Tetrodotoxin-sensitive Na_vs contribute to early and delayed afterdepolarizations in long QT arrhythmia models

Cardiac arrhythmias, a leading cause of death in the United States, are often precipitated by premature heartbeats (Kong et al., 2011). Such premature electrical activity, dubbed triggered activity, is a hallmark of pathologies stemming from defects in Ca²⁺ cycling proteins. Catecholaminergic polymorphic ventricular tachycardia (CPVT) is one such syndrome, caused by genetic defects in the RYR2 Ca²⁺ release channel and its complex (Priori et al., 2001; Knollmann et al., 2006). Such dysfunctional RYR2 gating, in the setting of catecholamine stimulation (Bovo et al., 2017), permits the RYR2 channels to reopen during diastole. The resulting aberrant Ca²⁺ release can, in turn, activate depolarizing membrane currents and prompt pro-arrhythmic delayed afterdepolarizations (DADs). These form the basis for the bifocal or multifocal aberrant electrical activity that underlies narrow QRS complex CPVT (Cerrone et al., 2007; Györke, 2009). It is important to note that the RYR2 dysfunction by itself, without the aid of catecholamines, is insufficient to induce triggered activity or the resulting ventricular tachycardias (VTs; Györke, 2009; Radwański et al., 2016). On the other hand, in arrhythmic syndromes such as long QT (LQT), prolongation of the action potential drives triggered activity, specifically early afterdepolarizations (EADs), which in turn lead to reentrant polymorphic VTs marked by a wide QRS complex, dubbed torsades de pointes (TdP; Volders et al., 1997, 2000; Varró and Baczkó, 2011).

Recent evidence suggests that these two types of triggered arrhythmias share mechanistic similarities (Burashnikov and Antzelevitch, 2006; Shiferaw et al., 2012). Just as with CPVT, various LQT syndromes have been linked to altered RYR2 function (Volders et al., 1997; Katra and Laurita, 2005; Cheng et al., 2011; Terentyev et al., 2014). Furthermore, LQT syndromes can also induce triggered activity via DADs, which result from SR Ca²⁺ release, and the electrogenic Na⁺/Ca²⁺ exchange (NCX; Volders et al., 1997; Burashnikov and Antzelevitch, 2000; Gallacher et al., 2007; Radwański et al., 2010; Radwański and Poelzing, 2011). However, outside of dysfunctional RYR2 and increased SR Ca²⁺ load, the mechanisms underlying EADs and abnormal Ca²⁺ handling–mediated DADs in LQT are not well understood. Surprisingly, despite the central role of intracellular Ca²⁺ in LQT-promoted arrhythmias, Na⁺ channel blockade has proven to be effective in suppressing triggered activity in these models (January et al., 1988; Radwański et al., 2013).

Multiple Na⁺ channel isoforms have been identified in the heart. The most abundant, the cardiac-type Na⁺ channels (Na_v1.5), are primarily distributed on the outer membrane of myocytes and are responsible for electrical excitability (Haufe et al., 2007). The other pool of Na⁺ channels, the neuronal Na⁺ channels (nNa_vs), are localized to membrane invaginations known as transverse tubules (Maier et al., 2002, 2004; Westenbroek et al., 2013; Radwański et al., 2015, 2016; Veeraraghavan et al., 2017) and modulate excitation–contraction coupling (Lipp and Niggli, 1994; Larbig et al., 2010; Torres et al., 2010). Although Na_v1.5 mutations have been linked to LQT syndrome (Wang et al., 1995), the role of noncardiac-type Na_v isoforms in LQT syndrome remains unclear. Recently, we demonstrated that physiological Na⁺ influx through nNa_v, in a setting of RYR2 dysfunction, as evidenced during catecholamine stimulation, can trigger arrhythmias (Radwański et al., 2015, 2016). Here, we propose that Na⁺ influx through nNa_v, along with dysfunctional RYR2, and enhanced SR Ca²⁺ load can conspire to prompt arrhythmias in LQT-driven pathologies as well as those resulting directly from Ca²⁺ cycling defects (i.e., CPVT). To this end, we demonstrate a key role for nNa_Vs in triggered activity (EADs and DADs) during pharmacologically induced prolongation in repolarization and/or augmentation of global Na_v or L-type Ca²⁺ channel activity. Notably, the modulation of the arrhythmia risk by nNa_vs occurs independently of changes in action potential duration (APD) and QT interval.

All animal procedures were approved by The Ohio State University Institutional Animal Care and Use Committee and conformed to the Guide for the Care and Use of Laboratory Animals published by the U.S. National Institutes of Health (NIH Publication No. 85-23, revised 2011).

Ventricular myocytes were obtained by enzymatic isolation from 3–11-mo-old cardiac calsequestrin (CASQ2)-null mice (on mixed background; Radwański et al., 2016) as well as WT (on C57BL/6 background) mice of both genders. Mice were anaesthetized with isoflurane, and once a deep level of anesthesia was reached, the hearts were rapidly removed and Langendorff perfused as previously described (Radwański et al., 2015, 2016). Whole-cell patch-clamp recordings of action potentials were performed with an Axopatch 200B amplifier using external solution that contained (in mM) 140 NaCl, 5.4 KCl, 1.8 CaCl₂, 0.5 MgCl₂, 10 HEPES, and 5.6 glucose, pH 7.4. Patch pipettes were filled with a solution that contained (in mM) 90 potassium aspartate, 50 KCl, 5 MgATP, 5 NaCl, 1 MgCl₂, 0.1 Tris GTP, 10 HEPES, and 0.1 EGTA, pH 7.2. Action potentials were evoked by injection of brief stimulus current 1.5–4 pA, 0.5–1 ms. Late and peak sodium currents (I_Nas) were recorded using internal solution containing (in mM) 10 NaCl, 20 TEACl, 123 CsCl, 1 MgCl₂, 0.1 Tris GTP, 5 MgATP, 10 HEPES, and 1 EGTA while free Ca²⁺ was maintained at 100 nmol/L with CaCl₂, pH 7.2. The extracellular bathing solution for late I_Na recordings contained (in mM) 140 NaCl, 4 CsCl, 1 CaCl₂, 2 MgCl₂, 0.05 CdCl₂, 10 HEPES, 10 glucose, 0.03 niflumic acid, 0.004 strophanthidin, and 0.2 NiCl₂. pH was maintained at 7.4 with CsOH. For peak I_Na recordings, extracellular bathing solution was altered by reducing NaCl to 10 mM, increasing CsCl to 123 mM, and adding 20 mM TEACl. Whole-cell capacitance and series resistance compensation (≥60%) was applied along with leak subtraction. Signals were filtered with a 10-kHz Bessel filter, and I_Na was then normalized to membrane capacitance. Late I_Na was estimated by integrating I_Na between 50 and 450 ms. Intracellular Ca²⁺ cycling was monitored by either Nikon A1R or Olympus FluoView 1000 laser scanning confocal microscope equipped with 60× 1.4-NA oil objectives. For intact, field-stimulated myocytes exposed to 4-aminopyridine (4AP) and anemone toxin II (ATX-II), we used the cytosolic Ca²⁺-sensitive indicator Fluo-3 AM (Molecular Probes) to assess SR Ca²⁺ load. Cells were electrically stimulated at 2 Hz using extracellular platinum electrodes for at least 10 s, and 20 mM caffeine was rapidly applied. To more accurately assess SR Ca²⁺ load in cardiomyocytes exposed to Bay K 8644 (BayK), we used low-affinity cytosolic Ca²⁺-sensitive indicator Fluo-4FF AM (Molecular Probes). The fluorescent probes were excited with the 488-nm line of an argon laser, and emission was collected at 500–600 nm. Fluorescence was recorded in the linescan mode of the confocal microscope.

For simultaneous whole-cell patch-clamp recordings of action potentials and Ca²⁺ waves, patch pipette solution was supplemented with Fluo-3 pentapotassium (0.03 mM; Molecular Probes). Myocytes were paced at 1 Hz to obtain Ca²⁺ wave frequency. Any aberrant Ca²⁺ release event (i.e., a wave or wavelet) that increased cell-wide fluorescence intensity >10% of the signal generated by the preceding stimulated Ca²⁺ transient was included in the analysis. The fluorescence emitted was expressed as F/F₀, where F is the fluorescence at time t and F₀ represents the background signal. Recordings were made 1 min after solution change, except for ATX-II, which required a 10-min wash-in to achieve steady state. EAD was defined as transient slowing or reversal of membrane potential during repolarization. DAD was defined as a fluctuation from the resting membrane potential of more than 4 mV. After making the baseline recordings, in the majority of experiments, we induced arrhythmogenic events (4AP + isoproterenol [ISO], BayK, or ATX-II), which were then followed by sodium channel blocking interventions. We attempted to alternate the order of sodium channel blockers between experiments. All experiments were performed at room temperature (26°C).

Continuous electrocardiographic (ECG) recordings (PL3504 PowerLab 4/35; ADInstruments) were obtained from mice anesthetized with isoflurane (1–1.5%) as previously described (Radwański et al., 2015, 2016). Briefly, after baseline recording (5 min), a subset of animals received either intraperitoneal ATX-II (30 µg/kg; Alomone Labs) or BayK (1 mg/kg; Tocris). After 10 min, animals received vehicle, riluzole (15 mg/kg; Sigma), or 4,9-anhydro-tetrodotoxin (4,9-ah-TTX; 750 µg/kg; Focus Biomolecules). After an additional 5–10 min, animals were exposed to an intraperitoneal epinephrine (1.5 mg/kg; Sigma) and caffeine (120 mg/kg; Sigma) challenge, and ECG recording continued for 20 min. Because increased heart rate has been linked to reduced arrhythmia inducibility in CPVT (Faggioni et al., 2013), and because BayK is a potent positive chronotrope (Seifen and Kennedy, 1986), animals exposed to BayK were pretreated with ivabradine (3 mg/kg; Sigma) 10 min before BayK exposure. ECG recordings were analyzed using the LabChart 7.3 program (ADInstruments). QT interval was measured before catecholamine challenge from the beginning of the QRS complex to the isoelectric baseline for entirely positive T waves or to the trough of any negative T wave (Zhang et al., 2014). Heart rate–corrected QT (QTc) intervals were then obtained using the formula QTc = QT/(RR/100)^1/2 (Mitchell et al., 1998). VT was defined as three or more premature ectopies with prolonged QRS interval (≥5 ms relative to the one originating during sinus rhythm) and was categorized as either narrow or wide QRS (≥29 ms).

Isolated ventricular myocytes were prepared for immunofluorescence as described previously (Radwański et al., 2015, 2016). Briefly, cells were plated on laminin-coated glass coverslips, fixed with 4% paraformaldehyde for 5 min, permeabilized with 0.1% Triton X-100, and washed with PBS. Endogenous immunoglobulin was blocked using a 2% BSA PBS solution for 1 h at room temperature and subsequently incubated with primary antibodies. We immunolabeled for nNa_vs (Na_v1.1, 1.3, and 1.6; Alomone) and RYR2 (Pierce Antibodies) overnight at 4°C. After washing, goat secondary antibodies (anti–mouse and anti–rabbit) conjugated to Alexa Fluor 488 or 549 (Life Technologies) were added for 1 h.

Action potential and I_Na analysis was performed using pCLAMP9 and pCLAMP10.3 software (Molecular Devices). ECG recordings were analyzed using the LabChart 7.3 program (ADInstruments). Line scanning images of Ca²⁺ were normalized for baseline fluorescence (Radwański et al., 2015, 2016). Ca²⁺ imaging data were processed using ImageJ (National Institutes of Health) and Origin software. Statistical analysis of the data was performed using a Wilcoxon signed rank and Friedman rank sum test or Kruskal–Wallis one-way ANOVA for paired and unpaired data, respectively. The Conover correction with further adjustment by the Holm family-wide error rate method was applied to adjust for multiple comparisons. Data are presented as median with 25th and 75th percentiles (box) and 10th and 90th percentiles (whiskers). Fisher’s exact test was used to test differences in nominal data. P < 0.05 was considered statistically significant.

Fig. S1 shows the effect of Na⁺ channel blockade on peak Na⁺ current and action potential. Fig. S2 shows the effect of Na⁺ channel blockade on SR Ca²⁺ load during prolonged repolarization. Fig. S3 shows the effect of APD on triggered activity during prolonged repolarization. Fig. S4 shows the effect of nNa_V blockade during concomitant exposure to ATX-II on SR Ca²⁺ load. Fig. S5 shows the effect of nNa_V blockade during concomitant exposure to BayK on SR Ca²⁺ load. Table S1 shows the impact of nNa_v inhibition on RR and QT interval in ATX-II– and BayK-treated mice.

We first investigated the role of Na⁺ channel blockade on triggered arrhythmias induced by prolonged repolarization. Specifically, we used 4AP to inhibit transient outward current (I_to), the major repolarizing current in mouse heart. Notably, exposure of patch-clamped ventricular cardiomyocytes isolated from WT mice to 4AP (4 mM) and ISO (100 nM) resulted in frequent triggered electrical activity in the form of EADs and DADs (Fig. 1, a and b, red traces and bars). Here we examined the effect of nonselective, pan-Na⁺ channel block with lidocaine (10 µM; Theile and Cummins, 2011), nNa_v inhibition with riluzole (10 µM; Song et al., 1997) or Na_v1.6 inhibition with 4,9-ah-TTX (300 nM; Rosker et al., 2007). At these concentrations, the agents achieved similar levels of I_Na reduction (Fig. S1 a), while having no significant effect on APD measured at 90% (APD₉₀) repolarization (Fig. S1 b). Next, we examined the impact of these Na_v-blocking agents on 4AP-promoted cellular triggered activity. Addition of lidocaine (10 µM) to 4AP-ISO–exposed patch-clamped WT cardiomyocytes did not significantly reduce the frequency of triggered activity, whether EADs or DADs (Fig. 1, a and b, green traces and bars). On the other hand, 4,9-ah-TTX (300 nM) and riluzole (10 µM) significantly reduced triggered activity in 4AP-ISO–exposed WT cardiomyocytes (Fig. 1, a and b, blue and purple traces and bars), suggesting that nNa_v blockade and not necessarily the extent of total I_Na reduction contributes to the antiarrhythmic action of Na_v blockers during prolonged repolarization achieved with 4AP. Based on this, lidocaine was not used in the investigation of the role of nNa_vs in the models of LQT-CPVT overlap phenotypes.

To investigate the effects of Na_v blockade on aberrant Ca²⁺ handling, we examined the response of aberrant Ca²⁺ release to various Na_v blockers during concomitant exposure to 4AP and ISO. Exposure of WT myocytes to 4AP-ISO induced frequent aberrant Ca²⁺ release events (Fig. 1, a and b, red traces and bars). Notably, addition of lidocaine (10 µM) had no significant effect on aberrant Ca²⁺ release (Fig. 1, a and b, green trace and bar), whereas 4,9-ah-TTX (300 µM) significantly reduced aberrant Ca²⁺ release (blue trace and bar). Treatment with riluzole exhibited a trend toward reduction of aberrant Ca²⁺ release relative to myocytes solely exposed to 4AP-ISO and those concomitantly exposed to 4AP-ISO and lidocaine (P = 0.2). In line with these observations, addition of 4,9-ah-TTX and riluzole resulted in a significant increase in SR Ca²⁺ load relative to lidocaine-treated cardiomyocytes (Fig. S2). Collectively, these findings strongly suggest that nNa_v blockade, and not necessarily the extent of I_Na blockade, contributes to attenuation of aberrant Ca²⁺ handling evidenced during concomitant prolonged repolarization and exposure to β-adrenergic stimulation.

Slowed repolarization and the corresponding triggered arrhythmias have been attributed to prolongation of cardiac APD (Wang and Duff, 1997). Specifically, prolonged APD elevates the risk of EADs (Varró and Baczkó, 2011), while at the same time promoting aberrant Ca²⁺ handling that underlies DADs (Noble and Noble, 2006). Therefore, we quantified the effects of 4AP during β-adrenergic stimulation and subsequent Na_v inhibition on APD. 4AP+ISO significantly prolonged APD measured at 50% (APD₅₀) and APD₉₀ repolarization (Fig. 2 a, red trace and bar) relative to control. As expected, the extent of APD₉₀ prolongation correlated with frequency of EADs, but not DADs (Fig. S3). Notably, neither lidocaine (10 µM), 4,9-ah-TTX (300 nM), nor riluzole (10 µM) had a significant effect on APD (Fig. 2 b, green, blue and purple traces and bars). Despite this, Na_v blockade abolished the correlation between APD and EAD frequency (Fig. S3). These findings suggest that although nNa_vs may contribute to triggered arrhythmias, they do not contribute substantially to APD prolongation evidenced during concomitant prolonged repolarization by 4AP and β-adrenergic stimulation.

We investigated the role of nNa_V in triggered arrhythmias induced in a recently established murine model of LQT, where nonselective augmentation of Na⁺ influx with ATX-II caused APD prolongation (Lowe et al., 2012). Because LQT has been linked to altered RYR2 function (Katra and Laurita, 2005; Radwański et al., 2010; Cheng et al., 2011; Radwański and Poelzing, 2011; Terentyev et al., 2014), in our study we used a mouse with dysfunctional RYR2 gating caused by an ablation of a key regulatory protein, CASQ2 (Radwański et al., 2016). These experiments were conducted in the absence of ISO. As expected, ATX-II (1 nM) elicited a substantial late I_Na (Fig. 3, a and b, red trace and bar). To confirm the nonselective nature of ATX-II, we applied 4,9-ah-TTX and riluzole. Accordingly, only a fraction of ATX-II–induced late I_Na was sensitive to selective nNa_v blocker 4,9-ah-TTX (300 nM; Fig. 3, a and b, blue trace and bar). Although blockade with riluzole (10 µM) reduced ATX-II–induced late I_Na to a greater extent than 4,9-ah-TTX, it did not abolish this persistent Na⁺ influx induced by ATX-II (Fig. 3, a and b, purple trace). Functionally, the considerable augmentation of late I_Na by 1 nM ATX-II translated into significant prolongation of APD₅₀ and APD₉₀ (Fig. 4, a and b, red trace and bar). Notably, both 4,9-ah-TTX (300 nM) and riluzole (10 µM) shortened APD₉₀ while having a negligible effect on APD₅₀ (Fig. 4, a and b, blue and purple traces and bars). Analogous to a graded response of ATX-II–induced late I_Na to riluzole and 4,9-ah-TTX (Fig. 3, a and b, blue and purple traces and bars), riluzole abbreviated APD₉₀ to a greater extent than 4,9-ah-TTX (Fig. 4, a and b, purple trace and bar).

Next, we examined the effect of ATX-II–induced APD prolongation on QT interval in vivo. As expected based on cellular data, ATX-II (30 µg/kg) elicited a marked QTc prolongation in CASQ2-null mice (Fig. 5, a and b; and Table S1, red trace and lines). Pretreatment of mice with 4,9-ah-TTX (750 µg/kg), did not significantly affect QTc interval (Fig. 5, a and b; and Table S1, blue trace and lines). In contrast, riluzole (15 mg/kg) significantly abbreviated the QTc interval relative to CASQ2-null mice treated with ATX-II alone (Fig. 5, a and b; and Table S1, purple trace and lines).

Increased late I_Na and APD prolongation are both associated with triggered arrhythmias (Ackerman and Mohler, 2010). Therefore, we quantified the effects of ATX-II (1 nM) and concomitant nNa_v inhibition on aberrant Ca²⁺ waves and EAD and DAD frequency. Late I_Na augmentation by ATX-II (1 nM) induced frequent aberrant Ca²⁺ waves, which coincided with EADs and DADs (Fig. 6, a and b, red traces and bars). Despite a large remaining late I_Na (Fig. 3), both 4,9-ah-TTX (300 nM) and riluzole (10 µM) significantly reduced the frequency of aberrant Ca²⁺ waves and triggered activity in the form of EADs or DADs (Fig. 6, a and b, blue and purple traces and bars). Notably, both 4,9-ah-TTX and riluzole evidenced larger SR Ca²⁺ load relative to control condition (Fig. S4), reflecting Ca²⁺ loading achieved with ATX-II. Furthermore, riluzole evidenced greater Ca²⁺ load relative to 4,9-ah-TTX, suggesting that increased load may parallel greater reduction of cellular arrhythmia burden achieved with riluzole. Collectively, these data suggest that inhibition of nNa_Vs ameliorates triggered arrhythmias during augmentation of global Na⁺ influx with ATX-II.

Mitigation of aberrant Ca²⁺ cycling by nNa_v blockade suggests close proximity between nNa_v and SR Ca²⁺-release machinery. To examine this possibility, we performed confocal immunolocalization experiments in ventricular myocytes isolated from CASQ2-null hearts. Notably, nNa_v (Na_v1.1, Na_v1.3, and Na_v1.6) immunofluorescence (Fig. 7, red) coincided with RYR2 labeling (Fig. 7, green) and exhibited a periodic pattern consistent with T-tubular localization (Fig. 7, yellow; Maier et al., 2004; Noujaim et al., 2012; Radwański et al., 2015, 2016). These results suggest that nNa_vs colocalize with RYR2 in the same subcellular regions in the CASQ2-null mouse ventricle, and in part may explain the nNa_v contribution to aberrant Ca²⁺ handling.

We investigated the role of nNa_V in triggered arrhythmias induced in a model of LQT through augmentation of L-type Ca²⁺ channel activity. Addition of BayK (500 nM) to cardiomyocytes isolated from CASQ2-null mice induced frequent aberrant Ca²⁺ release and the accompanying EADs and DADs (Fig. 8, a and b, orange traces and bars). Next, we examined the contribution of nNa_Vs to BayK-promoted cellular triggered activity. Addition of 4,9-ah-TTX (300 nM) significantly reduced the frequency of aberrant Ca²⁺ waves and triggered activity (Fig. 8, a and b, blue traces and bars). Analogous to 4,9-ah-TTX, riluzole (10 µM) significantly reduced aberrant Ca²⁺ handling and triggered activity in BayK-exposed cardiomyocytes (Fig. 8, a and b, purple traces and bars). Notably, BayK-treated CASQ2-null cardiomyocytes exposed to riluzole evidenced greater SR Ca²⁺ content as assessed by rapid application of caffeine relative to control or those exposed to 4,9-ah-TTX (Fig. S5). Collectively, these data suggest that nNa_vs contribute to aberrant Ca²⁺ release and triggered activity during augmentation of L-type Ca²⁺ channel activity.

We quantified the effects of BayK (500 nM) and subsequent nNa_v inhibition on APD. Augmentation of L-type Ca²⁺ channel activity with BayK significant prolonged APD₅₀ and APD₉₀ (Fig. 9, a and b, orange trace and bar). Notably, both 4,9-ah-TTX (300 nM) and riluzole (10 µM) did not attenuate the effects of BayK on APD (Fig. 9, a and b, blue and purple traces and bars). In addition, APD prolongation has been demonstrated to underlie LQT interval, which promotes VT (El-Sherif et al., 2017). Here we examined whether APD prolongation associated with augmentation of L-type Ca²⁺ channel activity can give rise to LQT and whether nNa_V blockade can reverse this phenotype. Augmentation of L-type Ca²⁺ channel activity with BayK (1 mg/kg) elicited marked QTc prolongation in CASQ2-null mice (Fig. 10, a and b, orange trace and lines; and Table S1). As expected, pretreatment with neither 4,9-ah-TTX (750 µg/kg) nor riluzole (15 mg/kg) significantly affected QTc interval (Fig. 10, a and b, blue and purple traces and lines; and Table S1), thereby supporting the notion that nNa_V inhibition is insufficient to attenuate prolonged APD and QT interval elicited by BayK.

Last, we assessed whether inhibition of nNa_v is able to alleviate LQT-promoted VT. Because mice with dysfunctional RYR2 gating caused by an ablation of CASQ2 do not evidence in vivo arrhythmias without catecholamines (Radwański et al., 2016), we exposed them to a catecholamine challenge composed of epinephrine (1.5 mg/kg) and caffeine (120 mg/kg). As previously described (Radwański et al., 2016), catecholamine challenge induced bidirectional, narrow QRS complex VT with beat-to-beat alternations in the polarity of QRS, characteristic of CPVT, in 20% of animals tested (Fig. 11, a and b, black trace and bar). None of these animals evidenced the wide QRS complex polymorphic VT often associated with LQT (Lowe et al., 2012). On the other hand, BayK induced QT prolongation upon catecholamine challenge, which elicited sustained VTs in >70% of animals tested (Fig. 11 b, orange bar). Importantly, a third of these animals evidenced polymorphic VT similar to that shown in Fig. 11 (orange ECG), with a typical short-long-short initiating ventricular cycle leading to a wide (>29 ms) QRS complex polymorphic VT that resembled human TdP (Fig. 11, a and c, orange trace and bar). Of note, this TdP-like VT was absent from animals pretreated with nNa_vs, whether 4,9-ah-TTX (750 µg/kg) or riluzole (15 mg/kg; Fig. 11, a and c, blue trace and green bars, respectively), whereas this strategy reduced the overall VT incidence by >50% (Fig. 11 b, green bar). To further investigate whether inhibition of nNa_vs can reduce TdP-like VT during global augmentation of Na⁺ influx, we pretreated animals with ATX-II (30 µg/kg; Fig. 11 a, red trace). Recently, Lowe et al. (2012) demonstrated that disordered Na_v function leading to TdP is readily replicated in mice with ATX-II. This LQT prolongation strategy during catecholamine challange induced VT in >70% of animals tested (Fig. 11 b, red bar), and TdP-like VT was observed in more than half of the subjects (Fig. 11 c, red bar). Consistent with observations in isolated cardiomyocytes, ATX-II–promoted TdP-like VT was sensitive to nNa_v inhibition, as pretreatment with 4,9-ah-TTX or riluzole reduced TdP-like VT by half and overall VT by 10-fold (Fig. 11, b and c, green bars). Collectively, these data highlight that targeting nNa_vs can be a common antiarrhythmic therapy for LQT-CPVT overlap phenotypes.

Cardiac arrhythmias are often induced by ectopic triggered activity. Two mechanisms for triggered arrhythmias have been proposed. The first argues that prolonged APD facilitates EAD formation and is thought to underlie reentrant TdP in LQT syndromes (Varró and Baczkó, 2011). The second hinges on aberrant Ca²⁺ release through dysfunctional RYR2, which in turn activates depolarizing membrane currents, triggering DADs and repetitive bifocal or multifocal aberrant depolarizations that underlie CPVT (Cerrone et al., 2007; Belevych et al., 2013). Notably, triggered arrhythmias, independent of etiology, have been reported to be sensitive to nNa_v blockade (Radwański et al., 2013, 2015, 2016), suggesting that nNa_v-mediated Na⁺ influx may serve as a common trigger to both types of triggered activity. Here, we demonstrate that nNa_Vs play a role in the generation of aberrant Ca²⁺ release that contributes to EADs and DADs during loss of repolarization reserve (block of repolarizing current with 4AP) and APD prolongation. Furthermore, we demonstrate that, during augmentation of L-type Ca²⁺ or Na⁺ channel function, nNa_vs contribute to triggered activity and, consequently, to the genesis of both narrow-QRS VT, associated with genetic defects in RYR2, and LQT-associated, wide-QRS, TdP-like VT.

Despite the initial perceived mechanistic divergence between the two types of triggered arrhythmias, emerging evidence suggests a link between pathologies directly driven by Ca²⁺ cycling defects and those arising from QT prolongation. Abnormal RYR2 function (Fig. 12) and the resultant aberrant Ca²⁺ release have been linked to both types of triggered arrhythmias (Volders et al., 1997; Katra and Laurita, 2005; Radwański et al., 2010; Cheng et al., 2011; Radwański and Poelzing, 2011; Terentyev et al., 2014). However, abnormal RYR2 function by itself is insufficient to induce Ca²⁺ leak and triggered activity in vivo (Györke, 2009; Radwański et al., 2016). In the case of genetic defects in RYR2, augmentation of Na⁺ entry via nNa_v and enhancement of SR Ca²⁺ refill, secondary to β-adrenergic receptor stimulation by catecholamines, are necessary contributors to arrhythmogenesis (Fig. 12; Radwański et al., 2016). Accordingly, in the absence of catecholamines, triggered activity was infrequent in cardiomyocytes isolated from hearts with compromised RYR2 function caused by ablation of CASQ2 (Figs. 6 and 8, black traces). On the other hand, pharmacologically induced LQT by either augmentation of L-type Ca²⁺ channels with BayK (Mika et al., 2015) or augmentation of Na⁺ channels with ATX-II (Viatchenko-Karpinski et al., 2014) has been linked to activation Ca²⁺/CaM-dependent protein kinase II, which in turn renders RYR2 dysfunctional (Fig. 12). Against this backdrop of altered RYR2 function and increased intracellular Ca²⁺ loading, a disorder marked by prolonged APD and LQT can facilitate aberrant Ca²⁺ release and triggered activity (Figs. 1, 6, and 8, red, orange, and red traces, respectively). Notably, despite minor effects on pharmacologically elicited APD and QT prolongation (Figs. 2, 4, and 9, red, orange, and red traces, respectively), inhibition of nNa_vs effectively suppressed aberrant Ca²⁺ release and the accompanying triggered activity. These data are consistent with nNa_vs contributing to arrhythmogenesis associated with LQT and LQT-CPVT overlap phenotypes (Fig. 12). Furthermore, overall modulation of Na⁺ current with lidocaine (Figs. 1 and S1) or ATX-II (Fig. 6), did not completely account for modulation of cellular arrhythmias. It is important to note here that the ATX-II–treated cardiomyocytes evidenced a greater degree of APD prolongation accompanied with a greater EAD burden relative to BayK-treated myocytes, which exhibited more DADs. Given that treatment with either 4,9-ah-TTX or riluzole during exposure to ATX-II increased SR Ca²⁺ load (Fig. S4), this difference may reflect greater intracellular Ca²⁺ loading with ATX-II.

What differentiates CPVT and LQT-promoted VTs in our model? Consistent with previous results (Knollmann et al., 2006; Radwański et al., 2016), catecholamine challenge induced narrow-QRS complex polymorphic VT, often with QRS polarity alternating beat to beat. This reflects a bifocal or multifocal nature of type of arrhythmias, which stems from Ca²⁺-mediated triggered activity at multiple myocardial sites (Cerrone et al., 2007). On the other hand, in LQT induced by augmentation of L-type Ca²⁺ channel or Na_v activity, catecholamine challenge often resulted in wide-QRS polymorphic VTs that resembled human TdP (Lowe et al., 2012). Importantly, nNa_v blockade not only abolished these wide-QRS complex VTs, but also reduced the overall VT incidence during combined CPVT and LQT (Fig. 11 c). Thus, our results suggest that nNa_vs may be an integral part of the arrhythmogenic trigger in both CPVT and LQT.

How do nNa_Vs contribute to triggered activity? It is well established that nNa_Vs are localized within transverse tubules and along the cell periphery in both WT (Maier et al., 2002, 2004; Lin et al., 2011; Noujaim et al., 2012; Westenbroek et al., 2013) and CPVT (Radwański et al., 2015, 2016) mice. We recently demonstrated that nNa_Vs, along with NCX and RYR2, are organized into nanodomains along the transverse tubule, providing a structural basis for the functional interplay between cardiac Na⁺ and Ca²⁺ handling (Radwański et al., 2016). Therefore, the results from the present study in WT and CASQ2-null cardiomyocytes suggest that the impact of a specific Na_V isoform on arrhythmogenesis may depend on its localization relative to Ca²⁺ cycling proteins (Fig. 7).

Viewed in this structural context, Na⁺ influx via nNa_v may occur preferentially near Ca²⁺-cycling proteins (RYR2 and NCX), enhancing NCX-dependent local Ca²⁺ accumulation in the space between the sarcolemma and the RYR2. This nanodomain Ca²⁺ accumulation may prompt Ca²⁺ release via RYR2, resulting in arrhythmogenic triggered activity (Radwański et al., 2015, 2016; Veeraraghavan et al., 2017). The evidence that the antiarrhythmic effects of nNa_V inhibition result from prevention of aberrant Ca²⁺ release is compelling; however, possible direct effects on the triggered activity threshold should also be considered (Fig. S1). Regardless, our results provide impetus for future work correlating the nanoscale structural organization of specific nNa_V isoforms relative to Ca²⁺ cycling proteins with their functional roles in health and disease.

Our results suggest that nNa_vs contribute to the substrate for LQT-associated fatal polymorphic VT. Further, our results suggest that this is based on modulation of Ca²⁺ cycling by nNa_Vs rather than via global APD and QT prolongation, the classic substrates for reentrant VTs. However, nNa_vs may contribute to regional heterogeneities in late I_Na (Sakmann et al., 2000) and, consequently, heterogeneous APD prolongation, in turn precipitating reentrant arrhythmias such as TdP (El-Sherif et al., 2017). This intriguing hypothesis merits further investigation. In summary, we demonstrate the antiarrhythmic impact of nNa_v inhibition in LQT and CPVT-LQT overlap phenotypes and provide a basis for a mechanistically driven antiarrhythmic strategy using nNa_V-selective inhibitors.

This work was supported by National Institutes of Health grants R01-HL074045, R01-HL063043, and R01-HL138579 (to S. Györke) and R00-HL127299 (to P.B. Radwański).

The authors declare no competing financial interests.

Author contributions: S. Györke and P.B. Radwański conceived/designed the work; M. Koleske, I. Bonilla, J. Thomas, N. Zaman, S. Baine, and P.B. Radwański performed research; B.C. Knollmann contributed the murine transgenic model; M. Koleske, N. Zaman, R. Veeraraghavan, and P.B. Radwański analyzed data; M. Koleske, R. Veeraraghavan, S. Györke, and P.B. Radwański drafted the work or revised it critically for important intellectual content.

Eduardo Ríos served as editor.