Receptor-independent modulation of cAMP-dependent protein kinase and protein phosphatase signaling in cardiac myocytes by oxidizing agents

The contraction and relaxation of the heart is controlled by stimulation of the b 1 adrenoceptor (AR) signaling cascade, which leads to activation of cAMP-dependent protein kinase (PKA) and subsequent cardiac protein phosphorylation. Phosphorylation is counteracted by the main cardiac protein phosphatases, PP2A and PP1. Both kinase and phosphatases are sensitive to intramolecular disulfide formation in their catalytic subunits that inhibits their activity. Additionally, intermolecular disulfide formation between PKA type I regulatory subunits (PKA-RI) has been described to enhance PKA’s affinity for A-kinase anchoring proteins, which alters its subcellular distribution. Nitroxyl donors have been shown to affect contractility and relaxation, but the mechanistic basis for this effect is unclear. The present study investigates the impact of several nitroxyl donors the experimental nitroxyl donors and the thiol-oxidizing agent diamide on cardiac myocyte protein phosphorylation and oxidation. Although all tested compounds equally induced intermolecular disulfide formation in PKA-RI, only 1-nitrosocyclohexalycetate (NCA) and diamide induced reproducible protein phosphorylation. Phosphorylation occurred independently of b 1 -AR activation, but was atbolished after pharmacological PKA inhibition and thus potentially attributable to increased PKA activity. NCA treatment of cardiac myocytes induced translocation of PKA and phosphatases to the myofilament compartment as shown by fractionation, immunofluorescence and proximity ligation assays. Assessment of kinase and phosphatase activity within the myofilament fraction of cardiac myocytes after exposure to NCA revealed activation PKA and phosphatase in kinase/phosphatase Phosphorylation of cMyBP-C (Ser282) and PLN (Ser16) in lysates of stimulated ARVMs was investigated by IB using the respective phospho-specific antibodies. Chemiluminescence signals detected after a short and a long exposure are shown. Protein loading was examined using an anti- α -actinin antibody. Blots are representative of four independent experiments. D Following incubation with the β 1 -AR antagonist atenolol (AT, 1 µ mol/L), ARVMs were treated with vehicle (control), NCA (100 µ mol/L, 30 min) or ISO (10 nmol/L, 10 min) and cMyBP-C phosphorylation at Ser273, Ser282 and Ser302 analyzed by IB. Blots are representative of six independent experiments. FOR: forskolin, IBMX: 3-isobutyl-1-methylxanthine okadaic acid (OA; 10 nmol/L) and the substrate DiFMUP (500 µ mol/L). Fluorescence reflecting phosphatase activity was monitored over time (representative traces). For each treatment, a phosphatase-free blank sample was included in the measurement. The table shows the activity of active recombinant PP2A-C at each condition, calculated from the slope of the linear regression of averaged RFU values and expressed as Δ RFU/min or % of the activity calculated under control conditions (n = 4-5 experiments). Measured fluorescence was normalized to blank values and expressed as % of the control signal. C The data from B were reanalyzed to reflect the HNO donor incubation times in cell culture experiments. The scatter plots represent average phosphatase activity expressed as Δ RFU/min calculated from the slope of a linear curve fit after 15 min of CXL-1020 or 30 min of NCA treatment (n = 4-5 experiments) to reflect the cell culture conditions. P<0.01 two-tailed Student’s t-test for comparison of each time point with the corresponding vehicle control;

Introduction b1-Adrenoceptor (AR)-mediated activation of cAMP-dependent protein kinase (PKA) is the main pathway that regulates cardiac contractility. Through phosphorylation of substrate proteins that mediate cardiac excitation-contraction coupling, PKA enhances cardiac myocyte inotropy and lusitropy (1). PKA is a heterotetrameric kinase comprising two regulatory and two catalytic subunits. Binding of two molecules of cAMP to each of the two regulatory subunits (PKA-R) induces the release of the active catalytic subunits (PKA-C). PKA kinase activity can be modulated by oxidation (2). Recently, oxidant-mediated formation of antiparallel interdisulfide bonds between Cys 17 and 38 in neighboring PKA-RI subunits was described and shown to induce subcellular translocation of the cytosolic holoenzyme and a substrate-induced release of the catalytic subunits at the target site (3-6). Mice that cannot form the interdisulfide upon stimulation due to Cys17 replacement by non-oxidizable serine were deficient in VEGF-stimulated angiogenesis, supporting the physiological importance of this modification (5). Furthermore, Sglutathiolation at Cys199 or intradisulfide formation between Cys199 and C343 within one catalytic subunit was shown to inhibit kinase activity (7,8). Kinase-mediated phosphorylation is counteracted by phosphatases (9). Thereby, the serine/threonine phosphatase protein phosphatase 2A (PP2A) represents one of the most abundant and important phosphatases in cardiac myocytes. It composes of a scaffold and a catalytic subunit (PP2A-C), which assemble to form the core enzyme. The combination with one out of at least 18 regulatory subunits that confer cellular targeting and thus compartmentalized phosphatase action constitutes the active holoenzyme (10-12), which catalyses the dephosphorylation of various PKA substrates such as cardiac myosin-binding protein C (cMyBP-C) (13), cardiac troponin I (cTnI) (14) and to a lesser extent phospholamban (PLN) (15). Equally to kinases, phosphatases are also susceptible to oxidation with a reported inhibition of phosphatase activity (16-18). Thereby, intradisulfide formation between Cys266 and 269 in PP2A-C (19) as well as between Cys39 and Cys127 in protein phosphatase 1a (PP1a) (12,16,20) were reported as inhibitory redox switches in phosphatases. Pharmacological compounds that release nitroxyl (HNO) have attracted attention for their beneficial effects on cardiac function (21-23). HNO donors combine vasorelaxing properties with positive cardiac inotropy and lusitropy and are currently tested clinically for the treatment of heart failure (24-28). Various chemically distinct HNO donor compounds with different release-kinetics have been developed in recent years (26). The cardiac effects of HNO-releasing compounds are mainly attributed to HNO-induced oxidative interdisulfide formation in cardiac myofilament and sarcoplasmic reticulum proteins (29) with positive effects on cardiac myocyte function (30,31). Importantly, protein kinases and phosphatases that transduce extracellular signals into a contractile response via phosphorylation and dephosphorylation of substrate proteins, have also been described to be oxidant-susceptible. Whether HNO donor compounds impact on contraction and relaxation parameters through alteration of protein kinase and phosphatase activity and thus cardiac myocyte protein phosphorylation, has not been reported before.

Effect of HNO donors on cardiac myocyte contractility
We evaluated the impact of an experimental and a clinically relevant HNO donor, 1nitrosocyclohexyl acetate (NCA) and CXL-1020, respectively, on cardiac myocyte contractile function. Contractility of single adult rat ventricular myocytes (ARVMs) was recorded in response to vehicle (DMSO; control; Figure 1A), NCA (100 µmol/L; Figure 1B) or CXL-1020 (300 µmol/L; Figure 1C). Upon exposure to NCA, ARVMs displayed a significant but transient increase in sarcomere shortening and relaxation velocity, which reversed to baseline after approximately 4.5 min, reaching a novel plateau state ( Figure 1B,  Suppl Figure 1B). The NCA-induced significant prolongation of the time to peak was visible after 1 min of treatment and this effect was maintained during the new plateau state ( Figure 1D). Importantly, a decrease in diastolic length and decreased response to the subsequent isoprenaline (ISO) application was observed. Significantly increased sarcomere shortening as well as maximal contraction and relaxation velocities were also elicited by CXL-1020, but, in contrast to NCA, persisted after a plateau was reached, (Figure 1C, Suppl Figure 1C). Compared to baseline measurements ( Figure 1A, Suppl Figure 1A), CXL-1020 additionally displayed significantly accelerated relaxation as reflected by a decreased time to baseline50% ( Figure 1D). Exposure to CXL-1020 has been described previously to increase cellular cGMP levels via activation of soluble guanylate cyclase (sGC) with subsequent activation of cGMP-dependent protein kinase (PKG) (32). To rule out a contribution of CXL-1020-mediated PKG activity on the observed effect, experiments were repeated after 1H-(1,2,4)oxadiazolol(4,3-a)quinoxaline-1-one (ODQ) pretreatment to inhibit sGC (Suppl Figure 2). There was no difference in any contractile parameter detectable between ARVMs treated with CXL-1020 alone or after pretreatment with ODQ (Suppl Figure  2), excluding an effect of PKG on the CXL-1020-mediated positive inotropic effect.

Impact of oxidants on cardiac myocyte protein phosphorylation
To investigate whether NCA and CXL-1020 impact on the phosphorylation status of cardiac myocyte proteins at the concentration that was observed to induce increased contractility, ARVMs were exposed to NCA (100 µmol/L) and CXL-1020 (300 µmol/L) as well as to the HNO donor Angeli's salt (AS; 500 µmol/L), hydrogen peroxide (H2O2; 100 µmol/L) or the disulfide-inducing agent diamide (DIA; 500 µmol/L). Samples were resolved by sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE) and phosphoproteins stained by Pro-Q Diamond. In ARVMs exposed to the various oxidants, bands of increased intensity were observed at molecular weights of 150 and 25 kDa (Figure 2A; arrows). Comparable signals were detected in response to the PKA-activating β1-AR agonist isoprenaline (ISO), suggesting an impact of oxidant-mediated phosphorylation of PKA substrate proteins. Exposure of ARVMs to NCA led to enhanced phosphorylation of the 150 kDa PKA substrate protein cMyBP-C at Ser273, 282 and 302, accounting for 28.1 ± 23.6%, 59.5 ± 25.4% and 58.2 ± 39.9% of the response mediated by ISO, respectively ( Figure 2B). These phosphorylation sites in cMyBP-C, in particular Ser282, have been shown to regulate actin-myosin interaction and contribute to positive inotropy and lusitropy in response to β1-AR stimulation (33)(34)(35).
NCA exposure induced phosphorylation of PLN at Ser16 that measured 23.3 ± 14.7% of the signal induced by ISO. This phosphorylation site in PLN regulates activity of the sarcoplasmic reticulum Ca 2+ ATPase (SERCA) activity and contributes to positive inotropy and lusitropy in response to β1-AR stimulation (1). DIA mediated enhanced phosphorylation of cMyBP-C at Ser282 (33.6 ± 18.1% of ISO response) and Ser16 of PLN (11.0 ±11.5% of the ISO response). Surprisingly, there was no significant impact on the phosphorylation of cTnI in response to oxidants detectable. The data suggest that NCA and DIA induce protein phosphorylation potentially by modulation of the PKA signaling pathway. Phosphorylation induced by exposure to AS, CXL-1020 was lower and the extent of the phosphorylation signal varied between experiments and batches, potentially attributed to the fast release kinetics of HNO by these donors. Exposure of ARVMs to increasing concentrations of the oxidants revealed concentration-dependent phosphorylation of cMyBP-C at Ser282 and PLN at Ser16 that was most pronounced for NCA and DIA. However, there was no impact on the phosphorylation of cTnI at Ser22/23 detectable apart from a slight increase in response to the highest NCA concentration that remained statistically nonsignificant ( Figure 2B, C). ISO was used to yield the maximal phosphorylation response for all cardiac proteins tested here.

PKA activation in response to HNO donors in adult cardiac myocytes
Whether NCA and DIA exposure of ARVMs directly activates PKA was substantiated by Förster resonance energy transfer (FRET) experiments in adenovirally transduced ARVMs expressing the A-kinase activity reporter AKAR3-NES. Application of NCA and DIA induced a change in biosensor fluorescence with 34.1 ± 29.7% and 31.5 ± 30.5% of the maximal response, respectively, which was elicited by subsequent application of forskolin/3isobutyl-1-methylxanthine (FOR/IBMX) ( Figure 3A). To investigate whether the mode of NCAmediated PKA activation involves RI interdisulfide bond formation, the oxidation status of PKA-RI was assessed following oxidant treatment. The suitability of the PKA-RI antibody to unveil changes in the oxidation state of PKA-RIa was confirmed in isolated adult mouse ventricular myocytes from wildtype (WT) or knock-in mice that constitutively express a redox-deficient PKA-RIa Cys17Ser mutant exposed to H2O2 or ISO (Suppl Figure 3). By western immunoblot analyses under non-reducing conditions, enhanced formation of RI dimer was detected. Interdisulfide formation occurred to a similar extent following exposure to all tested oxidant stimuli ( Figure 3B). Notably, RI interdisulfide formation did not correlate with the previously shown phosphorylation of PKA substrate proteins (Figure 2). Addition of a reducing agent revealed loss of PKA-RI dimer in favor of monomeric PKA-RI and confirmed the oxidative nature of dimer formation. Interestingly, only after exposure to AS or CXL-1020, a proportion of the PKA-RI dimer reproducibly remained under reducing conditions. Pretreatment of ARVMs with the pharmacological PKA inhibitor H89 prior to oxidant exposure revealed attenuation of the phosphorylation of cMyBP-C at Ser282 and PLN at Ser16 in response to NCA, AS and ISO, suggesting oxidant-mediated PKA activation involved in oxidant-mediated cardiac myocyte protein phosphorylation. In contrast, CXL-1020-mediated phosphorylation remained unaffected by H89 pretreatment suggesting a distinct mode of action ( Figure 3C). The involvement of a panel of other kinases (Ca 2+ -calmodulindependent kinase, protein kinase C, p90 ribosomal S6 kinase, Rho-kinase) that have been described previously to phosphorylate cMyBP-C at Ser282 (36) was excluded by the use of pharmacological inhibitors (Suppl Figure 4). Importantly, ISO-but not NCAmediated phosphorylation of cMyBP-C was abolished by pretreatment of ARVMs with the b-AR antagonist atenolol, confirming direct and receptor-independent PKA activation by NCA ( Figure 3D).

Oxidant-mediated PKA and phosphatase translocation to the cardiac myofilament compartment
Time-response experiments in ARVMs exposed to NCA revealed a quick onset of PKA-RI dimer formation, which was followed by phosphorylation of cMyBP-C at Ser282 occurring with a timely delay. Phosphorylation was maintained for up to 100 min ( Figure 4A). To investigate whether oxidant exposure induces accumulation of PKA in the myofilament compartment, ARVMs were exposed to NCA or ISO, harvested under non-reducing conditions and the myofilament-containing Triton-insoluble fraction was isolated. Successful fractionation was confirmed by detection of the fraction-specific marker proteins GAPDH in the cytosolic, sodium-potassium ATPase (NKA) in the membrane and cTnI in the myofilament fraction (Suppl Figure 5). Exposure of ARVMs to NCA led to a significant accumulation of PKA-RI and PKA-C in the Triton-insoluble fraction ( Figure 4B). The catalytic and regulatory subunit of the PKA counteracting phosphatase PP2A-C and B56a were also found to accumulate after exposure to NCA in the Triton-insoluble fraction, while ISO treatment did not induce myofilament enrichment, neither of the kinase nor phosphatase subunits ( Figure 4B). Analysis of subcellular fractionation following timecourse treatment with NCA revealed a fast onset of myofilament translocation for PKA subunits within 10-30 min and PP2A-C and B56a within 3-10 min, culminating in cMyBP-C Ser282 phosphorylation ( Figure 4B). Subcellular compartmentalization of PKA and PP2A is regulated by their respective regulatory subunits. NCA-induced PKA-RI and B56a myofilament translocation was supported by immunofluorescence staining of ARVMs. Exposure to NCA resulted in a diffuse signal for PKA-RI with occasional appearance of transversal striations located nearby the characteristic doublet signals representing cMyBP-C ( Figure 5A; left panel). The signal obtained for B56a in response to NCA was more distinct and clearly located B56a at the M-band and to a lesser extent also at the Z-disc ( Figure 5A; right panel). A spatial rapprochement between PKA-RI within the myofilament lattice was further investigated by proximity ligation assay (PLA)-Duolink technology in ARVMs. The antibodies for cMyBP-C and a-actinin were tested prior to PLA experiments by immunofluorescence. This showed as expected the localization of cMyBP-C in characteristic doublets in the C-zone of sarcomeric A-bands at both sides of the Mline and a-actinin at the Z-discs. Respective IgG negative controls for the secondary antibodies did not show any signal ( Figure  5B). The number of PLA fluorescence signals suggested oxidant-mediated PKA-RI localization close to the C-zone of the A-band with a distinct colocalization of PKA-RI with cMyBP-C. In contrast, the presence of PKA-RI in the Z-disc was decreased as shown by a significantly reduced colocalization of PKA-RI with the Z-disc marker a-actinin ( Figure  5C). This observation was further corroborated by cAMP pull-down experiments. ARVMs were treated with vehicle, NCA, CXL-1020 or ISO and lysates were subjected to cAMP-agarose pull-down and analysis by LC-MS and western immunoblot analysis (Figure 6). As expected, cAMP-agarose captured PKA-RI with high affinity, which was present in all samples (Figure 6). When the protein content within the pull-down fractions was analyzed by unbiased LC-MS analysis, cMyBP-C was among the proteins detected in the samples derived from ARVMs that were exposed to NCA ( Figure 6A; Suppl Table 1; ProteomeXchange PXD019808). This observation was subsequently confirmed by western immunoblot analysis. Interestingly, cMyBP-C was only co-precipitated in response to HNO donor exposure when pulldown experiments were performed under non-reducing conditions ( Figure 6B). These data confirmed oxidant-mediated colocalization of PKA-RI and its substrate cMyBP-C. Furthermore, when cell fractionation was performed in the presence of a reducing agent, neither PKA nor PP2A were enriched in the Triton-insoluble fraction, suggesting indeed oxidation-dependent translocation and trapping of kinase and phosphatase in the myofilament compartment upon oxidant exposure ( Figure 7A). Importantly, oxidantmediated myofilament enrichment also applied to PP1 and calcineurin ( Figure 7B).

PKA and PP2A oxidation in response to oxidants in ARVMs
To investigate whether oxidant exposure alters the oxidation status of PKA and PP2A in intact cardiac myocytes, ARVMs were treated with vehicle (DMSO) or oxidants and subjected to biotin-or PEG-switch analyses (Figure 8). For biotin-switch analysis, ARVMs were exposed to vehicle (DMSO) or NCA, oxidative posttranslational modifications on cysteines switched to biotin-labelling and biotinylated proteins were subsequently enriched by streptavidinpull-down. Equal protein content in the samples was assured by Coomassie and Sypro Ruby staining and enhanced oxidation of cellular proteins in response to NCA reflected by increased biotinylation was detected by western immunoblotting using streptavidin-HRP ( Figure 8A). Immunoblot analysis of the eluted fractions revealed enrichment of PKA-RI, PKA-C, B56a and PP2A-C in NCA-treated samples compared to vehicle controls, suggesting that NCA induced oxidation of PKA and PP2A subunits in intact ARVMs. An alternate methodology that requires lower protein input for analysis of protein oxidation, and thus allowed inclusion of all oxidant stimuli, is PEG-switch analysis that we have previously optimized for the detection of protein oxidation in cardiac myocytes (37). ARVMs were exposed to vehicle (DMSO), NCA, AS, CXL-1020, H2O2, DIA or ISO and PEG-switch analysis performed allowing detection of oxidized proteins due to a shift in molecular mass caused by linkage of the PEG-tag to a previously oxidized cysteine ( Figure 8C). Prior to PEG-switch, input samples were analyzed by western immunoblotting under non-reducing or reducing conditions for PKA and PP2A subunits. Under non-reducing conditions, only the interdisulfide dimer of PKA-RI was detectable as previously observed. Interestingly, in samples treated with NCA and DIA, no band was detectable with the B56a antibody, suggesting that the loss in signal intensity could be attributable to oxidative modification of the protein. Under reducing conditions, PKA-RI was detectable as a monomer. Importantly, the loss in band intensity for B56a in response to NCA and DIA observed under non-reducing conditions revealed a signal, again suggesting the regaining of affinity of the antibody for its target under reducing conditions. PKA-C and PP2A-C were detectable in their monomeric state regardless of the experimental conditions. After PEG-switch, a shift to its dimer form was detectable for PKA-RI. Interestingly, a mass shift was also detectable for PKA-C and B56a in samples exposed to NCA, AS and DIA suggesting oxidation in intact ARVMs (Figure 8C; black arrows), confirming the results obtained by biotinswitch. In contrast to the results from the biotin-switch analysis, no mass shift was detectable for PP2A-C, potentially reflecting the lack of access of the large PEG-tag to the cysteines in PP2A-C. Taken together, the data obtained by two different methods demonstrate that oxidation of PKA and PP2A subunits occurs in intact cardiac myocytes.

Effects of HNO donors on PKA and PP2A catalytic activity
To investigate whether direct exposure of purified PKA-C to NCA modulates kinase activity, in vitro kinase assays were performed using recombinant His6-tagged C1-M-C2 domain of cMyBP-C. This substrate contains the phosphorylation sites Ser273, Ser282 and Ser302 that are targeted by PKA. Under control conditions, addition of PKA-C induced robust phosphorylation of the substrate protein ( Figure 9A). Pretreatment with NCA completely abolished substrate phosphorylation, supporting previous reports that oxidation inhibits PKA-C activity (7). Pretreatment of PKA-C with ATP prior to oxidation rescued kinase activity as shown by significant substrate phosphorylation. As expected, addition of H89 to the reaction mixture inhibited PKA-C activity. Interestingly, when immunoblotting was performed under non-reducing conditions with an antibody recognising PKA-C, a double band was detectable, revealing intradisulfide formation via Cys199 and 343 as previously reported (38). Equal substrate presence in all samples was shown by immunoblotting with the anti-His antibody and Coomassie staining of the membrane.
Inhibition of PKA-C by NCA does not explain the increased phosphorylation that was observed. Therefore, the effect of HNO exposition on PP2A-C activity was investigated. Purified PP2A-C was subjected to NCA or CXL-1020 and dephosphorylation of the fluorogenic substrate 6,8-difluoro-4methylumbelliferyl phosphate (DiFMUP) was assessed ( Figure 9B). The extent of phosphatase inhibition by HNO donors was compared to the effect observed in response to okadaic acid (OA), a prototypical serine/threonine phosphatase inhibitor that achieved approximately 70% PP2A-C inhibition in our experiments. Exposure to CXL-1020 significantly reduced PP2A-C activity by approximately 35% compared to the vehicle control. NCA reduced PP2A-C activity by approximately 8% compared to the vehicle control. To match the exposure time of ARVMs to HNO donors in cell culture experiments, data were reanalyzed ( Figure 9C). The bar chart reflects the phosphatase activity following 15 or 30 min of exposure to CXL-1020 or NCA, respectively. In both cases, phosphatase activity was significantly inhibited by the HNO donors.
Ultimately, the activity status of kinase and phosphatase within the myofilament compartment upon NCAmediated translocation was explored ( Figure  10). ARVMs were pretreated with NCA and fractionation was performed under NR or R conditions. In vitro kinase assays were performed by addition of the PKA substrate C1-M-C2 of cMyBP-C. Only under NR conditions, substrate phosphorylation was detectable, which is in accordance with the previous results demonstrating oxidative modification as a prerequisite for translocation. Addition of DTT to the NR IVK reaction mixture significantly abolished substrate phosphorylation, suggesting reactivation of a phosphatase and subsequent dephosphorylation of the substrate in the sample. This was corroborated by addition of 100 nmol/L OA to the NR IVK reaction mixture in order to inhibit both, PP1a and PP2A. OA addition significantly increased substrate phosphorylation ( Figure 10A). The investigations on localized kinase activity were paralleled by experiments assaying myofilament phosphatase activity. ARVMs were subjected to NCA treatment and fractionation as described above under NR or R conditions. Phosphatase activity within the myofilament fraction was investigated by addition of DiFMUP to the myofilament samples. Increased fluorescence due to dephosphorylation of the fluorogenic substrate was detectable in samples harvested under both, NR and R conditions ( Figure  10B). Addition of DTT to reduce the oxidative modification of the phosphatase and achieve reactivation revealed significantly increased phosphatase activity in the NR samples compared to the R samples.
Taken together, substrate phosphorylation conveyed by HNO donors results from a combination of localized kinase activation and phosphatase inhibition.

Discussion
In the treatment of chronic heart failure (HF), desensitization of β-ARs and poor prognosis in response to long-term treatment of patients with inotropic drugs are major obstacles that have to be addressed in order to achieve restoration of adequate cardiac contraction (39). Both, experimental and clinical data suggest therapeutic benefit of HNO donors in HF patients with reduced ejection fraction. This novel class of pharmacological agents has been shown to enhance cardiac contractility independently of β-ARstimulation (30), without development of tolerance or obvious adverse side effects, emphasizing their potential for clinical application. To date, cardiac inotropic and lusitropic effects of HNO donors have been entirely attributed to direct oxidative modification of protein components of the contractile machinery and those involved in Ca 2+ -cycling in cardiac myocytes (29). Here, we report a previously undescribed property of HNO donor compounds to indirectly impact on cardiac contractility by inducing oxidation of functionally important cysteines in PKA and PP2A thus modulating kinase and phosphatase activity and consequently prolonging basal substrate phosphorylation. Information regarding the impact of phosphorylation on sarcomeric oxidation and vice versa is scarce. We showed previously that in human heart failure, increased Sglutathiolation of sarcomeric proteins was accompanied by reduced phosphorylation, while phosphorylation prevented subsequent oxidation. This suggested a negative crosstalk between different posttranslational modifications (37). Thus far, the induction of disulfide bonds between actin-tropomyosin and myosin light chain 1-myosin heavy chain accounted for the positive inotropic effects observed in response to HNO donors (29). In heart failure, hypophosphorylation of sarcomeric proteins is considered as a major contributing factor for contractile dysfunction evoked by desensitization of the PKA signaling pathway in combination with increased expression and activity of protein phosphatases (37,40,41). This suggests that the contribution of HNO-mediated partial restoration of sarcomeric protein phosphorylation by an orchestrated modulation of receptor-independent kinase translocation and activation combined with phosphatase inhibition would assume greater significance under heart failure conditions with the potential for rescuing the positive inotropy by increasing cross-bridge cycling kinetics (33)(34)(35). The main observation of our study was that HNO released by various donor compounds induced simultaneous and orchestrated spatiotemporal oxidation of multiple signaling components with a net result of increased cardiac protein phosphorylation. The order of events supported by our data is summarized schematically in Figure 11 and suggests that HNO donors induce interdisulfide formation and translocation of PKA-RI and PKA-C to the myofilament compartment in the absence of receptoractivation, which as we show occurred fast upon exposure to HNO. Thereby, a substrateinduced local release of the catalytic subunits might trigger phosphorylation of target proteins such as cMyBP-C. The lack of cTnI phosphorylation observed in the present study was surprising and suggests that either distinct spatiotemporal signaling upon oxidant exposure or more likely the inability of the antibody to detect cTnI phosphorylation under native pro-oxidative conditions might serve as an explanation. In line with previous reports (7,8,42), our in vitro data demonstrated oxidant-mediated inactivation of the catalytic subunit by intradisulfide formation, which cannot fully explain the increased phosphorylation of substrate proteins. However, a limitation of this in vitro experiment was the lack of the fully functional heterotetrameric PKA complex, but instead the catalytic PKA subunit was exposed to a pro-oxidant environment. This result provided the rationale to investigate the impact of HNO donor compounds on activity and subcellular localization of one of the main cardiac myocyte protein phosphatases PP2A. We reported previously that PP2A dynamically redistributed between myofilament and cytosolic compartment upon stimulation (43) and therefore hypothesized that HNO donors might affect this. Indeed, we found both, catalytic and targeting subunit of PP2A, coincidentally accumulating with the kinase subunits within the myofilament compartment upon oxidant exposure. These in vitro observations prompted us to monitor the actual activity of both, kinase and phosphatases in their native myofilament environment upon oxidant exposure. While PKA activity increased moderately, pronounced inhibition of the myofilamentlocalized phosphatases was detectable, which could be reactivated by the reductant DTT. Combined with our in vitro observation that HNO donors inhibited PP2A activity, these findings strongly support the seminal observations made by the Taylor group (42) that the model-oxidant DIA impacted on kinase and phosphatase activity, suggesting this as a general phenomenon. Thereby, basal or kinase-mediated substrate phosphorylation would accumulate upon HNO compound exposure and would be maintained over a longer time period, which the present study confirmed in time course experiments. Of note, also PP1 has been shown to form an inactivating intradisulfide between Cys39 and Cys127 upon exposure to oxidants (12,16,20). Importantly, the present study detects both main cardiac protein phosphatases accumulating in the myofilament fraction upon oxidant exposure. Overlap has been described regarding the spectrum of cardiac proteins impacting on cardiac myocyte contractility such as cMyBP-C (44) and cTnI (45) that are targeted by both, PP1a and PP2A. Therefore, despite the focus of the present manuscript on the contribution of PP2A in this experimental context, it is likely that HNO-mediated intradisulfide formation and subsequent inhibition of PP1a activity might also contribute to the increase in cardiac myocyte protein phosphorylation observed in response to HNO donors. This is also supported by the results obtained with OA that was on purpose applied at a concentration that inhibits both types of phosphatases. When positive inotropy and lusitropy were described for HNO for the first time and also in studies that reported HNO-mediated oxidative modification of SERCA (46), PLN (31) and the ryanodine receptor (47), the prototypical donor compound AS was employed. Assuming the observed effects to rely entirely on released HNO, results were translated to chemically distinct HNO donor compounds without further testing. In fact, AS, NCA and CXL-1020 share the positive effects on vasorelaxation (32,48,49) and cardiac myocyte contractility (47,50,51). Nonetheless, there are considerable differences, as shown by deviating effects of NCA and AS on sarcomeric protein oxidation (29). In line with these latter observations, our data show a distinct potential of the experimental NCA and AS and the clinical CXL-1020 HNO donor compound to modulate cardiac myocyte protein phosphorylation. Interestingly, significant differences in ARVM contractility were observed in response to NCA and CXL-1020. With its short t1/2 value of 2.3 min, CXL-1020 is expected to produce higher concentrations of HNO than the slowly hydrolyzing donor NCA (t1/2: 800 min), rendering HNO release kinetics unlikely to explain the responses exclusively observed with NCA. The unexpectedly fast effect in RI dimerization, PKA translocation and ARVM contractility suggested an additional direct interaction of NCA with protein thiols independent from HNO release to underlie these effects, as it had been previously suggested (52). The finding that NCA effects were shared by DIA, which acts by inducing disulfide bonds, further supported the potential of NCA to mediate protein oxidation by a direct mechanism in addition to HNO release. Although PKA-RI dimer formation was induced by all tested oxidants, it did not correlate with the extent of substrate phosphorylation evoked by the different compounds. Thus, the role of oxidative RI dimerization as a direct readout for oxidant-mediated PKA activity remains ambiguous. Our study suggests that cardiac myocyte protein phosphorylation in response to oxidants depends on the direct oxidative modification of susceptible cysteines impacting on the activity and translocation of the catalytic subunits of both, PKA and PP2A. A similar observation was reported previously on thiol-disulfide inhibition of PKA and PP2A by DIA with impact on kinase and phosphatase activity (42). This combined inhibitory impact of oxidants may serve as a potential explanation of the NCAmediated cardiac myocyte protein phosphorylation.

Conclusion
The present study demonstrates a role for HNO donors in the regulation of kinase and phosphatase signaling with impact on cardiac myocyte protein phosphorylation and function. Homogenates from adult mouse ventricular myocytes isolated from wildtype or Cys17Ser PKA knock-in mice exposed to hydrogen peroxide (100 µmol/L 10 min) or isoprenaline (10 nmol/L 10 min) were kindly provided by Prof. Philip Eaton from Queen Mary University London, UK.

Isolation of ARVMs
The isolation of primary ARVMs from male Wistar rats was performed as described previously (37) and executed in compliance with the German law for the protection of animals and the Guide for the Care and Use of Laboratory Animals issued by the National Research Council (US) Committee (2011). ARVMs were cultured for 24 h in laminincoated 6-well plates containing Medium 199 supplemented with 5 mmol/L taurine, 2 mmol/L carnitine, 2 mmol/L creatine and 100 U/mL penicillin-streptomycin.
Oxidant treatment of ARVMs and sample analysis ARVMs were exposed to vehicle DMSO Lysates were subjected to SDS-PAGE and phosphoproteins visualized with Pro-Q Diamond using a Typhoon 9400 imager (GE Healthcare, Chicago, USA). Total protein content was visualized by re-staining with SYPRO Ruby and Coomassie brilliant blue. Alternatively, samples were analyzed by western immunoblotting. For time course treatment, ARVMs were exposed to vehicle DMSO (0.1% (v/v)), NCA (100 µmol/L; 3, 10, 30, 100 min) or ISO (10 nmol/L; 10 min) and analyzed by immunoblot analysis.

Förster Resonance Energy Transfer (FRET) measurements in ARVMs
Isolated ARVMs were seeded onto glass slides and transduced at MOI 300 with an adenovirus encoding the AKAR3-NES biosensor that was provided for the present study and had been extensively characterized by Vandecasteele and colleagues (53). The ratio of light emitted by CFP and YFP of this biosensor changes upon phosphorylation by PKA. After 48 h of cultivation, glass slides were transferred into a microscopy cell chamber and baseline fluorescence recorded in FRET buffer [in mmol/L: NaCl 144, HEPES 10, KCl 5.4, MgCl2 1; pH 7.3], before NCA (100 µmol/L) or DIA (500 µmol/L) were applied. When a plateau was reached, maximal sensor activation was achieved by combined addition of FOR (10 µmol/L) / IBMX (100 µmol/L). Measurements were performed using a DMI3000b inverted microscope with a 40x objective (Leica, Wetzlar, Germany). Signals from CFP and YFP were simultaneously recorded using a DV2 DualView emission splitting system (Photometrics, Tucson, USA) and an optiMOS™ Camera (QImaging, Surrey, Canada). CFP excitation was induced by a Light source pE-100 (440 nm; CoolLED, Andover, UK). Data processing was performed with ImageJ µManager with a customized plugin and Microsoft Excel. FRET changes in response to the applied stimuli were normalized to baseline values and expressed as % of the effect induced by FOR/IBMX.

Subcellular fractionation
Following the treatment as described above, ARVMs were scraped into 180 µL/well NR fractionation buffer [in mmol/L: Tris-HCl 50 pH 7.5, NaF 100, EGTA 5, EDTA 2, 0.05% (w/v) digitonin, protease inhibitor] or R fractionation buffer supplemented with dithiothreitol (DTT) (5 mmol/L) and snap frozen in liquid N2. After thawing, ARVMs were harvested and subjected to subcellular fractionation as previously described (43). In brief, lysates were centrifuged at 10 000 xg for 2 min at 4 °C and the supernatant representing the cytosolic fraction was collected. The pellet was resuspended in fractionation buffer containing 1% Triton X-100 and after centrifugation, the supernatant comprising membrane-associated proteins was collected. The remaining myofilament containing detergent-insoluble pellet was resolved in sample buffer. Cytosolic and membrane fractions were supplemented with sample buffer before analysis by western immunoblotting.

cAMP-agarose pull-down
After treatment of ARVMs, cells were harvested in 200 µL lysis buffer (with or without DTT)/well of a 6-well plate [in mmol/L: HEPES 30 pH 7.4, EDTA 2, EGTA 2, NaCl 150, 1% (v/v) Triton X-100, NaF 2, protease inhibitors]. Lysates were centrifuged at 10 000 xg for 10 min at 4 °C and the supernatant collected, supplemented with 30 µL 8-AHA-cAMP-agarose and incubated under rotation at 4 °C for 4 h. Beads were washed twice with lysis buffer supplemented with detergent and protease inhibitors and twice without.

Tryptic in-gel digestion
In-gel digestion was done following Shevchenko et al. (54). Shrinking and swelling was performed with 100% ACN and 100 mmol/L NH4HCO3. In-gel reduction was achieved with 10 mmol/L dithiothreitol (dissolved in 100 mmol/L NH4HCO3). Alkylation was performed with 55 mmol/L iodacetamide (dissolved in 100 mmol/L NH4HCO3). Proteins in the gel pieces were digested by covering them with a trypsin solution (8 ng/µL sequencing-grade trypsin, dissolved in 50 mmol/L NH4HCO3 containing 10% ACN) and incubating the mixture at 37°C for overnight. Tryptic peptides were yielded by extraction with 2% FA, 80% ACN. The extract was evaporated.

LC-MS/MS data analysis
With Proteome Discoverer 2.0 (Thermo Scientific, Bremen, Germany) the LC-MS/MS data were processed. Identification of the proteins from the MS/MS spectra were performed with the search engine Sequest HT using the Rattus norvegicus SwissProt database (release June 2016, 7961 entries; www.uniprot.org) and a contaminant database. For the searches, the following parameters were applied: Precursor mass tolerance: 10 ppm; Fragment mass tolerance: 0.2 Da. Fully tryptic digestion. Two missed cleavages were allowed. Carbamidomethylation on cysteine residues as a fixed modification and oxidation of methionine residues as a variable modification was used for the search. Peptides with an FDR of 1% using Percolator were identified. At least two unique peptides per protein were used as a condition for a reliable identification.

In vitro kinase assay
For the in vitro kinase (IVK) assays, 1250 IU/sample active PKA catalytic subunit were diluted in 5 µL assay buffer (30 mmol/L Tris, pH 7.4, 15 mmol/L MgCl2) and pre-incubated for 10 min at 37°C with either 100 µmol/L ATP, 25 µmol/L H89 or the equivalent volume of assay buffer (Ø). This was followed by a 30-min treatment with vehicle DMSO (control) or NCA (100 µmol/L) at 37°C. Subsequently, ATP (100 µmol/L) was added to the H89-or Ø-pretreated samples and the appropriate volume of assay buffer was added so that the final reaction volume would be 50 µL/sample. The kinase reaction was initiated by adding 100 pmol/sample recombinant His6-tagged C1-M-C2 domain (amino acid residues 153-450) of human cMyBP-C as substrate (prepared as described in (37)), proceeded for 30 min at 30°C and was terminated by addition of 25 µL/sample 3x non-reducing sample buffer. For each condition, a blank sample devoid of kinase was also prepared.

In vitro PP2A-C activity assay
The PP2A-C activity was analyzed in vitro by a fluorescence assay in which fluorogenic DiFMUP was used as phosphatase substrate. The substrate was initially prepared as a 10 mmol/L stock solution in 50 mmol/L Tris-HCl buffer (pH 7.0) and stored at -20 °C. Before each experiment, this stock was further diluted to 1 mmol/L with phosphatase assay buffer (40 mmo/L Tris-HCl, pH 7.5, 34 mmol/L MgCl2, 4 mmol/L EDTA, 0.005% (w/v) BSA). The initial stock solutions of NCA (100 mmol/L), CXL-1020 (300 mmol/L) and the serine/threonine protein phosphatase inhibitor OA (10 µmol/L) were prepared in DMSO. These were then diluted in phosphatase assay buffer to acquire solutions of 250 µmol/L NCA, 750 µmol/L CXL-1020 and 25 nmol/L OA. For each reaction, 40 µL compound solution were added to 50 µL phosphatase assay buffer containing DiFMUP in a black-walled 96well plate. The reaction started by adding 10 µL/well of 12.1 µU/µL recombinant human PP2A-C solution prepared in phosphatase assay buffer (final concentration 121 µU/well). This resulted in a final volume of 100 µL per sample and final concentrations of 500 µmol/L DiFMUP, 100 µmol/L NCA, 300 µmol/L CXL-1020 and 10 nmol/L OA. As a blank for each sample, a phosphatase-free control was prepared and measured in parallel. The dephosphorylation of the DiFMUP substrate to fluorescent DiFMU was measured in a Tecan Safire II microplate reader (Männerdorf, Switzerland) at excitation/emission wavelengths of 358/455 λ for 40 min (1 read/min) at 30 °C. Average activities expressed as ΔRFU/min were calculated from the slope of a linear curve fit through the origin of the coordinate system from averaged normalized values of 4-5 independent assays. For the reanalysis of the data to reflect cell culture conditions, phosphatase activity in response to CXL-1020 exposure was calculated involving datapoints from 15 to 40 min and for NCA from 30 to 40 min.

Myofilament kinase and phosphatase assay
ARVMs were cultured in 6-well plates and 4 wells were combined per intervention. After pharmacological treatment, ARVMs were lysed in 200 µL fractionation buffer per well [in mmol/L: Tris-HCl 50 pH 7.4, EGTA 5, EDTA 2, NaF 100, supplemented with 1% (v/v) Triton X-100 and Complete protease inhibitors]. Harvesting under reducing conditions occurred with fractionation buffer containing 5 mmol/L DTT. The crude lysates were then aliquoted in two sets of tubes (labeled A and B; 350 µL lysate per tube) and centrifuged for 5 min, at 3000 xg and 4 °C. After removal of the supernatant, the pellets that contained the Triton X-100 insoluble myofilament fraction were washed with assay buffer [in mmol/L: Tris-HCl 40 pH 7.5, MgCl2 34, EDTA 4, supplemented with 0.005% (w/v) BSA] and centrifuged again as described above. Each myofilament pellet was then resuspended in 120 µL assay buffer without DTT (set A) or with 5 mmol/L DTT (set B). For the kinase assays, each reaction was prepared in a final volume of 60 µL and contained 50 µL myofilament suspension, 100 µmol/L ATP, 100 pmol substrate (recombinantly expressed His6-tagged C1-M-C2 domain) and either vehicle (DMSO) or 100 nmol/L of the pan-phosphatase inhibitor OA. The reactions were carried out at 30°C for 30 min and terminated by addition of 30 µL 3x Laemmli sample buffer. For the phosphatase assays, 25 µL myofilament suspension were mixed with DiFMUP (500 µmol/L final concentration) in assay buffer without (set A) or with 5 mmol/L DTT (set B) at a final volume of 100 µL per reaction. Measurements were performed as described in the "in vitro PP2A-C activity assay" section. For each condition measured, a linear regression was used to calculate the slope from the available number of measurements.

SDS-PAGE and western immunoblot analysis
Protein samples in reducing or non-reducing sample buffer were resolved by SDS-PAGE and analyzed by western immunoblotting as described previously (55). In brief, gels were prepared using acrylamide/bis acrylamide (Bio-Rad; Feldkirchen, Germany; #1610156). Running gels contained a buffer composed of (in mmol/L) 375 Tris base pH 8.0, 3.5 SDS, 4.38 ammonium persulfate (APS), 0.1% (v/v) N,N,N',N'tetramethylethylenediamine (TEMED). Thereby, 7.5% (v/v) polyacrylamide gels were used for the analysis of cMyBP-C phosphorylation, α-actinin and NKA1; 10% (v/v) for the analysis of PKA-RI, PKA-C, PP2A-C, B56α, PP1, calcineurin, GAPDH and His6-tag; 15% (v/v) for pSer16 PLN and pSer23/24 cTnI. Stacking gels were composed of 3.5% (v/v) polyacrylamide in a buffer comprising (in mmol/L): 125 Tris base pH 6.8, 3.5 SDS, 4.38 APS and 0.1% (v/v) TEMED. As secondary antibodies, HRPlinked sheep anti-mouse IgG or the HRPlinked donkey anti-rabbit IgG at a dilution of 1:2000 was used. Band density quantification was performed using the GelQuant.NET software provided by biochemlabsolutions.com. Validation of the respective phosphospecific antibodies recognizing cMyBP-C, cTnI and PLN phosphorylation has been described previously (35,(55)(56)(57)(58)(59)(60)(61). For analysis of protein phosphorylation in ARVMs, all signals were normalized to the respective loading control (α-actinin) from the same blot and data were presented as % fold-change over the isoprenaline positive control. For the analysis of the interdisulfideinduced PKA-RI dimer formation, after normalization to the loading control, the ratio of dimer to monomer signal density of the same lane was calculated and expressed as fold-change over the control value for each experiment. For analysis of protein translocation, signals in the insoluble fraction were normalized to the respective input signals and data were expressed as fold change over the control value for each experiment. In the IVK assays, the phosphorylation signals were normalized to the total substrate loading and expressed as % of the vehicle control after ATP preincubation.

Proximity ligation assay (PLA)
The proximity between PKA-RI and cMyBP-C or α-actinin was assessed by PLA using the Duolink ® In Situ Orange Starter Kit Mouse/Rabbit (Merck Millipore, Billerica, USA). ARVMs were exposed to DMSO (0.1% (v/v), 30 min) or NCA (100 µmol/L, 30 min) and the plasma membrane removed with 1% Triton X-100 in PBS before fixing in 4% PFA in PBS. Unspecific sites were blocked with 5% NGS in BSA/Gold buffer and PLA was performed according to the manufacturer's instructions. Fluorescence signals indicating a distance of maximally 40 nm between PKA-RI and cMyBP-C or αactinin were detected by Z-stack imaging at 2 µm intervals with a confocal laser scanning microscope LSM 800 with a Plan-NEOFLUAR 40x oil immersion objective and a Cy3 filter (Carl Zeiss, Oberkochen, Germany). PLA counts were obtained from the maximum intensity projection of each Zstack experiment and thresholding performed by the 'Intermodes' function of ImageJ. To allow better image visualisation, digital image processing was performed using GIMP version 2.10.14. Specifically, brightness was adjusted in all confocal images in an identical manner. For the images obtained after measuring "PLA counts", the color mode was changed to grayscale, brightness was then adjusted, and color was finally inverted. All "PLA count" images were processed in the same way to allow comparison between images. The brightfield images were adjusted regarding brightness and contrast in an independent fashion, which does not affect the results but allows better visible cells. Adjustments were applied on the entire field of each image.

Biotin-switch analysis
The biotin-switch protocol was adapted from the PEG-switch protocol that was previously described (63). In brief, cultured ARVMs were treated with vehicle (DMSO; 30 min) or NCA (100 µmol/L, 30 min). After a washing step with 1 mL ice-cold Tris-HCl buffer (100 mmol/L, pH 7.4), 70 µL ice-cold biotinswitch harvesting buffer (in mmol/L: NaCl 150, N-ethylmaleimide 100, Tris-HCl 100 pH 7.4, supplemented with 1% (v/v) Triton X-100 and protease inhibitors) was pipetted on the cells and the cell culture plate frozen in liquid N2. The ARVMs were thawed, harvested and incubated at room-temperature for 10 min. SDS (1%; applied as 20% solution) was added to the lysates and samples were incubated at 50 °C and 300 rpm for 25 min to block free thiol groups by alkylation with N-ethylmaleimide in the harvesting buffer. Subsequently, a sample volume of 130 µL was desalted by a Zeba™ Spin desalting column (Thermo-Fischer Scientific, Waltham, USA) to remove Nethylmaleimide. The residual volume was kept as input. Oxidized protein thiol groups of desalted samples were reduced by the addition of 50 mmol/L DTT. Following 20 min of incubation at room-temperature, DTT was removed by another desalting step. The biotinylation reagent EZ-Link ® Maleimide-PEG2-Biotin (1 mmol/L; #21901BID; Thermo-Fischer Scientific, Waltham, USA) and 0.5% SDS (applied as 20% solution) were added sequentially and samples incubated on a laboratory agitator at room-temperature for 2 h in the dark. To remove unbound Maleimide-PEG2-Biotin, four volumes of acetone (-20 °C) were added to the samples and proteins precipitated at -20 °C for 1 h. Proteins were pelleted by centrifugation at 16 000 g and 4 °C for 10 min. The supernatant was removed, and the pellets washed twice with 731 µL 80% acetone (-20 °C) by detaching the pellet from the tube, centrifugation and discarding the supernatant. Residual acetone was left to evaporate for 2 min, then 400 µL Tris-HCl buffer (100 mmol/L, pH 7.4) were added to the protein pellets. Resuspension was supported by vortexing and ultrasonic homogenization. After a quick centrifugation step, 40 µL of the supernatant were kept as input (biotinylated). The residual volume was added to 100 µL streptavidin-agarose beads (50% slurry) and incubated overnight on a laboratory agitator at 4 °C to bind biotinylated proteins. The beads were washed three times with 1 mL RIPA buffer (in mmol/L: NaCL 150, Tris-HCl 25 pH 7.4, SDS 0.1% (w/v), sodium deoxycholate 0.1% (w/v), Triton X-100 1% (v/v)) for 5 min at room-temperature on the agitator, followed by two washes with Tris-HCl buffer (100 mmol/L, pH 7.4). After each washing step, the beads were sedimented by centrifugation (1 min, 3000 g). Eventually, 50 µL elution buffer was added to the beads and the samples incubated at roomtemperature and at 95 °C for 15 min each. The beads were pelleted by centrifugation at 16 100 g for 10 min. Subsequently, the supernatant was decanted and supplemented with 6x reducing Laemmli sample buffer. The precipitation of proteins, which indicated the biotinylation of cysteine residues as a consequence of protein oxidation in response to the initial treatment, was analyzed by SDS-PAGE and Western immunoblotting.

PEG-switch analysis
The PEG-switch protocol was performed as described before (37,63). This method has the advantage over the previously described Biotin-switch method that less starting material is needed. In brief, cultured ARVMs were treated with vehicle DMSO (30 min), NCA (100 µmol/L, 30 min), AS (500 µmol/L, 15 min), CXL-1020 (300 µmol/L, 15 min), H2O2 (100 µmol/L, 10 min), DIA (500 µmol/L, 10 min) or ISO (10 nmol/L, 10 min). After a washing step with 2 mL ice-cold Tris-HCl buffer (100 mmol/L, pH 7.4), 100 µL ice-cold PEGswitch harvesting buffer (in mmol/L: maleimide 100, Tris-HCl 100 pH 7.4, SDS 34.7, supplemented with protease inhibitors) was pipetted on the cells and the cell culture plate frozen in liquid N2. The ARVMs were thawed, collected and 130 µL of each lysate was incubated at 50 °C and 300 rpm for 25 min to block free thiol groups by alkylation with maleimide contained in the harvesting buffer. The residual sample volume was kept as input and one half each supplemented with 3x non-reducing or reducing Laemmli sample buffer. Maleimide from the harvesting buffer was removed by desalting with Zeba™ Spin desalting columns (Thermo-Fischer Scientific, Waltham, USA). Desalted samples were supplemented with 50 mmol/L DTT, causing the reduction of oxidized protein thiols. Followed by 20 min incubation at RT, another desalting step was performed to remove DTT. After sequential addition of the thiol labeling reagent PEG-maleimide (2 mmol/L, 5 kDa; #63187; Sigma Aldrich, St. Louis, USA) and 0.5% SDS (applied as 20% solution), samples were incubated on a laboratory agitator at room-temperature for 2 h in the dark. Finally, each sample was supplemented with 3x non-reducing Laemmli sample buffer and subjected to Western immunoblot analysis. The detection of a shift in molecular weight of a protein indicated PEG-labeling and is considered as a surrogate for the oxidation of cysteine residues in response to the initial treatment.

Single cell contractility measurements
After isolation, ARVMs were maintained in IonOptix buffer [in mmol/L: NaCl 135, glucose 20, HEPES 10 pH 7.46, KCl 4.7, CaCl2 1.5, MgSO4 1.2, KH2PO4 0.6, Na2HPO4 0.6] for at least 1 h. During pacing at 1 Hz (15 V, pulse duration: 4 ms), diastolic sarcomere length (in µm), sarcomere shortening (as % of diastolic sarcomere length), maximal contraction and relaxation velocities reached (maximal change in sarcomere length over time; dL/dt max), as well as the time needed to reach maximal contraction (time to peak, in s) and the state of 50% maximal relaxation (time to baseline50%; in s) were assessed under basal conditions or after pretreatment with ODQ (0.3 µmol/L 10 min) during exposure to DMSO (0.001% (v/v)), NCA (100 µmol/L) or CXL-1020 (300 µmol/L). ISO (10 nmol/L) was added at the end of the treatment protocol to ensure cell viability. Contractile parameters were recorded using MyoPacer EP, MyoCam-S and a Fluorescence System Interface unit (IonOptix, Westwood, USA). The IonWizard software was used for data recording and analysis.

Statistical analysis
All data were analyzed by GraphPad PrismSoftware Inc. San Diego, CA, USA) apart from the data concerning FRET measurements, which were analyzed by Origin (OriginLab Corporation, Northampton, MA, USA). Data from the contractility measurements, immunoblot analyses and activity assays were statistically compared by one-way or two-way ANOVA followed by Dunnett's or Bonferroni posthoc test, as indicated in the respective Figure legends. Two-tailed Student's t-test was used to compare data obtained from PLA assays, from the cAMP-agarose pull-down experiments and the PP2A in vitro assay. For each experiment, N numbers reflect biological replicates and are mentioned in the respective Figure legend. Quantitative data are given as mean±S.D. and P < 0.05 was considered as significant.

Data availability
All data are located within this manuscript. The mass spectrometry proteomics dataset has been deposited to the ProteomeXchange Consortium via the PRIDE (64) partner repository with the dataset identifier PXD019808 and 10.6019/PXD019808. The protein identifications are presented in Suppl Table  1.

14.
Jideama      : Oxidation-mediated effects on PKA and PP2A-C activity in vitro. A Active PKA catalytic subunit (PKA-C) was preincubated for 10 min with either ATP (100 µmol/L), H89 (25 µmol/L) or assay buffer (Ø) and then exposed to vehicle (control) or 100 µmol/L NCA for 30 min. Then, the samples that initially did not contain ATP were supplemented with 100 µmol/L ATP and the in vitro kinase reaction was initiated by adding 100 pmol recombinant His6-tagged C1-M-C2 cMyBP-C to each sample. Samples that did not contain kinase served as additional controls. Substrate phosphorylation and content was assessed under reducing (R) conditions (sample reduction with 10% (v/v) β-mercaptoethanol) by immunoblotting (IB) using antibodies against pSer282-cMyBP-C and the His6-tag. The presence and oxidation of PKA-C was monitored under non-reducing (NR) conditions by IB using an anti-PKA-C antibody. Coomassie stain of the substrate blot is also shown as additional loading reference.