Mode shift of human K_v12.1 channels was recently demonstrated (Li et al., 2015), but a detailed characterization is currently not available. In order to identify exclusive properties of human K_v12.1 channels, we thus set out with detailed biophysical analysis of these channels in an overexpression system. In CHO cells, activation of K_v12.1 channels through depolarizing voltage steps produced robust outwardly rectifying currents (Figure 1A) (Zou et al., 2003; Li et al., 2015; Dai and Zagotta, 2017). Channel deactivation at hyperpolarized potentials was best described by double exponential kinetics, and deactivation slowed down with more depolarized pre-potentials (Figures 1B,C). When we activated K_v12.1 channels at 0 mV for different time intervals, tail current amplitudes increased (Figures 1D,E) and deactivation slowed down with the duration of the pre-pulse (Figure 1F). Tail current increase and slowing of deactivation saturated at a pulse duration of about 200 ms and 400–500 ms, respectively.

We then analyzed voltage dependence of human K_v12.1 channels with voltage protocols established previously to study mode shift of related K_v11.1 (Tan et al., 2012). We applied depolarizing holding potentials (conditioning potentials; 200 ms) before a series of activating voltage steps (pulse potentials from -140 mV to +10 mV; 600 ms) (Figure 1G and Supplementary Figures 1A–C). To minimize time intervals at hyperpolarized potentials that may counteract mode shift (c.f. Villalba-Galea, 2017), we at start recorded tail currents at correspondingly depolarized potentials (Figures 1G–I). In these experiments, amplitudes of K_v12.1-mediated outward currents were similar irrespective of conditioning potentials. This indicated that comparable steady-state channel activation was reached with all voltage protocols (Supplementary Figures 1A–C). After a conditioning potential of -60 mV, voltage dependence of K_v12.1 channels also did not change relevantly upon extension of the activating steps to 4000 ms (Figures 1J,K). This additionally demonstrated that steady-state activation of K_v12.1 channels was already reached by activating pulses as short as 600 ms. For 600 ms activating pulses, half-maximal voltages of activation (V_h) were -29.3 ± 2.3 mV and -24.1 ± 1.6 mV for negative conditioning potentials of -120 mV or -60 mV, respectively (n = 7; 600 ms activating pulses; Figures 1H,I). When cells were held at depolarized conditioning pulses of 0 mV or +40 mV, V_h was -85.3 ± 0.9 mV and -87.6 ± 0.8 mV, respectively (Figures 1H,I; 600 ms activating pulses). In these experiments, slope factors derived from Boltzmann fits to the recordings changed from -15.7 ± 0.7 mV (conditioning pulse of -120 mV) and -16.4 ± 0.9 mV (-60 mV) to -8.6 ± 0.1 mV (0 mV) and -8.9 ± 0.6 (+40 mV) (n = 7; Supplementary Figure 1D; 600 ms activating pulses). Accordingly, depolarizing conditioning potentials induced a large shift of voltage dependence by about -60 mV, and the full shift occurred across a potential range of 60 mV (between holding potentials of -60 and 0 mV). In analogous experiments, the same conditioning voltages shifted V_h of related K_v11.1 channels from -6.8 ± 4.4 mV (conditioning voltage -60 mV) to -62.4 ± 1.5 (conditioning voltage +40 mV; n = 7; Figures 2A,B), consistent with a previous report (Tan et al., 2012).

We next tested whether mode shift was sensitive to the employed voltage protocol. Therefore, we again applied 200 ms depolarized conditioning pulses, but this time we extended the activating pulses to 4000 ms (Figures 1J,K). When in these experiments cells were held at conditioning pulses of -60 and 0 mV, V_h was -43.3 ± 2.5 mV and -66.8 ± 1.5 mV, respectively (Figures 1J,K). Thus, mode shift of K_v12.1 channels was readily induced also when activating the channels for 4000 ms. However, the extent of mode shift was significantly reduced in these experiments compared to experiments with 600 ms activating pulses (Figure 1K; P ≥ 0.01), i.e., increased time intervals at hyperpolarized holding potentials during these protocols presumably counteracted development of mode shift. Similarly, the extent of mode shift was reduced when we activated channels for 600 ms after conditioning pulses of -60 and 0 mV, but recorded tail currents at hyperpolarized holding potentials (-120 mV; Supplementary Figures 1E,F), i.e., we introduced additional hyperpolarizing potentials after every activating pulse. Thus, also in these experiments hyperpolarizing holding potentials between the conditioning pulses reduced the expression of mode shift.

Taken together, human K_v12.1 channels exhibited significant voltage-dependent mode shift that in response to depolarized holding potentials manifested by slowed channel deactivation and by a large hyperpolarizing shift of voltage dependence. This mode shift of K_v12.1 channels can be induced robustly employing different voltage protocols, but the extent of mode shift significantly varies with duration of hyperpolarized holding potentials.

Inhibition of K_v11 channels by extracellular Na⁺ is well established (Numaguchi et al., 2000; Sturm et al., 2005) and a hallmark used to identify K_v11-mediated currents in neurons (e.g., Hardman and Forsythe, 2009). In control experiments, replacement of extracellular Na⁺ with NMDG without altering extracellular K⁺ concentration slightly increased K_v11.1-mediated outward currents (Figure 2A), as previously reported (Numaguchi et al., 2000). Upon removal of extracellular Na⁺, activation voltage range of K_v11.1 channels conditioned at -60 and +40 mV shifted to hyperpolarized potentials by -24.4 ± 1.5 mV and -13.9 ± 0.7 mV, respectively (n = 7; Figures 2B,C). Accordingly, overall mode shift of voltage dependence was attenuated from -55.7 ± 4.4 mV under control conditions to -45.1 ± 3.5 mV in absence of extracellular Na⁺ (n = 7; P ≤ 0.05). We then tested whether related K_v12.1 channels also exhibited such Na⁺ sensitivity. We found that neither current amplitudes nor voltage dependence of K_v12.1 channels were affected by removal of extracellular Na⁺ (Figures 2D–F). Consequently, the extent of mode shift was also insensitive to changes of the Na⁺ concentration. To conclude, despite the high similarity in mode shift behavior, the absence of Na⁺ sensitivity in K_v12.1 channels distinguishes K_v11.1-mediated currents from K_v12.1.

We then evaluated whether K_v12.1 channels were sensitive to channel inhibitors that are widely used to attribute neuronal K⁺ currents to particular channel families. At concentrations generally applied to inhibit established target channels, human K_v12.1 channels were insensitive to both E-4031 (20 μM), a specific inhibitor of K_v11 channels (Supplementary Figures 2A,B) (Trudeau et al., 1995), and XE991 (10 μM), a specific antagonist of neuronal K_v7 (KCNQ) channels (Supplementary Figures 2D,E) (Wang et al., 1998). K_v12.1-mediated currents were also insensitive to E-4031 and XE991 concentrations up to 100 μM (Supplementary Figures 2C,F). K_v12.1 channels also were not affected by the broad-spectrum K⁺ channel inhibitor tetraethylammonium (TEA) at a concentration that completely inhibited K_v7.2 channels (5 mM; Supplementary Figures 2G,H) (Hadley et al., 2000). However, K_v12.1 channels were slightly inhibited by 50 and 100 mM TEA (I_{50mM TEA}/I_Start = 0.90 ± 0.01; I_{100mM TEA}/I_Start = 0.83 ± 0.01; n = 9; Supplementary Figure 2I). Of note, low sensitivity of K_v12.1 channels to E-4031 and TEA has been shown in an early report (Shi et al., 1998). In summary, E-4031, XE991 and TEA cannot be used to inhibit K_v12.1 channels in native tissue, but may be used to isolate K_v12.1 channel activity through inhibition of other K⁺ channels.

We then tested whether human K_v12.1 channels were sensitive to 4-AP, another established antagonist of several K_v families (Gutman et al., 2005). Surprisingly, 4-AP at a concentration well in the range often used to isolate neuronal K⁺ currents (3 mM; e.g., Marcotti et al., 2003) increased K_v12.1-mediated steady-state and tail currents within seconds (Figures 3A–C). Current potentiation was the same for conditioning pulses of -60 and 0 mV, and 4-AP potentiated currents at all holding potentials positive to -60 mV (Figure 3C). 4-AP-dependent current increase was reversible within 2–3 min after washout of the drug (Figure 3D). 4-AP (3 mM) not only potentiated K_v12.1-mediated currents, but also significantly shifted the voltage dependence of K_v12.1 channels to hyperpolarized potentials (Figure 3E). Compared to control recordings measured before application of the substance, 3 mM 4-AP shifted V_h by -19.7 ± 1.7 mV (n = 7; P ≤ 0.001) and by -10.4 ± 1.5 mV (n = 7; P ≤ 0.001) after condition voltage pulses of -60 and 0 mV, respectively. Thus, the shift of V_h was significantly more pronounced for currents conditioned at -60 mV (P ≤ 0.01; c.f. Figure 3E).

We then analyzed the dose-response relationship of 4-AP action on K_v12.1 channels. As 5 and 10 mM 4-AP increased pH of the solution to about 8.1 and 9.0, respectively, we adjusted pH of these solutions to 7.4 after addition of 4-AP. Of note, 3 mM 4-AP or lower 4-AP concentrations did not alter the pH of the solution relevantly and thus no adjustment of pH was necessary. Under these experimental conditions, 4-AP potentiated K_v12.1 currents in a concentration-dependent manner with an EC₅₀ of about 2.1 and a Hill coefficient of about 0.8 (Figure 3F, red trace). In contrast, K_v11.1 channels were inhibited by 4-AP with an IC₅₀ of approximately 2.6 mM and a Hill coefficient of about 0.7 (Supplementary Figure 3), consistent with previous reports (e.g., Ridley et al., 2003). Thus, despite inverse effects of 4-AP on K_v12.1 and K_v11.1, the sensitivity of both channels to 4-AP was quite similar. When we applied 5 mM or 10 mM 4-AP without adjusting pH, we found that these concentrations activated K_v12.1 channel even stronger than at physiological pH (Figures 3F,G). Without adjusting pH at higher concentrations, the EC₅₀ was about 3.1 and the Hill coefficient was 0.9 (Figure 3F, black trace). As in line with a previous study (Kazmierczak et al., 2013) increasing pH of the extracellular solution from 7.4 to 8.1 or 9.0 without addition of 4-AP did not potentiate K_v12.1-mediated steady-state currents (Figure 3G), these data suggested that 4-AP activated K_v12.1 channels more efficiently at more alkaline pH. Indeed, increasing pH of the extracellular solution containing 3 mM 4-AP to 8.1 further increased K_v12.1-mediated steady-state outward currents (Figure 3H).

Several other aminopyridines have been shown to inhibit voltage-gated K⁺ channels, albeit antagonistic efficiency of these substances was lower than that of 4-AP (Robertson and Nelson, 1994; Sedehizadeh et al., 2012; Strupp et al., 2017). We thus wondered whether we could identify isomeric aminopyridines that activated K_v12.1, at best without affecting other K⁺ channels. We found that 2-aminopyridine (2-AP) and 3-aminopyridine (3-AP) potentiated K_v12.1-mediated currents at a concentration close to the EC₅₀ of 4-AP (3 mM) by about 20% (P ≤ 0.001) and 7% (P ≤ 0.001), respectively (Figure 4). At the same concentration 3,4-diaminopyridine (3,4-DAP; 3 mM) was ineffective. Hence, 2-AP (P ≤ 0.01) and 3-AP (P ≤ 0.001) activated K_v12.1 channels significantly less than 4-AP mirroring efficacy of inhibition of other K⁺ channels by these substances (Robertson and Nelson, 1994; Sedehizadeh et al., 2012; Strupp et al., 2017).

We then turned to NS1643, a partial agonist of the K_v11 channel family (Casis et al., 2006). As shown earlier (c.f. Hansen et al., 2006), NS1643 (30 μM) slowed channel deactivation and potentiated K_v11.1-mediated outward currents (Figures 5A–C). In contrast, the same concentration of NS1643 (30 μM) completely inhibited K_v12.1 channels with a time constant of 16.8 ± 2.6 s (n = 7; Figures 5A–C). When we applied only 10 μM NS1643, a concentration close to the reported EC₅₀ of NS1643 for activation of K_v11.1 channels (Casis et al., 2006), K_v12.1-mediated currents were reduced to 37.1 ± 4.1% of initial current amplitudes. Inhibition of currents by 10 μM was much slower than when 30 μM NS1643 was applied (n = 6; compare Figure 5B and Supplementary Figure 4). We then analyzed voltage dependence of residual K_v12.1 currents in the presence of 10 μM NS1643 (Figures 5D–F): Application of NS1643 (10 μM) shifted V_h by +8.8 ± 2.9 mV (n = 6; P ≤ 0.05) and by +21.7 ± 5.2 mV (n = 6; P ≤ 0.01) after condition voltage pulses of -60 and 0 mV, respectively (Figures 5D–F). At the same time, slope factors changed by +8.0 ± 1.5 mV (conditioning pulse of -60 mV; n = 6; P ≤ 0.05) and -1.8 ± 0.9 mV (conditioning pulse 0 mV; n = 6; P ≤ 0.01). Furthermore, NS1643 (10 μM) significantly accelerated deactivation of K_v12.1 channels at hyperpolarized potentials (Figures 5G,H; P ≤ 0.05; n = 6).

Our results suggested that voltage-clamp protocols designed to detect mode shift in combination with pharmacology using 4-AP or NS1643 should provide a robust approach for isolation of K_v12.1-mediated currents in native cell types. As proof of principle, we sought to isolate K_v12.1 channel activity in cells expressing different K⁺ channels (Figure 6). To this end, we co-expressed K_v12.1 channels together with K_v11.1 and typical neuronal K⁺ channels (K_v7.2, K_v7.3 and Kir2.1). In these experiments, CHO cells were transiently transfected with equal amounts of plasmid DNA encoding the channel subunits (see section “Materials and Methods” for details). We first analyzed whether we could isolate Kir2.1- and K_v7-mediated currents in those cells. In all cells tested, we found large inward currents at hyperpolarized potentials and XE991-sensitive outward currents at depolarized potentials demonstrating expression of functional Kir2.1 and K_v7 channels, respectively (Supplementary Figure 5). As measure for abundance of K_v11.1 and K_v12.1 channels, we then tested whether we could detect mode shift of voltage dependence in the mix of K⁺ currents. As determined from whole cell currents, V_h was -6.7 ± 1.3 mV and -80.0 ± 2.1 mV after conditioning potentials of -60 and 0 mV, respectively (n = 7; Figures 6A–C). Thus, mode shift of channels expressed in these cells (K_v11.1 and K_v12.1 channels) was readily detectable even among a complement of different voltage-dependent K⁺ channels. We next attempted to isolate K_v12.1-mediated currents among the mixture of K⁺ channel by making use of the pharmacological profile established above. 4-AP (3 mM) shifted the voltage dependence of whole cell currents by -8.5 ± 0.9 mV (n = 7; P ≤ 0.001) and by -12.3 ± 2.3 mV (n = 7; P ≤ 0.05) to hyperpolarized potentials after conditioning voltages of -60 and 0 mV, respectively (Figures 6B,C). Thus, although being less pronounced than for K_v12.1 channels alone (Figure 6C), the 4-AP-induced shift of voltage dependence was readily detectable in the mixed K⁺ current situation. At the same time, 4-AP (3 mM) potentiated whole cell currents elicited at -60 and 0 mV to about 210% and 140% of baseline amplitudes, respectively (Figures 6A,D–F). Finally, we applied 30 μM NS1643 (on top of 4-AP) to the same cells and found that the substance inhibited outward currents and inward tail currents (Figures 6A,D,E). NS1643 also slowed deactivation kinetics of remaining currents (c.f. Figure 6A) indicating that NS1643-sensitive and functional K_v11.1 channels were expressed in these cells.

Taken together, using abovementioned experimental protocols we could unequivocally identify properties of K_v12.1 channels (mode shift, 4-AP and NS1643 sensitivity) among a mixture of K⁺ currents. These results therefore suggest that the same protocols may be used to assign K⁺ current components to K_v12.1 channels in complex (native) cellular/neuronal settings.

Mode shift of activation was recently demonstrated for K_v12.1 channel isoforms of humans and zebrafish heterologously expressed in Xenopus laevis oocytes (Li et al., 2015; Dai and Zagotta, 2017). However, mode shift of these channels has not been characterized in mammalian cells in detail yet. We found that mode shift of human K_v12.1 was readily induced by depolarized holding potentials between -60 and 0 mV and that it manifested by slowed channel deactivation and a striking shift of voltage dependence to hyperpolarized potentials. Deceleration of deactivation is generally considered a biophysical hallmark of mode shift, as channels undergo additional (time-consuming) transitions from this “relaxed” (metastable) open state into deactivation (Bezanilla et al., 1982; Villalba-Galea et al., 2008; Corbin-Leftwich et al., 2016; Villalba-Galea, 2017). Similar to K_v11.1 (Tan et al., 2012) and zebrafish K_v12.1 (Dai and Zagotta, 2017), mode shift of human K_v12.1 channels manifested by a striking -60 mV shift of voltage dependence. Thus, mode shift-dependent hyperpolarizing shifts of voltage dependence were qualitatively and quantitatively comparable for these three ion channels. Of note, mode shift of K_v12.1 channels was readily induced using different voltage protocols, albeit the extent of the hyperpolarizing shift of activation potentials varied considerably with duration of hyperpolarized voltage steps. This demonstrated high sensitivity of K_v12.1 channels to the holding potential, but also highlighted that voltage dependence of native K_v12.1 currents might also vary with the used voltage protocols. In contrast to zebrafish K_v12.1 (Dai and Zagotta, 2017), activation of human K_v12.1 channels did not exhibit prominent double exponential kinetics that might indicate transition into a more stable open conformation. However, human K_v11.1 channels that beyond doubt exhibit mode shift do not display such kinetics neither (Tan et al., 2012; Goodchild et al., 2015). This indicates that either kinetics were masked by channel inactivation, or transition was too fast in the human channel isoforms.

Time course of transition into the relaxed state varies considerably between channels requiring depolarization for minutes in Na_v (Bezanilla et al., 1982), seconds in Shaker (Olcese et al., 1997) and some hundreds of milliseconds in K_v11.1 channels (Piper et al., 2003). For human K_v12.1, we found that also some hundreds of milliseconds of conditioning depolarization was sufficient for significant alterations of voltage dependence and kinetics. Thus, time course of development of K_v12.1 mode shift was well in the range of that published for K_v11.1 channels (Piper et al., 2003). As K_v12.1 channels are highly sensitive to changes of the holding potential, voltage dependence of the channels strongly depends on the employed voltage protocols. In fact, such protocol differences could account for considerable variations in voltage dependence of K_v12.1 channels reported in different studies (V_h close to 0 mV in Shi et al., 1998;V_h of about -60 mV in Zou et al., 2003).

4-AP is a rather selective blocker of voltage-gated K⁺ channels: At micromolar concentrations, it reversibly inhibits activity of K_v1 and K_v3 family members, but at higher concentrations (in the millimolar range) 4-AP also blocks other K_v channels (e.g., K_v11) (Gutman et al., 2005; Alexander et al., 2015). Thus, it was somewhat surprising to find that 4-AP activates K_v12.1 channels. As an early study mentioned potentiation of K_v12.3 through 4-AP without, however, showing any recordings (named rELK1; Engeland et al., 1998), such 4-AP sensitivity may constitute a general feature of the K_v12 family.

Earlier, it was shown that currents through K_v2.1 channels that are normally inhibited by 4-AP were potentiated by the substance, but only when K_v6.4 subunits were co-expressed (Stas et al., 2015). As K_v2.1 and K_v6.4 co-assemble into functional channels (reviewed in Bocksteins, 2016), this suggested that K_v6.4 largely determined altered 4-AP sensitivity of the resulting heteromeric channels. 4-AP suppressed closed state inactivation of K_v2.1/K_v6.4 resulting in exclusive current potentiation of currents through those heteromers (Stas et al., 2015). In contrast, K_v12.1 channels did not inactivate in our experiments, and thus 4-AP probably does not potentiate K_v12.1-mediated currents through a similar mechanism. As 4-AP, in contrast to K_v2.1/K_v6.4 heteromers, also modulated voltage dependence of K_v12.1, actions of 4-AP are probably even more complex for these channels. Interestingly, a recent study demonstrated 4-AP-dependent activation of K_v7.4 channels (Khammy et al., 2017) indicating that 4-AP-dependent activation of voltage-gated K⁺ channels may constitute a more frequent phenomenon than expected. However, further work is needed to elucidate whether other members of the K_v channel superfamily also exhibit this special 4-AP sensitivity.

Yet, we do not know whether 4-AP directly activates K_v12.1 channels or whether an auxiliary subunit endogenously expressed in CHO cells confers 4-AP activation to K_v12.1. Unfortunately, at present nothing is known about physiologically relevant interaction partners of K_v12.1 channels. As K_v channels (with K_v2.1 as exception) normally do not form functional channels with members of other K_v families, heteromerization of K_v12.1 channels with another pore forming α subunit apart from K_v12.2 or K_v12.3 is quite unlikely (Zou et al., 2003). Accordingly, a mechanism as shown for K_v2.1 is rather implausible, and it is hard to imagine how a non-pore-forming auxiliary subunit could reverse 4-AP sensitivity from inhibition to activation. Furthermore, any endogenously expressed auxiliary subunit would need to be expressed at high abundance to saturate overexpressed K_v12.1 channels. Hence, a straightforward model proposes direct activation of K_v12.1 channels through 4-AP. However, we want to point-out that we cannot exclude indirect actions on the channels at the moment.

NS1643 is a well-characterized partial agonist of K_v11 channels that slows deactivation, increases tail-current amplitude, and shifts voltage dependence of activation to hyperpolarized potentials and voltage dependence of C-type inactivation to depolarized potentials (Casis et al., 2006; Hansen et al., 2006). At the same time, NS1643 exhibits weak antagonistic effects on K_v11 channels as evident by an attenuation of drug-induced current increase at higher concentrations (Casis et al., 2006; Schuster et al., 2011). K_v11.3 channels are even inhibited by higher concentrations of NS1643. For K_v12.1 channels, NS1643 accelerated deactivation, inhibited outward and tail currents and shifted voltage dependence of activation to depolarized potentials (c.f. Figure 5). Thus, NS1643 affected K_v11 and K_v12.1 channels exactly in opposite directions. Similar to K_v11.3 channels, NS1643 increased slope factor of voltage dependence of K_v12.1 channels, even though already at lower concentrations. This suggests that NS1643 similarly affects K_v11 and K_v12.1 channels, but antagonistic effects might dominate over activation for K_v12.1. In line, the concentration range of NS1643 effects was similar for these channel isoforms (Casis et al., 2006). However, so far we cannot tell whether NS1643 also exhibits weak agonistic effects on K_v12.1 channels or whether NS1643 binds to homologous regions in K_v11 and K_v12.1 channels.

As pharmacological tools and appropriate mouse models are currently missing, identification of native K_v12.1-mediated currents critically depends on identification of unique biophysical and pharmacological properties. Indeed, native currents were successfully attributed to related K_v10 and K_v11 channels by exploiting their unique activation kinetics (c.f. Cole-Moore shift; e.g., Meyer and Heinemann, 1998) or exclusive Na⁺ sensitivity and pharmacology (Hirdes et al., 2005, 2009; Hardman and Forsythe, 2009), respectively. Here, we present distinctive pharmacological properties, and straightforward experimental protocols that may be employed to isolate K_v12.1 channel activity in native tissue. As mode shift readily manifested in cells expressing various neuronal K⁺ currents, this experimental protocol may be used to demonstrate expression of channels with mode shift in native cell types. In neurons expressing various K⁺ current components, such changes of voltage dependence may be easier to detect than associated changes of deactivation kinetics. Such experiments may not provide definite proof for K_v12 channel expression. However, expression of mode shift may be employed together with expression analyses, Na⁺ sensitivity, activation through 4-AP and inhibition by NS1643 to narrow down (or exclude) contribution of K_v12.1 channels to whole cell currents. Identification of the combination of these properties would provide strong evidence for expression and thus potential physiological relevance of K_v12.1 channels.

Chinese hamster ovary (CHO) dhFR^- cells were maintained as previously reported (Leitner et al., 2016). Cells were kept in MEM Alpha Medium supplemented with 10% fetal calf serum (FCS) and 1% penicillin/streptomycin (pen/strep) (all Invitrogen GmbH, Darmstadt, Germany) in a humidified atmosphere at 5% CO₂ and 37°C. Transient transfection of cells was done with jetPEI transfection reagent (Polyplus Transfection, Illkirch, France). All experiments were performed 24–48 h after transfection at room temperature (22–25°C). The expression vectors used were: K_v7.2-pBK-CMV (gene: human KCNQ2; UniProt accession number: O43526), K_v7.3-pBK-CMV (human KCNQ3; O43525), K_v7.4-pBK-CMV (human KCNQ4; P56696) K_v11.1 (Erg1)-pcDNA3.1 (rat KCNH2; O08962), K_v12.1(Elk1)-pcDNA3.1-IRESeGFP (human KCNH8; Q96L42), Kir2.1-pBK-CMV (human KCNJ2; P63252), and pEGFP-C1 (Addgene, Teddington, United Kingdom). For recordings presented in Figure 6 and Supplementary Figure 5, CHO cells were transiently transfected with identical amounts of plasmids encoding K_v7.2, K_v7.3, K_v11.1, K_v12.1, and Kir2.1 (0.6 μg of plasmid DNA for every subunit).

Electrophysiological recordings were performed in the whole cell configuration with an Axopatch 200B amplifier (Molecular Devices, Union City, CA, United States) or an HEKA EPC10 USB patch clamp amplifier HEKA (Elektronik, Lambrecht, Germany) in voltage-clamp mode (Leitner et al., 2011). All recordings were low-pass filtered at 2 kHz and sampled at 5 kHz. Currents were elicited by voltage protocols indicated in the figures. Dashed lines in representative recordings highlight zero current. Borosilicate glass patch pipettes (Sutter Instrument Company, Novato, CA, United States) were used with an open pipette resistance of 2–3 MΩ after back-filling with intracellular solution. Liquid junction potentials were not compensated (approximately -4 mV). Series resistance (R_s) typically was below 6 MΩ and compensated throughout the recordings (80–90%). Whole cell currents are presented normalized to the cell capacitance (current density; pA/pF) or as normalized to baseline current amplitude (I/I_Start). Extracellular solutions for most experiments contained (mM) 144 NaCl, 5.8 KCl, 1.3 CaCl₂, 0.9 MgCl₂, 0.7 NaH₂PO₄, 10 HEPES and 5.6 D-glucose, pH 7.4 (with NaOH), 305–310 mOsm/kg. In some experiments, NaCl in the extracellular solution was substituted by N-methyl-D-glucamine (NMDG; Sigma–Aldrich) (c.f. Figure 2). The standard intracellular solution contained (mM) 135 KCl, 2.41 CaCl₂ (100 nM free Ca²⁺), 3.5 MgCl₂, 5 HEPES, 5 EGTA, 2.5 Na₂ATP, pH 7.3 (with KOH), 290–295 mOsm/kg (Wilke et al., 2014).

Tetraethylammonium (TEA, Sigma), NMDG (Sigma), 4-aminopyridine (≥99%; Sigma and Tocris Bioscience, Bristol, United Kingdom), 2-aminopyridine (2-AP; Sigma), 3-aminopyridine (3-AP; Sigma), 3,4-diaminopyridine (3,4-DAP; Sigma), 1,3-Bis-(2-hydroxy-5-trifluoromethyl-phenyl)-urea (NS 1643, Tocris), 10,10-bis(4-Pyridinylmethyl)-9(10H)-anthracenone dihydrochloride (XE991, Tocris), and N-[4-[[1- [2-(6-Methyl-2-pyridinyl)ethyl]-4-piperidinyl]carbonyl]phenyl] methanesulfonamide dihydrochloride (E-4031, Tocris) were diluted in extracellular solution to concentrations indicated in “Results.” All substances were applied locally via a glass capillary through a custom-made application system.

4-AP did not significantly change the pH of the extracellular solution at concentrations below 3 mM. After dilution of the substance, pH of the extracellular solution containing 5 mM or 10 mM 4-AP typically was about 8.1 or 9.0, respectively. As indicated in “Results,” in some experiments the pH of solutions containing 5 and 10 mM 4-AP was adjusted to 7.4 after dilution of 4-AP. At the concentration applied in the present study (3 mM), isomeric aminopyridines did not change pH of the extracellular solution (c.f. Figure 4).

Patch clamp recordings were analyzed with PatchMaster (HEKA) and IgorPro (Wavemetrics, Lake Oswego, OR, United States). Voltage dependence of activation was derived from tail current amplitudes using voltage protocols indicated: Tail currents were fitted with a two-state Boltzmann function with I = I_min + (I_max - I_min)/(1 + exp((V -V_h)/s)), where I is current, V is the membrane voltage, V_h is the voltage at half maximal activation, and s describes the steepness of the curve. Results are shown as conductance-voltage curves, obtained by normalizing to (I_max -I_min), obtained from fits to data of individual experiments. Time constants of deactivation were derived from double-exponential fits to deactivating current components at indicated potentials. For dose-response relations, current potentiation at 0 mV (normalized to baseline) was fitted to a Hill equation with I/I_b = I_b + (I_max-I_b)/(1 + (EC₅₀/[S])^n_H), where I is the (normalized) current, I_b and I_max denote minimal and maximal currents at low and high drug concentrations, EC₅₀ is the concentration at the half maximal effect, [S] is the drug concentration and n_H is the Hill coefficient (Leitner et al., 2012).

Isolated cells under investigation were randomly assigned to different treatment groups. Data recordings and analysis for experiments presented were not performed in a blinded manner. For some experiments, single recordings were normalized to base line values individually to account for baseline variations between cells. Statistical analysis was performed using two-tailed Student’s t-test/Wilcoxon–Mann–Whitney test, and when appropriate comparisons between multiple groups were performed with ANOVA followed by Dunnett test. Significance was assigned at P ≤ 0.05 (^∗P ≤ 0.05, ^∗∗P ≤ 0.01, ^∗∗∗P ≤ 0.001). Data subjected to statistical analysis have n over 5 per group and data are presented as mean ± SEM. In electrophysiological experiments, n represents the number of individual cells and accordingly the number of independent experiments (no pseudo-replication).