed in the absence of calmodulin. To further examine the specific lipid-protein interactions captured in the modeled closed Kv7.2, we also performed additional lipid-binding simulations of a recent cryo-EM structure of Kv7.2 (PDB ID: 7CR3) after removing calmodulin44. All the missing loops were constructed using MODELLER76 and the entire tetrameric structure was embedded in an explicit lipid bilayer containing POPC and 2.2% of PIP2 lipids. Three independent 500-ns simulations were performed on this system.

Molecular dynamics simulation protocols

All the simulations were performed under periodic boundary conditions using NAMD280,81 and CHARMM36m force field parameters82,83 for protein and lipid. During the initial equilibration, the protein’s backbone atoms were harmonically restrained to their initial positions with a force constant of k = 1 kcal/mol/Å2. The restraints were released at the start of the production run. All the non-bonded forces were calculated with a cutoff of 12 Å and a switching distance of 10 Å. Long-range electrostatic forces were calculated using the particle mesh Ewald (PME) method84. A Langevin thermostat using γ = 1 ps−1 was used to maintain the system temperature at 310 K. The pressure was maintained at 1 bar using a Nosé Hoover Langevin piston method85. During pressure control, the simulation box was allowed to fluctuate in all the dimensions with constant ratio in the xy (lipid bilayer) plane. An integration time step of 2 fs was used in all the simulations.

Simulation analysis

To characterize lipid-protein interactions and potential lipid binding sites, occupancy maps of the PIP2 headgroup were calculated using the Volmap plugin in VMD86. Based on our previous experience35, a 4-Å heavy-atom distance cutoff was chosen to define contacts between the phosphate groups of PIP2 lipids and protein residues, while a 3.5-Å cutoff was used to define contacts with the hydroxyl groups of PIP2. The role of PIP2 lipids in stabilizing the open conformation of Kv7.2 was determined by performing dynamical network analysis using the Network-View plugin87 in VMD. In a network, all Cα atoms are defined as nodes connected by edges if they are within 4.5 Å of each other for at least 75% of the MD trajectory. Pearson correlation was used to define the communities (the set of residues that move in concert) in the network.

DNA construct and mutagenesis

Plasmid pcDNA3 carrying KCNQ2 cDNA (GenBank: Y15065.1) encoding Kv7.2 (GenBank: CAA 75348.1) was previously described14,25,88,89. This short isoform of Kv7.2 lacks 2 exons compared to the reference Kv7.2 sequence (GenBank: NP_742105.1). Our lab has previously shown that currents through this Kv7.2 variant can be potentiated by PIP2 increase and are sensitive to Dr-VSP activation14,25. Plasmid pIRES-dsRed-PIPKIγ9090 was a kind gift from Dr. Anastasios Tzingounis (University of Connecticut). Selected mutations (R214Q, K219N, R325Q, R353Q, R214Q/K219N double mutation, R214Q/K219N/R353Q triple mutation) were generate using the Quik Change II XL Site-Directed Mutagenesis Kit (Agilent) and verified by sequencing the entire cDNA construct. The following primers were used for mutagenesis: R214Q (sense-5′GAT CCG CAT GGA CCG GCA GGG AGG CAC CTG G3′, antisense-5′CCA GGT GCC TCC CTG CCG GTC CAT GCG GAT C3′), K219N (sense-5′GGA GGC ACC TGG AAC CTG CTG GGC TCT GTG3′, antisense-5′CAC AGA GCC CAG CAG GTT CCA GGT GCC TCC3′), R325Q (sense- 5′GGT TCA GGA GCA GCA CCA GCA GAA GCA CTT TGA GAA G3′, antisense- 5′CTT CTC AAA GTG CTT CTG CTG GTG CTG CTC CTG AAC C3′), R353Q (sense- 5′GCC ACC AAC CTC TCG CAG ACA GAC CTG CAC TCC3′, antisense- 5′GGA GTG CAG GTC TGT CTG CGA GAG GTT GGT GGC3′).

Electrophysiology

Whole cell patch clamp recording was performed in Chinese ovary cells (CHO hm1) at room temperature (22–24 °C) as described14,25. Cells were plated on 12 mm glass coverslips (Warner Instrument, 1 ×105 cells per coverslip) treated with poly D-lysine (0.1 mg/mL) (Sigma-Aldrich). To express Kv7.2 channels and PIP5K, cells were transfected with pEGFPN1 (0.2 μg), pIRES-dsRed-PIPKIγ90 (0.45 μg) and pcDNA3-Kv7.2 WT or mutant (0.45 μg). For the control experiment of PIP5K, cells were transfected with pEGFPN1 (0.65 μg) and pcDNA3-Kv7.2 WT or mutant (0.45 μg). At 24–48 h post-transfection, GFP-positive cells were recorded in extracellular solution containing (mM): 138 NaCl, 5.4 KCl, 2 CaCl2, 1 MgCl2, 10 D-glucose, and 10 HEPES (pH 7.4, 307–312 mOsm). Patch pipettes (3–4 mΩ) were filled with intracellular solution containing (mM): 140 KCl, 2 MgCl2, 10 EGTA, 10 HEPES, 5 Mg-ATP (pH 7.4 with KOH, 290–300 mOsm). To record K+ currents, cells were held at −80 mV. Currents were evoked by depolarization for 1.5 s from −100 mV to +20 mV in 10-mV increments, followed by a step to 0 mV for 300 ms. Leak-subtracted current densities (pA/pF), normalized conductance and channel biophysical properties were calculated as described25. Briefly, leak current was defined as non-voltage-dependent current through GFP-transfected CHO hm1 cells. Current density was calculated by dividing leak-subtracted current (pA) by capacitance (pF). V1/2 and the slope factor (k) were calculated by fitting the points of G/Gmax to a Boltzmann equation in the following form: G/Gmax = 1/{1+exp(V1/2−V)/k}.

To examine the decline of Kv7.2 current upon activation of Dr-VSP, CHO hm1 cells were transfected with pDrVSP-IRES2-EGFP (0.5 μg) and pcDNA3-Kv7.2 WT or mutant (0.5 μg). The pDrVSP-IRES2-EGFP plasmid was a gift from Yasushi Okamura (Addgene plasmid # 80333). Voltage-clamp recording of Kv7.2 current upon depolarization-induced Dr-VSP activation was performed as described91 with an external solution containing 144 mM NaCl, 5 mM KCl, 2 mM CaCl2, 0.5 mM MgCl2, 10 mM glucose and 10 mM HEPES (pH 7.4). Patch pipettes (3–4 MΩ) were filled with intracellular solution containing 135 mM potassium aspartate, 2 mM MgCl2, 1 mM EGTA, 0.1 mM CaCl2, 4 mM ATP, 0.1 mM GTP and 10 mM HEPES (pH 7.2). Cells were held at −70 mV and 10 s step depolarizations were applied in 20-mV steps from −20 to +100 mV with 2 min inter-step intervals to allow PIP2 regeneration. The extent of Kv7.2 current decay upon Dr-VSP activation during 10 s depolarization was measured as the ratio of current at 10 s over peak current at each voltage step.

Western blot

CHO hm1 cells were plated on 35 mm tissue culture dishes (Corning, 3 × 105 cells per well). Next day, cells were transfected with pcDNA3-Kv7.2 WT or mutant (0.5 μg) using FuGENE6 transfection reagent (Promega). At 24 h post-transfection, cells were lysed, and lysates were analyzed by immunoblotting as described14. Briefly, cells were collected by cell scraper in 300 μL ice-cold lysis buffer containing (mM): 150 NaCl, 50 Tris, 2 EGTA, 1 EDTA, 1% Triton-X, 0.5% deoxycholic acid (pH 7.4), supplemented with Halt protease inhibitor cocktail (Thermo Fisher Scientific). Lysates were harvested by 15 min incubation on ice, followed by 15 min centrifugation at 14,000 × g in 4 °C. Samples were mixed with SDS sample buffer in 1:5 ratio and heated at 75 °C for 30 min. The SDS sample buffer contained (mM): 75 Tris, 50 TCEP, 0.5 EDTA, 10% SDS, 12.5% glycerol, 0.5 mg/mL Bromophenol Blue. The samples were then run on 4%–20% gradient SDS-PAGE gels (Bio-Rad) and transferred to a methanol-treated polyvinyl difluoride (PVDF) membrane (Millipore). The membranes were blocked with blocking buffer (5% nonfat milk/0.1% Tween-20 in Tris-buffered saline/TBS containing 150 mM NaCl, 50 mM Tris, pH 7.5) for 1 h followed by overnight incubation of primary antibodies in washing buffer (1% nonfat milk/0.1% Tween-20 in TBS) in 4 °C. After 1 hr incubation with horse radish peroxidase (HRP)-conjugated secondary antibodies in washing buffer, membranes were treated with Enhanced Chemiluminescence substrate (ECL, Thermo Fisher Scientific, #32106) and immediately imaged with the iBright CL1000 imaging system (Thermo Fisher Scientific). Acquired images were analyzed using ImageJ software (National Institutes of Health). GAPDH, α-tubulin and β-tubulin were used as loading controls. Background-subtracted intensities of each immunoblot band was measured and the Kv7.2/loading control ratio of WT or mutants was computed. The Kv7.2/loading control of WT was used as 100% and mutants were normalized to WT as described14. Antibodies used include anti-GAPDH (Cell Signaling #2118, 1:1000 dilution), anti-α-tubulin (Cell Signaling #2144, 1:1000 dilution), anti-β-tubulin (Cell Signaling #2146, 1:1000), anti-Kv7.2 (Neuromab, N26A/23, 1:200 dilution), donkey anti-rabbit and donkey anti-mouse HRP secondary antibodies (The Jackson Laboratory, 711-035-152; 715-035-150).

Surface biotinylation

CHO hm1 cells were plated on 60 mm culture dishes (Corning, 8 × 105 cells per well). Next day, cells were transfected with pcDNA3-Kv7.2 WT or mutant (0.8 μg) using FuGENE6 transfection reagent (Promega). At 24 h post-transfection, the cells were subjected to surface biotinylation as previously described92. The culture dishes containing transfected cells were placed on ice and washed with 1X PBS twice. To biotinylate surface proteins, the cells were then incubated with Sulfo-NHS-SS-Biotin (1 mg/mL, Pierce) in ice-cold PBS (3 mL) for 20 min. The cells were then washed with 1X PBS twice and 1X TBS once. Cells were collected using cell scraper in 400 μL ice-cold lysis buffer containing: (mM): 150 NaCl, 50 Tris, 2 EGTA, 1 EDTA, 1% Triton-X, 0.5% deoxycholic acid, supplemented with Halt protease inhibitor cocktail (Thermo Fisher Scientific). Lysates were harvested by 15 min incubation on ice, followed by 15 min centrifugation at 14,000 x g at 4 °C. 40 μL lysates were saved for western blotting. The remaining 360 μL of lysates were incubated with 50% NeutraAvidin agarose beads (Pierce, 100 μL of 1:1 slurry) for overnight at 4 °C to isolate biotinylated surface proteins from the lysate. After washing with the lysis buffer, biotinylated surface proteins were eluted by heating in 1x SDS sample buffer containing 50 mM TCEP at 75 °C for 30 min. Eluted biotinylated surface proteins and lysates were examined by immunoblotting for Kv7.2 and α-tubulin or β-tubulin as described in the previous section. ImageJ software (NIH) was used to measure background-subtracted intensity of each immunoblot band, and surface or total Kv7.2 intensity was normalized to total α-tubulin or β-tubulin band intensity. To confirm that cell membranes were intact and intracellular proteins were not biotinylated during surface biotinylation, immunoblot of biotinylated surface proteins was also performed with anti-α-tubulin or anti-β-tubulin.

Statistics and reproducibility

All the measurements were taken from distinct samples. All electrophysiology and immunoblotting data are reported as mean ± SEM. Origin 9.1 (OriginLab, Inc) was used for Student’s t-test and one-way ANOVA with post-hoc Fisher’s multiple comparison tests. Specifically, one-way ANOVA with post-hoc Fisher’s test was used for surface biotinylation and electrophysiology figures with the exception that Student’s unpaired t-test was used when comparing the results before and after PIP5K cotransfection in Supplementary Fig. 4a–b. Listed sample sizes of electrophysiology indicate number of cells successfully recorded, while the sample sizes of surface biotinylation studies represent the numbers of experimental replica. Statistical significance was assessed at a priori value (p) < 0.05.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

The datasets generated and analyzed in the current study and presented as the main and supplementary figures are available in the Figshare repository with the identifier https://doi.org/10.6084/m9.figshare.1518103893. All other data that support the findings of this study will be available from the corresponding authors upon reasonable request.

Code availability

Simulation trajectories were collected using the simulation program NAMD. Visualization and analysis were performed using VMD and Python. All of these software packages are publicly available.

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Acknowledgements

This research was supported by the National Institutes of Health under awards R01-GM123455 and P41-GM104601 from the National Institute of General Medical Sciences (to E.T.) and R01-NS083402 and R01-NS097610 from the National Institute of Neurological Disorders and Stroke (to H.J.C.). S.P. acknowledges receiving support from the Beckman Institute Graduate Fellowship. We would like to also acknowledge the computing resources provided by Blue Waters of National Center for Supercomputing Applications (to E.T.) and by Extreme Science and Engineering Discovery Environment (XSEDE award MCA06N060 to E.T.) and TACC Frontera. We would like to thank Tao Jiang for her assistance with simulations on Frontera. All authors declare no competing interests.

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S.P., J.Z., H.J.C. and E.T. conceived of the study and participated in its design and coordination. S.P., K.L. and E.T. performed and analyzed MD simulation. E.C.K., J.Z. and H.J.C. generated and characterized selected mutations. S.P., J.Z., E.C.K., H.J.C. and E.T. wrote the manuscript. All authors read and approved the final manuscript.

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Correspondence to Hee Jung Chung or Emad Tajkhorshid.

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Pant, S., Zhang, J., Kim, E.C. et al. PIP2-dependent coupling of voltage sensor and pore domains in Kv7.2 channel. Commun Biol 4, 1189 (2021). https://doi.org/10.1038/s42003-021-02729-3

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