Transgenic (TG) mice with heart-specific overexpression of murine PR72 were generated by the transgenic animal and genetic engineering models core facility at the University of Münster. The αMHC-PR72 transgene was injected into the pronuclei of fertilized FVB/N mouse eggs. Southern blotting was used to identify six transgenic founder mice, which were then cross-bred with wild-type (WT) mice. The founder mouse that produced TG offspring with the highest levels of overexpression was selected for breeding littermates for subsequent experiments. All experiments were performed on 20- to 24-week-old mice. Balanced proportions of females and males were always used and animals were bred on a FVB/N strain background. All animal handling and maintenance procedures were conducted in accordance with approved protocols by the animal welfare committee of the University of Munster and the LANUV (NRW, Germany; ID 81-02.04.2021.A151). These protocols also complied with the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals.

Isolation of ventricular cardiomyocytes followed an approved protocol (17). Mice were killed by cervical dislocation, and the heart was immediately removed. The cannulated heart was then connected to a modified Langendorff apparatus and retrogradely perfused for 5 min with a perfusion buffer (17) at a flow rate of 2.5 ml/min. For another 7.25 min, digestion was performed with an enzyme solution containing Liberase DH. The aorta and atria were separated from the heart. The remaining heart was minced in a 5 ml enzyme stop solution. After 10 min of incubation, the sedimented pellet was transferred into 10 ml of an enzyme stop solution, filtered through nylon gauze, and centrifuged at 500 rpm for 1 min. The cell pellet was then resuspended in 10 ml of perfusion buffer and the Ca²⁺ concentration was raised stepwise to 1 mM. Then, centrifugation was performed again for 1 min at 500 rpm, and the cell pellet was resuspended in perfusion buffer containing 1 mM CaCl₂. Cardiomyocytes were kept at room temperature and used within 6 h of isolation.

Isolated mouse cardiomyocytes were immobilized by a 10 min incubation with a 4% formaldehyde/PBS solution, followed by a PBS wash, placed on poly-Lysine-coated slides to adhere for a 30-min incubation and permeabilized with a solution containing PBS and 0.2% Tergitol 15-S-9 (NeoFroxx GmbH) for 10 min. The cells were then incubated in a blocking solution consisting of 2% goat serum (Sigma Aldrich) and 1% bovine serum albumin (BSA; Sigma Aldrich) in PBS-T (PBS with 1% Tween 20) for 1 h. This was followed by treatment with an antibody against human PR72 (1:200, HPA-035829, Sigma Aldrich) in the blocking solution for 1 h. After thorough washing (typically three times for 10 min each in PBS), the cardiomyocytes were incubated for 1 h at room temperature with a secondary antibody, Alexa Fluor 488 (1:500, Alexa Fluor® 488 chicken anti-rabbit IgG, Invitrogen). After another washing step, the cells were blocked with unconjugated Fab fragments [100 µg/ml goat anti-mouse IgG (H + L) unconjugated; Dianova] for 1 h. Antibodies specific for GAPDH (1:300, AM4300, Thermo Fisher) or sarcomeric α-actinin (1:200, A7732, Sigma Aldrich) was then applied for 1 h, followed by washing and incubation for 1 h with a secondary antibody, Alexa Fluor 594 (1:500, Alexa Fluor® 594 goat anti-mouse IgG, Invitrogen). Fluorescence signals were acquired using a confocal laser scanning microscope (LSM 710; Carl Zeiss AG) equipped with laser lines at 488 nm and 543 nm, operated at 2% power with 16-fold line averaging and a 33-μm pinhole. Imaging was performed with a Plan Neofluar 25x/0.8 objective.

See supplementary methods section.

Cells were cultured (37°C and 5% CO₂) in Dulbecco's modified Eagle's medium (DMEM), supplemented with FCS (10% v/v, Cytogen) and Penicillin Streptomycin (100 U/ml, Sigma-Aldrich). DMEM low glucose (Sigma-Aldrich) was used for HEK293 cells (ATCC).

To investigate the protein-protein interaction of PR72 with the catalytic PP2A subunit, pull-down assays were performed. HEK293 cells were transiently transfected with the N-terminal RGS6xHis-tagged PR72 cDNA or the empty RGS6xHis-pcDNA3.1 vector as a control in 75-cm² culture flasks. Xtreme gene-HP DNA (DNA ratio 1:1; Roche Diagnostics) was used for transfection according to the manufacturer's protocol. After 48 h, the cells were washed with PBS, scraped in 4 ml PBS, centrifuged at 100 × g and lysed with 600 µl lysis buffer (50 mM NaH₂PO₄, 300 mM NaCl, 1% NP40, pH 8.0, supplemented with protease and phosphatase inhibitors). Lysates were sonicated four times for 15 s, incubated on ice for 20 min and centrifuged at 21,000 × g for 10 min. The protein concentration of the supernatant corresponding to the crude lysate was measured by Bradford assay and adjusted to 4 mg/ml with lysis buffer; imidazole was added to a final concentration of 20 mM. For the pull-down assay, 100 µl of bead solution corresponding to 5 µl of magnetic beads (SERVA Ni-NTA Magnetic Beads) were used for each probe according to the manufacturer's protocol. Raw lysates and eluates from RGS6xHis-tagged PR72 and control-transfected HEK293 cells were loaded on a SDS polyacrylamide gel at a 5 to 1 ratio.

Subcellular fractions were prepared from whole heart homogenates by differential centrifugation as described (18). This protocol allows the preparation of cardiomyocyte-derived fractions enriched in different cellular organelles. The supernatant obtained after the first centrifugation of heart homogenates represents the cytosol. It can be suggested that most of the fibroblasts, constituting the main fraction of non-cardiomyocyte cells, are enriched in the pellet after the first centrifugation. Thus, the cytosol fraction should contain mostly proteins derived from cardiomyocytes. Moreover, this preparation protocol enables the preparation of purified cardiac contractile myofibrils that are virtually free of contamination by mitochondrial, sarcolemmal, and sarcoplasmic reticulum (SR) membranes (19). The membrane-enriched fraction contains sarcolemmal as well as SR proteins. However, it cannot be excluded that fractions contain small quantities of non-cardiomyocyte cells.

As described previously (20), TG and WT hearts were immersed into 4% saline-buffered formaldehyde and embedded in paraffin. Transversal sections at a thickness of 5 μm were mounted on glass slides. For estimation of interstitial fibrosis, sections were stained with Masson–Goldner's Trichrome. To estimate cell diameters, tissue sections were subjected to hematoxylin-eosin (HE) staining. Within each HE-stained section, 100 longitudinally oriented cells were randomly selected. The transverse diameters, which were orthogonal to the longitudinal cell axes, were subsequently quantified at the widest region of each selected cell. This measurement was performed using the two-point measurement tool available in the NIS-Elements AR software (Nikon Instruments).

The preparation of [³²P]-labeled phosphorylase α is based on an established protocol (21). For the assay, heart samples obtained from Langendorff experiments (section “Heart tissue samples”) were mixed with a buffer that contained EDTA resulting in complexation of free Ca²⁺ and Mg²⁺ ions, which completely inhibits the catalytic activity of PP's, except for PP1 and PP2A. The dilution of the samples was chosen to convert no more than 18% of the maximum releasable [³²P]. The dephosphorylation reaction was performed for 20 min at 30°C by adding 20 μl of phosphorylase α. Subsequently, an equal volume of 50% trichloroacetic acid was added to the mixtures which stopped the reaction. Moreover, 20 μl of ultrapure water and 30 μl of bovine serum albumin solution were added. The mixtures were centrifuged at 14,000 × g and 4°C for an additional 10 min after incubation on ice for 10 min. Thereafter, 50 μl of the supernatant was analyzed in the scintillation counter. The specific activity of the phosphorylase α substrate was calculated from the difference between total counts and background radioactivity. To distinguish between PP1 and PP2A, samples were incubated in the absence and presence of 3 nM okadaic acid. At this concentration, okadaic acid completely inhibits PP2A activity, leaving PP1 activity unaffected (22).

The cAMP-concentration of heart samples obtained from Langendorff experiments (section “Heart tissue samples”) were measured by Gs Dynamic HTRF Kit (CISBIO). According to manufacturer instructions, 10 µg protein for each sample were used. After 1 h of incubation fluorescence signals were detected with Mithras LB 940 Microplate Analyzer (Berthold Technologies, Bad Wildbad, Germany). cAMP-concentrations were calculated by a cAMP-standard curve in Origin 2022b software.

Hemodynamic measurements were performed according to an approved protocol (23). Experimental animals were anesthetized with an i.p. dose of 400 mg/kg of a 2% solution of tribromoethanol. For data acquisition, a 1.4 Fr pressure-volume catheter (model SPR-839) was inserted into the left ventricle. The catheter was optimally placed when sinusoidal pressure curves were recorded using the MPVS-400 system and LabChart software (ADInstruments). After a 10 min stabilization period, heart rates and left ventricular pressures were recorded and analyzed. At the end of the experiment, animals were killed by a repeated dose of tribromoethanol, followed by cervical dislocation in deep anesthesia.

Contractile function and intracellular Ca²⁺ cycling of cardiomyocytes were studied by use of a dual emission photometry system combined with a CCD camera (Myocyte Calcium and Contractility Recording System; IonOptix Ltd.) attached to an epifluorescence microscope (Nikon). Cardiomyocytes were initially incubated with 23.3 μM Indo-1/AM (Molecular Probes) for 10 min in a small perfusion chamber mounted on the microscope. This chamber was perfused with bath solution (composition in mM: 140 NaCl, 5.8 KCl, 0.5 KH₂PO₄, 0.4 Na₂HPO₄, 0.9 MgSO₄, 10 HEPES, 1 glucose, and 2 CaCl₂, pH = 7.45), and cardiomyocytes were field-stimulated with 0.5 Hz under basal conditions as well as in the presence of 1 μM isoprenaline or 3 nM okadaic acid. The emitted fluorescence was recorded at wavelengths of 405 and 495 nm. The ratio of both wavelengths was taken as an index of cytosolic Ca²⁺ concentration. The maximum peak amplitude of Ca²⁺ transients was related to baseline (Δ Indo-1 ratio). The shortening of sarcomere length (SL) was detected simultaneously. The maximum SL shortening was normalized to resting sarcomere length (Δ SL shortening).

Whole-cell patch-clamp recordings were performed in voltage clamp configuration to record voltage gated L-type Ca²⁺ currents (I_CaL) in isolated ventricular myocytes. Measurements were conducted using the ruptured patch method and were performed at room temperature (22 ± 1°C, RT). Borosilicate glass capillaries (GB150TF-8P, Science Products, Hofheim, Germany) were pulled to a resistance of 3.5 ± 1 MΩ. Data was acquired and filtered at 10 kHz using an EPC-800 amplifier and sampled with an 18bit A/D converter InstruTech ITC-18 under the control of the PatchMaster software (HEKA Elektronik, Lambrecht, Germany). Series resistance was compensated for at least 60%. The bath solution contained (in mM): 136 NaCl, 5.4 KCl, 1.8 CaCl₂, 1 MgCl₂, 5 HEPES, 0.33 NaH₂PO₄, 10 TEA-Cl, 0.1 BaCl₂, and 10 glucose, pH 7.4. Pipettes were filled with a solution containing (in mM): 120 CsCl, 1 MgCl₂, 5 Na₂ATP, 10 TEA-Cl, 10 EGTA, and 10 HEPES, pH 7.2. Current density (pA/pF), current-voltage (IV) relations and voltage dependence of activation and inactivation were calculated as described before (24, 25).

Measurement of SR Ca²⁺ release was performed according to a revised protocol (26). Briefly, isolated cardiomyocytes were perfused with Tyrode's solution and at least 10 Ca²⁺ transients were recorded at 0.5 Hz using the IonOptix system as described for the detection of Ca²⁺ transients. After a 10 s pause in stimulation, cells were superfused with 10 mM caffeine for 45 s using a rapid application aid. Within a few seconds, the cardiomyocytes responded immediately with a complete SR Ca²⁺ release and increased contraction (27). Data were recorded and analyzed, whereby the SR Ca²⁺ content corresponds to the maximum (peak) amplitude of the caffeine-induced Ca²⁺ transient.

Cardiomyocytes were incubated with 4 μM Fluo-4 in the perfusion chamber of a confocal microscope (LSM710; Zeiss). Excess dye was removed, and cells were then perfused with Tyrode's solution. During a pause in stimulation, a line scan (X-T scan) was performed (excitation: 488 nm; emission range: 505–622 nm). Detection and analysis of Ca²⁺ sparks were performed using ImageJ software (https://imagej.nih.gov/ij/) and the Sparkmaster plugin (28).

Samples of cardiac tissues were collected from TG and WT mice. The mice were either left untreated or treated with an intraperitoneal injection of 4 mg isoprenaline per kg body weight. Two minutes after injection, the hearts were removed from euthanized mice, washed, and rapidly frozen in liquid nitrogen. Additional samples were obtained from spontaneously beating TG and WT hearts, retrogradely perfused in a Langendorff apparatus. After a stabilization period of 25 min, the hearts were either left untreated or stimulated with 1 µM isoprenaline for 10 min. The beating hearts were promptly frozen in liquid nitrogen.

Quantitative real-time PCR of WT and TG heart tissue samples was performed as previously described with minor modifications (20). Addionally, mRNA levels of Gapdh (Glyceraldehyde 3-phosphate dehydrogenase) were determind using AAC TTT GGC ATT GTG GAA GG as a forward and GGA TGC AGG GAT GAT GTT CT as a reverse primer sequence.

Heart homogenates were prepared following the previously described method (29). For immunoblot analysis, the heart homogenates were mixed with reducing Laemmli loading buffer to achieve a final SDS concentration of 2.5%. The samples were then incubated at 30°C for 10 min and separated on either 10% SDS polyacrylamide gels or 4%–15% Criterion TGX Precast gels (BioRad). The proteins were transferred from the gels to nitrocellulose membranes using an electric current of 3 or 6 Ah. The membranes were incubated overnight at 4°C with specific antibodies against the following proteins: PLN (1:2000, Badrilla), PLN phospho-Ser¹⁶ (1:1000, Badrilla), PLN phospho-Thr¹⁷ (1:1000, Badrilla), RyR (1:1000, Thermo Fisher Scientific), RyR phospho-Ser²⁸⁰⁸ (1:1000, Badrilla), RyR phospho-Ser²⁸¹⁴ (1:5000, Badrilla), TNI (1:1000, Cell Signaling), TNI phospho Ser^23/24 (1:1000, Cell Signaling), MyBP-C (1:1000, Proteintech Group), MyBP-C phospho Ser²⁸² (1:1000, Enzo), PKA-Cα (1:1000, Cell Signaling), CaMKII (1:1000, Thermo Fisher Scientific), Cav1.2 (1:500, Alomone), SERCA2a [1:1000, (30)], NCX (1:1000, Swant Inc.), PPP2R3A/PR72 (1:1000, Sigma Aldrich), PPP2CA (1:2000, Proteintech Group), CSQ2 (1:2500, Thermo Fisher Scientific). The bound antibodies were detected by incubating the membranes with secondary antibodies (ECL rabbit/mouse IgG, HRP-linked whole antibody, GE Healthcare) for 2 h. The signals were visualized using the SERVALight Helios CL HRP WB Substrate kit (Serva) and a ChemicDoc XRS detection system. The antibody signals were quantified using the TotalLab TL120 software.

The results are presented both in terms of individual data points and as means with corresponding standard deviations (+SD), if not indicated otherwise. In the context of single-cell measurements, the data points represent individual cardiomyocytes (“n”), whereas in the context of multicellular measurements, the data points correspond to individual hearts or mice (“N”). Statistical analyses were conducted using either GraphPad Prism (Graphpad Software Inc.) or SigmaPlot software (Systat Software GmbH). To assess the normality of residuals, the following tests were employed: Anderson-Darling (A2*), D'Agostino-Pearson omnibus (K2), Shapiro-Wilk (W), and Kolmogorov-Smirnov (distance) tests. Depending on the specific research question, the comparison groups being examined, and the normality characteristics of the residuals, various statistical tests were utilized. The respective tests are indicated in the figure captions. Statistical significance was determined by a threshold of P < 0.05, denoting group differences that were considered to be statistically significant.

First of all, we examined the expression level of PR72 in different tissues. We found that PR72 is mainly expressed in the myocardium, including the ventricles and atria, and represents the more abundant PPP2R3A splice variant in the mouse heart (Figure 1A). After clarifying the expression of PR72, the question arose as to whether this regulatory B'' subunit interacts with other subunits of the PP2A holoenzyme. Biochemical experiments revealed a direct interaction between PR72 regulatory and catalytic subunits by immobilizing 6xHis-tagged PR72 on Ni columns, which resulted in pulldown of PP2Ac (Figure 1B). Recombinant PR72 was expressed in HEK293, and the total cell lysate for the assay was also prepared. After investigating the organ-specific expression of PR72 and its interaction with PP2Ac, we examined the intracellular localization of PR72 in cardiac tissue. PR72 was mostly detected in the cytosolic fraction of the dissected heart tissue (Figure 1C). The purity of the cytosolic fraction was demonstrated using two different marker proteins (GAPDH and myoglobin) in the immunoblot analyses (Figure 1C, Supplementary Figure 1). The enrichment of membrane preparation and contractile filaments were also examined using specific marker proteins (Supplementary Figure 1).

To test the functional relevance of PR72, we generated a heart-directed transgenic mouse model under the control of the well-established αMHC promoter. Southern blot analysis identified a total of six founders (Supplementary Figure 2A), with one dying and another producing only wild-type descendants. Founder line 4 exhibited the highest level of overexpression (2.5-fold) compared to wild-type (WT) littermates (Figure 2A). This line was used for further characterization of the transgenic overexpression model because the expression level of PR72 is comparable to that measured in human heart failure (12). Of the remaining three founders, only line 3 showed weak overexpression (Supplementary Figure 2B). Overexpression of PR72 was accompanied by unchanged expression of PP2Ac and calsequestrin, a myocardial marker protein (Figure 2A). Under basal conditions, there was no difference in PP2A activity between TG and WT littermates (Figure 2B). Stimulation of Langendorff-perfused hearts with 1 μM isoprenaline increased PP2A activity, but there was no difference between genotypes (Figure 2B). Interestingly, the ratio of PP2A to total PP activity changed between genotypes after isoprenaline application (Figure 2C); in other words, the initially increased proportion of PP2A normalizes after β-adrenergic stimulation.

Overexpression was confirmed in immunocytotological measurements (Figure 3). In WT cardiomyocytes, the expression of PR72 was only minimally higher than in negative controls. In a study conducted by DeGrande et al. (12), expression of PR72 along the Z line was detected for the first time in cardiomyocytes. Our study also used confocal microscopy and revealed a striated pattern, that was more pronounced in TG, indicating a sarcomeric organization of PR72 at the Z line (Figure 3). Detailed analysis of PR72 and sarcomeric α-actinin localization showed a match of both proteins (see white arrows), suggesting direct colocalization. A similar pattern was also demonstrated for the detection of GAPDH (a marker of the cytosol) and PR72 (Figure 3). Previous work showed no staining of any specific structure by PR72 (31) in cross-sections of cardiac muscle fibers, suggesting a cytosolic localization. In our immunocytological studies in cardiomyocytes, we detected a diffuse distribution of PR72 in addition to the striated pattern (Figure 3). This may hind to a cytosolic localization of this regulatory subunit of PP2A. In contrast, the negative control showed only autofluorescence.

Overexpression of PR72 was associated with a mild cardiac hypertrophy (Figure 4A), which was not sex-specific. This was not associated with comparable changes at the level of single cardiomyocytes (Figure 4B). Signs of cardiac decompensation could be excluded because both lung and liver weights were unchanged (Supplementary Figure 2C), and there were no signs of developing fibrosis (Figure 4C). However, mRNA levels of skeletal muscle α-actin (ACTA1), which marks the earliest appearance in embryonic cardiomyocytes and, therefore, represents a marker of fibrogenic cell activity, were enhanced in TG compered to WT hearts (Figure 4D). Other marker proteins of hypertrophy and fibrosis tested were unchanged between genotypes.

To study the functional effects of PR72 overexpression on contractility, we first conducted in vivo left ventricular catheterization. We found that basal maximal ventricular pressure (P_max) increased by 8% in TG animals compared to WT (Figure 5A). There were no chronotropic effects on this increase (Figure 5B). The contraction and relaxation velocities (dP/dt_max and dP/dt_min) showed a tendency towards an increase compared to WT (Figures 5C,D).

To determine whether the increased left ventricular pressure measured in the whole animal by catheterization also occurs at the single-cell level, we isolated cardiomyocytes by enzymatic digestion and electrically stimulated them at 0.5 Hz in a perfusion chamber (Figure 6A). Under basal conditions, sarcomere length (SL) shortening was increased by 97% in TG cardiomyocytes compared to WT (Figure 6B). Application of 3 nM okadaic acid had no further effect on either genotype (Figure 6B). Under β-adrenergic stimulation, the increase in SL shortening observed under basal conditions in TG cardiomyocytes was abolished (Figure 6B). The relaxation of transgenic cardiomyocytes, measured as the time to 50% SL relengthening, was 26% faster compared to WT (Figure 6C). The improved relaxation in TG seemed to be influenced by PP2A because the administration of 3 nM okadaic acid further shortened this parameter (Figure 6C). The administration of 1 μM isoprenaline resulted in an unchanged SL relengthening between TG and WT cardiomyocytes (Figure 6C).

Concomitant with the increased SL shortening, there was an 25% increase in the peak amplitude of Ca²⁺ transients under basal conditions in TG cardiomyocytes compared to WT (Figures 7A,B). Inhibition of PP2A increased the peak amplitude of Ca²⁺ transients only in WT cardiomyocytes, while it remained unchanged in TG (Figure 7B). Administration of isoprenaline demonstrated an unchanged peak amplitude of Ca²⁺ transients between the two genotypes (Figure 7B), as observed for SL shortening (Figure 6B). The faster decay in the relaxation time in TG cardiomyocytes compared to WT (Figure 6C) was paralleled by corresponding effects for the Ca²⁺ transients (Figure 7C). This effect was sensitive to PP2A inhibition, as the administration of 3 nM okadaic acid resulted in an enhancement of the Ca²⁺ transient decay only in TG (Figure 7C). Isoprenaline augmented the Ca²⁺ transient decay in both genotypes to different extents compared to basal conditions but towards comparable final effects (Figure 7C).

The increased amplitude and hastened decay of Ca²⁺ transients were accompanied by an unchanged SR Ca²⁺ load under both basal conditions and β-adrenergic stimulation (Figure 8A). To test whether an altered trigger was responsible for the increased Ca²⁺ transient amplitude in TG, we determined the Ca²⁺-induced Ca²⁺ release gain by measuring I_CaL. However, the peak current density was comparable between TG and WT (−10.2 ± 2.7 vs. −10.0 ± 2.8 pA/pF, respectively, n/N = 26–27 cardiomyocytes/5 hearts, unpaired Student's t-test). In addition, the I_CaL inactivation kinetics were unchanged in TG compared to WT (τ₁: 10.8 ± 7.7 vs. 10.6 ± 5.2 ms, respectively; τ₂: 71.7 ± 8.6 vs. 72.7 ± 6.4 ms, respectively, n/N = 26–27 cardiomyocytes/5 hearts, Mann–Whitney U-test).

To investigate the influence of potentially altered release characteristics of the junctional SR complex on Ca²⁺ transients, we determined various Ca²⁺ spark parameters (Figure 8B). The Ca²⁺ spark frequency, which most closely reflects the number of individual or clustered RyRs, was correspondingly unchanged between TG and WT cardiomyocytes (Figure 8C). Ca²⁺ spark amplitude tended to be increased in TG cardiomyocytes but did not reach significance (Figure 8C). The full duration and the full width of Ca²⁺ sparks were unchanged between both genotypes (Figure 8C). However, the spatial spread or size of Ca²⁺ sparks, measured as the full duration at half maximum (FDHM) and the width at half maximum (FWHM), were increased in TG compered to WT cardiomyocytes (Figure 8C). Lastly, the decay of Ca²⁺ sparks was unaffected between TG and WT cells (Figure 8C).

To determine whether the observed contractile and intracellular Ca²⁺ cycling effects were accompanied by corresponding biochemical changes, we investigated the phosphorylation state of key proteins involved in contractility and Ca²⁺ regulation (Figure 9A). Our findings indicated that, under basal conditions, the phosphorylation of phospholamban at threonine-17 was increased by 74% in TG compared to WT (Figure 9B). However, phosphorylation of phospholamban at serine-16, ryanodine receptor type 2 at serine-2808 and serine-2814, myosin-binding protein C at serine-282, and troponin inhibitor at serine-23/24 remained unchanged (Figure 9B). Following β-adrenergic stimulation, the phosphorylation of both phospholamban at threonine-17 and myosin-binding protein C increased in TG compared to WT (Figure 9B). Notably, phosphorylation levels for the other proteins remained unchanged between genotypes under isoprenaline. To investigate whether changes in the β-adrenergic signaling are responsible for the phosphorylation effects, we measured cAMP levels and the protein expression of the catalytic subunit of cAMP-dependent protein kinase. Both parameters remained unchanged between measured groups (Figure 9C). Ca²⁺/calmodulin-dependent protein kinase II expression also showed no differences between groups (Figure 9C). Interestingly, the expression of the Na⁺/Ca²⁺ exchanger (NCX) was decreased in TG compared to WT under basal and ISO-stimulated conditions, whereas no differences were detectable for both Cav1.2 and the SR Ca^2 +ATPase (Figure 9D).