|Home | About | Journals | Submit | Contact Us | Français|
Antagonists of L-type Ca2+ channels (LTCCs) have been used to treat human cardiovascular diseases for decades. However, these inhibitors can have untoward effects in patients with heart failure, and their overall therapeutic profile remains nebulous given differential effects in the vasculature when compared with those in cardiomyocytes. To investigate this issue, we examined mice heterozygous for the gene encoding the pore-forming subunit of LTCC (calcium channel, voltage-dependent, L type, α1C subunit [Cacna1c mice; referred to herein as α1C–/+ mice]) and mice in which this gene was loxP targeted to achieve graded heart-specific gene deletion (termed herein α1C-loxP mice). Adult cardiomyocytes from the hearts of α1C–/+ mice at 10 weeks of age showed a decrease in LTCC current and a modest decrease in cardiac function, which we initially hypothesized would be cardioprotective. However, α1C–/+ mice subjected to pressure overload stimulation, isoproterenol infusion, and swimming showed greater cardiac hypertrophy, greater reductions in ventricular performance, and greater ventricular dilation than α1C+/+ controls. The same detrimental effects were observed in α1C-loxP animals with a cardiomyocyte-specific deletion of one allele. More severe reductions in α1C protein levels with combinatorial deleted alleles produced spontaneous cardiac hypertrophy before 3 months of age, with early adulthood lethality. Mechanistically, our data suggest that a reduction in LTCC current leads to neuroendocrine stress, with sensitized and leaky sarcoplasmic reticulum Ca2+ release as a compensatory mechanism to preserve contractility. This state results in calcineurin/nuclear factor of activated T cells signaling that promotes hypertrophy and disease.
Voltage-gated L-type Ca2+ channels (LTCCs) are the primary source of Ca2+ influx to initiate cardiac excitation-contraction coupling (ECC) (1, 2). The molecular composition of the LTCC in cardiomyocytes includes the pore-forming α1C subunit (Cacna1c; referred to herein as α1C), the β subunit, the α2δ subunit, and the more recently suggested γ subunit (3–6). The auxiliary subunits β, α2δ, and γ are involved in trafficking the pore-forming α1C subunit to the sarcolemma and modulating the voltage dependence of channel gating (1). Genetic deletions of either the α1C or the β2 encoding genes results in embryonic lethality with heart dysfunction (7, 8).
Ca2+ influx via α1C triggers a much larger Ca2+ release from the sarcoplasmic reticulum (SR) via ryanodine receptors (RyRs) to promote myocyte contraction (9). In addition to regulating cardiomyocyte contraction, Ca2+ influx from LTCCs is also involved in intracellular signaling and gene regulatory events that underlie cardiac hypertrophy and disease (10–13). For example, multiple studies have suggested that enhanced Ca2+ influx via the LTCC is responsible for cardiac hypertrophy and pathological remodeling of the ventricles (14–16). Consistent with these observations, LTCC blockers effectively inhibit cardiac remodeling and disease after pressure overload stimulation in animal models (17–19). However, α1C is also widely expressed in the vasculature, and the net beneficial cardiovascular effects of LTCC inhibitors in animal models could result from changes in vascular compliance and/or function to secondarily benefit the heart (15). In human clinical trials, LTCC blockers appear to either have no effect or to negatively impact the survival and cardiovascular event profile in patients with heart failure with systolic dysfunction (20). Thus, it remains unclear how LTCC inhibitors might be of clinical importance in heart failure as well as the cell type that mediates beneficial compared with detrimental effects.
Many studies have documented an association between increased Ca2+ influx during cardiac stress states and the induction of hypertrophy. Thus, we tested the hypothesis that a genetic reduction in LTCC current would reduce Ca2+ influx and thereby protect the heart during disease-causing stress stimulation, consistent with animal models treated with LTCC blockers that show less hypertrophy. However, we observed the opposite effect and show that reductions in cardiomyocyte LTCCs cause a compensatory increase in neuroendocrine stimulation and RyR2 Ca2+ leak toward increasing the gain in ECC, resulting in pathologic cardiac hypertrophy and failure through calcineurin/nuclear factor of activated T cells (calcineurin/NFAT) signaling.
Given the protective effects associated with LTCC inhibitors in animal models of heart failure and hypertrophy, we hypothesized that α1C–/+ mice would be protected from heart failure secondary to cardiac injury. Cardiac protein levels of α1C were reduced by approximately 40% in α1C–/+ mice compared with those in control mice at 10 weeks of age (Figure (Figure1A),1A), which correlated with roughly 25% less whole-cell L-type Ca2+ current (ICa-L) measured in freshly isolated adult ventricular myocytes (Figure (Figure1B).1B). No changes in cell capacitance were observed (data not shown). Ca2+ photometry experiments showed that the maximal amplitude of electrically evoked (Figure (Figure1,1, C, D, and F) and caffeine-evoked (Figure (Figure1,1, C, D, and H) Ca2+ transients was significantly reduced in α1C–/+ adult cardiomyocytes compared with that in WT cardiomyocytes, with no noticeable changes in diastolic Ca2+ or the decay time constant for Ca2+ reuptake and extrusion (Figure (Figure1,1, E and G). Associated with these reductions in Ca2+ handling, myocyte shortening (Figure (Figure1I)1I) and ventricular fractional shortening (FS) were also reduced in α1C–/+ mice compared with those in WT mice (Figure (Figure2A),2A), as was cardiac +dP/dt, measured ex vivo with a Millar pressure transducing catheter (Figure (Figure2B).2B). This mild reduction in cardiac performance in α1C–/+ mice was also associated with increased left ventricular chamber size in systole at 10 and 32 weeks of age (Figure (Figure2C),2C), which eventually resulted in a small but significant induction of cardiac hypertrophy by 32 weeks of age, as assessed by measurement of heart weight normalized to body weight (HW/BW; Figure Figure2D). 2D).
To further examine the cardiac effects associated with a reduction in LTCC current, α1C–/+ mice at 10 weeks of age, which is prior to an increase in heart weight, were subjected to pathologic and physiologic hypertrophic stimulation. Again, since increased Ca2+ influx has been associated with cardiac hypertrophy and pathological remodeling, we initially hypothesized that reduced whole-cell LTCC current would be cardioprotective in mice subjected to pressure overload by transverse aortic constriction (TAC). However, α1C–/+ mice subjected to TAC for 2 weeks exhibited enhanced cardiac remodeling, demonstrated by increased HW/BW (Figure (Figure3A),3A), reduced cardiac ventricular performance (Figure (Figure3B),3B), and ventricular chamber dilation, compared with that in α1C+/+ mice (Figure (Figure3C).3C). To extend these observations, we used a model of catecholamine overload-induced disease with 2 weeks of isoproterenol (Iso) infusion. Consistent with those in the TAC experiments, α1C–/+ mice showed a minor but significant reduction in ventricular performance and increase in cardiac hypertrophy compared with that of α1C+/+ littermates treated with Iso (Figure (Figure3,3, D and E). Finally, and unexpectedly, α1C–/+ mice also showed a significant reduction in ventricular performance and increased cardiac hypertrophy after exercise stimulation for 21 days by forced swimming (Figure (Figure3,3, F and G). Collectively, these results indicate that decreased ICa-L does not protect against cardiac hypertrophy after either pathologic or physiologic stimulation, but, to the contrary, it exacerbates disease.
To exclude the known effects of LTCC antagonists on the vasculature, we crossed α1C-loxP–targeted (floxed [fl]) heterozygous mice with transgenic mice expressing Cre recombinase under control of the α-myosin heavy chain (α-MHC) promoter (only 1 allele would be specifically deleted in cardiomyocytes). Echocardiographic analysis of α1C+/fl-Cre mice after 2 weeks of TAC showed a significant decrease in cardiac function and enhanced propensity for myocardial remodeling compared with that of controls (α1C+/fl) (Supplemental Figure 1; supplemental material available online with this article; doi: 10.1172/JCI58227DS1). Heart-specific α1C+/fl-Cre mice also showed greater decompensation and hypertrophy after swimming exercise for 21 days compared with that of controls, an effect that was not altered with an LTCC blocker, diltiazem (Supplemental Figure 1). Finally, we also crossed α1C-loxP heterozygous mice with α-MHC-MerCreMer transgenic mice to permit tamoxifen-inducible deletion of 1 α1C allele in the adult heart. Ten-week-old α1C+/fl-MerCreMer mice were given tamoxifen (25 mg/kg/d) for 5 days and then subjected to TAC 4 weeks later for 2 weeks. Importantly, these mice also showed a significant reduction in ventricular performance and ventricular dilation and an increase in hypertrophy compared with those in controls, again suggesting that myocyte-specific reduction in LTCC activity worsens cardiac disease after pathologic stimulation (Supplemental Figure 1).
We were also interested in achieving an even greater reduction in LTCC current to better investigate the mechanism of disease predisposition observed in α1C heterozygous mice. We analyzed double α1C-loxP–targeted mice (2 floxed alleles) with the α-MHC-Cre transgene as well as single allele α1C-loxP mice (with Cre) over an α1C-null allele to achieve an even more complete knockdown of LTCC activity in the heart, which was verified by Western blotting (Figure (Figure4A)4A) and careful assessment of Ca2+ current from isolated myocytes of each of the genotypes (Supplemental Figure 2). Indeed, α1C–/fl-Cre mice had the greatest reduction in Ca2+ current, followed by α1Cfl/fl-Cre mice, and then α1C–/fl mice (Supplemental Figure 2). We were unable to examine standard, fully deleted α1C–/– mice, because they are embryonically lethal (7). Use of the α-MHC-Cre transgene bypassed embryonic lethality in α1Cfl/fl and α1C–/fl mice, due to its predominant postnatal expression characteristics.
Consistent with the degree of α1C protein and current reduction, α1Cfl/fl-Cre mice and α1C–/fl-Cre mice showed partial adult lethality as they aged to 80 days (Figure (Figure4B).4B). Both α1Cfl/fl-Cre and α1C–/fl-Cre mice showed a significant reduction in cardiac ventricular performance at 10 weeks of age compared with that of control mice (Figure (Figure4C).4C). The reduction in performance was significantly greater in α1C–/fl-Cre mice, which had the greatest reduction in α1C protein and current. Consistent with these results, α1Cfl/fl-Cre and α1C–/fl-Cre mice each showed dramatic increases in left ventricular chamber dimensions and cardiac hypertrophy, which, again, were even more pronounced in the α1C–/fl-Cre genotype (Figure (Figure4,4, D, E, and G). This dramatic increase in cardiac hypertrophy in α1C–/fl-Cre mice was also associated with heart failure, as assessed by increases in lung weight normalized to body weight (Figure (Figure4F).4F). These results suggest that reductions in Ca2+ influx induce cardiac hypertrophy and heart failure in young adult mice.
We originally hypothesized that a genetic reduction in α1C current would reduce the total amount of Ca2+ available to activate calcineurin/NFAT signaling, a pro-hypertrophic regulatory pathway in the heart, especially given previous work suggesting that enhanced LTCC activity mediates cardiac hypertrophy through Ca2+-dependent signaling pathways (21). Thus, we first crossed NFAT-luciferase transgenic reporter mice with α1C–/fl-Cre mice, expecting to observe less activity. However, consistent with the unexpected and profound induction of hypertrophy seen in α1C–/fl-Cre mice, we observed a large induction in cardiac NFAT activity and calcineurin phosphatase activity compared with that in control hearts (Figure (Figure5,5, A and B). This suggested that a reduction in Ca2+ influx from the LTCC might lead to a secondary increase in Ca2+ that is important for activating calcineurin/NFAT signaling. Indeed, α1C–/fl-Cre mice treated for 3 weeks with cyclosporine A (CsA), a calcineurin inhibitor, showed partial but significant regression in ventricular dilation and hypertrophy and significantly (albeit minor) better cardiac function compared with that of untreated controls (Figure (Figure5,5, C–E). These results suggest that reduced LTCC current leads to calcineurin/NFAT activation and cardiac hypertrophy with pathology.
Considering the results of our study to this point, we revised our hypothesis to indicate that there must be an increase in SR Ca2+ leak or greater resting Ca2+ levels in the cleft microenvironment to compensate for less trigger Ca2+. This hypothetical increase in Ca2+ could then serve as a potent stimulus for reactive signaling pathways, such as calcineurin (22, 23). To assess this revised hypothesis, we measured Ca2+ handling in adult cardiomyocytes from α1C–/fl-Cre mice. As expected, the maximal amplitude of electrically evoked Ca2+ transients was significantly reduced in myocytes from α1C–/fl-Cre hearts and almost significantly reduced again in myocytes from standard heterozygous hearts (α1C–/fl; compare with a larger sampling in Figure Figure1F)1F) compared with that in controls (Figure (Figure6A).6A). The time constant of decay was significantly longer in myocytes from α1C–/fl-Cre hearts compared with that in control myocytes (Figure (Figure6B).6B). These changes were accompanied by a concomitant reduction in the SR Ca2+ load and peak release in both deficient genotypes (Figure (Figure6C).6C). However, the frequency of SR Ca2+ sparks measured in α1C–/fl-Cre myocytes was significantly higher than that in control myocytes (Figure (Figure6D),6D), despite a lower SR Ca2+ content. Consistent with this, independent measurements of tetracaine-sensitive diastolic SR Ca2+ leak (normalized for SR Ca2+ content) were also higher in α1C–/fl-Cre myocytes (Figure (Figure6E).6E). These data suggest that the SR Ca2+ release channel is sensitized, perhaps in compensation for the reduced Ca2+ entry via LTCC current. Indeed, in voltage-clamped myocytes in which SR Ca2+ load was matched among the groups (via longer loading pulses in α1C–/fl-Cre myocytes; Figure Figure6F)6F) the Ca2+ transient peaks were not different among groups, despite the reduced Ca2+ trigger current in α1C–/fl-Cre myocytes (Figure (Figure6G).6G). More importantly, when normalized for Ca2+ current amplitude, there was increased gain of ECC in α1C–/fl-Cre hearts compared with that in control hearts (Figure (Figure6H).6H). This again suggests that the SR Ca2+ release channel is more sensitive to Ca2+ in the α1C–/fl-Cre myocytes.
The increase in ECC gain and increased SR Ca2+ leak can be regulated by β-adrenergic receptor signaling through PKA and Ca2+/calmodulin-dependent kinase II (CaMKII) (24–26). Such a profile is associated with hyperphosphorylation of RyR2 at serine 2808 and 2814, with less calstabin2 (FKBP12.6) binding to the channel. Indeed, hearts from α1C–/fl-Cre mice showed significantly greater phosphorylation of RyR2, with less calstabin2 binding, and profound oxidative nitrosylation (with 2,4-dinitrophenyl [DNP]) compared with that of α1C heterozygous controls (Figure (Figure7,7, A–D). Given these results, we further reasoned that β-receptor blockade might limit the compensatory neurohumoral response that attempts to increase the gain in ECC, in part by phosphorylating RyR2, secondarily reducing this detrimental pool of Ca2+ that induces hypertrophy and disease. Hence, we treated α1C+/+ and α1C–/+ mice with the β1-adrenergic receptor selective antagonist metoprolol and then subjected the mice to pressure overload. Metoprolol-treated α1C–/+ mice maintained better cardiac ventricular performance and showed less ventricular dilation and less cardiac hypertrophy compared with vehicle-treated α1C–/+ mice (Figure (Figure7,7, E–G). Activation of CaMKII is also a consequence of β-receptor and neurohumoral activation that can cause increased RyR2 Ca2+ leak. Indeed, the CaMKII inhibitory drug KN93 restored the detriment to the Ca2+ transient and SR Ca2+ content and prevented SR Ca2+ leak in myocytes from α1C–/fl-Cre hearts (Figure (Figure7,7, H and I, and data not shown). These results again suggest that the neurohumoral response is engaged in α1C–/fl-Cre hearts in an attempt to compensate for less trigger Ca2+ but, in so doing, causes Ca2+ leak from RyR2 that induces pathological signaling and hypertrophy/heart failure.
We also crossed α1C–/fl-Cre mice with transgenic mice expressing the α1G subunit of the T-type Ca2+ channel (TTCC) to assess rescue in hypertrophic disease by enhancing trigger Ca2+ independent of the LTCC (27). Remarkably, α1G overexpression essentially rescued the secondary hypertrophy in α1C–/fl-Cre mice by providing more Ca2+ for acute systolic release (Supplemental Figure 3). α1G overexpression also mildly reduced ventricular remodeling in hearts from α1C–/fl-Cre mice, which was associated with a restoration in the Ca2+ transient and peak SR Ca2+ load (Supplemental Figure 3). Collectively, these results further suggest that a reduction in trigger Ca2+ is responsible for a compensatory increase in diastolic Ca2+ to increase ECC gain. Increased diastolic Ca2+ is a potent means of activating calcineurin.
Increased Ca2+ influx via LTCCs has been implicated in the development of hypertrophy and pathological remodeling of the heart with select stress stimuli (15, 28). For example, studies in animal models of heart disease have shown that antagonizing LTCC activity with pharmacologic inhibitors can prevent or reverse pathological cardiac remodeling and hypertrophy (17–19). We previously showed that upregulation of LTCC activity in the hearts of transgenic mice due to overexpression of the β2a subunit induced cardiomyopathy (29). Total LTCC currents were increased by nearly 50% in adult myocytes from these hearts, which enhanced cardiac contractility and eventually caused necrotic death of myocytes, leading to dilated failure, with slightly heavier hearts. However, one perplexing observation from these mice was the relative lack of bona fide hypertrophy associated with enhanced Ca2+ influx, especially in mice that were less than 3 months old, as we initially hypothesized that such a dramatic increase in Ca2+ would induce calcineurin activation (29). Similarly, α1C-overexpressing transgenic mice did not show cardiac hypertrophy until 8 months of age, and then it was mild and possibly secondary to neurohumoral effects (30, 31). Thus, increasing the peak or acute release of Ca2+ appears to be a weak inducer of hypertrophic disease and pathologic signaling, though it can lead to mitochondrial Ca2+ overload and cellular necrosis, leading to dilated heart failure, with slight increases in heart weight that are likely dilatory in nature (sarcomeres added in series) and secondary to neurohumoral status.
In contrast to the above interpretation, downregulation of the β2 subunit using a gene transfer strategy in aortic banded rats showed prevention of the hypertrophic response (32). However, this knockdown approach of β2 did not reduce systolic function in these animals, and α1C protein was not reduced compared with that in sham viral-injected hearts. Hence, it remains uncertain how these data relate to our observations in α1C–/+ and α1Cfl/fl-Cre mice that did show reduced current, α1C protein, and cardiac function. Indeed, Rosati et al. recently showed that heart-specific deletion of α1C resulted in reduced cardiac function and early postnatal lethality, although heterozygous deleted mice showed no baseline phenotype in their analysis (33). These disparities notwithstanding, perhaps the most important observations are from clinical trials with Ca2+ channel antagonists, which failed to show protective effects in patients with heart failure with systolic dysfunction and, in some cases, showed signs of worsening disease and increased mortality (34, 35). Thus, reducing Ca2+ influx through LTCCs in cardiomyocytes may not be a desirable therapeutic strategy for systolic heart failure, consistent with our observations of worsening disease due to graded deletion of α1C in our multiple genetic strategies.
We did not anticipate that a genetic-based reduction in LTCC current would induce spontaneous cardiac hypertrophy or dramatically enhance hypertrophy after pressure overload stimulation. Our new working hypothesis is that, in the absence of sufficient LTCC current, SR Ca2+ release is sensitized in an attempt to maintain cardiac contractility, leading to hypertrophic remodeling through calcineurin/NFAT activation. The reduction in cardiac function likely causes a compensatory neuroendocrine response through β-adrenergic receptors that in turn mediates PKA and CaMKII activation to phosphorylate nodal Ca2+ handling and contractile proteins in an attempt to augment contractility (36, 37). In this manner, phosphorylation of the LTCCs and RyR2s would attempt to compensate for the reduced Ca2+-induced Ca2+ release relationship by allowing the RyR2 to leak and open with substantially less trigger Ca2+. This would tend to elevate Ca2+ in the cleft microenvironment in diastole, which may be a more potent Ca2+ pool in activating calcineurin and CaMKII (22, 38, 39). Indeed, the T-tubule/RyR2 junctions align with sarcomeric Z-discs in which calcineurin/NFAT are anchored through calsarcin and α-actinin, suggesting they could directly sample Ca2+ in this microenvironment (40). Moreover, leaky RyRs in the heart associated with PKA/CaMKII activation and oxidation and nitrosylation, downstream of β-adrenergic signaling, are known disease determinants for hypertrophy and worsening heart failure (25, 41). This contention is entirely consistent with a recent report from Wehrens and colleagues, in which they made a RyR2 knockin mouse model that leaks Ca2+, leading to greater cardiac hypertrophy and calcineurin/NFAT activation (42). Thus, enhanced β-receptor signaling and elevated resting Ca2+ in the cleft microenvironment due to RyR2 leak (or Ca2+ from another source) likely serve as interrelated effects, leading to a greater hypertrophic response in α1C-deleted mice. In addition, decreased Ca2+ influx via α1C could decrease Ca2+-dependent inactivation of ICa-L and thereby also contribute to increasing junctional cleft Ca2+ concentration. Indeed, we observed a significant elevation in the fast component of inactivation (τf) in both α1C–/+ and α1C–/fl-Cre myocytes (Supplemental Figure 2).
The proposed elevation in resting Ca2+ in the RyR2 cleft microenvironment should render these channels more likely to open with less LTCC influx. Another means of accomplishing this compensatory alteration could be to change NCX1 activity. Indeed, deletion of NCX1 from the mouse heart led to a compensatory reduction of nearly 60% in LTCC current, while increased NCX1 activity in the heart, due to transgene-mediated overexpression, produced more LTCC current (43, 44). Moreover, LTCC activity directly influences NCX1 activity in the cleft microenvironment, leading to the hypothesis that reduced LTCC activity should produce less NCX1 activity, leading to increased cleft Ca2+ and increased gain in ECC (45). While we did not directly observe a change in NCX1 current from isolated adult cardiomyocytes from α1C–/fl-Cre mice, in vitro conditions are probably not appropriate to observe the proposed compensatory alterations that might result in less Ca2+ extrusion by NCX1 in vivo (Supplemental Figure 4). Finally, another compensatory alteration that might occur with reduced LTCC activity is through increased transient receptor potential canonical (TRPC) channel activity. Indeed, TRPC3 or TRPC6 overexpression in the heart induces Ca2+ influx, calcineurin activation, and mild hypertrophy (46, 47), and TRPC channels can functionally “couple” with LTCCs to alter each others’ activity (48–50). TRPC channels were also shown to couple to RyR1 in skeletal muscle to induce Ca2+ release (51). Despite these relationships, we did not observe an increase in store-operated Ca2+ entry in cardiomyocytes from α1C–/fl-Cre mice, suggesting that TRPC channel activity was not affected, at least as suggested by the surrogate measure of store-operated entry (Supplemental Figure 4). It should also be noted that we did not observe an increase in TTCC expression in hearts from α1C-deleted mice (data not shown). Despite a lack of changes in these and other Ca2+ handling proteins and currents in α1C-deleted hearts, we did observe a mild increase in heart rate and a reduction in mean arterial blood pressure compared with those of control mice (Supplemental Figure 5).
While the ability to directly measure Ca2+ in the cleft microenvironment is lacking, many additional indirect lines of evidence support a mechanism whereby Ca2+ in this compartment is elevated and functions as the primary disease effector in α1C-deleted mice. First, metoprolol reversed manifestations of heart disease in α1C–/+ mice, likely by antagonizing the neuroendocrine signaling machinery that enhances Ca2+ leak from RyR2 or that otherwise contributes to Ca2+ dysregulation secondary to CaMKII and PKA signaling. Second, augmentation in TTCC current with the α1G transgene rescued hypertrophy in α1C-deleted mice by providing more systolic Ca2+ (presumably then reducing diastolic Ca2+), although these mice still developed heart failure and perished prematurely (Supplemental Figure 3). The simplest interpretation of these observations is that the additional peak Ca2+ release from TTCCs obviated the need for compensatory increases in cleft Ca2+, though because the TTCC is not coupled in the same manner as the LTCC, it ultimately still resulted in lethality and arrhythmia. Third, despite dramatically reduced SR Ca2+ levels, direct measures of relative SR leak and SR Ca2+ sparks were enhanced in α1C–/fl-Cre mice, suggesting that conditions are appropriate for RyR2 leak. Fourth, calcineurin/NFAT signaling was dramatically increased in the hearts of α1C–/fl-Cre mice. Fifth, deletion of NCX1 in the heart, which also leads to a dramatic compensatory reduction in LTCC current (60%) (43), similarly led to much greater cardiac hypertrophy and dysfunction in young adult mice after TAC stimulation (45).
There may also be physiologic relevance to our results, such that LTCC current reductions might occur in the natural course of heart failure disease etiology. While it remains controversial whether LTCC current is reduced in cardiomyocytes from rats or humans with heart failure (15), there is general agreement that myocytes from failing human hearts show less β-adrenergic LTCC reserve (52). In the failing mouse heart, we reliably observed a significant (P < 0.05) reduction in α1C protein after 8 weeks of pressure overload stimulation (Supplemental Figure 6). Myocytes from these same WT hearts showed hypertrophy, a reduction in the Ca2+ transient, and a significant reduction in LTCC current (Supplemental Figure 6). Thus, a reduction in LTCC “function” (actual or reserve activity) may indeed be a physiologic consequence of heart failure, leading to the same increase in resting cleft Ca2+ through RyR2 leak, leading to secondary hypertrophy signaling. Interestingly, BAY K8644-induced maximal ICa-L in α1C–/fl-Cre myocytes was significantly (P < 0.001) blunted compared with that of control myocytes (data not shown), similar to what has been shown in failing human ventricular myocytes (52). At the very minimum, our data at least suggest caution in applying LTCC antagonists for the treatment of heart failure, given the prominent disease-predisposing pathway that arose in the mouse with reduced cardiomyocyte-specific LTCC activity.
α1C gene-targeted mice were described previously as were α1C mice with targeted loxP sites to permit conditional gene deletion with Cre recombinase expression (7, 53). Transgenic mice expressing Cre recombinase or MerCreMer from the α-MHC promoter were described previously (54, 55). NFAT-luciferase, tetracycline transactivator, and cardiac-specific α1G overexpressing transgenic mice were described previously (27, 56, 57). Use of animals in this study was approved by the IACUC at the Cincinnati Children’s Hospital Medical Center.
Western blot analysis was performed using mouse ventricles snap frozen in liquid nitrogen and stored at –70°C. Ventricles were homogenized in modified RIPA buffer, containing 50 mM Tris (pH 7.4), 150 mM NaCl, 0.25% sodium deoxycholate, 1 mM EDTA, and protease inhibitors. Homogenates were centrifuged at 20,817 g for 10 minutes, and the supernatants were used for blotting. Six to twenty-five micrograms of protein were loaded on 6%–15% SDS polyacrylamide gels suitable for detecting specific proteins relative to their molecular weights. Antibodies against the α1C subunit (Alamone Labs), α1G (NeuroMab), α1H (NeuroMab), α1D (NeuroMab), and GAPDH (Fitzgerald Industries International) were used. Chemifluorescence detection was performed with the Vistra ECF reagent (Amersham Pharmacia Biotech) and scanned with a Bio-Rad Gel Documentation center. For assessing RyR modifications, hearts were isotonically lysed in 1.0 ml of a buffer containing 50 mM Tris-HCl (pH 7.4), 20 mM NaF, 1.0 mM Na3VO4, and protease inhibitors. RyR2 was immunoprecipitated from the sample with a RyR2 antibody (4 μg Ab5029) in 1.0 ml of a modified RIPA buffer (50 mM Tris-HCl [pH 7.2], 0.9% NaCl, 5.0 mM NaF, 1.0 mM Na3VO4, 1% Triton X-100, and protease inhibitors) for 1 hour at 4°C. The immune complexes were incubated with protein A Sepharose beads (Sigma-Aldrich) at 4°C for 1 hour, and the beads were washed 3 times with RIPA buffer. To determine channel oxidation, the carbonyl groups in the protein side chains within the immunoprecipitate were derivatized to 2,4-dinitrophenylhydrazone (DNP-hydrazone) by reaction with 2,4-dinitrophenylhydrazine (Oxyblot Protein Oxidation Detection Kit, Millipore). Proteins were separated by SDS-PAGE (6% for RyR, 15% for calstabin) and transferred onto nitrocellulose membranes for 1 hour at 200 mA (SemiDry transfer blot, Bio-Rad). After incubation with blocking solution (LICOR Biosciences) to prevent nonspecific antibody binding, immunoblots were developed with an anti-RyR antibody (Affinity Bioreagents), RyR phospho-specific antibodies (RyR2-P2808 or RyR2-P2815; 1:5,000), or anti-calstabin (Santa Cruz Biotechnology Inc.). All immunoblots were developed and quantified using the Odyssey Infrared Imaging System (LICOR Biosystems) and infrared-labeled secondary antibodies.
Ca2+-tolerant cardiomyocytes were selected after a standard isolation procedure from whole hearts placed in Tyrode solution with additives (120 mM NaCl, 5.4 mM KCl, 1.2 mM NaH2PO4, 5.6 mM glucose, 20 mM NaHCO3, 1.6 mM MgCl2, 10 mM 2,3-butanedione monoxime [BDM], and 5 mM taurine; buffer A). Hearts were then perfused with buffer containing liberase blendzyme (Roche) at 37°C (10–14 minutes total). After perfusion, the ventricles were disassociated into individual myocytes, filtered, and incubated with 2 μM Fura-2 acetoxymethyl ester (Invitrogen) and pluronic acid for 15 minutes in M199 media with BDM at room temperature. After loading, the cells were washed and resuspended in Ringer’s solution. The photometry measurements were made in Ringer’s solution using a DeltaRam spectrofluorophotometer (Photon Technology International), operated at an emission wavelength of 510 nm, with excitation wavelengths of 340 and 380 nm. The stimulating frequency for Ca2+ transient measurements was 0.5 Hz. Baseline amplitude (estimated by a 340 nm/380 nm ratio) of the Ca2+ signal was acquired, and data were analyzed using Felix and Clampfit software. SR Ca2+ leak and Ca2+ spark measurements were made as previously described (58, 59). Hearts were quickly excised and washed in chilled Ca2+-free Tyrode solution, followed by retrograde perfusion with buffer containing collagenase II (Worthington) at 37°C for 6 to 10 minutes. The ventricles were subsequently dissociated into single myocytes and filtered, and Ca2+ was increased to reach a final level of 1.8 mM. Ca2+-tolerant cells were then loaded with 1 μM Fluo4-AM and 20% pluronic acid (Invitrogen) for 30 minutes at room temperature. The myocytes were plated on glass cover slips coated with laminin (Invitrogen) and washed for 20 minutes to get rid of excess dye and to allow deesterification used for Ca2+ leak or spark measurements. For Ca2+ leak measurements, the coverslips were mounted on an inverted Nikon microscope and electrically stimulated at 0.5 Hz at least 20 times in 1.8 mM Ca2+ normal tyrode to reach steady state. Cells were excited at 490 ± 5 nm, and emission was recorded at 530 ± 20 nm. After stopping the stimulation, the solution was rapidly switched to a Na+- and Ca2+-free solution (140 mM LiCl, 1 mM MgCl2, 4 mM CsCl, 10 mM HEPES, 1 mM EGTA, 10 mM glucose) for 30 seconds, so the Na+-Ca2+ exchanger was blocked, and little or no Ca2+ could leave or enter the cell. One millimolar tetracaine (Sigma-Aldrich) was added to block the RyRs and thus inhibit leak from the SR. After 20 seconds, the solutions were switched back to Na+- and Ca2+-free tyrode for a duration of 10 seconds before 10 mM caffeine was added to deplete the Ca2+ stores. The difference in levels of baseline Ca2+ with and without tetracaine is considered Ca2+ leak, and the difference in levels between basal and peak total cytosolic Ca2+ transient in the presence of caffeine was an index of SR Ca2+ content. Ca2+ values were calculated based on the assumption of a diastolic Ca2+ value of 0.12 μM.
Ca2+ sparks were recorded with an Olympus Fluoview FV 1000 in line scan mode. The cells were stimulated at 0.5 Hz at least 20 times before stimulation was stopped, and sparks were recorded at rest. The fluorophore was excited with an argon laser at 488 nm, and emission maximum was recorded at 516 nm. Sparks were counted using a custom made algorithm (IDL, ITT Visual Information Solutions) and calculated as sparks/s/100 μm.
Patch clamp experiments were used to measure ICa-L in myocytes. To isolate myocytes, after perfusion, the ventricles were separated from the atria, minced, and gently agitated in low Cl–, high K+ Kraft-Bruhe (KB) solution consisting of 50 mM glutamic acid, 40 mM KCl, 20 mM taurine, 20 mM KH2PO4, 3 mM MgCl2, 10 mM glucose, 1 mM EGTA, and 10 mM HEPES (pH 7.4). The dissociated cells were filtered through a nylon mesh and stored at 4°C in KB solution until use. Only Ca2+-tolerant cells with clear cross-striations and without spontaneous contractions or substantial granulation were selected for the experiments. All patch clamp experiments were conducted at room temperature (20°C–23°C) using a patch clamp amplifier (Axopatch200A; Axon Instruments). The recorded currents (ICa-L) were filtered at 2 kHz through a 4-pole low-pass Bessel filter and digitized at 5 kHz. The experiments were controlled using pClamp software (Axon Instruments) and analyzed using Clampfit. Current recordings were performed in bath solution superfused with the following Na+-free solution: 2 mM CaCl2, 5 mM 4-aminopyridine, 136 mM tetraethylammonium-Cl (TEA-Cl), 1.1 mM MgCl2, 25 mM HEPES, and 22 mM glucose (pH 7.4 with TEA-OH). The pipette solution contained 100 mM cesium aspartate, 20 mM CsCl, 1 mM MgCl2, 2 mM Mg-ATP, 0.5 mM Na2-GTP, 5 mM EGTA, 5 mM HEPES (pH 7.3 with 1 N CsOH). ICa-L was measured by applying depolarizing voltage steps (380 ms) –40 mV to +60 mV, respectively, in 10-mV increments.
ECC efficiency in both control and α1C-/fl-Cre myocytes was evaluated by simultaneous recording of ICa-L and [Ca2+]I with a whole-cell voltage clamp technique. Myocytes were placed in a chamber mounted on an inverted Nikon microscope and perfused with normal Tyrode salt solution containing 150 mM NaCl, 5.4 mM KCl, 1 mM CaCl2, 1.2 mM MgCl2, 10 mM glucose, 2 mM sodium pyruvate, and 5 mM HEPES (pH 7.4) at 35°C. The pipette solution was composed of salts: 120 mM Cs-aspartate, 10 mM NMDG, 20 mM TEA-Cl, 2.5 mM Tris-ATP, 0.05 mM Tris-GTP, and 5.4 mM KCl (pH 7.2 with CsOH). Na+ currents were eliminated by holding the cell at –40 mV, and K+ currents were suppressed by low intracellular K+ and extracellular Cs+ and intracellular TEA+. ECC efficiency was studied in myocytes with matched SR Ca2+ load. For control myocytes, 10 consecutive depolarizations to +10 mV for 100 ms at 1 Hz were applied before each test potential, and, for α1C–/fl-Cre myocytes, 10 consecutive depolarizations to +10 mV for 400 ms were applied before each test potential. SR Ca2+ load was measured with a caffeine spritz, and peak caffeine-induced Ca2+ transient amplitude was considered as the index of SR Ca2+ content. Only cells with comparable SR load were used for data analysis. The gain of ECC was calculated by dividing the amplitude of Ca2+ transient by the amplitude of ICa-L at each voltage.
To measure INCX, myocytes were depolarized to –45 mV from a holding potential of –90 mV to inactivate Na+ current. The voltage was then stepped to +60 mV and ramped down to –90 mV to induce remaining currents. When steady state was reached, the protocol was repeated in the presence of 5 mm NiCl2. INCX is defined as the nickel-sensitive current induced during the ramped potential. Pipette solutions contained 45 mM CsCl, 55 mM Cs-methanesulfonic acid, 10 mM ATP-Tris, 0.3 mM GTP-Tris, 20 mM HEPES, 5 mM BAPTA, 10.8 mM MgCl2, 2.21 mM CaCl2, and 14 mM NaCl (pH 7.3 with CsOH). Bath solution contained 137 mM NaCl, 1.0 mM MgCl2, 5.4 mM CsCl, 1.0 mM CaCl2, 10 mM dextrose, and 10 mM HEPES (pH 7.4 with NaOH) plus 10 μm nifedipine.
Mice from all genotypes or treatment groups were anesthetized with isoflurane, and echocardiography was performed using a Hewlett Packard 5500 instrument with a 15-MHz microprobe. Echocardiographic measurements were taken on M-mode in triplicate for each mouse. For pressure overload, 10-week-old mice of each genotype were subjected to a TAC or sham surgical procedure as previously described (56). Pressure gradients across the constriction were measured by Doppler echocardiography as previously described (27). Alzet osmotic minipumps (no. 2002; Durect Corp.) containing Iso (60 mg/kg/d) or PBS were surgically inserted dorsally and subcutaneously in 2-month-old mice under isoflurane anesthesia. The β-adrenergic receptor antagonist metoprolol (2 g/ml, Sigma-Aldrich), diltiazem (450 mg/l, Sigma-Aldrich), and verapamil (400 mg/l, Sigma-Aldrich) were given in drinking water as previously reported (19, 29, 60). Swimming exercise protocol, which called for 21 days of exercise to induce physiological hypertrophy, was described previously (56). CsA (Sandimmune, Novartis) was administered subcutaneously twice daily at 10 mg/kg for 21 days.
Phosphatase activity was measured by using the calcineurin assay kit (Enzo Life Sciences) according to the manufacturer’s instructions. Cardiac lysates from either control or α1C–/fl-Cre mice were isolated, and calcineurin activity was measured as the dephosphorylation rate of a synthetic phosphopeptide substrate (RII peptide) in the presence or absence of EGTA. The amount of PO4 release was determined photometrically by using the Biomol Green reagent (Enzo Life Sciences).
All results are presented as mean ± SEM. Statistical analysis was performed with unpaired 2-tailed t test (for 2 groups) and 1-way ANOVA with Bonferroni correction (for groups of 3 or more). P values of less than 0.05 were considered significant.
This work was supported by grants from the NIH (to J.D. Molkentin, D.M. Bers, A.R. Marks, and S.R. Houser). J.D. Molkentin was also supported by the Howard Hughes Medical Institute. S.A. Goonasekera was supported by a local affiliate of the American Heart Association, R.N. Correll was supported by an NIH postdoctoral fellowship, and M. Auger-Messier was supported by Heart and Stroke Foundation of Canada.
Conflict of interest: The authors have declared that no conflict of interest exists.
Citation for this article: J Clin Invest. 2012;122(1):280–290. doi:10.1172/JCI58227.