The SERCA2 knock out mouse (KO) has previously been described and studied in detail as a model of contractile failure (Andersson et al., 2009; Liu et al., 2011; Li et al., 2012; Swift et al., 2012; Land et al., 2013). Briefly, cardiac-specific SERCA2 KO excision was achieved by tamoxifen activation of Cre-recombinase (α-MHC driven MerCreMer) via a single tamoxifen injection at 8–12 weeks of age. Here we studied these animals near to their mean age of death at 7 weeks after tamoxifen injection, when they exhibit pronounced contractile dysfunction, and markedly reduced cardiac output (Andersson et al., 2009). Age-matched flox/flox mice (FF) were used as controls.

Telemetry transmitters (Physiotel ETA-F10, Data Sciences International, St. Paul, MN, United States) were used to record electrocardiograms in freely moving animals. Implantation was performed 6 weeks after tamoxifen injection, and as previously described (Manotheepan et al., 2016). Intraperitoneal Xylazine hydrochloride and ketamine hydrochloride were administered in combination with isoflurane inhalation (1–4%) during the procedure, and subcutaneous buprenorphine was given toward the end of the procedure for postoperative analgesia. The transmitters were placed subcutaneously in the dorsal thoracolumbar region, and stabilized by ligatures to the dorsal muscles. Leads were attached to the pectoral muscles by ligatures in the upper left and lower right pectoral regions. Electrocardiograms were recorded after 7 days recovery from surgery, and recordings were analyzed for heart rate as well as ECG parameters during baseline conditions, and after a subsequent intraperitoneal injection of adrenalin (Epi, 0.5mg/kg). Analysis was performed manually and in fully-blinded fashion by the same experienced technician using both proprietary software (Matlab version 2013b, The Mathworks, Natick, MA, United States), and Ponemah (Data Sciences International, St. Paul, MN, United States). Steady state ECG analyses were performed by applying conventional definitions for all intervals. Of note, because the mouse often exhibits a pronounced J-wave we have defined QRS width (duration) as the time from first negative deflection after the P-wave to the first time after the S-wave minimum at which the ascending voltage signal crosses the isoelectric line. Rate dependent QT correction was performed by the Bazzet formula:

Premature ventricular complexes (PVCs) were manually identified, and episodes of ventricular tachycardia (VT) were defined as 4 or more PVCs occurring in sequence. Runs of VT were defined as non-sustained (NSVT) if they lasted less than 20-s, and as sustained (SVT) if longer.

Cardiomyocytes were isolated from ventricles of FF and KO mice similarly to our prior studies (Ottesen et al., 2015; Manotheepan et al., 2016). Briefly, mice were isoflurane-anesthetized (98% oxygen, 2.0% isoflurane), and euthanized by cervical dislocation. Excised hearts were first rinsed in ice-cold isolation buffer containing (mM): 130 NaCl, 25 Hepes, 22 D-glucose, 5.4 KCl, 0.5 MgCl₂, 0.4 NaH₂PO4 (pH 7.4). Each heart was then retrograde-perfused under constant flow conditions (3 ml/min) with warmed (37°C) isolation buffer for 4 min, and then with the same buffer supplemented with 400 U/ml collagenase Type II (Worthington Biochemical Corporation, Lakewood, NJ, United States) and 0.015 mmol/L Ca²⁺. After 10 min of enzyme perfusion, hearts were cut down and the left ventricle was removed, diced, and triturated in collagenase-free isolation buffer including 1% BSA and 0.02 U/L deoxyribonuclease I (Worthington), again at 37°C. The resulting cell suspension was filtered (200 μm nylon mesh) and sedimented, the cell pellet was washed in isolation buffer supplemented with 1% BSA, and Ca²⁺ was progressively reintroduced (0.05, 0.1, 0.2, and 0.5 mmol/L). The Ca²⁺-tolerant cardiomyocytes were stored at room temperature until use, which occurred within 8 h of isolation.

Single rod-shaped cardiomyocytes with clear striations were patch-clamped in whole-cell configuration using Axoclamp 2A and 2B amplifiers. Borosilicate patch-pipettes had resistances of 2–3 MOhms with the corresponding internal and external solutions. We used identical solutions for current clamp AP recordings and end-pulse K⁺ currents in voltage clamp, where the internal solution contained (in mM: 120 K-Asp, 25 KCl, 0.5 MgCl₂, 6 NaCl, 4 K₂-ATP, 0.06 EGTA, 10 HEPES, and 10 D-Glucose, pH corrected to 7.2 with KOH), and external (in mM): NaCl 140, HEPES 5, KCl 5.4, MgCl₂⋅6H₂O 0.5, Glucose 5.5, NaH₂PO₄H₂O 0.4, CaCl₂ 1. For voltage clamp recordings of the transient outward potassium current (I_to), the internal solution contained (in mM) 110 K-Asp, 20 KCl, 0.5 MgCl₂, 4 K₂-ATP, 5 EGTA, 5 HEPES, and 10 D-Glucose - pH 7.2 with KOH, with a Na⁺ and Ca²⁺ free external solution (mM): 140 CholineCl, 1 CdCl₂, 0.5 MgCl₂, 5 HEPES, 5.5 Glucose, 5.4 KCl, pH 7.2 with KOH. For I_Ca,L the internal solution was (mM) 130 CsCl, 0.33 MgCl₂, 4 Mg-ATP, 0.06 EGTA, 10 HEPES and 20 tetraethylammonium chloride - pH 7.2 with CsOH, and the bathing solution included (mM) 135 NaCl, 20 CsCl, 1 MgCl₂, 10 glucose, 10 HEPES, 1 CaCl₂, and 4 4-AP (pH 7.4).

Cells were superfused with external solution in all experiments, and all current measurements were made during step-pulse protocols at 1 Hz. Following ten 50-ms conditioning pulses (also 1 Hz) to 0 mV from a holding potential of −70 mV, I_CaL was elicited by a 200-ms depolarizing voltage step (Mørk et al., 2009). Test potentials ranged from −40 to +50 mV with 10-mV increments from a post-train holding potential of −40 mV (to inactivate I_Na). Current amplitude was measured as the difference between peak inward current and steady-state current at the end of the test pulse. I_to (peak I_K) was elicited by 300-ms steps to test potentials between −40 and +60 mV in 10-mV increments from a holding potential of −70 mV (Louch et al., 2010a). Peak I_K was calculated as the peak outward current less the steady-state end pulse current from these 300-ms steps. The pedestal or plateau component of I_K (I_Kp) was calculated as the mean of the final 20-ms of 500-ms test-pulse steps between −100 and +60 mV, again at 10 mV increments from a –70 mV holding potential. All currents were normalized to capacitance, which was calculated as the integral of the transient current response to a 10-mV hyperpolarizing step (150 ms) from a holding potential of −70 mV.

Action potential waveforms were also recorded at 1 Hz in all experimental conditions, and at least 20 sequential beats were allowed to achieve steady state. In experiments assessing EAD and DAD susceptibility, Iso-containing superfusate (1 μM) was applied after 30-s of baseline recording, and maintained for at least 3 min further.

The computational models used here are modifications of those published by Li et al. (2012). Briefly, both the FF and KO models were constructed on the basis of extensive data collected in the FF and KO mice at the same age and time after SERCA gene excision as studied here. Our modifications here were made sequentially to establish the requirements for the observed AP changes accompanying SERCA-KO. Thus, we describe them in that sequence in the results section. All parameter changes and details for code access are provided in the Supplementary Material.

Steady state electrocardiographic parameters were assessed by repeated measures ANOVA (RMANOVA: Genotype × Epi) with repeated measures for Epi. ECG arrhythmia outcomes (PVC and NSVT frequency) were non-normally distributed, and interrogated by Kruskal-Wallis rank sum test after being separated into baseline and Epi recordings. Similarly, skewed data for EAD frequency were assessed via the Kruskal-Wallis test, and the Fisher exact test was used to determine differences in EAD incidence. Peak I_K, I_Kp, I_CaL, and AP durations at 20, 50, 70, and 90% repolarization were all tested by RMANOVA where step potential was repeated for current measurements, and treatment conditions (Ca²⁺ concentration, Nifedipine, or Isoproterenol) were repeated for AP recordings.

For all RMANOVAs, the Holm-Sidak test was used to identify pairwise effects post hoc. In the event that the sphericity assumption was not valid, RMANOVA was replaced by the Mann-Whitney-Wilcoxon test. Significant effects were defined at p < 0.05, and p-values are explicitly stated for all marginal results (0.05 ≤ p ≤ 0.1). All statistical analyses were performed either with SigmaPlot (Systat Software Inc., CA, United States), or R software (version 3.2.1, The R Foundation of Statistical Computing).

Seven weeks after gene excision, KO mice exhibit a range of electrophysiologic dysfunction as measured by telemetered ECG. The most pronounced effects are slowed ventricular activation observed as increased QRS width (p < 0.01, Figure 1A, left), and a much more prominent and positive-going T-wave than is normally measurable via standard lead II recordings in the mouse (p < 0.0001), particularly during epinephrine challenge (Figures 1B,C). Interestingly, these changes exist in the absence of significant rate-corrected QT prolongation (p = 0.07, Figure 1A, right), thus suggesting that terminal repolarization is not delayed in the KO mice. These outcomes will be discussed in more detail later, but are consistent with a markedly altered AP morphology in the KO myocytes.

The combined influence of these electrocardiographic abnormalities did not result in a higher frequency of either isolated PVCs or runs of non-sustained (or sustained) VT (p > 0.1). In fact, during Epi challenge we observed reduced frequency of PVCs (p < 0.05) and a tendency for reduced NSVT frequency in the KO group (p = 0.09, Figure 1D). Together, these observations suggest that while the KO mice exhibit clear disturbances to global cardiac electrophysiology, they remain resilient to arrhythmia.

Figure 2 shows the effect of SERCA2 loss on the cardiomyocyte AP and underlying currents. At the level of the AP (Figure 2A), the outstanding feature is a brief early plateau in KO myocytes, which is reminiscent of the phase 2 plateau in large mammals albeit much shorter (∼20 ms). Mechanistically, this plateau indicates that the balance of currents in early repolarization is shifted inwardly, particularly in the region between +20 and −20 mV. This effect was pronounced, and can be seen as a 273% longer APD₅₀ and 230% longer APD₇₀ in KO myocytes (both p < 0.01). Panels B and C show that this effect is due to modest potentiation of peak I_CaL (39% increase at 0 mV, p < 0.001), and dramatically slowed I_CaL inactivation (50% relaxation time at 0 mV is increased by 178% in KO, p < 0.001), which together result in a 200% increase in Ca²⁺ influx, measured as the I_CaL integral during the square-pulse voltage clamp protocol (p < 0.001). Exacerbating this gain in I_CaL was a moderate reduction of end-pulse potassium current (I_Kp) at positive potentials (p < 0.05). While we did not attempt to experimentally dissect the different components of this current, we used a computational model to assess the ability of those components to contribute to the observed changes in AP morphology. These analyses are described in further detail below.

In Figure 3 we elaborate on the role of I_CaL in slowing early repolarization either by acutely changing superfusate Ca²⁺ concentration, or applying I_CaL blockade by Nifedipine (Nif, 1 μM). Figure 3A shows that the difference in APD₅₀ and APD₇₀ is removed when Ca²⁺ is excluded from the bath, and slightly exaggerated when it is increased to 1.8 mM (both p < 0.05). These alterations in extracellular [Ca²⁺] did not have statistically discernible effects on the kinetics of early repolarization in FF myocytes (p > 0.1). Because field-screening effects and non LCC-specific Ca fluxes²⁺ (such as Na⁺/Ca²⁺ exchange) will be altered by modulation of extracellular Ca²⁺, we also used Nifedipine to pharmacologically antagonize I_CaL (Figure 3C). As for removal of extracellular Ca²⁺, this maneuver eliminated virtually all of the delay in early repolarization present in the KO myocytes – APD₅₀ and APD₇₀ were almost completely normalized (both p < 0.01 vs. control superfusion). This suggests that inward I_CaL is the major contributor to the small phase 2 plateau in KO cells. Finally, to determine whether maneuvers that ordinarily modulate Ca²⁺-dependent I_CaL inactivation (via SR Ca²⁺ release) are ineffective in KO mice, we also assessed the frequency-dependence of peak I_CaL and I_CaL inactivation (Supplementary Figure 1). In moving from 0.1 to 1 Hz FF animals exhibit slowed I_CaL inactivation due to well-known Ca²⁺-dependent facilitation, and this property is dependent on cytosolic Ca²⁺ (Fauconnier et al., 2003). In KO animals this frequency-dependent modulation of I_CaL inactivation is completely absent (given the already very slow inactivation in KO myocytes), further indicating the loss of SR-dependent I_CaL control in KO animals.

Importantly, this shift in balance during early repolarization is not mirrored by slowed terminal repolarization. That is, the differences present at APD₅₀ and APD₇₀ are lost by 90% (APD₉₀) repolarization. The dominant inward currents modulating this late phase of repolarization in the mouse are I_NaCa and recovering I_Na (Edwards et al., 2014; Morotti et al., 2014). As described further below these inward currents compete with several components of I_K, particularly the inward rectifier K⁺ current (I_K1) to shape terminal repolarization. While we have not directly assessed the balance of these currents during late repolarization in KO myocytes, the near complete absence of triggered Ca²⁺ release (Andersson et al., 2009) drastically reduces the potential for inward I_NaCa. The slower early repolarization would also be expected to limit I_Na recovery and the potential for I_Na reactivation.

In larger mammals, the increases in I_CaL peak current and 50% relaxation time would be expected to promote repolarization instabilities and EADs. To assess EAD susceptibility we challenged KO and FF myocytes with Isoproterenol (1 μM) for at least 3 min while pacing at 1 Hz in current clamp, and defined EADs as any upward deflection in the voltage signal exceeding 3 mV. Figure 4A shows that under these conditions EAD incidence and frequency were not increased in KO cells (both p > 0.1), rather EAD frequency tended to be higher (albeit non-significantly, p > 0.05) in the FF (9.3 ± 4.7%) than KO group (2.1 ± 1.8%). In FF cells, EADs exhibited signature properties identified in previous work (Sato et al., 2009; Tran et al., 2009). These features included periods with very high APD variability, indicating variable timing of terminal repolarization, but also intermittent beats that did not exhibit EADs. Furthermore, analysis of EAD take-off potentials (Figure 4B) suggests that EADs in the FF group exhibited very similar underlying dynamics to those we have described previously in the mouse, and different to those in larger mammals (Edwards et al., 2014). That is, they initiated at potentials too negative (−55 ± 7 mV) to be carried by I_CaL, and occurred too soon after stimulation (34.6 ± 2.7 ms; approximately coinciding with the peak of the bulk cytosolic Ca²⁺ transient) to result from subcellular Ca²⁺ waves. Thus, it is very likely that EADs in FF myocytes were carried almost entirely by non-equilibrium reactivation of the fast component of the sodium current (I_Na) (Edwards et al., 2014). In contrast, the EADs present in KO myocytes initiated at more positive take-off potentials (−41.8 ± 2 mV, p < 0.001), were much larger in amplitude (KO = 26.4 ± 9 mV vs. FF = 7.8 ± 2.6 mV; p < 0.001), and lasted much longer than EADs in the FF myocytes (KO = 124 ± 24 ms vs. FF = 55 ± 27 ms; p < 0.001). While these EADs initiated later than those in FF cells, because spontaneous SR Ca²⁺ release in KO cells is virtually negligible (Supplementary Figure 2), it is very unlikely that discoordinated SR Ca²⁺ release made any contribution to these events. Even after accounting for leftward shifts in I_CaL activation due to β-adrenergic activation (Bean et al., 1984; Tiaho et al., 1991; Kumari et al., 2018), the −40 mV take-off potentials are still only just approaching the lower limit of the I_CaL activation range, thus the ability for I_CaL to have directly contributed to initiation of these EADs is somewhat limited. However, once initiated, the large amplitude excursions of these EADs (see for example Figure 4B bottom left) suggest that I_CaL reactivation is pronounced and likely to be the dominant contributor to EAD dynamics, as it is in ventricular myocytes of large mammals (Weiss et al., 2010).

In considering the balance of currents shaping the trajectory of repolarization in the mouse, an important quantitative aspect is how rapid early repolarization favors I_CaL deactivation and limits the opportunity for Ca²⁺-dependent I_CaL inactivation, which is prominent in larger species. Combining this with the known redundancy and different kinetic characteristics among the various K⁺ currents active in this early phase of repolarization, it becomes much more difficult to predict or account for how changes in I_CaL regulation can contribute to modulating early repolarization. To interrogate these dynamics, we employed a published computational model of the mouse cardiomyocyte, which is specifically parameterized to incorporate the reduced SR Ca²⁺ handling and altered Na⁺ balance in failing KO myocytes [16]. This model already incorporates the increased Cav1.2 and NCX1 expression in KO cells, which leads to exaggerated peak I_CaL and I_NaCa. However, it does not fully capture the pronounced slowing of I_CaL inactivation we have observed in KO myocytes at 7 weeks of age, and incorporating this characteristic was our first alteration. Figure 5A shows the behavior of the reparameterized I_CaL model in square-pulse voltage-clamp protocols identical to our experiments (all parameter changes are provided in full in Supplementary Table 1). Even for its larger peak current and much slower inactivation, this I_CaL model was still overwhelmed by the fast component of the transient outward current (I_to,f) during early repolarization in both the FF and KO models, and had relatively little impact on the trajectory of early repolarization (APD₂₀, Figure 5C).

As mentioned above, KO myocytes also exhibit a reduction in the measurable total I_K at the end of our 500 ms test steps. The molecular identity of K⁺ channels contributing to this slowly inactivating current are not fully reconciled, although it is clear that the slow component of the transient outward current (carried by the Kv1.4 alpha subunit) is minimally expressed in mice, and that currents carried by Kv1.5 and Kv2.1 make important contributions (Nerbonne, 2014). Together these channels carry the slowly inactivating current (I_Kslow), also commonly referred to as the ultrarapidly activating delayed rectifier (I_Kur) K⁺ current. I_Kslow is a dominant contributor to the end-pulse K⁺ current in mice. It combines with the steady state K⁺ current (I_Kss, primarily carried by TASK1 and TREK1 channels (Nerbonne, 2014), and I_K1 at more negative potentials, to comprise the compound end-pulse I_K that we term the “pedestal” or “plateau” I_Kp. Due to difficulties in separating the activation kinetics of the Kv1.5 and Kv2.1 contributions to I_Kslow in cardiac cells, most models have assumed that this current is a single functional entity, with very rapid activation and slow inactivation. One relatively recent model separated the currents for the purpose of implementing differing phosphoregulation and inactivation kinetics (Morotti et al., 2014). To simultaneously fit intermediate AP prolongation (APD₅₀ and APD₇₀) and measured end pulse I_K with the KO model, we also had to employ the approach of Morotti et al. (2014) and separate I_Kslow into Kv1.5-(I_Kslow,1) and Kv2.1-(I_Kslow,2) specific components. However, unlike Morotti et al. (2014) we implemented slower activation kinetics for I_Kslow,2, as shown in Figure 5B, and which can be measured for Kv2.1 in heterologous systems (Gordon et al., 2006). These slower activation kinetics of Kv2.1 slowed the onset of I_Kslow enough to permit I_CaL modulation of intermediate repolarization, while also matching end-pulse I_K and permitting stable overall repolarization, as observed in our experiments.

We employed mathematical models specific to the KO and FF myocytes (Li et al., 2012) and challenged them with a simple model implementation of β-adrenergic stimulation, which is closely analogous to our experimental challenge. This strategy involves implementing established effects of β-adrenergic regulation at I_CaL (2–3 fold increased whole-cell permeability, P_CaL, and 5–10 mV left-shifted steady state activation), SERCA (50–60% reduction in the K_m for cytosolic Ca²⁺ binding), the Na⁺-K⁺ ATPase (25–30% reduction in the K_m for cytosolic Na⁺ binding), and I_Kur (15–20% increase in whole cell conductance) (Edwards et al., 2014). The steady-state baseline models for the FF and KO myocytes were each simulated for 3 min at 1 Hz after applying these β-adrenergic parameter changes. Via this approach, EADs could be initiated in both the FF and KO models with identical changes to these key parameters, which remained wholly within the physiologic range. The EADs that result share several of the key features of EADs observed in our experiments during saturating Isoproterenol challenge. First the KO EADs exhibit greater I_CaL reactivation, and their dynamical characteristics (oscillation period and amplitude) are increased similarly to the experimental recordings (Figure 4B). These dynamic characteristics are similar to those of I_CaL-dominated EADs in large mammals (Sato et al., 2010). However, importantly, and somewhat surprisingly, even though the model EADs occurred at relatively positive potentials (approximately −38 mV) and relatively late (80 to 120 ms after the AP stimulus), they were initiated in both models by non-equilibrium reactivation of I_Na (Edwards et al., 2014) rather than I_CaL (Figure 6C). This suggests that even for EADs that initiate 120 ms after the AP peak, recovery from inactivation of I_Na is sufficient to permit an arrhythmogenic current reactivation.