A comparison of currents carried by HERG, with and without coexpression of MiRP1, and the native rapid delayed rectifier current. Is MiRP1 the missing link?

Abstract

Although it has been suggested that coexpression of minK related peptide (MiRP1) is required for reconstitution of native rapid delayed-rectifier current (I_Kr) by human ether-a-go-go related gene (HERG), currents resulting from HERG (I_HERG) and HERG plus MiRP1 expression have not been directly compared with native I_Kr. We compared the pharmacological and selected biophysical properties of I_HERG with and without MiRP1 coexpression in Chinese hamster ovary (CHO) cells with those of guinea-pig I_Kr under comparable conditions. Comparisons were also made with HERG expressed in Xenopus oocytes. MiRP1 coexpression significantly accelerated I_HERG deactivation at potentials negative to the reversal potential, but did not affect more physiologically relevant deactivation of outward I_HERG, which remained slower than that of I_Kr. MiRP1 shifted I_HERG activation voltage dependence in the hyperpolarizing direction, whereas I_Kr activated at voltages more positive than I_HERG. There were major discrepancies between the sensitivity to quinidine, E-4031 and dofetilide of I_HERG in Xenopus oocytes compared to I_Kr, which were not substantially affected by coexpression with MiRP1. On the other hand, the pharmacological sensitivity of I_HERG in CHO cells was indistinguishable from that of I_Kr and was unaffected by MiRP1 coexpression. We conclude that the properties of I_HERG in CHO cells are similar in many ways to those of native I_Kr under the same recording conditions, and that the discrepancies that remain are not reduced by coexpression with MiRP1. These results suggest that the physiological role of MiRP1 may not be to act as an essential consituent of the HERG channel complex carrying native I_Kr.

The delayed rectifier current (I_K) plays a critical role in cardiac repolarization (Zeng et al. 1995). Abnormalities in channel subunits encoding the rapid (I_Kr) and slow (I_Ks) components of I_K underlie the most common forms of the congenital long-QT syndrome. I_Ks is formed by the co-assembly of the LQT1 α-subunit and the minK β-subunit (Sanguinetti et al. 1996; Barhanin et al. 1996). Currents carried by homomeric KvLQT1 channels are small and have much more rapid activation kinetics than native I_Ks (Sanguinetti et al. 1996; Barhanin et al. 1996). MinK expression by itself does not result in macroscopic currents in the absence of KvLQT1, but in the presence of KvLQT1 produces robust currents that are typical of native I_Ks. The human ether-à-go-go related gene (HERG) encodes K⁺-channel subunits that carry currents (I_HERG) resembling native I_Kr; however, discrepancies have been noted between I_HERG and native I_Kr (Sanguinetti et al. 1995). Recently, it has been suggested that coexpression of HERG with a MinK-related peptide (MiRP1) reconstitutes native I_Kr (Abbott et al. 1999). In particular, MiRP1 coexpression accelerated the E-4031 pulse-dependent block of I_HERG in Xenopus oocytes and increased the blocking potency, moving both responses towards those reported for native cardiomyocytes. The role of MiRP1 in I_Kr was considered to be analogous with that of minK in I_Ks - just as minK reconstitutes I_Ks upon coexpression with the α-subunit KvLQT1, MiRP1 was considered to reconstitute I_Kr upon coexpression with HERG (Abbott et al. 1999).

A variety of properties of currents in Xenopus oocytes make their response to blocking drugs difficult to compare directly with effects on native currents. The vitelline membrane and viscous yolk of oocytes act as sinks for drugs, slowing their action and reducing their potency. The conditions used to record currents in Xenopus oocytes are different from those for voltage-clamp studies of native currents. The present study was designed to compare I_HERG in oocytes and CHO cells (with and without MiRP1 coexpression) with I_Kr in native cardiomyocytes. We were particularly interested in comparing the properties of I_Kr-blocking drug actions among HERG and HERG/MiRP1 expressed in CHO cells and native I_Kr using comparable study conditions, since pharmacological observations were a major part of the evidence used to argue that MiRP1 coexpression with HERG reconstitutes I_Kr.

METHODS

Oocyte isolation and cRNA injection

All animal handling procedures were approved by the Animal Research Ethics Committee of the Montreal Heart Institute and conformed to the guidelines of the Canadian Council on Animal Care. Female Xenopus laevis were anaesthetized in 0.13 % w/v tricaine (Sigma Chemicals, St Louis, MO, USA) for 30 min at 4 °C. Segments of the ovarian lobe were removed through a small abdominal incision. Up to four collections were made from each frog with adequate time allowed for healing between each. After the final collection, frogs were killed by exsanguination following a lethal overdose of the anaesthetic. The follicular layer was removed by digestion with 2 U ml⁻¹ collagenase type V (Sigma) in Ca²⁺-free Barth's solution (mmol l⁻¹: NaCl, 88; KCl, 1; NaHCO₃, 2.4; MgSO₄, 0.82; Hepes, 5; pH 7.6; 10 mg ml⁻¹ penicillin- streptomycin solution). The oocytes were incubated at 17 °C in L-15 medium (50 % v/v Leibovitz L-15 medium, 0.4 g l⁻¹ glutamine, 8 mmol l⁻¹ Hepes, 40 mg l⁻¹ gentamycin, pH 7.6). For in vitro transcription, HERG cDNA subcloned into pSP64 plasmid vector was linearized with EcoR1 (New England BioLabs, Mississauga, ON, Canada) and transcribed with SP6 RNA-polymerase (Ambion Inc., Austin, TX, USA) for 1.5–2 h at 37 °C. Human MinK-related peptide (hMiRP1) cDNA subcloned into pCI-neo vector was linearized with Not 1 and transcribed with T7 RNA-polymerase for 1 h at 37 °C. Stage IV and V oocytes were injected with 25–50 nl of HERG cRNA (2–4 ng per oocyte) alone or in combination with hMiRP1 cRNA (at ∼4:1 MiRP1:HERG molar ratio) 24 h after isolation.

Expression of HERG and MiRP1 in CHO cells

HERG was stably transfected into a CHO-K1 cell line with the use of Lipofectamine-Plus and selected with 600 μg ml⁻¹ G418 (Life Technologies). Cells stably expressing HERG were transiently transfected with 2 μg MiRP1 cDNA (subcloned into pCI-neo vector) and 0.4 μg green fluorescent protein cDNA (GFP, a 5:1 ratio of MiRP1 to GFP) 36 − 48 h prior to recording. GFP-positive cells were selected for recording of HERG/MiRP1 currents. Cells transfected with only GFP and the vehicle were used to record HERG currents alone.

Cardiomyocyte isolation

Male guinea-pigs were killed by blunt trauma to the head and the hearts were rapidly excised and mounted on a Langendorff apparatus. The hearts were perfused for 3–5 min with Tyrode solution containing (mmol l⁻¹): 140 NaCl, 5.4 KCl, 1.8 CaCl₂, 1 MgCl₂, 0.34 NaH₂PO₄, 10 glucose and 10 Hepes (pH adjusted to 7.4 with NaOH). The perfusion was then switched to nominally Ca²⁺-free Tyrode solution for 5 min, followed by 20–30 min perfusion with 50 μmol l⁻¹ CaCl₂ Tyrode solution containing 0.4–0.5 mg ml⁻¹ collagenase (Type II, Worthington Biochemical Corp., Lakewood, NJ, USA) and 0.2 mg ml⁻¹ protease (Type XIV, Sigma). Ventricular free walls were removed and gently agitated in low-Ca²⁺ solution. Harvested cardiomyocytes were maintained at room temperature in 200 μmol l⁻¹ CaCl₂ Tyrode solution. I_Kr was recorded from rod-shaped left or right ventricular cardiomyocytes.

Electrophysiology and data analysis

Currents were recorded from oocytes with the two-electrode voltage-clamp technique 1–2 days after injection of cRNA. Voltage command pulses were delivered via a GeneClamp 500 amplifier with pCLAMP 6 software (Axon Instruments Inc., Foster City, CA, USA). Currents were recorded at room temperature in ND96 (mmol l⁻¹: NaCl, 96; KCl, 2; CaCl₂, 1.8; MgCl_2, 1; Hepes, 5; pH adjusted to 7.5 with NaOH). Glass microelectrodes (borosilicate with filament) were pulled (Flaming/ Brown micropipette puller) to resistances of 1–3 MΩ (voltage-sensing electrode) and 0.1–0.5 MΩ (current-injecting electrode) when filled with 3 mol l⁻¹ KCl. The tips of the current-injecting electrodes were back-filled with 1 % agarose in 3 mol l⁻¹ KCl to prevent KCl leakage.

All recordings in cardiomyocytes and CHO cells were performed at 34–36 °C (TC-324B Single channel heater controller, Warner Instrument Corp., Hamden, CT, USA) with an Axopatch 200B amplifier and pClamp-7 software (Axon Instruments). The external solution contained (mmol l⁻¹): 145 NaCl, 5 KCl, 1.8 CaCl₂, 1 MgCl₂ and 10 Hepes (pH adjusted to 7.4 with NaOH). Patch pipettes (2–2.5 MΩ) were filled with (mmol l⁻¹): 140 KCl, 1 MgCl₂, 5 K₂ATP, 5 EGTA and 10 Hepes (pH adjusted to 7.2 with KOH). All junction potentials were zeroed prior to formation of the seals. After obtaining whole-cell access, series resistance was compensated to minimize the duration of the capacitive transient. Capacitive transients were recorded by applying 10 mV depolarizing steps from the holding potential V_H (V_H = −80 mV for CHO cells and 0 mV for cardiomyocytes). Membrane capacitances averaged 30.2 ± 2.3 pF (n = 19) and 223 ± 16 pF (n = 24) for CHO cells and cardiomyocytes respectively. Before compensation, series resistances (R_s) averaged 8.2 ± 1.3 MΩ and 10.3 ± 1.6 MΩ, and after compensation 5.1 ± 0.5 MΩ and 2.7 ± 0.2 MΩ for CHO cells and cardiomyocytes respectively. Maximum HERG step currents in CHO cells averaged 510 pA, whereas maximum tail currents averaged 690 pA, providing an average maximum voltage drop across the series resistance of 2.6 mV for HERG step current and 3.5 mV for tail current. In cardiomyocytes, maximal I_Kr amplitudes rarely exceeded 200 pA, and the maximum voltage error due to R_s was therefore less than 1 mV.

For I_Kr recordings in native cardiomyocytes, 50 μmol l⁻¹ chromanol 293B and 1 μmol l⁻¹ nimodipine were added to block I_Ks and I_Ca.L, respectively, and a V_H of −40 mV was used to ensure complete I_Na inactivation. In some experiments in CHO cells, 293B and nimodipine were also added to ensure comparability with cardiomyocyte recording conditions. All extracellular solutions were delivered via a fast perfusion system (Warner Instrument Corp.) that allowed for solution changes within 5–8 s.

Stock solutions of E-4031 (10 mmol l⁻¹, Eisai Pharmaceuticals, Ibaraki, Japan) and dofetilide (1 mmol l⁻¹, Pfizer Pharmaceuticals, Sandwich, UK) were prepared in distilled water and quinidine-Cl (10 mmol l⁻¹, Sigma) in 40 % ethanol. Chromanol 293B (100 mmol l⁻¹, Aventis Pharmaceuticals, Frankfurt, Germany) and nimodipine (10 mmol l⁻¹, Sigma) were dissolved in DMSO. DMSO concentrations in the external solutions did not exceed 0.06 % v/v. The vehicles for drug administration had no effect on I_Kr or I_HERG at maximum concentrations.

A previously developed protocol was used to evaluate kinetics of block onset and drug potency without repeated cycling through various channel states (Weerapura et al. 2002) A V_H of 0 mV was applied to oocytes or CHO cells expressing HERG or HERG/MiRP1, and very brief (25 ms) hyperpolarizing sampling pulses were applied to −110 mV to remove inactivation and reveal I_HERG inactivating current upon subsequent return to 0 mV. A similar protocol was applied in cardiomyocytes but the hyperpolarizing step of each sampling pulse was to −40 (rather than −110) mV. This voltage was selected because it was sufficiently positive to prevent recovery of I_Na from inactivation but allowed for I_Kr recovery from inactivation as shown previously for ferret atrial I_Kr (Liu et al. 1996). When direct pharmacological comparisons were conducted between I_Kr and HERG or HERG/MiRP1 currents, the sampling pulse hyperpolarizing step was changed to −40 mV in CHO cells (i.e. the same voltage protocol was used as for I_Kr in native myocytes). To determine the kinetics of block onset, sampling pulses were applied three times under drug-free conditions to ensure steady-state activation at 0 mV and at 15, 30, 45, 60, 90, 120, 180, 240 and 300 s after drug application. Upon exposure to an I_HERG or I_Kr blocker, the degree of block was identical with this protocol whether sampling pulses were applied to define the time course or whether only a single sampling pulse was applied at the end of the exposure period. For example, dofetilide (0.5 μmol l⁻¹) reduced I_HERG expressed in oocytes by 92 ± 2 % after 5 min of exposure with the repeated sampling pulse protocol, compared to 95 ± 1 % (n.s. versus repeated sampling pulses) when a single sampling pulse was used after the same exposure period. This finding indicates that intermittent sampling pulses themselves do not affect current inhibition. To determine the concentration dependence of drug block, sampling pulses were applied three times under drug-free conditions to ensure steady-state activation and after 5–7 min (for oocytes) or 3–4 min (for cardiomyocytes and CHO cells) perfusion with successively larger drug concentrations.

Clampfit (Axon Instruments) and/or Origin (Microcal Corp., Northampton, MA, USA) were used for data analysis. Non-linear curve-fitting was performed with algorithms in Origin. Data are presented as means ± s.e.m. and statistical comparisons were obtained with ANOVA or two-tailed t tests (where only two groups were included in the comparison).

RESULTS

Block of I_HERG in oocytes versus I_Kr block in cardiomyocytes

Prior to evaluating the sensitivity and kinetics of drug-induced block of I_HERG in Xenopus oocytes, we confirmed functional effects of coexpression by observing currents during and after 2 s depolarizing pulses from a V_H of −80 mV. Abbott et al. (1999) have shown accelerated deactivation of oocyte I_HERG inward current in the presence of MiRP1. Tail current deactivation kinetics at −80 mV were accelerated upon MiRP1 coexpression. Biexponential fits yielded τ_fast and τ_slow in nine oocytes of 211 ± 23 and 683 ± 41 ms, respectively, for I_HERG, significantly larger than in the presence of MiRP1 coexpression (123 ± 4 and 431 ± 16 ms, respectively; n = 7, P < 0.001vs. I_HERG alone for each).

Figure 1A and B shows, respectively, the concentration dependence and kinetics of I_HERG block in oocytes compared with results for native I_Kr determined using the sampling pulse protocol applied at a V_H of 0 mV as described in Methods. The IC₅₀ values for I_HERG block in oocytes were 16.8 ± 2.2 μmol l⁻¹ (n = 4 per concentration), 0.17 ± 0.03 μmol l⁻¹ (n = 4 per concentration) and 0.11 ± 0.01 μmol l⁻¹ (n = 6 per concentration) for quinidine, E-4031 and dofetilide, respectively. The IC₅₀ values upon MiRP1 coexpression were 20.1 ± 3.1 μmol l⁻¹ for quinidine (n = 4 per concentration), 0.14 ± 0.04 μmol l⁻¹ for E-4031 (n = 3 per concentration) and 0.09 ± 0.01 μmol l⁻¹ for dofetilide (n = 4 per concentration), i.e. were not significantly altered. In contrast, the IC₅₀ values for I_Kr block in cardiomyocytes were: 1.4 ± 0.2 μmol l⁻¹ (quinidine, n = 6 per concentration), 10.3 ± 0.3 nmol l⁻¹ (E-4031, n = 4 per concentration) and 8.7 ± 2.0 nmol l⁻¹ (dofetilide, n = 4 per concentration; P < 0.05versus oocytes for each agent).

Figure 1. Inhibition of native I_Krversus I_HERG with or without MiRP1 coexpression in Xenopus oocytes by quinidine, E-4031 and dofetilide.

A, sampling pulse protocol (left panel inset) described in Methods was used to evaluate the concentration-dependent current inhibition. Curves are best fit equations of the form I/I_max = 1/(1+[IC₅₀/C]ⁿ), where I/I_max represents the normalized current, IC₅₀ and n are constants and C represents the drug concentration. B, time-dependent inhibition of I_HERG with or without MiRP1 coexpression and I_Kr. The drug concentrations used to evaluate kinetics of current block were approximately 5 times the IC₅₀ for each drug reported in panel A in the respective systems: i.e. 100 μmol l⁻¹ quinidine, 1 μmol l⁻¹ E-4031 and 0.5 μmol l⁻¹ dofetilide for I_HERG with or without MiRP1 recorded from oocytes and 5 μmol l⁻¹ quinidine, 50 nmol l⁻¹ E-4031 and 50 nmol l⁻¹ dofetilide for native I_Kr. The normalized data were fitted to monoexponential functions to determine the time constants (τ) of block. All data are presented as means ± s.e.m.

The kinetics of block onset were evaluated at concentrations approximately 5 times the IC₅₀ for each drug in the respective system (Fig. 1B). The time constants for I_HERG block in Xenopus oocytes averaged 17.4 ± 1.6 s (n = 6), 130 ± 11 s (n = 5) and 114 ± 18 s (n = 5) for 100 μmol l⁻¹ quinidine, 1 μmol l⁻¹ E-4031 and 0.5 μmol l⁻¹ dofetilide, respectively. Blocking kinetics were not significantly altered by MiRP1 coexpression: 11.7 ± 2.0 s (n = 4), 112 ± 9 s (n = 4) and 131 ± 8 s (n = 5) for quinidine, E-4031 and dofetilide, respectively. Quinidine (5 μmol l⁻¹) blocked I_Kr with a time constant of 18.1 ± 2.0 s (n = 6), similar to that for I_HERG block in oocytes. For 50 nmol l⁻¹ E-4031 and 50 nmol l⁻¹ dofetilide, the I_Kr blocking time-constants averaged 32.2 ± 1.1 s (n = 4) and 36.6 ± 2.1 s (n = 5), respectively, i.e. were several times faster than the time constants for I_HERG or I_HERG/MiRP1 block in oocytes (P < 0.05 for each). These results confirm previous observations indicating important differences between the pharmacological responses of I_HERG in oocytes and those of I_Kr in cardiomyocytes (Sanguinetti et al. 1995; Abbott et al. 1999).

Voltage and time dependence of I_Krversus I_HERG in CHO cells with and without MiRP1 coexpression

Figure 2A shows I_HERG recordings from CHO cells in the absence and presence of MiRP1 coexpression, during 0.4 s hyperpolarizing pulses following 1 s depolarizing pulses to +40 mV. As shown by the continuous curves, deactivation was well fitted by a single exponential function. The mean deactivation time constants (Fig. 2B) indicate that MiRP1 significantly accelerated deactivation of inward current negative to the reversal potential, as occurs in oocytes. We exploited this difference to obtain a functional index of successful MiRP1 coexpression with HERG (in addition to visualization of GFP expression) in CHO cells. A deactivation time constant of less than 20 ms at −110 mV was taken as evidence of functional coexpression of MiRP1 in each cell used to study the properties of HERG/MiRP1. In contrast, deactivation time constants were always greater than 25 ms in stable HERG-expressing CHO cells co-transfected with GFP alone.

Figure 2. MiRP1 coexpression in CHO cells accelerates I_HERG deactivation at hyperpolarized potentials.

A, representative current recordings in cells expressing HERG alone or in combination with MiRP1 during 400 ms hyperpolarizing steps following 1 s depolarizations to +40 mV. Bold lines are monoexponential fits to deactivation phases. Horizontal bars show zero current levels. B, deactivation time constants as a function of test potential of hyperpolarizing pulse (means ± s.e.m., n = 8 and 9 for HERG and HERG/MiRP1, respectively. P < 0.01 for HERG vs. HERG/MiRP1, ANOVA).

Under physiological conditions, I_Kr deactivation occurs at voltages positive to the reversal potential, i.e. deactivating current is outward. Figure 3 shows the time course of I_Kr and I_HERG (with or without MiRP1) deactivation at −50 mV following 2 s depolarizing pulses to +20 mV. As shown in Fig. 3A, deactivation of outward tail currents occurred at similar rates for I_HERG in the absence and presence of MiRP1, whereas deactivation of I_Kr appeared faster. Mean deactivation kinetic data determined from biexponential fits for I_Kr, I_HERG alone and in combination with MiRP1 are shown in Fig. 3B (n = 21, 20 and 21 cells, respectively). The deactivation time constants (τ) were similar between I_HERG with or without MiRP1 coexpression. This is in contrast to the significantly different deactivation kinetics at potentials negative to the reversal potential (Fig. 2). The deactivation time constants for I_Kr were similar to those of HERG and HERG/MiRP1 (Fig. 3B); however, a larger proportion of I_Kr deactivation was attributable to the rapid phase in comparison with HERG or HERG/MiRP1 (Fig. 3C). This change in the relative magnitude of rapid deactivation was responsible for the faster overall deactivation of native I_Kr. In summary, MiRP1 coexpression accelerates I_HERG tail current deactivation at potentials negative to −90 mV. At more positive potentials, a significant difference is not evident and deactivation remains significantly slower than that of native I_Kr.

Figure 3. Native I_Kr outward current deactivation is faster than I_HERG expressed alone or in combination with MiRP1 in CHO cells.

A, representative examples of tail currents recorded from a guinea-pig cardiomyocyte (I_Kr) and CHO cells expressing HERG or HERG/MiRP1 during 2 s steps to −50 mV following 2 s depolarizing steps to +20 mV. Horizontal bars show zero current levels. B, I_Kr, HERG or HERG/MiRP1 tail current deactivation time constants as estimated by biexponential fits. C, proportion of deactivation attributable to faster phase, I_f /(I_s + I_f), for I_Kr, HERG or HERG/MiRP1. All data are presented as means ± s.e.m. (*P < 0.001 versus HERG or HERG/hMiRP1).

We next sought to evaluate the voltage dependence of I_HERG and I_Kr activation. Two-second depolarizing pulses in 10 mV increments were applied from V_H values of −40 mV and −80 mV for cardiomyocytes and CHO cells, respectively. Tail currents from both cell types were recorded by applying 2 s repolarizing steps to −50 mV (see protocol insets in top panels of Fig. 4). Figure 4A and B shows I_HERG recordings with steps to four voltages in the absence and presence of MiRP1, respectively. Figure 4C shows I_Kr recordings from a guinea-pig cardiomyocyte. Figure 4D shows mean normalized step current-voltage relations for guinea-pig I_Kr (n = 21, ○, dashed line), I_HERG alone (n = 20, □) and I_HERG coexpressed with MiRP1 (n = 21, ▪). Similar to corresponding results in oocytes (Fig. 4D inset), I_HERG alone activated at voltages more positive than when coexpressed with MiRP1. I_Kr activated at voltages more positive than I_HERG, with or without MiRP1 coexpression. Maximum step current density for HERG alone in 20 CHO cells averaged 17.3 ± 2.8 pA pF⁻¹, which was not significantly different from the maximum value for HERG/MiRP1, 14.2 ± 1.9 pA pF⁻¹ (n = 21). On the other hand, maximum tail current density following 2 s steps to +40 mV was smaller in cells expressing HERG/ MiRP1 (14.9 ± 2.1 pA pF⁻¹), compared to HERG alone (22.9 ± 3.1 pA pF⁻¹; P < 0.01). The voltage dependence of activation is analysed in Fig. 4E, based on mean normalized tail currents. I_HERG activation occurred at voltages (−22.1 ± 2.0 mV, n = 21) ∼10 mV more positive in the absence of MiRP1 than in its presence (−32.1 ± 0.6 mV, n = 20; P < 0.05versus I_HERG alone). The I_Kr half-activation voltage (−16.1 ± 2.0 mV, n = 21) was ∼6 mV more positive than that of I_HERG alone (P < 0.05) and ∼16 mV more positive than that of I_HERG coexpressed with MiRP1 (P < 0.05). The inset in Fig. 4E shows that qualitatively similar results were obtained upon HERG and HERG/MiRP1 coexpression in oocytes: the half-activation voltage of I_HERG (−19.5 ± 1.0 mV, n = 9) was shifted in the hyperpolarizing direction upon MiRP1 coexpression (−25.8 ± 1.5 mV, n = 7; P < 0.01vs. HERG alone). We saw no evidence of I_Ks in cardiomyocyte studies performed in the presence of 293B. Nevertheless, we verified the voltage dependence of native I_Kr activation with the use of currents sensitive to 5 μmol l⁻¹ E-4031. In six cardiomyocytes studied in the absence and presence of E-4031, the half-activation voltage of E-4031 sensitive I_Kr tails averaged −14.7 ± 3.1 mV, which was not significantly different from the value obtained with overall current as illustrated in Fig. 4E.

Figure 4. Voltage dependence of HERG, HERG/MiRP1 and guinea-pig I_Kr activation.

Representative recordings are shown from CHO cells expressing HERG alone (A), HERG/MiRP1 (B) or I_Kr in a guinea-pig ventricular cardiomyocyte (C). Horizontal bars show zero current levels. D, normalized step current amplitudes measured at the end of the 2 s depolarizing pulses. Inset, normalized step current amplitudes recorded from oocytes expressing HERG alone (n = 9, open symbols) or HERG/MiRP1 (n = 7, filled symbols). E, normalized tail current amplitudes determined by fitting the deactivation phases to monoexponential functions and extrapolating back to the beginning of the repolarizing step. Inset: normalized tail current amplitudes recorded from oocytes expressing HERG alone (n = 9, open symbols) or HERG/MiRP1 (n = 7, filled symbols). Curves are Boltzmann fits of the form I/I_max = (1+exp[(V_1/2 - V_T)/k])⁻¹, where V_1/2 is half-activation voltage, V_T is test voltage and k is slope factor. All data are presented as means ± s.e.m.

Response of guinea-pig I_Kr, and HERG or HERG/MiRP1 in CHO cells, to I_Kr-blocking agents

To study the response of HERG and HERG/MiRP1 in CHO cells to I_Kr-blocking antiarrhythmic drugs, the sampling pulse protocol was used (inset in Fig. 5A). In the absence of blockers, I_HERG was stable throughout the observation period (Fig. 5A, left). In the presence of 5 μmol l⁻¹ quinidine, block was extremely rapid, with steady-state block achieved before the first sampling pulse at 15 s. Current was completely blocked by 100 μmol l⁻¹ quinidine and this drug effect was largely reversible upon washout (WO). Results in the presence of MiRP1 (Fig. 5A, right) were qualitatively similar to those in its absence (Fig. 5A, middle). Figure 5B shows mean data for the onset of quinidine-induced I_HERG block, measured both on the basis of steady-state current prior to the hyperpolarizing pulse (corresponding to vertical arrows in Fig. 5A, open symbols in Fig. 5B) and inactivating current following the hyperpolarization (filled symbols). Because the rapid onset of block could not be resolved with the sampling pulse protocol, we repeated the experiments with quinidine applied during a maintained depolarization to 0 mV and studied changes in steady-state current, as illustrated in the inset of Fig. 5B. The kinetics of I_HERG block by 5 μmol l⁻¹ quinidine were similar in the absence (n = 5) and presence (n = 4) of MiRP1, with monoexponential time constants of 2.05 ± 0.45 and 2.27 ± 0.53 s, respectively (n.s.).

Figure 5. Kinetics of block onset of I_HERG by quinidine, E-4031 and dofetilide are unaltered by MiRP1 coexpression in CHO cells.

A left, currents recorded using sampling pulse protocol (inset) from a cell expressing HERG over a 5 min drug-free recording period. A centre and right, recordings from cells expressing HERG or HERG/MiRP1 at 15 s, 1 min and 1.5 min after application of 5 μmol l⁻¹ quinidine. Current was completely blocked by subsequent application of 100 μmol l⁻¹ quinidine and block was largely reversed after 4 min of washout (WO). Horizontal bars show zero current levels. B, normalized steady-state (vertical arrows in A) or inactivating current amplitudes decreased rapidly in the presence of 5 μmol l⁻¹ quinidine. To resolve the kinetics of block, 5 μmol l⁻¹ quinidine was applied during continued depolarization to 0 mV (inset). Monoexponential fits to current decay yielded time constants of 2.05 ± 0.45 and 2.27 ± 0.53 s (n.s.) for HERG and HERG/MiRP1. Block onset by 50 nm E-4031 (C) and 50 nm dofetilide (D) were also evaluated using both the steady-state and inactivating current amplitudes. All data are presented as means ± s.e.m.

Figure 5C indicates the response of I_HERG to 50 nmol l⁻¹ E-4031. I_HERG block by E-4031 occurred slowly, taking several minutes to reach maximal inhibition, and was essentially irreversible. Monoexponential fits provided time constants of 58.7 ± 7.3 s (n = 3) and 67.2 ± 14.9 s (n = 4) for HERG and HERG/hMiRP1 (based on steady-state current) and 54.5 ± 2.9 and 55.1 ± 2.8 s (with inactivating current; n.s. for HERG versus HERG/MiRP1; n.s. for results with steady-state versus inactivating current). Figure 5D shows data for 50 nmol l⁻¹ dofetilide. Block time constants averaged 103.6 ± 5.6 s (n = 5) and 96.7 ± 4.4 s (n = 4) (steady-state current) and 100.7 ± 6.5 and 96.3 ± 4.1 s (inactivating current) for HERG and HERG/MiRP1, respectively (n.s.). Block of I_HERG by dofetilide was also irreversible.

Figure 6A illustrates the concentration-dependent effects of quinidine on HERG and HERG/MiRP1. Complete block was achieved at 100 μmol l⁻¹ and block was reversible. Figure 6B shows mean concentration-response relations for quinidine block of steady-state (open symbols) and inactivating (filled symbols) I_HERG. IC₅₀ values were 1.0 ± 0.2 μmol l⁻¹ (n = 5) and 0.96 ± 0.1 μmol l⁻¹ (n = 4) for HERG and HERG/MiRP1, respectively, based on steady-state current (n.s. for HERG versus HERG/MiRP1). IC₅₀ values based on inactivating current were 1.1 ± 0.4 and 1.2 ± 0.3 μmol l⁻¹ (n.s.).

Figure 6. Sensitivity of I_HERG to quinidine, E-4031 and dofetilide are unaltered by MiRP1 coexpression in CHO cells.

A, representative recordings with the sampling pulse protocol (inset) before and after various quinidine concentrations in two cells expressing HERG or HERG/MiRP1, as well as after washout. Horizontal bars indicate zero current levels. Normalized steady-state (vertical arrows in A) or inactivating current amplitudes plotted as a function of quinidine (B), E-4031 (C) or dofetilide (D) concentrations. All data are presented as means ± s.e.m.

Figure 6C illustrates logistic concentration-response fits for I_HERG inhibition by E-4031. IC₅₀ values for steady-state current were 5.0 ± 0.6 nmol l⁻¹ (n = 3) and 6.0 ± 0.1 nmol l⁻¹ (n = 4) for HERG and HERG/MiRP1, respectively (n.s.). Fits to inactivating current data provided respective IC₅₀ values of 6.1 ± 1.0 and 10.0 ± 0.8 nmol l⁻¹ (n.s.). The concentration-response relationships for dofetilide are illustrated in Fig. 6D. Mean IC₅₀ values for HERG and HERG/MiRP1 were 10.0 ± 2.0 nmol l⁻¹ (n = 5) and 8.4 ± 1.1 nmol l⁻¹ (n = 4), respectively, based on steady-state currents and 12.5 ± 3.1 and 9.7 ± 2.3 nmol l⁻¹ based on inactivating currents (n.s. for HERG versus HERG/MiRP1).

In summary, inhibition of I_HERG with or without MiRP1 coexpression can be evaluated during sampling pulse protocol application by measuring either the steady-state current or inactivating current amplitudes, and coexpression of MiRP1 did not significantly alter the kinetics or the IC₅₀ of I_HERG block by quinidine, E-4031 or dofetilide.

The data shown in Fig. 5 and Fig. 6 suggest that MiRP1 coexpression does not significantly alter the pharmacological response of I_HERG in CHO cells. We wished, however, to verify the blocking actions of quinidine, dofetilide and E-4031 on I_HERG (with or without MiRP1 coexpression) and I_Kr under comparable conditions. We therefore recorded I_HERG in CHO cells and I_Kr in cardiomyocytes with the same voltage protocol (Fig. 7A inset) and superfusate contents (including 1 μmol l⁻¹ nimodipine and 50 μmol l⁻¹ 293B). Figure 7A illustrates concentration-dependent dofetilide block of I_Kr (left), HERG (middle) and HERG/ MiRP1 (right panel). There are no major discrepancies apparent in dofetilide block among I_Kr, HERG and HERG/ MiRP1. Mean data for concentration-dependent block by quinidine, E-4031 and dofetilide are shown in Fig. 7B, C and D, respectively. All data are based on normalized steady-state holding currents (vertical arrows in Fig. 7A), since inactivating I_Kr currents tended to be small. The IC₅₀ values were 1.4 ± 0.2 μmol l⁻¹ (n = 6), 1.1 ± 0.1 μmol l⁻¹ (n = 4) and 1.5 ± 0.2 μmol l⁻¹ (n = 4) for I_Kr, HERG and HERG/ MiRP1 block, respectively (n.s. for I_Krversus HERG and HERG/MiRP1) by quinidine. Corresponding values for E-4031 were 10.3 ± 0.3 nmol l⁻¹ (n = 4), 7.8 ± 0.4 nmol l⁻¹ (n = 5) and 7.3 ± 0.7 nmol l⁻¹ (n = 4) (n.s.), and values for dofetilide averaged 8.7 ± 2.0 nmol l⁻¹ (n = 5), 10.2 ± 0.9 nmol l⁻¹ (n = 4) and 8.6 ± 1.5 nmol l⁻¹ (n = 5) (n.s.). There were no significant differences in IC₅₀ between I_Kr and I_HERG with or without MiRP1 for any drug.

Figure 7. Inhibition of I_Kr, HERG or HERG/MiRP1 recorded under identical conditions shows only minor pharmacological differences.

The sampling pulse protocol (inset in A) was used to evaluate the sensitivity and kinetics of block by quinidine, E-4031 and dofetilide. A, representative recordings before and after 3–4 min perfusion with successively larger dofetilide concentrations. Horizontal bars indicate zero current levels. B-D, normalized steady-state holding current amplitudes (vertical arrows in A) plotted as a function of drug concentration. I_Kr data is the same as that shown in Fig. 1A. E-G, time-dependent block of the steady-state current amplitudes by 5 μm quinidine, 50 nm E-4031 and 50 nm dofetilide. Because steady-state block of HERG and HERG/MiRP1 was evident during the first pulse for quinidine, block onset was quantified by studying the reduction in current as a function of time during a single maintained depolarization to 0 mV, as shown in inset of E. All data are presented as means ± s.e.m. Dashed lines in B-G join results for I_Kr, whereas continuous lines join data for HERG and HERG/MiRP1.

Mean data for the kinetics of block onset are shown in Fig. 7E, F and G. I_HERG block onset for quinidine could not be resolved on a pulse-dependent basis; therefore, the quantitative results for quinidine were obtained by studying the onset of steady-state current block as in the inset of Fig. 7E. Mean blocking time constants for quinidine (5 μm) were 18.1 ± 2.0 s (n = 6), 2.3 ± 0.1 s (n = 4) and 2.4 ± 0.3 s (n = 4), respectively for I_Kr, HERG without and with MiRP1 (P < 0.01 for I_Krversus HERG and HERG/MiRP1; n.s. for HERG versus HERG/MiRP1). Corresponding results for 50 nm E-4031 were 32.2 ± 1.2 s (n = 4), 35.3 ± 1.3 s (n = 4) and 28.0 ± 3.2 s (n = 7, n.s. among I_Kr, HERG and HERG/MiRP1). For 50 nm dofetilide, time constants averaged 36.6 ± 2.1 s (n = 5), 89.6 ± 8.2 s (n = 4) and 85.5 ± 4.6 s (n = 4, P < 0.05 for I_Krversus HERG and HERG/MiRP1; n.s. for HERG versus HERG/MiRP1). Mean time constants for block onset based on HERG or HERG/MiRP1 inactivating current were 34.5 ± 1.0 s (n = 4) and 28.9 ± 0.9 s (n = 7, n.s.) for E-4031 and 88.0 ± 3.5 s (n = 4) and 91.3 ± 5.2 s (n = 4, n.s.) for dofetilide, respectively. In summary, the pharmacological responses of I_HERG expressed alone in CHO cells were very similar to that of native I_Kr and the relatively minor differences in blocking kinetics of quinidine and dofetilide were not eliminated upon MiRP1 coexpression.

DISCUSSION

To the best of our knowledge, our study constitutes the first attempt at directly comparing the biophysical and pharmacological properties of HERG, HERG coexpressed with MiRP1 in CHO cells and native I_Kr using similar electrophysiological conditions. Coexpression with MiRP1 did not alter the pharmacological responses of I_HERG to three I_Kr-blocking antiarrhythmic drugs. The concentration dependence of I_HERG block was indistinguishable from that of I_Kr. There were some differences in blocking kinetics between I_HERG in CHO cells and I_Kr, but the differences were not reduced by coexpression with MiRP1. MiRP1 coexpression similarly failed to diminish differences between I_Kr and I_HERG in activation voltage dependence and deactivation kinetics of outward tail currents. In the oocyte system, I_HERG was much less sensitive to drug block than was I_Kr in native myocytes, and block onset was much slower.

Relationship between present findings regarding MiRP1 and previous studies

The potential role of MiRP1 as a β-subunit for HERG was first described by Abbott et al. (1999). Based on several lines of evidence, the authors argued that MiRP1 and HERG constitute native I_Kr in much the same way that KvLQT1 and minK form native I_Ks. The data supporting this contention included the relative biophysical and pharmacological properties of currents resulting from the expression of HERG and HERG with MiRP1, along with results indicating the ability of the subunits to associate physically. Most of the functional data were obtained in Xenopus oocytes, and comparisons with the behaviour of I_Kr were based on results in the literature; no direct studies of I_Kr were performed. The results of our studies of HERG expression alone in Xenopus oocytes (Fig. 1) are consistent with previous work in the literature (Sanguinetti et al. 1995; Abbott et al. 1999) indicating important differences in pharmacological response and sensitivity from native I_Kr.

On the other hand, our studies of CHO cells stably expressing I_HERG showed that, under comparable conditions, the sensitivity of I_HERG to quinidine, E-4031 and dofetilide is indistinguishable from that of native I_Kr, and is not affected by coexpression with MiRP1. We observed acceleration by coexpression with MiRP1 of inward I_HERG tail currents at hyperpolarized potentials (Fig. 2), qualitatively similar to results reported by Abbott et al. (1999), but the more physiologically relevant I_HERG outward tail currents were not affected by MiRP1 and remained slower than those of I_Kr (Fig. 3). Of note, although Abbott et al. (1999) do not provide kinetic data for outward I_HERG tail currents; outward tail currents are present under the experimental conditions shown in their Fig. 7 and do not suggest acceleration upon coexpression with MiRP1. Like Abbott et al. (1999) we found that coexpression with MiRP1 shifted the voltage dependence of activation; however, the voltage shift we observed was in the hyperpolarizing direction and did not make the properties of I_HERG more similar to those of native I_Kr. Abbott et al. (1999) described a 9 mV depolarizing shift in I_HERG inactivation upon coexpression with rat MiRP1, but only a 4 mV depolarizing shift of unstated statistical significance when HERG was coexpressed with human MiRP1, and the I_HERG current-voltage relation was not altered by coexpression with human or rat MiRP1. Zhang et al. (2001) did not observe a significant activation voltage shift or change in I_HERG deactivation kinetics upon coexpression with MiRP1 at a 6.6:1 (MiRP1:HERG) molar ratio. A recent study by Mazhari et al. (2001) showed no change in voltage dependence of HERG upon coexpression with MiRP1 in HEK cells and obtained qualitatively similar results to ours regarding changes in deactivation kinetics and current amplitude.

Other recent studies are compatible with the notion that the physiological function of MiRP1 may not be limited to the reconstitution of native I_Kr upon co-assembly with HERG. Zhang et al. (2001) have shown that MiRP1 can associate with Kv4.2, and modify the kinetics and voltage dependence of Kv4.2 and Kv4.3. MiRP1 coexpression can also affect the current amplitude and gating kinetics of KvLQT1, conferring background characteristics to the channel (Tinel et al. 2000a). There is also evidence that MiRP1 can associate with the brain K⁺ channel subunits KCNQ2 and KCNQ3 and accelerate their deactivation kinetics (Tinel et al. 2000b). Finally, coexpression of HERG and MiRP1 can alter the modulation of I_HERG by cyclic AMP, favouring current enhancement by a hyperpolarizing shift in activation due to direct binding with a cyclic nucleotide binding domain over inhibition caused by channel phosphorylation (Cui et al. 2000). In combination with our findings, these results suggest that MiRP1 may act by subtly modifying the function of a number of ion channels, rather than by specifically co-assembling with HERG to reconstitute I_Kr.

Comparison between I_Kr properties in the present study and previous results in the literature

The activation V_1/2 we estimated for I_Kr (∼-16 mV) is similar to the values of −13.7 and −16.5 mV reported for rabbit ventricular cardiomyocytes (Carmeliet 1992, 1993), −10.8 mV in rabbit AV nodal cells (Mitcheson & Hancox, 1999), −21.5, −14.9 and −17.1 mV in guinea-pig ventricular cardiomyocytes (Sanguinetti & Jurkiewicz, 1990; Heath & Terrar, 1996; Bosch et al. 1998), −16.3 mV in fetal mouse ventricular cardiomyocytes (Wang & Duff, 1996) and −13 mV in ferret atrial cardiomyocytes (Liu et al. 1996). Thus, the V_1/2 values for I_Kr in a range of species is comparable and distinctly more positive than the result we obtained for HERG/MiRP1 coexpression.

Data in Fig. 3 indicate that though the fast and slow deactivation time constants were similar among I_Kr, HERG and HERG/MiRP1, I_Kr deactivation appeared faster due to a larger contribution of the fast component. In previous studies, I_Kr deactivation time constants at −40 or −50 mV have generally been estimated by monoexponential curve fits. Values have ranged from 358 ms in fetal mouse heart (Wang & Duff, 1996) to ∼350–400 ms in ferret atrium (Liu et al. 1996) and rabbit ventricle (Clay et al. 1995) and 140 ms in guinea-pig ventricle (Sanguinetti and Jurkiewicz 1990). When we applied monoexponential curve fitting to our data, we obtained results similar to these values in the literature, and the time constants for HERG (788 ± 88 ms) and HERG/MiRP1 (745 ± 46 ms) were significantly greater than those for I_Kr (319 ± 17 ms, P < 0.001vs. HERG or HERG/MiRP1). However, biexponential analyses provided better fits to the data and suggested that the differences between I_Kr and HERG with or without MiRP1 were due to the relative importance of fast-phase deactivation rather than in the intrinsic rates of deactivation per se.

The sensitivity of I_Kr to quinidine that we observed (IC₅₀ about 1.4 μmol l⁻¹) is similar to values (1.0 and 0.9 μmol l⁻¹) reported for AT-1 (atrial tumour line-1) cells (Yang & Roden 1996; Yang et al. 1997). The sensitivity to E-4031 (IC₅₀ ∼10 nmol l⁻¹) is similar to that reported for ferret atrial cardiomyocytes (10 nmol l⁻¹; Liu et al. 1996), but greater than the first report for guinea-pig I_Kr (397 nmol l⁻¹, Sanguinetti & Jurkiewicz, 1990). Dofetilide sensitivity (IC₅₀ ∼10 nmol l⁻¹) was also comparable with values of 12 and 11 nmol l⁻¹ reported in AT-1 cells (Yang & Roden, 1996; Yang et al. 1997) and results (4 nmol l⁻¹) in rabbit ventricular cardiomyocytes (Carmeliet, 1992).

Novel aspects and potential significance

I_Kr plays a crucial role in cardiac repolarization. An accurate appreciation of its molecular basis is critical for an understanding of the basis of cardiac repolarization and for the development of antiarrhythmic drug therapy. Since the report of Abbott et al. (1999), it has been widely believed that co-assembly of HERG and MiRP1 is required to reconstitute the properties of native I_Kr, in the same way that the combination of minK and KvLQT1 is believed to produce I_Ks. Our findings raise doubts about this contention. We were unable to document any properties of native I_Kr that were closer to those of currents resulting from the coexpression of HERG and MiRP1 than those carried by HERG expression alone.

There is clear evidence that MiRP1 mutations are associated with ventricular tachyarrhythmic phenotypes (Abbott et al. 1999; Sesti et al. 2000). In the light of our findings, which suggest that MiRP1 does not function to recapitulate native I_Kr upon coexpression with HERG, what potential mechanisms could be involved in MiRP1 mutation-related arrhythmias? First, MiRP1 may affect cardiac repolarization by modulating ion channels other than I_Kr. Potentially important interactions have been reported for MiRP1 and Kv4.2 and 4.3 (Zhang et al. 2001) as well as MiRP1 and KvLQT1 (Tinel et al. 2000a). Second, the fact that MiRP1 coexpression does not improve the concordance between the properties of I_HERG and I_Kr does not exclude the possibility that HERG and MiRP1 co-assemble. MiRP1 may play a primary role in governing the response of I_HERG/I_Kr to changes in intracellular cyclic AMP (Cui et al. 2000; Cui et al. 2001). Mutations in MiRP1 might alter the three-dimensional structure of the HERG/ MiRP1 complex, impairing channel function (Abbott et al. 1999; Sesti et al. 2000), even if binding of wild-type MiRP1 per se is not necessary for normal I_Kr function. A recent modelling analysis suggests that acceleration of I_HERG deactivation by an arrhythmogenic MiRP1 mutation may account for associated repolarization changes (Mazhari et al. 2001).

Our findings may have practical importance for the evaluation of drugs for I_Kr-blocking properties. There is presently considerable concern about the potential risks of acquired long-QT syndrome induction by all medications, and the Food and Drug Administration requires screening of all new compounds for such risks. Effective screening for I_Kr blockade is difficult because of the low throughput and technical difficulty of assays employing isolated cardiomyocytes. The heterologous expression of cloned channel subunits provides a much more practical solution, but requires that the system used has a drug blocking sensitivity comparable to that of native I_Kr. Our results suggest that the expression of HERG in a mammalian system may provide such a screening tool because of its very similar sensitivity to that of native I_Kr (Figs 7B, C and D). The study of I_HERG in oocytes would not seem to be adequate, because of its much lower sensitivity to I_Kr blockers, and the coexpression of MiRP1 would appear to be unnecessary.

Potential limitations

The biophysical properties of I_HERG in CHO cells were not identical to those of native I_Kr. In addition, the kinetics of onset of quinidine block were clearly faster, and dofetilide slightly slower, for HERG in CHO cells than for native I_Kr. Therefore, additional factors must be invoked to account for the discrepancies. Regulatory elements or specific membrane constituents may differ between CHO cells and guinea-pig cardiomyocytes, modifying HERG channel function and accessibility to block by quinidine and dofetilide. Splice variants of HERG are known to be present in the heart and to show differences in deactivation kinetics (London et al. 1997; Lees-Miller et al. 1997). The HERG construct we expressed was the longer variant, HERG A; the coexpression of HERG B in native cardiomyocytes could account for some of the differences between I_Kr and I_HERG that we noted. There is, however, evidence in human and rat hearts that only HERG A is expressed at the protein level (Pond et al. 2000).

I_HERG was studied with the use of a stably expressing cell line and MiRP1 was added when desired by transient transfection. With this type of approach, it is essential to ensure adequate MiRP1 expression. We assessed the effective expression of MiRP1 in two ways in each cell studied: (1) visualization of coexpressed green fluorescent protein; (2) demonstration of accelerated deactivation kinetics at −110 mV. Qualitatively similar results for the HERG versus HERG/MiRP1 phenotypes were also obtained with direct injection of HERG and MiRP1 cRNA in Xenopus oocytes.

We used a sampling pulse protocol for our pharmacological studies, because we have found the protocol to be well-tolerated and to allow for detailed evaluation of drug blocking actions without cycling the cell repeatedly through different states (Weerapura et al. 2002). This protocol differs from that in many previous studies, in which cells are held at a negative potential (e.g. −80 mV) and repeatedly depolarized for shorter intervals (e.g. 2 s). We have examined the onset and extent of block with 50 nm E-4031, 50 nm dofetilide and 5 μm quinidine, with the use of 2 s pulses to 0 mV every 10 s from a holding potential of −80 mV. The results (not shown) indicate that MiRP1 coexpression also fails to alter HERG responses to these agents under repeated pulse conditions. Unlike Abbott et al. (1999), we failed to observe a tonic or closed state block when cells were pre-equilibrated for 1 min at −80 mV with E-4031 or dofetilide prior to repetitive pulsing.

We used 2 s depolarizing pulses to determine the half-activation voltage. At negative voltages (e.g. −40 mV), 2 s are not sufficient to achieve steady state activation (Weerapura et al. 2002). This phenomenon may result in errors in estimating the true half-activation voltage. However, other studies in the literature have used similar protocols. For example, Abbott et al. (1999) used 1–2 s, Mazhari et al. (2001) used 3.5 s and Cui et al. (2001) used 4 s pulses. All these studies were performed at room temperature, at which activation is much more than two-fold slower than in our studies at 34–36 °C. Even at physiologically relevant recording temperatures, it is still not technically feasible to use long enough pulses to reach steady state at all voltages required to determine half-maximal activation voltage. We have therefore used protocols similar to those in the literature to facilitate comparison. Since the same protocol was applied under each condition, inter-group comparisons should be valid providing comparable activation kinetics.