Abstract
Key points
Augmented inositol 1,4,5‐trisphosphate (IP3) receptor (IP3R2) expression has been linked to a variety of cardiac pathologies.
Although cardiac IP3R2 function has been in the focus of research for some time, a detailed understanding of its potential role in ventricular myocyte excitation–contraction coupling under pathophysiological conditions remains elusive.
The present study focuses on mechanisms of IP3R2‐mediated sarcoplasmic reticulum (SR)‐Ca2+ release in ventricular excitation–contraction coupling under IP3R2‐overexpressing conditions by studying intracellular Ca2+ events.
We report that, upon IP3R2 overexpression in ventricular myocytes, IP3‐induced Ca2+ release (IP3ICR) modulates the SR‐Ca2+ content via “eventless” SR‐Ca2+ release, affecting the global SR‐Ca2+ leak. Thus, IP3R2 activation could act as a SR‐Ca2+ gateway mechanism to escape ominous SR‐Ca2+ overload.
Our approach unmasks a so far unrecognized mechanism by which “eventless” IP3ICR plays a protective role against ventricular Ca2+‐dependent arrhythmogenicity.
Abstract
Augmented inositol 1,4,5‐trisphosphate (IP3) receptor (IP3R2) function has been linked to a variety of cardiac pathologies including cardiac arrhythmias. The functional role of IP3‐induced Ca2+ release (IP3ICR) within ventricular excitation–contraction coupling (ECC) remains elusive. As part of pathophysiological cellular remodelling, IP3R2s are overexpressed and have been repeatedly linked to enhanced Ca2+‐dependent arrhythmogenicity. In this study we test the hypothesis that an opposite scenario might be plausible in which IP3ICR is part of an ECC protecting mechanism, resulting in a Ca2+‐dependent anti‐arrhythmogenic response on the cellular scale. IP3R2 activation was triggered via endothelin‐1 or IP3‐salt application in single ventricular myocytes from a cardiac‐specific IP3R type 2 overexpressing mouse model. Upon IP3R2 overexpression, IP3R activation reduced Ca2+‐wave occurrence (46 vs. 21.72%; P < 0.001) while its block increased SR‐Ca2+ content (∼29.4% 2‐aminoethoxydiphenyl borate, ∼16.4% xestospongin C; P < 0.001), suggesting an active role of IP3ICR in SR‐Ca2+ content regulation and anti‐arrhythmogenic function. Pharmacological separation of ryanodine receptor RyR2 and IP3R2 functions and two‐dimensional Ca2+ event analysis failed to identify local IP3ICR events (Ca2+ puffs). SR‐Ca2+ leak measurements revealed that under pathophysiological conditions, “eventless” SR‐Ca2+ efflux via enhanced IP3ICR maintains the SR‐Ca2+ content below Ca2+ spark threshold, preventing aberrant SR‐Ca2+ release and resulting in a protective mechanism against SR‐Ca2+ overload and arrhythmias. Our results support a so far unrecognized modulatory mechanism in ventricular myocytes working in an anti‐arrhythmogenic fashion.
Keywords: excitation‐contraction coupling, calcium cycling, cardiac arrhythmias, heart failure
Key points
Augmented inositol 1,4,5‐trisphosphate (IP3) receptor (IP3R2) expression has been linked to a variety of cardiac pathologies.
Although cardiac IP3R2 function has been in the focus of research for some time, a detailed understanding of its potential role in ventricular myocyte excitation–contraction coupling under pathophysiological conditions remains elusive.
The present study focuses on mechanisms of IP3R2‐mediated sarcoplasmic reticulum (SR)‐Ca2+ release in ventricular excitation–contraction coupling under IP3R2‐overexpressing conditions by studying intracellular Ca2+ events.
We report that, upon IP3R2 overexpression in ventricular myocytes, IP3‐induced Ca2+ release (IP3ICR) modulates the SR‐Ca2+ content via “eventless” SR‐Ca2+ release, affecting the global SR‐Ca2+ leak. Thus, IP3R2 activation could act as a SR‐Ca2+ gateway mechanism to escape ominous SR‐Ca2+ overload.
Our approach unmasks a so far unrecognized mechanism by which “eventless” IP3ICR plays a protective role against ventricular Ca2+‐dependent arrhythmogenicity.
Introduction
Sudden cardiac death due to arrhythmia is responsible for 10–30% of deaths among patients with hypertrophic cardiomyopathy (Ullal et al. 2016). In cardiac hypertrophy, alterations in Ca2+‐induced Ca2+ release (CICR) and excitation–contraction coupling (ECC) have been associated with cardiac instability (Gusev et al. 2009). Thus understanding the underlying mechanisms essentially contributes to the development of novel treatment strategies.
Ca2+ release from the sarcoplasmic reticulum (SR) in ventricular cardiomyocytes is mediated by two Ca2+ release channels: the ryanodine receptor type 2 (RyR2) and the inositol 1,4,5‐trisphosphate receptor type 2 (IP3R2; Bers, 2002; Hohendanner et al. 2014). While RyR2s play a central role in ventricular ECC, the contribution of functionally expressed IP3R2s under physiological and pathophysiological conditions is less well understood. In contrast to RyR2s, which are exclusively activated by Ca2+, IP3R2s are sensitive to Ca2+ and IP3. The latter SR‐Ca2+ release mechanism has been established as IP3‐induced Ca2+ release (IP3ICR; Zima & Blatter, 2004; Harzheim et al. 2009). Intracellular IP3 synthesis is triggered by a variety of vaso‐ and cardioactive hormones (e.g. endothelin‐1, angiotensin II) by stimulating G protein‐coupled receptors (GPCRs) that couple to phospholipase C, liberating IP3 from phosphatidylinositol 4,5‐bisphosphate (Proven et al. 2006; Berridge, 2009).
Although the ratio of functionally expressed RyR2s versus IP3R2s is approximately 100:1, a modulatory function of IP3ICR in ventricular myocyte ECC has been suggested and is the subject of an ongoing controversial discussion (Lipp et al. 2000; Kockskämper et al. 2008). In contrast to the physiological condition, a more pronounced role of IP3ICR in the framework of cardiac pathophysiology seems to be reasonable. It has been found that remodelling of ECC under various cardiac pathologies (e.g. cardiac hypertrophy, ischaemic dilated cardiomyopathy, atrial fibrillation, failing myocardium and hypertension) is associated with IP3R2s overexpression and, in this context, enhanced IP3ICR has been repeatedly linked to arrhythmogenicity (Zhang et al. 2002; Harzheim et al. 2009, 2010; Nakayama et al. 2010; Signore et al. 2013). Supporting evidence comes from the observation that endothelin‐1 and angiotensin II – both GPCR agonists – have similar positive inotropic effects in ventricular myocytes, suggesting a role of IP3ICR in the regulation of cytosolic Ca2+ homeostasis (Zima & Blatter, 2004). In addition, RyR2–IP3R2 local bi‐directional crosstalk mechanisms in cardiomyocytes have been recently described and may be involved in Ca2+‐dependent arrhythmogenicity at the cellular level (Wullschleger et al. 2017). Moreover, these studies support the concept of distinct RyR2‐IP3R2 Ca2+ signalling compartments within cardiomyocytes – either in the cytosol and/or in the nucleus – during pathophysiological signalling driven by IP3 and IP3R2 (Wu & Bers, 2006; Drawnel et al. 2010). In contrast, it has also been suggested that increased synthesis of IP3 as well as IP3R2 expression, may be a common feature of cardiac pathologies but without having functional significance for the ECC mechanism (Cooley et al. 2013).
At the subcellular level, synchronized and functionally clustered RyR2 openings result in “Ca2+ sparks”, the building blocks of global Ca2+ transients in ventricular myocytes; analogously coordinated openings of clustered IP3R2s give rise to “Ca2+ puffs” (Cheng et al. 1993; Yao et al. 1995). Although similar, the distinct spatio‐temporal characteristics of Ca2+ puffs and Ca2+ sparks and their pharmacological profile can be used for classification and separation of SR‐Ca2+ release events (Hohendanner et al. 2015). Openings of singular and/or “misplaced” RyR2s or IP3R2s that do not belong to channel clusters may not generate detectable SR‐Ca2+ release events but can still contribute to the intracellular Ca2+ homeostasis (Lipp & Niggli, 1996; Sobie et al. 2006; Brochet et al. 2011). Analogously to recently described “rogue” RyR2s, “rogue” IP3R2s may generate an “eventless” SR‐Ca2+ release flux which might have a significant impact on the total SR‐Ca2+ leak in ventricular myocytes without giving rise to localized Ca2+ events (Sobie et al. 2006).
Among other factors, the open probability (P o) of both IP3R2s and RyR2s and thus the SR‐Ca2+ leak, is critically controlled by SR luminal Ca2+ (Nunn & Taylor, 1992; Terentyev et al. 2002). Spontaneous increased RyR2 P o under SR‐Ca2+ overload results in asynchronous SR‐Ca2+ release (e.g. Ca2+ waves) and may lead to delayed after‐depolarizations, a major cause of ventricular tachyarrhythmias and sudden cardiac death (Keller et al. 2007; Liu et al. 2013). Although largely outnumbered by RyR2s, overexpression of IP3R2s within cardiac pathology may render the latter receptor a more significant SR‐Ca2+ regulatory function. The present study focuses on potential mechanisms by which IP3R2 may have a Ca2+‐dependent anti‐arrhythmogenic function at the cellular level.
Our data in the present study reveal that stimulation of IP3ICR in freshly isolated ventricular myocytes from a cardiac‐specific IP3R2‐overexpressing mouse model leads to a decrease in asynchronous SR‐Ca2+ release, a decline in the SR‐Ca2+ content and subsequent decrease in pro‐arrhythmic Ca2+ wave occurrence, suggesting a potential anti‐arrhythmic role of IP3ICR. Although under cellular remodelling conditions the SR‐Ca2+ content appears to be under active control of IP3ICR, no additional IP3R2‐mediated local SR‐Ca2+ events (e.g. Ca2+ puffs) were identified. Interestingly, study of the SR‐Ca2+ leak revealed a more prominent IP3R2‐dependent component that controls the SR‐Ca2+ content. Thus, these results support the view that IP3ICR contributes to the total SR‐Ca2+ leak via “eventless” SR‐Ca2+ release. The underlying mechanism actively stabilizes the SR‐Ca2+ content and limits luminal RyR2 sensitization below the critical threshold for spontaneous Ca2+ release thereby offering a protective mechanism against arrhythmias in ventricular myocytes.
Methods
Ethical approval
All experiments were approved by the State Veterinary Office of Bern (Switzerland), in accordance with Swiss Federal Animal Protection Law and were approved by the in‐house ethical committee (Ref. BE96/16, Ref. EFWW0443). For single myocyte isolation, mice were killed by cervical dislocation, hearts were extracted and perfused using the Langendorff perfusion technique. For transthoracic echochardiography measurements, mice were anaesthetized with 1–1.5% isoflurane, maintaining heart rate at 450–500 beats min−1. Following data acquisition, mice were monitored until recovered from anaesthesia.
Cardiac‐specific IP3R2‐overexpressing mouse model
Mouse lines FVB‐Tg(Myh6/tet0‐Itpr2)3.11Jmol/J (The Jackson Laboratory, Bar Harbor, ME, USA) and FVB.CG‐Tg(MyH6‐tTA)6Smbf/J (Jacksons) were crossbred to generate the cardiac‐specific IP3R type 2 (IP3R2) overexpressing mouse model (TG; Nakayama et al. 2010). In the following, wild‐type (WT) will be a synonym for ventricular myocytes isolated from WT mice and transgene (TG) for cells from IP3R2‐overexpressing mice. Genotype was assessed via conventional polymerase chain reaction. Primers used include: for IP3R2 5′‐CTTTCATCGGATGCCTGCCT‐3′ and 5′‐TTGGGAGGCCAAGGTGGGTA‐3′ forward and reverse, respectively; and for tetracycline transactivator (tTA) 5′‐CGCTGTGGGGCATTTTACTTTAG‐3′ and 5′‐CATGTCCAGATCGAAATCGTC‐3′ forward and reverse, respectively. Heart and body weights (mg g−1) were determined at 5 months of age. All mice were fed with doxycycline‐containing food (0.2 g kg−1, Envigo, Füllinsdorf, Switzerland) during pregnancy and until 4 weeks of age.
Echocardiography
Transthoracic echocardiography was performed using a 30 MHz probe and the Vevo 2100 Ultrasound machine (VisualSonics, Bothell, WA, USA). All mice were anaesthetized with 1–1.5% isoflurane. The heart was imaged in 2‐D mode on a parasternal long‐axis view from where an M‐mode cursor was positioned perpendicular to the interventricular septum and the posterior wall of the left ventricle. Echocardiographies were performed at the Cardiovascular Assessment Facility (CAF, Lausanne, Switzerland).
RNA and protein analysis
Animals were killed by cervical dislocation and tissues from ventricle, atria, aortic smooth muscle, femoral skeletal muscle and brain were excised, washed and snap‐frozen in liquid nitrogen. For western blot analysis, ventricular tissue was ground in extraction buffer (1x Protease Inhibitor (04693159001, Sigma‐Aldrich Chemie, Buchs, Switzerland) in NP40 extraction buffer, Invitrogen, Thermo Fisher Scientific, Reinach, Switzerland) for 30 min. Samples were centrifuged at 1400 g for 10 min at 4°C to obtain the supernatant. Following quantification by the bicinchoninic acid assay (BCA) method, 25 μg per lane of protein in loading solution (10% glycerol, 5% β‐mercaptoethanol, 43% SDS, 62.5 mM Tris, 6 M urea) were loaded in 4–20% pre‐cast gel (Bio‐Rad, Basel, Switzerland) and run for 55 min at 200 V. Samples were transferred to PVDF membranes at 100 mA and 4°C overnight. After PBS washing steps, membranes were blocked for 1 h at room temperature (RT) in blocking solution (5% milk in PBS). Membranes were incubated with primary antibodies anti‐IP3R2 (1:5000 KM1083, kindly provided by Prof. Dr K. Mikoshiba; Sugiyama et al. 1994), anti‐RyR2 (1:2000 ab2827, Abcam, Cambridge, UK) or anti‐GAPDH (1:100,000 10R‐G109A, Fitzgerald, Acton, MA, USA) in primary antibody solution (PBS‐Tween, 0.5% milk). Secondary antibody anti‐mouse Alexa 790 (1:100,000, ThermoFisher Scientific, Reinach, Switzerland) was applied in secondary antibody solution (PBS‐Tween, 0.5% milk, 0.01% SDS) for 1 h at RT. Imaging was performed on an Odyssey set‐up and images were analysed using GelAnalyzer. Full western blot images are available upon request.
Total RNA was extracted from snap‐frozen mice tissues with NucleoSpin RNA Plus kit (Macherey‐Nagel, Oensingen, Switzerland) and concentrations were determined with a NanoDrop 2000c (ThermoFisher Scientific). RT‐qPCR was performed by loading 10 ng of sample in an Eco Real‐Time PCR system (Illumina, Cambridge, UK) using KAPA SYBR FAST One‐Step Universal kit (LabGene, Châtel‐Saint‐Denis, Switzerland) in accordance with the manufacturers’ instructions. Primers used include: ITPR2 primer pair (M_Itpr2_3, Sigma) and β‐2‐microglobulin primers (forward FM1_B2m, and reverse RM1_B2m, Sigma). ITPR2 gene relative expression was normalized to reference gene β‐2‐microglobulin. ΔΔCq was calculated and used for statistical analysis. TG data are shown normalized to their respective WT tissue (norm.).
Ventricular myocyte isolation
Animals were killed at 5 months of age by cervical dislocation. Ventricular cardiomyocytes were isolated in Tyrode Ca2+‐free solution with the Langendorff perfusion technique with collagenase type II (Worthington) and Protease type XIV (Sigma). Extracellular Ca2+ was slowly increased to physiological concentration (1.8 mM).
Solutions and chemicals
Ventricular cardiomyocytes were permeabilized with 0.05% escin (Sigma). Cells were kept in final solution containing (mM): 120 potassium aspartate (KAsp), 3 K‐ATP, 3.734 MgCl2, 0.5 EGTA, 10 phosphocreatine, 10 HEPES and 0.05 Fluo‐3 5K (Biotium), with the addition of 5 U L−1 creatine phosphokinase (pH 7.4, 310 mOsm L−1). Free‐[Ca2+] was measured using Indo‐1 AM (Molecular Probes, Eugene, OR, USA) in a NanoDrop fluorospectrometer and adjusted to 70 nM. Cells were plated using Cell‐Tak (Corning, Fisher Scientific, Schwerte, Germany)‐coated coverslips. Pharmacological interventions include 10 μM IP3‐salt (IP3, SiChem, Bremen, Germany), 2 μM 2‐aminoethoxydiphenyl borate (2‐APB, Calbiochem) and 20 mM caffeine (Sigma). Reports suggest that 2‐APB at higher concentrations than that used in the present study may have secondary effects in other intracellular components besides blocking IP3R (Maruyama et al. 1997; Ascher‐Landsberg et al. 1999; Peppiatt et al. 2003). In our hands, application of 2 μM 2‐APB in WT cells – in the absence of IP3 stimulation – returned no unspecific effects. To ensure that the responses seen with 2‐APB were due to IP3R block, 3 μM xestospongin C (XeC, Focus Biomolecules) was used as an alternative IP3R blocker. We are aware that both blockers used in our study do not have a 100% selectivity; however, the combination of both is currently the best option for a pharmacological separation of IP3 effects.
Intact myocytes were loaded at RT with 5 μM Fluo‐3 AM (Biotium, Fremont, CA, USA) for 30 min, plated in Cell‐Tak‐coated coverslips and kept in extracellular solution containing (mM): 140 NaCl, 5 HEPES, 1.1 MgCl2, 5.4 KCl, 10 glucose and 1.8 CaCl2. For Na+ and Ca2+ free solution, NaCl was replaced by 140 LiCl and 0.5 EGTA. To reach steady state cells were field stimulated for 30 s at a frequency of 1 Hz. The following pharmacological agents were used: 100 nM endothelin‐1 (ET‐1, Sigma), 2 μM 2‐APB, 3 μM XeC, 1 mM tetracaine (tet, Sigma) and 10 mM caffeine (Sigma).
[Ca]2+ measurements and protocols
For the analysis of the SR‐Ca2+ leak/load relationship, intact cardiomyocytes were kept at 1.8 mM Ca2+ and paced for 30 s at 1 Hz to reach steady state. External solution was rapidly exchanged to a Na+ and Ca2+ free solution containing 1 mM NiCl2 to ensure Na+/Ca2+ exchanger block (Kimura et al. 1987). After 20 s, 1 mM tetracaine was added to block RyR2. After 30 s, 10 mM caffeine was rapidly applied to determine the SR‐Ca2+ content. ΔCa2+ cyt is estimated as the difference (ΔCa2+ cyt = F tet – F 0) between fluorescence before (F 0) and after RyR2 block (F tet); as more Ca2+ is released to the cytosol via SR‐Ca2+ leak, the difference (ΔCa2+ cyt) between baselines shrinks and the SR‐Ca2+ leak becomes more positive.
For single cell arrhythmogenicity studies, intact cells were kept at 1.8 mM Ca2+ external solution and were field stimulated for 30 s at 1 Hz prior to a 10 s rest period to study Ca2+ wave occurrence as an estimate of cell propensity to aberrant SR‐Ca2+ release. Finally, SR‐Ca2+ load was assessed by application of 10 mM caffeine. Ca2+ wave occurrence (%) is calculated as the portion of cells analysed that showed Ca2+ waves during the 10 s resting period.
Confocal Ca2+ imaging
To assess intracellular Ca2+ changes, Fluo‐3 was excited at 488 nm wavelength of an argon laser and fluorescence was collected at 525 nm in line‐scan mode (2 ms per line) with a photomultiplier tube (PMT) on a MicroRadiance laser scanning confocal microscope (Bio‐Rad, Watford, UK) or alternatively a FluoView 1000 (Olympus, Volketswil, Switzerland) with a solid‐state laser (Sapphire, Coherent, Santa Clara, CA, USA).
Data analysis and statistics
Raw line‐scan data from permeabilized myocytes were analysed using the ImageJ (NIH) plug‐in SparkMaster to obtain Ca2+ spark frequency (Ca2+ sparks (100 μm)–1 s–1), amplitude (Amp, change in fluorescence relative to the fluorescence background (ΔF/F 0)), full‐width at amplitude half‐maximum (μm, FWHM), full duration at amplitude half‐maximum (ms, FDHM), full duration (ms), exponential time constant of the spark decay (τdecay, ms) and time to peak (ms). Spark mass was calculated as Amplitude × 1.206 × (FWHM)3 (Hollingworth et al. 2001); Ca2+ released via Ca2+ sparks was estimated as the product of the spark frequency and spark mass. Spark frequency data are presented as 100 sparks μm−1 s−1 or normalized to the vector where indicated (norm.). Data analysis was performed using the softwares ImageJ (NIH, Bethesda, MD, USA) and Igor Pro (WaveMetrics, Lake Oswego, OR, USA).
Data are presented as box (25–75 percentiles) and whisker (10 and 90 percentiles) plots; outliers are highlighted. Data within tables are presented as mean ± standard deviation (SD). Normal distribution of the data was assessed by D'Agostino's K 2 test. Statistical comparison was performed via one‐way ANOVA for multi‐parametric comparison followed by Dunnett's test or paired or unpaired Student's t test. Binary data were tested by the χ2 test. Number of animals (N), cells (n), Ca2+ events (n e) and statistical results are indicated in the figure legends.
Results
Cardiac IP3R2 overexpression induces cardiac remodelling both at cellular and organ levels
A cardiac‐specific IP3R2‐overexpressing mouse model (TG; Nakayama et al. 2010) has been used to mimic cardiac hypertrophy occurring in patients (Harzheim et al. 2009). In addition, the strategy to tune one element within a complex cellular remodelling situation to extreme could unmask an otherwise potentially invisible regulatory mechanism. The TG mice display a significantly higher heart weight to body weight ratio when compared to their wild‐type littermates (WT, 7.75 ± 0.92 mg g−1 vs. TG 10.6 ± 1.69 mg g−1; N = 5, 9; P = 0.001; Fig. 1 A). A cardiac hypertrophic phenotype was confirmed via transthoracic echocardiography (Table 1), which revealed greater interventricular septum (IVS, 0.78 ± 0.08 mm WT vs. 0.93 ± 0.09 mm TG; N = 4, 6; P = 0.039) and left ventricular posterior wall thickness (LVPW, 0.78 ± 0.07 mm WT vs. 0.94 ± 0.11 mm TG; P = 0.033) in TG hearts when compared to WT.
Table 1.
WT | TG | |
---|---|---|
IVSd (mm) | 0.78 ± 0.08 | 0.93 ± 0.09* |
IVSs (mm) | 1.07 ± 0.11 | 1.27 ± 0.13* |
LVPWd (mm) | 0.78 ± 0.07 | 0.94 ± 0.11* |
LVPWs (mm) | 1.09 ± 0.07 | 1.27 ± 0.16 |
%FS | 33.35 ± 1.97 | 29.04 ± 4.06 |
%EF | 62.45 ± 2.78 | 55.94 ± 6.21 |
E‐wave deceleration time (ms) | 21.07 ± 6.84 | 12.38 ± 3.8* |
ET (ms) | 49.29 ± 3.6 | 58.81 ± 6.42* |
TG mice display mild cardiac hypertrophy at 5 months of age. Values are means ± SD. Abbreviations correspond to: diastole (d), systole (s), interventricular septum (IVS), left ventricular posterior wall (LVPW), fractional shortening (%FS), ejection fraction (%EF) and ejection time (ET) (* P < 0.05; N = 4, 7; Student's t test).
Figure 1 B shows an increase in the relative expression of ITPR2 mRNA (IP3R2 encoding gene) levels in both ventricular and atrial tissue. A minor ITPR2 expression increase was detected in smooth and skeletal muscle but was absent in brain tissue. Accordingly, western blot analysis revealed a 12.72 (±2.97)‐fold increase in IP3R2 protein synthesis (N = 4 each; P = 0.001; Fig. 1 C, left) in ventricle. In addition and in line with other reports (Zhao et al. 2007), RyR2 protein level was significantly decreased in TG ventricular tissue (61.73 ± 0.24%; N = 4 each; P = 0.012; Fig. 1 C, right).
Global Ca2+ wave occurrence in response to IP3R2 pathway stimulation
Spontaneous Ca2+ waves are triggers for cardiac arrhythmias at the cellular level. To assess Ca2+‐dependent arrhythmogenicity in individual cells, IP3 synthesis was elicited via activation of the phosphatidylinositol 4,5‐bisphosphate (PIP2)–phospholipase C (PLC)–IP3 signalling cascade by endothelin‐1 (ET‐1, 100 nM) in intact ventricular myocytes (Table 2). Cells were field stimulated for 30 s at 1 Hz to ensure comparable SR‐Ca2+ loading conditions prior to a 10 s rest period to assess the occurrence of Ca2+ waves (Fig. 2 A). In control conditions, without ET‐1 stimulation and when compared to WT, TG myocytes displayed higher Ca2+ transient amplitude (1.9 ± 0.4 ΔF/F 0 WT vs. 2.32 ± 0.54 ΔF/F 0 TG; n = 13, 14; P = 0.014; Fig. 2 B) under conditions of comparable SR‐Ca2+ load (3.5 ± 0.79 ΔF/F 0 WT vs. 3.53 ± 0.78 ΔF/F 0 TG; P = 0.46) and consequently the fractional SR‐Ca2+ release in TG cells was increased (0.56 ± 0.1 a.u. WT vs. 0.65 ± 0.12 a.u. TG; P = 0.025). Furthermore, TG myocytes displayed a higher spontaneous Ca2+ wave occurrence (5.5% WT vs. 46% TG; P < 0.001; Fig. 2 C).
Table 2.
WT | Ca2+ transient amplitude (ΔF/F 0) | τdecay (ms) | T50 (ms) | Fractional SR‐Ca2+ release (a.u.) | SR‐Ca2+ content (ΔF/F 0) |
---|---|---|---|---|---|
Control (13, 4) | 1.9 ± 0.4 | 156.92 ± 27.76 | 119.24 ± 19.25 | 0.56 ± 0.1 | 3.5 ± 0.79 |
ET‐1 (11, 3) | 3.25 ± 1.17** | 162.26 ± 20.73 | 124.88 ± 12.6 | 0.66 ± 0.14* | 5.06 ± 1.81* |
ET‐1 + 2‐APB (16, 3) | 2.69 ± 0.84 | 176.78 ± 40.56 | 132.62 ± 23.49 | 0.63 ± 0.11 | 4.17 ± 0.92 |
2‐APB (16, 3) | 2.09 ± 0.58 | 173.83 ± 42.17 | 129.04 ± 25.02 | 0.62 ± 0.15 | 3.47 ± 1.02 |
XeC (15, 3) | 2.12 ± 0.73 | 168.44 ± 31.59 | 131.1 ± 21.19 | 0.62 ± 0.15 | 3.57 ± 1.46 |
TG | Ca2+ transient amplitude (ΔF/F 0) | τdecay (ms) | T50 (ms) | Fractional SR‐Ca2+ release (a.u.) | SR‐Ca2+ content (ΔF/F 0) |
---|---|---|---|---|---|
Control (14, 4) | 2.32 ± 0.54‡ | 122.39 ± 22.3‡‡‡ | 97.01 ± 17.82‡‡ | 0.65 ± 0.12‡ | 3.53 ± 0.78 |
ET‐1 (16, 4) | 2.63 ± 0.77 | 135.01 ± 35.5 | 106.82 ± 26.29 | 0.65 ± 0.13 | 4.07 ± 0.9 |
ET‐1 + 2‐APB (14, 3) | 2.77 ± 0.86 | 126.68 ± 25.8 | 102.57 ± 20.22 | 0.71 ± 0.12 | 4.37 ± 0.65 |
2‐APB (14, 3) | 3.04 ± 1.28* | 131.9 ± 12.36 | 107.82 ± 13.74* | 0.65 ± 0.18 | 4.57 ± 1.18** |
XeC (11, 3) | 2.95 ± 0.78* | 164.2 ± 44.4 | 128.5 ± 24.78 | 0.72 ± 0.1 | 4.11 ± 1.08 |
Values are means ± SD. Spatio‐temporal characteristics analysed include Ca2+ transient amplitude (ΔF/F 0), tau decay (τdecay, ms), time to 50% amplitude (T50, ms), fractional SR‐Ca2+ release (a.u.) and SR‐Ca2+ content (ΔF/F 0) from WT and TG (* P < 0.05, ** P < 0.01 vs. control; ‡ P < 0.05, ‡‡ P < 0.01, ‡‡‡ P < 0.001 vs. WT). Number of animals (N) and cells (n) are indicated below each condition as (n, N).
Compared to control, ET‐1 (100 nM, after 7–10 min application) triggered an increase in Ca2+ transient amplitude (3.25 ± 1.17 ΔF/F 0 ET‐1; n = 11; P = 0.001; Fig. 2 B, upper left), SR‐Ca2+ content (5.06 ± 1.81 ΔF/F 0 ET‐1; P = 0.013; Fig. 2 B, upper right) and Ca2+ wave occurrence (27.78% ET‐1; P = 0.035; Fig. 2 C) in WT cardiomyocytes, but surprisingly this effect was absent in IP3R2‐overexpressing cells (Fig. 2 B and C). In contrast, ET‐1 induced a 21.72% decrease in Ca2+ wave occurrence only in TG myocytes (Fig. 2 C; n = 16; P < 0.001).
In WT cardiomyocytes, block of IP3R2 by 2‐aminoethoxydiphenyl borate (2‐APB, 2 μM) or xestospongin C (XeC, 3 μM) returned no detectable changes in the analysed Ca2+ responses. In contrast, under IP3R2‐overexpressing conditions, blockade of IP3R2 led to increases in Ca2+ transient amplitude (3.04 ± 1.28 ΔF/F 0 2‐APB and 2.95 ± 0.78 ΔF/F 0 XeC; n = 14, 11; P = 0.033, 0.018; Fig. 2 B, lower left) with concomitantly higher SR‐Ca2+ content (4.57 ± 1.18 ΔF/F 0 2‐APB and 4.11 ± 1.08 ΔF/F 0 XeC; P = 0.007, 0.08; Fig. 2 B, lower right). Finally, block of IP3R2 by 2 μM 2‐APB abolished ET‐1‐mediated effects on cellular arrhythmogenicity both in WT and TG myocytes (Fig. 3). In other words, ET‐1 stimulation fails to alter the Ca2+ wave occurrence while IP3R remains blocked (Fig. 3).
To further study ET‐1‐mediated effects, estimation of sarco(endo)plasmic reticulum Ca2+‐ATPase (SERCA) and Na+–Ca2+ exchanger (NCX) relative contributions to the total Ca2+ transient decay was approached (Fig. 4). Fitting of a single exponential function to the action potential (AP)‐induced Ca2+ transient decay was used as a reference of the overall Ca2+ extrusion capacity of the cell, and to the Ca2+ transient (T) decay in the presence of 10 mM caffeine to assess NCX contribution (Fig. 4 A, left panel). SERCA and NCX (orange) contributions to the twitch decline (%) were calculated as: ((1/T twitch) − (1/T caffeine))/(1/T twitch) and (1/T caffeine)/(1/T twitch), respectively. In control conditions (Fig. 4 A, right panel), TG myocytes display a higher SERCA contribution when compared to WT (** P < 0.01; N = 4 each, n = 13, 14). An increase in SERCA contribution to the twitch Ca2+ decline upon 100 nM ET‐1 stimulation was seen only in WT ventricular myocytes (Fig. 4 Ba; ‡‡ P < 0.01; N = 4, 3; n = 13, 11).
Taken together, under conditions of IP3R2 overexpression, ET‐1 stimulation unexpectedly, and in contrast to the wild‐type situation, led to a decrease in Ca2+ wave frequency without altering Ca2+ transient amplitude nor SR‐Ca2+ content. However, block of IP3R2 results in a significant increase in SR‐Ca2+ content, which indicates that IP3ICR may play an intrinsic role in the regulation of the SR‐Ca2+ content and the propensity to arrhythmias.
Reduced SR‐Ca2+ content cannot be explained by Ca2+ sparks
The Ca2+ wave is a fundamental phenomenon that arises from unsynchronized SR‐Ca2+ release. Ca2+ sparks are elementary Ca2+ release events that can explain, at least in part, both the initiation and propagation of Ca2+ waves via the summation of individual Ca2+ release events and CICR. As was shown for atrial cells (Wullschleger et al. 2017), local bi‐directional crosstalk of IP3R2 and RyR2 may trigger and boost ventricular CICR. Under this view, pronounced IP3ICR may lead to an increased local Ca2+ events occurrence. Therefore, the effect of IP3ICR under IP3R2‐overexpressing conditions on the incidence of Ca2+ sparks was examined next.
Figure 5 A shows spontaneous Ca2+ sparks recorded in permeabilized cardiomyocytes upon diverse pharmacological interventions targeting the IP3R2 pathway. Under resting conditions, IP3R2‐overexpressing cells exhibit a higher spontaneous Ca2+ spark frequency (CaSpF) than WT (3.67 ± 1.8 sparks (100 μm)−1 s−1 TG vs. 1.76 ± 1 sparks (100 μm)−1 s−1 WT; n = 25 each; P < 0.001; Fig. 5 C, left panel) under similar SR‐Ca2+ loading conditions (Fig. 5 B, right, and C). In line with the concept that IP3ICR can boost CICR (e.g. Ca2+ sparks), in WT ventricular cardiomyocytes IP3‐salt (10 μM) evoked a significant increase in CaSpF (24.2%; n = 22; P = 0.006; Fig. 5 Da, left) together with a statistically significant decrease of the SR‐Ca2+ content (3.73 ± 0.89 ΔF/F 0 control vs. 3.3 ± 0.69 ΔF/F 0 IP3; P = 0.041; Fig. 5 Da, right) with no further changes in Ca2+ spark mass (Table 3). This response was sensitive both to 2 μM 2‐APB and 3 μM XeC (Fig. 5 Da). In contrast to the native situation, in IP3R2 overexpressing cardiomyocytes, IP3 induced a 25.17% decrease of the CaSpF (P = 0.015; n = 25, 13; P = 0.015; Fig. 5 Db, right) along with a reduction of the SR‐Ca2+ content (4.17 ± 0.48 ΔF/F 0 control vs. 3.56 ± 0.57 ΔF/F 0 IP3; P = 0.012; Fig. 5 Db, left) 2–4 min after application. Both effects were sensitive to XeC and 2‐APB.
Table 3.
WT | Amplitude (ΔF/F 0) | FDHM (ms) | Full duration (ms) | FWHM (μm) | Spark mass (a.u.) | τdecay (ms) | Time to peak (ms) |
---|---|---|---|---|---|---|---|
Control (25, 8) | 1.33 ± 0.12 | 18.17 ± 1.35 | 45.62 ± 5.08 | 1.95 ± 0.16 | 13.41 ± 4.44 | 18.12 ± 2.9 | 10.8 ± 1.28 |
IP3 (22, 8) | 1.34 ± 0.15 | 18.22 ± 2.32 | 44.71 ± 7.26 | 1.96 ± 0.16 | 14 ± 4.26 | 19.82 ± 8.38 | 10.76 ± 1.14 |
2‐APB + IP3 (20, 3) | 1.37 ± 0.22 | 18.51 ± 2.15 | 48 ± 9 | 2.01 ± 0.14 | 15.3 ± 4.47 | 17.56 ± 4.54 | 11.5 ± 1.94 |
XeC + IP3 (16, 3) | 1.27 ± 0.11 | 22.75 ± 3.41††† | 50.69 ± 10.35††† | 2.04 ± 0.2 | 16.55 ± 5.37 | 24.67 ± 9.48† | 13.32 ± 3.31 |
2‐APB (12, 3) | 1.32 ± 0.13 | 19.5 ± 3.53 | 50.25 ± 10.07 | 1.98 ± 0.2 | 13.96 ± 3.63 | 20.81 ± 8.87 | 11.83 ± 2.28 |
XeC (16, 3) | 1.33 ± 0.25 | 22.54 ± 2.54*** | 55.28 ± 8.66*** | 2.03 ± 0.24 | 15.31 ± 5.02 | 22.34 ± 4.8** | 13.9 ± 2.88*** |
TG | Amplitude (ΔF/F 0) | FDHM (ms) | Full duration (ms) | FWHM (μm) | Spark mass (a.u.) | τdecay (ms) | Time to peak (ms) |
---|---|---|---|---|---|---|---|
Control (25, 8) | 1.32 ± 0.14 | 26.22 ± 6.53 | 62.13 ± 14.89 | 1.96 ± 0.22 | 13.89 ± 4.34 | 26.86 ± 8.38 | 13.6 ± 2.71 |
IP3 (13, 3) | 1.41 ± 0.15 | 24.79 ± 7.5 | 57.94 ± 13.57 | 1.98 ± 0.15 | 16 ± 5.22 | 32.49 ± 17.1 | 13.38 ± 5.01 |
2‐APB + IP3 (11, 3) | 1.32 ± 0.25 | 29.95 ± 12.8 | 68.63 ± 27.89 | 2 ± 0.12 | 15.04 ± 4.58 | 38.29 ± 27.57 | 15.64 ± 7.37 |
XeC + IP3 (10, 3) | 1.31 ± 0.08† | 27.16 ± 8.57 | 61.99 ± 16.85 | 1.91 ± 0.17 | 13.27 ± 4.76 | 36.91 ± 19.59 | 12.35 ± 1.57 |
2‐APB (17, 4) | 1.31 ± 0.17 | 26.1 ± 5.27 | 57.12 ± 10.58 | 2.05 ± 0.23 | 16.34 ± 6.52 | 30.12 ± 11.52 | 13.03 ± 2.58 |
XeC (14, 3) | 1.28 ± 0.16 | 26.93 ± 7.47 | 62.62 ± 14.65 | 2.02 ± 0.2 | 15.69 ± 5.17 | 42.3 ± 12.72 | 15.4 ± 4.73 |
Following cell permeabilization and plating, a representative line‐scan was recorded to assess Ca2+ events in control conditions. Pharmacological effects were assessed 2–4 min post‐intervention (10 μM IP3‐salt, 2 μM 2‐APB, 3 μM XeC). Parameters analysed include Ca2+ spark amplitude (ΔF/F 0), full duration at amplitude half‐maximum (FDHM, ms), full duration (ms), full width at amplitude half‐maximum (FWHM, μm), spark mass (a.u.), tau decay (τdecay, ms) and time to peak (ms) for WT and TG (** P < 0.01, *** P < 0.001 vs. control; † P < 0.05, ††† P < 0.001 vs. IP3‐salt; ANOVA followed by Dunnett's test). N and n are indicated below each condition as (n, N).
To elucidate how the SR‐Ca2+ content can be reduced even with a lower number of Ca2+ sparks, individual Ca2+ events were analysed. Spatio‐temporal Ca2+ event analysis revealed no differences in individual parameters between Ca2+ events observed in WT and TG cells upon direct IP3 stimulation (Table 3). Recently, we reported that multi‐parametric analysis of single Ca2+ events upon similar pharmacological interventions as presented in the current study can be used as a tool to differentiate among Ca2+ sparks and Ca2+ puffs in permeabilized atrial myocytes (Wullschleger et al. 2017). Hence, we repeated the analysis for the data presented here. Figure 6 shows single Ca2+ events in control conditions and upon IP3 application plotted according to their individual fluorescence amplitude and full duration at amplitude half‐maximum (FDHM). IP3 application did not induce any detectable differences either in WT (Fig. 6 Aa; n e = 195, 374) or TG myocytes (Fig. 6 Ab; n e = 443, 269). The Ca2+ event analysis suggests that only one population of Ca2+ events was detected and can be identified as Ca2+ sparks. In other words, a reduced SR‐Ca2+ content in TG cardiomyocytes cannot be explained by RyR2 activity alone, given the reduced Ca2+ sparks occurrence.
In addition, we estimated the Ca2+ released via Ca2+ sparks under the tested conditions as the product of the Ca2+ spark mass and the CaSpF. Figure 6 B shows that IP3 stimulation increased Ca2+ release via Ca2+ sparks in WT (Fig. 6 Ba), but evoked a reduction in TG cells (Fig. 6 Bb). Both effects are presumably due to the differences in CaSpF induced by the application of IP3.
The discrepancy between reduced SR‐Ca2+ content and the total amount of Ca2+ released observed during IP3 stimulation in TG cardiomyocytes can neither be explained by the number of detected events nor the spatio‐temporal characteristics of individual Ca2+ events.
Intrinsically active IP3ICR modulates SR‐Ca2+ content via SR‐Ca2+ leak
Enhanced luminal Ca2+ increases RyR2 channel open probability (P o; Györke & Györke, 1998). Thus aberrant SR‐Ca2+ overload may result in spontaneous SR‐Ca2+ release (Chen et al. 2014). Since a discrepancy between CaSpF and SR‐Ca2+ content was evident, we hypothesized a contribution of IP3ICR via “eventless” SR‐Ca2+ release. We performed a series of SR‐Ca2+ leak protocols (Fig. 7 A) in the presence of RyR2 antagonists and/or IP3R2 agonists/antagonists to assess a potential contribution of “eventless” IP3ICR on the total SR‐Ca2+ leak (Shannon et al. 2002). Figure 7 B shows zoomed‐in examples with the corresponding fluorescence profiles for WT and TG myocytes (left and right, respectively) showing the RyR2 contribution to the total SR‐Ca2+ leak estimated as the difference (ΔCa2+ cyt = F tet − F 0) between the fluorescence before (F 0) and after RyR2 block (F tet). In principle, an increase in the “tetracaine‐insensitive” SR‐Ca2+ leak (e.g. via IP3ICR) may result in an apparent elevation in cytosolic [Ca2+] despite the presence of tetracaine (F tet), thus the difference between baselines (ΔCa2+ cyt) may become smaller, implying an increase in the total SR‐Ca2+ leak. Upon NCX block, the apparent ΔCa2+ cyt significantly depends on the SR‐Ca2+ leak/load balance (e.g. SR‐Ca2+ release and re‐uptake) and steady state prior to RyR2 block (F 0). In other words, the expected ΔCa2+ cyt decrease may not fully reflect large increases in SR‐Ca2+ leak above steady state. This is seen in Figs 7 and 8.
Figure 7 C shows that compared to WT, TG cells display a lower ΔCa2+ cyt (−0.16 ± 0.06 ΔF/F 0 WT vs. −0.05 ± 0.07 ΔF/F 0 TG; n = 10, 16; P < 0.001), larger Ca2+ transients with enhanced fractional SR‐Ca2+ release and SR‐Ca2+ content (3.84 ± 1.62 ΔF/F 0 WT vs. 4.75 ± 1.39 ΔF/F 0 TG; P = 0.075).
Figure 8 A shows representative SR‐Ca2+ leak profiles in the tested conditions (control, ET‐1, ET‐1 + 2‐APB and 2‐APB) for WT and TG ventricular myocytes. Figure 8 Ba exhibits the aforementioned traces from the different conditions overlapped for each animal group (WT and TG, above and below, respectively). Finally, Fig. 8 Bb displays the fitted samples according to cell type (WT or TG, above and below, respectively) and pharmacological intervention (control vs. ET‐1, left and right, respectively) and indicates the SR‐Ca2+ leak portion corresponding to RyR or IP3R.
In WT myocytes, 100 nM ET‐1 induced a 31.25% decrease in ΔCa2+ cyt (n = 20; P = 0.011; Fig. 8 C, top left panel), which was sensitive to 2 μM 2‐APB. 2‐APB alone did not significantly affect any of the tested parameters. In addition, ET‐1 induced IP3‐independent increases of the SR‐Ca2+ content but not the Ca2+ transient amplitude (Fig. 8 C, top middle and right panels, respectively).
Surprisingly, in TG cells, ET‐1 application resulted in no detectable ΔCa2+ cyt changes (Fig. 8 C, bottom left panel) and non‐significant decreases of the Ca2+ transient amplitude and the SR‐Ca2+ content (Fig. 8 C, bottom middle and left panels, respectively). In contrast, IP3R2 block by 2‐APB resulted in a large decrease of the ΔCa2+ cyt both in the presence and absence of ET‐1 (Fig. 8 C, bottom left panel) together with increases in the SR‐Ca2+ content (Fig. 8 C, bottom right panel). No further changes were detected in Ca2+ transient amplitude (Fig. 8 C, bottom middle panel) upon IP3R2 block. However, the “hidden” IP3‐dependent ΔCa2+ cyt effect in the leak protocol is unmasked by IP3R2 block.
Taken together, in IP3R2‐overexpressing myocytes IP3ICR modulates the SR‐Ca2+ content via sustained SR‐Ca2+ leak driven by “eventless” IP3ICR rather than by localized Ca2+ release events, e.g. Ca2+ puffs. In addition, under IP3R2‐overexpressing conditions, IP3ICR is intrinsically active and thereby fine‐tunes the SR‐Ca2+ load and cytosolic Ca2+, subsequently modulating Ca2+ wave occurrence.
Discussion
Our results obtained in ventricular cardiomyocytes reveal that, upon IP3R2 overexpression, IP3ICR stimulation leads to a decrease in RyR2 activity together with a decline in SR‐Ca2+ content, indicating that IP3R2 activation can act as a SR‐Ca2+ gateway mechanism to escape ominous SR‐Ca2+ overload and subsequent unsynchronized SR‐Ca2+ release. Our approach unmasks a so far unrecognized mechanism by which IP3ICR modulates SR‐Ca2+ content via “eventless” SR‐Ca2+ leak and effectively restores cardiomyocyte Ca2+ homeostasis.
These observations are based on a cardiac‐specific IP3R2‐overexpressing mouse model that mimics at least in part cardiac hypertrophy in patients (Nakayama et al. 2010). Cardiac remodelling is a complex process that affects a broad range of intracellular Ca2+ signalling components. TG myocytes are characterized by increases in IP3R2 expression together with RyR2 down‐regulation, a phenomenon known for various cardiac diseases (Go et al. 1995; Domeier et al. 2012; Zima et al. 2014).
IP3 is known to significantly contribute to a variety of intracellular Ca2+ signalling mechanisms by functionally exercising as a secondary messenger that interconnects extracellular signals to numerous subcellular processes. In ventricular myocytes, IP3Rs are preferentially located within the nuclear envelope and are found at a lower extent in the SR membrane (Bare et al. 2005). While nuclear IP3ICR has been linked to the modulation of gene expression (Higazi et al. 2009), the contribution of SR‐IP3Rs in ventricular ECC under physiological conditions remains controversial (Zhang et al. 2002; Harzheim et al. 2009, 2010; Nakayama et al. 2010; Cooley et al. 2013; Signore et al. 2013). Whether and in which direction enhanced IP3ICR may effectively modulate cardiac ECC in pathophysiological conditions warrants further investigation.
Ca2+ wave occurrence is controlled by IP3ICR
In the myocardium, the amplitude of global Ca2+ signals is locally controlled by Ca2+ influx through L‐type Ca2+ channels (LTCCs) and subsequently amplified by SR‐Ca2+ release via CICR. Because of its positive feedback, CICR has an inherent tendency to oscillate, particularly under pathological conditions. Based on spontaneous SR‐Ca2+ release, these oscillations have been found to play a fundamental role in the generation of cardiac arrhythmias. At the cellular level, Ca2+ oscillations originating from spontaneous SR‐Ca2+ release channel openings often manifest as Ca2+ waves that travel from release junction to release junction in an unsynchronized manner. This process can be described as a “fire‐diffuse‐fire” mechanism (Keizer et al. 1998). In this scenario, aberrant SR‐Ca2+ releases can substantially increase cytosolic Ca2+ levels and trigger delayed afterdepolarizations, a major source for ventricular tachyarrhythmias (Lipp & Niggli, 1993). A contribution of IP3ICR to the genesis of Ca2+‐dependent arrhythmias has been suggested for some time (Zhang et al. 2002; Zima & Blatter, 2004; Kockskämper et al. 2008; Harzheim et al. 2009; Nakayama et al. 2010). Our observations in wild‐type cells (WT) are in agreement with this concept. Under control conditions we found an increase in SR‐Ca2+ release activity triggered by stimulation of the IP3 pathway in the shape of Ca2+ waves or Ca2+ sparks.
The use of G‐protein coupled receptor (GPCR) agonists (e.g. ET‐1) leads to the intracellular synthesis of IP3 together with activation of protein kinase C (PKC). Although the positive inotropic action of ET‐1 in ventricular myocytes is generally accepted, the underlying mechanisms are controversial, while numerous cellular targets are involved (Braz et al. 2004; Li et al. 2005; Domeier et al. 2012; Smyrnias et al. 2018). In our hands, the inotropic responses triggered by ET‐1 in WT cells (Figs 4 and 7) might be partially mediated by PKC during the ET‐1/GPCR pathway stimulation. In the presence of ET‐1, the SR‐Ca2+ leak increases together with the SR‐Ca2+ content; this discrepancy could be explained by the positive inotropic PKC‐related effects on the LTCCs. As a consequence, increased Ca2+ influx and SERCA activity during the pacing protocol may lead to the observed increased SR‐Ca2+ content (Fig. 2). This may also have some relevance for the leak protocol (Fig. 7). Interestingly, application of 2‐APB in the presence of ET‐1 restores the SR‐Ca2+ leak to basal levels, while the SR‐Ca2+ content remains elevated (Fig. 8). In other words, the leak effect trigged by ET‐1 is IP3R dependent. In addition, the ET‐1‐triggered arrhythmogenic responses on a cellular level can be reversed by IP3R2 block excluding significant PKC‐dependent side effects (e.g. LTCCs, NCX) and indicating that IP3ICR seems to be the main mechanism responsible for the induced arrhythmic responses.
Analogously, the increase in Ca2+ spark and Ca2+ wave occurrence upon IP3 stimulation may be partly explained by a functional bi‐directional interaction between IP3R2 and RyR2 (Wullschleger et al. 2017). Therefore, IP3ICR may directly trigger and/or amplify CICR resulting in the initiation and/or propagation of Ca2+ waves.
Under conditions of pathophysiological stress and the associated increased IP3R2 expression, the scenario given above may be fundamentally altered. Neurohumoral GPCR stimulation (e.g. via ET‐1) serves as a positive inotropic reserve that partially functions via IP3ICR activation. This may include sensitization of RyR2 to activating (“triggering”) cytosolic Ca2+ together with increases in SR‐Ca2+ load, events that cooperatively increase single RyR2 channel open probability (P o) and hence the tendency to develop arrhythmogenic Ca2+ waves (Domeier et al. 2008). Hence activation of GPCR via ET‐1 unfolds a positive inotropic reserve at the expense of increased spontaneous SR‐Ca2+ release events. In line with this scenario, under increased IP3R2 expression, an even higher probability of Ca2+ wave occurrence compared to the wild‐type situation would be expected. Surprisingly, the opposite response was evident: activation of IP3ICR in conditions of IP3R2 overexpression (TG) resulted in a decrease rather than an increase in Ca2+ wave occurrence. Furthermore, block of IP3R2 increased the SR‐Ca2+ content, suggesting an inherent regulatory role of IP3ICR in the control of the SR‐Ca2+ content. Thus, under conditions of enhanced IP3ICR, we hypothesized a secondary regulatory mechanism operating through IP3ICR activation and resulting in the reduction of Ca2+ wave occurrence.
Global cytosolic Ca2+ waves are composed of the temporal and spatial summation of individual CICR events (Ca2+ sparks) that occur by spontaneous but concerted openings of RyR2 clusters (Cheng et al. 1993). Analogously, IP3R2s are organized in clusters prerequisite to the generation of local Ca2+ events known as Ca2+ puffs (Yao et al. 1995). Analysis of individual Ca2+ events may reveal the contribution of IP3ICR (e.g. Ca2+ puffs) to the Ca2+ wave occurrence during IP3R stimulation. Under equal cytosolic Ca2+ levels, TG cells displayed a higher Ca2+ spark frequency (Fig. 5 C) than cells from WT littermates, reflecting a higher basal RyR2 sensitization. In agreement with others (Domeier et al. 2008), IP3 stimulation in WT cells induced an expected increase in RyR activity. Surprisingly, the effect was reversed in TG myocytes, in which IP3R2 stimulation returned not an increase but a decrease in Ca2+ event occurrence and SR‐Ca2+ content. The results in CaSpF in permeabilized myocytes are thus in line with those obtained in Ca2+ wave occurrence within intact ventricular myocytes (Fig. 2), suggesting that (1) cell permeabilization did not disrupt the mechanism by which IP3 may be altering SR‐Ca2+ release, and (2) discarding sarcolemmal components (e.g. LTCCs, Na+–Ca2+ exchanger) as the mechanisms directly involved in the reduction of Ca2+ wave occurrence.
IP3 stimulation of TG permeabilized ventricular myocytes induced a decrease in Ca2+ event frequency together with a reduction of the SR‐Ca2+ content. To explain such discrepancy, we hypothesized that IP3 may induce a change in single Ca2+ event properties, so that larger Ca2+ events carrying more Ca2+ may keep CaSpF steady but yet reduce the SR‐Ca2+ content. In this context, multi‐parametric analysis of Ca2+ event profiles has been recently used to distinguish between Ca2+ sparks and Ca2+ puffs (Hohendanner et al. 2015; Wullschleger et al. 2017). Recently we showed that, under similar experimental conditions, IP3 application to permeabilized atrial myocytes triggered the appearance of a group of Ca2+ events that can be identified and classified – via multi‐parametric analysis of single Ca2+ event properties – as IP3R2‐mediated Ca2+ puffs. In the present study, however, multi‐parametric analysis of Ca2+ event characteristics from permeabilized ventricular myocytes (Fig. 6 A) failed to identify and distinguish between two classes of events and thus the sole group detected was classified as RyR2 Ca2+ sparks. Hence, as no differences in Ca2+ event parameters were detectable and IP3 application triggered a decrease in Ca2+ event occurrence, RyR2 Ca2+ sparks cannot account for the decrease in SR‐Ca2+ content.
SR‐Ca2+ leak and “eventless” SR‐Ca2+ release
The local SR‐Ca2+ release carried via stochastic opening of SR‐Ca2+ release channels independent from the beat‐to‐beat ECC mechanism is known as SR‐Ca2+ leak. The luminal SR‐Ca2+ has been shown to be a major regulatory factor controlling Ca2+ release channel P o, hence the SR‐Ca2+ leak (Terentyev et al. 2002; Keller et al. 2007). Although SR‐Ca2+ release channels are grouped in clusters from which detectable Ca2+ events arise, a differential channel distribution has been proposed (Sobie et al. 2006). Individual RyR2s that do not belong in a cluster or small RyR clusters can still release Ca2+ from the SR in a manner that does not generate a Ca2+ event detectable by confocal microscopy (Lipp & Niggli, 1996; Brochet et al. 2011). Single or “rogue” RyR2s may modulate SR‐Ca2+ content with no apparent changes in Ca2+ event occurrence (Song et al. 2006). Analogously, singular or “rogue” IP3Rs may modulate SR‐Ca2+ content in an “eventless” manner. Thus the SR‐Ca2+ leak mechanism includes both SR‐Ca2+ release events (Ca2+ sparks and Ca2+ puffs) and non‐detectable or “eventless” SR‐Ca2+ release. Our data in CaSpF showing a decrease in Ca2+ spark occurrence with no detectable changes in Ca2+ event characteristics suggest that the triggered IP3ICR occurs in an “eventless” manner and thus contributes to the total SR‐Ca2+ leak without generating distinct Ca2+ events. In this scenario, we hypothesized that “eventless” IP3ICR reduces SR‐Ca2+ content and ultimately modulates the cardiomyocytes’ propensity to Ca2+ waves. Indeed, by approaching an adapted SR‐Ca2+ leak/load protocol (Shannon et al. 2002) we revealed a significant IP3R2‐dependent component of the SR‐Ca2+ leak suggesting that albeit being largely outnumbered by RyR2s, IP3R2s in wild‐type ventricular myocytes can still contribute to the total SR‐Ca2+ efflux when recruited by GPCR agonists.
Regarding the leak/load protocol, interpretation of the data is less straightforward than expected. Strong evidence of pronounced SR‐Ca2+ leak mediated by IP3ICR is shown by IP3R block (2‐APB) in TG cells, where 2‐APB triggers a pronounced decay of the total SR‐Ca2+ leak, especially when compared to the antagonists’ effect in WT (Fig. 8 C). In the latter, no differences to the control are detected whereas in the former a strong decrease in the basal SR‐Ca2+ leak is seen, indicative of increased IP3ICR under IP3R2‐overexpression conditions. Now, in TG cells, an intuitively expected SR‐Ca2+ leak increase was virtually absent upon ET‐1 stimulation. When compared to WT, in TG cells, tetracaine revealed a much smaller basal RyR2‐dependent SR‐Ca2+ leak (Fig. 6). Under these specific conditions SERCA works close to the cytosolic steady‐state [Ca2+] (F 0) and thus, additional or excess IP3R‐mediated SR‐Ca2+ leak is immediately compensated by SERCA activity. Because of the closed system (NCX block), cytosolic Ca2+ levels above the steady‐state (F 0) cannot be expected, as also shown by others (Shannon et al. 2002; Curran et al. 2007; Gonzalez et al. 2007; Santiago et al. 2010; Bers, 2014; Tzimas et al. 2017). Hence the ET‐1 effect is minor or not detectable in TG myocytes as it is masked by SERCA activity. On the contrary, in WT cells, as tetracaine application sharply decreases the cytosolic Ca2+ (F tet) far from steady‐state levels (F 0), extra SR‐Ca2+ release (triggered by ET‐1) is readily detectable. In other words, a direct comparison of ΔCa2+ cyt WT versus TG is inapplicable and only feasible in combination with IP3R2 blocking conditions, as suggested.
In addition, our data in TG cells show that the RyR2 protein down‐regulation detected concomitantly diminishes the receptors’ contribution to the basal SR‐Ca2+ leak. One would expect that the RyR2 down‐regulation seen under remodelling conditions goes in parallel with an increased ECC gain in order to stabilize the CICR as long as possible. Indeed, that was the case. The “remaining” RyR2 Ca2+ release units are more sensitive to the trigger Ca2+ resulting in a higher P o. That would imply a significantly higher RyR2 Ca2+ leak and/or a significant increase in Ca2+ spark occurrence. However, that was not observed under IP3R2 overxpression conditions and can be used as additional evidence that the observed SR‐Ca2+ leak was not due to increased RyR2 P o alone. Moreover IP3R block induces a large reduction of the total SR‐Ca2+ leak suggesting that IP3R2 P o compensates at least in part the loss of RyR2‐mediated SR‐Ca2+ leak. This view is supported by increases in the SR‐Ca2+ content only seen upon IP3R inhibition in IP3R2‐overexpressing cells.
Taken together our results indicate that the contribution of IP3ICR to the total SR‐Ca2+ release is mainly driven by “eventless” SR‐Ca2+ leak rather than by localized Ca2+ events. We hypothesize that this may be because the low expression of IP3R2s, even in pathologically high expressing conditions, may not suffice to group the receptors in clusters large enough to form discernible Ca2+ events (e.g. Ca2+ puffs). By this mechanism, activation of enhanced IP3ICR reduces the SR‐Ca2+ content below the threshold for Ca2+ spark generation, which prevents aberrant SR‐Ca2+ release and subsequently stabilizes intracellular Ca2+ homeostasis. In conclusion, the data presented reveal a novel mechanism by which activation of IP3R2s in a model of cardiac pathophysiology serves as an anti‐arrhythmic mechanism that fine‐tunes the SR‐Ca2+ content via “eventless” SR‐Ca2+ release.
Limitations of the study
At present, elucidating cardiac IP3‐dependent regulatory mechanisms is mainly limited to observations at the level of global Ca2+ events (e.g. of Ca2+ waves or Ca2+ transients) or by indirect proof of interaction by counting changes in Ca2+ event frequencies (Yamasaki et al. 2013). The reasons for this are: (1) in contrast to RyR Ca2+ signals, the smaller and rare IP3ICR events are difficult to identify and to discern among the larger Ca2+ responses that originate from CICR, and (2) identifying the specific contribution of IP3R2s in local and/or global Ca2+ responses is impeded by a deficiency of highly selective pharmacological tools (Gafni et al. 1997; Bootman et al. 2002). Although studies have fostered a contribution of IP3ICR on local Ca2+ signalling in cardiomyocytes, the observation of individual functional interactions of Ca2+ sparks and Ca2+ puffs was just recently successful (Wullschleger et al. 2017). In addition, immunocytochemistry largely failed to identify the precise localization of individual IP3R2s in mouse cardiac preparations, presumably because of limited antibody specificity. This limitation thus impedes the confirmation of the “rogue” IP3R2 function hypothesis (data not shown; Mohler et al. 2005; Kockskämper et al. 2008; Nakayama et al. 2010; Wullschleger et al. 2017).
Transgenic mouse models are sufficient and appropriate tools to examine regulatory mechanisms of CICR in cardiomyocytes. However, there is no doubt that a 1:1 extrapolation of findings emerging from animal studies to the human situation has to be critically evaluated. There are only a few studies comparing cellular remodelling situations in human cardiac disease with TG animal models used for specific aspects of cardiac remodelling both on global and cellular levels. Not only that, at present ECC under physiological conditions in human cardiomyocytes is only rudimentarily characterized (Signore et al. 2013). The TG mouse model used in the present study reveals cardiac hypertrophy with several similarities to a human pathology. However, it was used as an appropriate tool to study a potential regulatory function of IP3ICR in CICR. A modulatory mechanism based on IP3‐dependent SR‐Ca2+ leak has been unmasked by overexpression of IP3R2 which would otherwise have been potentially overlooked. Although several studies focus on IP3ICR as a potential pro‐arrhythmogenic factor (Mackenzie et al. 2002; Proven et al. 2006) this model shows that higher expression of the IP3R2 might actually be beneficial for the preservation of ECC within pathology.
Additional information
Competing interests
None declared.
Author contributions
All experiments were performed at the Department of Physiology, University of Bern, Switzerland. M.E. conceived and designed the experiments. J.B. collected and analysed the experimental data. M.E. and J.B. drafted and revised the article. All authors approved the final version for publication and agree to be accountable for all aspects of the work. All persons designated as authors qualify for authorship, and all those who qualify for authorship are listed.
Funding
This work was supported by the Swiss National Science Foundation (31‐149301) and Novartis Research Foundation to M.E.
Acknowledgements
The authors thank Ernst Niggli, Radoslav Janicek, Miguel Fernandez‐Tenorio, Jin Li and Ange Maguy for their helpful comments on the manuscript and Marianne Courtehoux for her technical support.
Biography
Joaquim Blanch i Salvador received his BS and MS degrees in Biomedical Sciences and Biotechnology from the University of Barcelona. After graduating, he joined Professor Dr M. Egger's group in the Department of Physiology of the University of Bern as a PhD student. Combining electrophysiology techniques with confocal Ca2+ imaging and molecular biology approaches, his research focuses on the study of intracellular Ca2+ oscillations within cardiomyocytes in a framework of cardiac pathology.
Edited by: Don Bers & Fernando Santana
Linked articles This article is highlighted in a Perspectives article by Dixon. To read this article, visit http://doi.org/10.1113/JP276793.
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