University of Groningen Understanding compartmentalized cAMP signaling for potential therapeutic approaches in cardiac disease Musheshe, Nshunge

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Understanding compartmentalized cAMP signaling for potential therapeutic approaches in

cardiac disease

Musheshe, Nshunge

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Publication date: 2018

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Musheshe, N. (2018). Understanding compartmentalized cAMP signaling for potential therapeutic

approaches in cardiac disease: Insights into the molecular mechanisms of the cAMP-mediated regulation of the cardiac phospholemman-Na+/K+ ATPase complex. University of Groningen.

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520589-L-bw-Nshunge 520589-L-bw-Nshunge 520589-L-bw-Nshunge 520589-L-bw-Nshunge Processed on: 26-6-2018 Processed on: 26-6-2018 Processed on: 26-6-2018

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Chapter 3 Submicroscopic compartmentalization

of cAMP/PKA signaling regulates ion

flux at the plasmalemma of cardiac

myocytes

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Processed on: 26-6-2018 PDF page: 63PDF page: 63PDF page: 63PDF page: 63 57

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Submicroscopic compartmentalization of

cAMP/PKA signaling regulates ion flux at the

plasmalemma of cardiac myocytes.

Musheshe N1,2_{., Surdo N.C}2_{., Schleicher K}2_{., Koschinski A}2_{., Schmidt M}1,3_., Zaccolo M2

1_{Department of Molecular Pharmacology, University of Groningen, The} Netherlands, 2_{Department of Physiology, Anatomy and Genetics, University of} Oxford, Oxford, UK, 3_{Groningen Research Institute for Asthma and COPD,} GRIAC, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands

Keywords: Adrenergic receptor, Na+_/K+ _{ATPase, Na}+_/Ca2+_exchanger, Phospholemman, AKAP79, cAMP, Fluorescence Resonance Energy Transfer (FRET), Phosphodiesterases, Protein Kinase A (PKA), phosphatases.

ABSTRACT:

The second messenger cAMP regulates the frequency and strength of heart contractions and dysregulation of this signaling pathway leads to cardiovascular disease. Within different subcellular compartments, cAMP signal propagation is spatially controlled by phosphodiesterases (PDEs) which, by degrading cAMP contribute to signal compartmentalization. This facilitates activation of distinct PKA subsets and results in unique responses to individual extracellular stimuli. In the heart cAMP-dependent PKA mediates the catecholaminergic control over

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the force and frequency of cardiac contraction via phosphorylation of proteins that are involved in excitation-contraction coupling (ECC). As part of the ECC machinery, catecholamine dependent activation of PKA leads to phosphorylation of the L-type Ca2+_{channels (LTCC) resulting in increased systolic Ca}2+_{. PKA} also phosphorylates phospholemman (PLM), a regulator of the cardiac Na+_/K+ ATPase (NKA), which leads to decreased systolic Ca2+_{content. Thus, PKA} activation appears to mediate opposing effects on intracellular Ca2+_{levels and} how cAMP/PKA signaling is coordinated at these two sites remains unclear. Using novel fluorescence resonance energy transfer (FRET)-based sensors that we targeted to unique multiprotein complexes at the plasmalemma, we show local heterogeneity of cAMP and PKA-dependent phosphorylation on β-adrenergic receptor stimulation that is dependent on local PDE and phosphatase activity. Our findings reveal a novel facet of the complex regulation of local cAMP signaling in cardiac myocytes and support tight coordination of PKA activity at the plasmalemma to optimally modulate Ca2+ _flux.

INTRODUCTION:

Cardiac cAMP-PKA signaling is compartmentalized. Compartmentalization is achieved via organization of the molecular components of this pathway into signaling complexes, which are distributed in various and discrete subcellular compartments within cardiomyocytes (Buxton I.L.O and Brunton L.L.,1983; Dodge-Kafka K.L et al., 2006). Multiple signaling complexes coexist within each single cell and operate under a strict spatial and temporal control that is dictated by localized changes in cAMP concentration (Surdo N.C et al., 2017; Barbagallo F et al., 2016; Zaccolo M and Pozzan T., 2002).

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PKA is the main effector of cAMP and it mediates a variety of cellular functions via direct phosphorylation of its targets and via cross talk with other signaling pathways (Dessauer, C.W et al., 1998). On β-AR stimulation, cAMP levels are increased. cAMP in turn binds to the regulatory subunits of PKA, leading to activation of the catalytic subunits and subsequent phosphorylation of serine and threonine residues on specific substrate proteins (Agnes R.S et al., 2010; Wang C.L et al.,2012). PKA is largely constrained to specific subcellular locations via binding to A Kinase anchoring proteins (AKAPs), a family of multiscaffolding proteins that tether PKA in proximity of its targets (Dodge-Kafka, K.L et al.,2006). In addition to PKA, multiprotein complexes organized by AKAPs may include adenyl cyclases, other kinases, phosphodiesterases (PDEs) and phosphatases (Dodge-kafka K.L et al., 2006). PDEs, the enzymes that hydrolyze cAMP into AMP, contribute to cAMP compartmentalization by confining cAMP changes to specific subcellular locations and by defining the amplitude of local cAMP signals. Phosphatases, by dephosphorylating PKA targets, may also contribute to defining the amplitude, duration and location of the signaling response (Redden J.M and Dodge-kafka K.L., 2011). Like for the other molecular components of the cAMP/PKA signaling pathway, the activity of phosphatases is also primarily controlled by their subcellular localization (Virshup and Shenolikar S., 2009, Ceulemans H et al., 2004).

A well-studied AKAP expressed in cardiac myocytes is AKAP79/150 (Carr D.W et al.,1992 and Coghlan V.M et al., 1995). AKAP79/150 localizes to the plasmalemma and forms a complex with the L-type Ca2+_{channel (LTCC), the} β-adrenergic receptor, adenyl cyclase 5/6 and the phosphatases PP1 (Le A.V et al., 2011) and PP2B (Dell’Acqua M.L et al., 2002; Efendiev R et al., 2010). On β-adrenergic receptor stimulation cAMP activates AKAP79/150-anchored PKA to phosphorylate the alpha1C subunit of the LTCC, leading to increased Ca2+_current (Gao T et al., 1997). Another plasmalemma protein that is phosphorylated by PKA on catecholamine activation is PLM – a regulator of the cardiac NKA

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(Crambert et al., 2002; Despa S et al., 2005; Pavlovic D et al., 2007, 2013b). NKA maintains cellular homeostasis by pumping Na+_{out of the cell in exchange for} K+_{. When phosphorylated at Ser68 by PKA, PLM enhances the activity of NKA} (Fuller W et al.,2004; Silverman B.Z et al., 2005) by increasing its affinity for Na+_{(Mishra et al.,2015). In turn, the enhanced activity of NKA sets the Na}+ gradient that promotes the Na+_/Ca2+_{exchanger (NCX) to pump Ca}2+_{out of the} cell in exchange for Na+_{. Thus, via regulation of PLM phosphorylation,} β-adrenergic receptor stimulation results in enhanced extrusion of Ca2+_{(Pavlovic D} et al.,2013b; Boguslavskyi et al., 2014). The mechanisms that allow coordinated regulation of these apparently opposing effects of PKA activation on systolic Ca2+ remain largely to be determined. Here, we investigate cAMP/PKA signaling at the AKAP79 and PLM/NKA complexes using selectively targeted FRET-based sensors for cAMP and PKA-dependent phosphorylation. Our findings show profound differences in local handling of cAMP levels and phosphatase activity at these two plasmalemma sites and reveal that adrenergic regulation of Ca2+_flux across the plasmalemma relies on submicroscopic compartmentalization of cAMP/PKA signals.

METHODS:

Isolation and culture of cardiomyocytes

Adult rat ventricular myocytes (ARVMs) were isolated from 350-375g male Sprague Dawley rats as detailed by Lomas O et al.,2015. The hearts were perfused on Langendorff apparatus at 37o_{C for about 5 minutes through the} coronary arteries with an oxygenated Ca2+_{-free Tyrode buffer solution (130mM} NaCl, 5mM Hepes, 0.4mM NaH2PO4, 5.6mM KCl, 3.5mM MgCl2, 20mM

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Taurine and 10mM Glucose). Afterwards, the hearts were digested with 0.06mg/ml Liberase TH (Roche Diagnostics Limited, UK) in a 100μM Ca2+ Tyrode buffer solution for another 20 minutes until the heart became flaccid. After perfusion the left ventricle was discarded, and the rest of the heart was minced and resuspended in equal volume of 1% BSA Ca2+_{- free Tyrode buffer} solution and centrifuged at 500rpm for 3 mins or left to settle. The pellet of cells was resuspended in extracellular Ca2+_{incrementally until a final concentration of} 1.0mM. Cells were cultured in minimum essential medium (MEM) (Sigma-Aldrich UK), supplemented with 2.5% foetal bovine serum (FBS), 1% penicillin-streptomycin, 1% L-Glutamine and 9mM NaHCO3, and then plated on laminin (40μg/ml) coated glass coverslips. Before replacing the medium with FBS-free MEM, ARVMs were left to adhere for 1.5-2hrs. Myocytes were infected at multiplicity of infection (MOI) of 10-100 with adenovirus encoding for FRET reporters for at least 3hrs. The medium was then replaced with fresh FBS-free MEM containing 0.5μM cytochalasin D (MP Biomedicals, UK). The infected cells were kept in culture for up to 36hrs for PLM-CUTie and MP-CUTie, and PLM-AKAR4 and for up to 48hrs for AKAP79-CUTie and AKAP79-AKAR4. Neonatal rat ventricular myocytes (NRVMs) from 1-3 days old Sprague Dawley rats were cultured and isolated as described by Zaccolo M and Pozzan T.,2002.

Pull down experiments and western blotting

ARVMs from one heart were plated onto 2x10cm dishes coated with laminin (20mg/ml) and infected with the respective adenovirus for 3hrs. After up to 36hrs, the cells were washed with 1xTyrode buffer solution and lysed for 5 mins on ice in Ripa buffer (Sigma Aldrich, UK) for the CUTie and AKAP79-AKAR4 and lysed in 0.1% of 20% Nonidet P40 (NP40) buffer (Company) supplemented with 50mM Tris PH 7.5 and 50mM NaCl for PLM-CUTie and

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PLM-AKAR4 samples respectively. All the respective lysis buffers were supplemented with complete EDTA-free protease inhibitor cocktail tablets (Roche Diagnostics Limited, UK) and phosphatase inhibitor tablets. The lysed cells were then collected and rotated for 20mins at 4o_{C. Samples were then} centrifuged for 10 mins at 10,000 rpm at 4o_{C to remove insoluble material. Total} protein was quantified using Micro BCA Protein Assay Kit (Pierce Biotechnology Inc.,IL,USA). 500μg of total protein was rotated for 2hrs at 4o_C with 25μl of agarose beads coated with a monoclonal anti-GFP antibody (GFP-Trp A,gta-10, ChromoTek GmbH, DE). Samples were then centrifuged at 2000 rpm for 1min and the supernatant was discarded. Beads were washed at least 4 times with ice cold Ripa or NP40 lysis buffer respectively. Bound proteins were eluted in 25μl of 2xSDS loading buffer (Life technologies) and denatured at 95o_C for 5mins. Pulled down proteins were run on Bolt 4-12% Bis-Tris Plus Gels in order to separate them by molecular weight. The proteins were then transferred onto hydrophobic PVDF membrane (Amersham hybond -P, 0.45μm, GE Healthcare Life Sciences,UK). After the transfer, the membranes were blocked for 1hr at room temperature in 5% skim milk (Sigma Aldrich, UK). They were then incubated overnight at 4o_{C with the respective antibodies: Adenyl cyclase} V/VI (C17) (Santa Cruz Biotechnology, TX, USA at 1:200), rabbit anti FXYD1(PLM) (Abcam ab76597at 1:1000), rabbit anti-Na+_/K+_{ATPase (Abcam,} ab185065 at 1: 10,000). After at least five washes with TBS-0.5% Tween20 (Alfa Aesar, MA, USA), membranes were incubated at room temperature for 1hr with appropriate horseradish peroxidase conjugated secondary antibodies (at 1:3000) and detected with ECL western blotting detection kit (Thermo Fisher Scientific, MA, USA). The blots were stripped with stripping buffer (Thermo Fisher Scientific, MA, USA) and reprobed with anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) antibody (sc-1666574, Santa Cruz Biotechnology, TX, USA, used at 1:3000) to detect protein contamination. In order to control for

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efficiency of the pull down, an anti-GFP antibody (sc-9996, Santa Cruz Biotechnology, TX, USA, at 1:1000) was used.

For NRVMs pull down experiments, 6x106_{cells were plated onto 2x10cm dishes} coated with laminin as described above. After about 36hrs, Infected cells were washed with 1xADS buffer (106mM NaCl, 20mM Hepes, 0.8mM NaH2PO4, 5.3mM KCl, 0.4mM MgSO4, 5mM glucose), and the same protocol as described for ARVMs above.

Western Blot Analysis

Freshly isolated ARVMs were resuspended in 1.4mM Ca2+_{1 x Tyrode solution} and left to stand for 2hrs at room temperature (RT) before treatment. 8 samples were then treated for 10 mins at RT with DMSO, H89 30μM (for PKA inhibition) alone, Bisindolylmaleimide I 1μM (for PKC inhibition) alone, Iso 0.1nM alone, Iso 1nM alone, Iso 1μM alone, IBMX 100μM alone, and saturating stimulus (SAT) of forskolin 25μM + IBMX 100μM as indicated. The pellets of an estimated 4-6mg by weight were directly lysed in 1 X SDS loading buffer (Life Technologies, UK). Sample proteins were denatured at 900_{C for 5 mins and} electrophoresed on 4-12% Bis Tris Plus gels. The proteins were then transferred onto hydrophobic PVDF membrane (Amersham hybond -P, 0.45μm, GE Healthcare Life Sciences,UK). After the transfer, the membranes were blocked for 1hr at room temperature in 5% (w/v) skim milk (Sigma Aldrich, UK) in tris-buffered saline (TBS) and 0.5% (v/v) Tween 20. They were then incubated overnight at 4o_{C with the respective antibodies: phospho-specific sheep} anti-phospholemman (PLM Ser68) antibody (Badrilla, UK at 1:2000), Rabbit anti FXYD1 (PLM) (Abcam ab76597 at 1:1000). The blots were washed and incubated with appropriate horseradish peroxidase-conjugated secondary antibodies for 2 h at room temperature. Immunoreactive bands were visualized

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by enhanced chemiluminescence (ECL) western blotting detection kit. Exposure times were adjusted to ensure that for each antibody the signal detected was within the linear range. Of note separate gels were run for blotting for phosphorylated PLM and total PLM to avoid cross signaling of the antibodies. Membranes were then stripped of bound antibodies by incubation with stripping buffer for 30 min at room temperature (RT). The blots were reprobed with rabbit anti α-actinin to sarcomeric (Abcam ab68167 at 1:1000). α-actinin was used as a loading control and phospho-PLM and total PLM were each normalized to this house keeping protein on the respective blots. Band intensity was quantified by densitometry using ImageJ software. Since PLM is phosphorylated by both PKA and PKC (Presti et al., 1958a, b) treatments, least phosphorylation was attained by treating cells with PKC inhibitor. Protein phosphorylation was measured as the ratio of the density of the band of phospho-PLM to the band of total PLM protein after normalization to the corresponding density of the band of α-actinin. All samples were normalized to DMSO. One-way ANOVAs with post hoc test or unpaired two-tailed Student’s t-tests were used as appropriate. Data is presented as mean ± s.e.m. Statistical signiﬁcance, when achieved, is indicated as *P≤0.05, **P≤0.01, ***P≤0.001.

FRET Imaging

FRET imaging experiments were performed 24–48h after infection with adenovirus carrying each targeted sensor, as described by Stangherlin A et al.,2011. Cells were maintained at room temperature in a modiﬁed Ringer solution (140mM NaCl, 3mM KCL, 2mM MgCL2(x 6 H2O), 1mM CaCl2(x 2 H2O), 15mM glucose, 10mM Hepes, pH 7.2). An inverted microscope (Olympus IX71) with a PlanApoN, 60, NA 1.42 oil immersion objective, 0.17/FN 26.5 (Olympus, UK), was used. The microscope was equipped with a CoolSNAP HQ2

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monochrome camera (Photometrics) and a DV2 optical beam-splitter (MAG Biosystems, Photometrics). Images were acquired and processed using MetaFluor 7.1, (Meta Imaging Series, Molecular Devices). FRET changes were measured as the ratio of acceptor ﬂuorescence emission (545nm) to donor emission (480nm) i.e. 545nm/480nm expressed as R/R0, where R is the intensity fluorescence ratio at time t and R0 is the average emission fluorescence intensity ratio value of the last 8 frames taken before addition of the respective stimulus. Information on stimuli (inhibitors) and concentrations used. Bay60-7550 100nM (PDE2 specific blocker from Cayman Chemicals, USA) (Boess F.G et al., 2004), Rolipram 10μM (PDE4 specific blocker from Cayman Chemicals, USA) (Souness J. E et al.,1996), PF-04957325 100nM (PDE8 specific blocker from Pfizer, USA) (Vang et al.,2011).

Data Analysis

MetaFluor software was used to extract the mean intensity values within the Regions of Interest (ROIs) drawn on the cell at the beginning of the experiment, from the CFP intensity image, the YFP intensity image and the mean values from the corresponding YFP/CFP ratio image. The data was exported to an Excel file as intensity values over time for data analysis. Statistical analysis was performed with GraphPad Prism 5.0. The number of technical and biological replicates is indicated in the ﬁgures and their legends. All groups that were statistically compared showed equal variance. One-way ANOVAs with post hoc test or unpaired two-tailed Student’s t-tests were used as appropriate. Data is presented as mean ± s.e.m. Statistical signiﬁcance, when achieved, is indicated as *P≤0.05, **P≤0.01, ***P≤0.001.

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RESULTS:

Expression of targeted CUTie sensors in Rat Ventricular Myocytes

To monitor cAMP changes selectively at AKAP79 and at PLM/NKA, we used targeted versions of the cAMP FRET-based reporter CUTie (Surdo N.C et al.,2017). The design of the CUTie sensor is such that its fusion to targeting domains minimizes steric hindrance of the targeting domain on the FRET module, allowing for direct comparison of FRET changes detected at the different targeting sites. In this study, three variants of the CUTie sensor targeted at the plasmalemma were used (Fig 1A): a fusion of CUTie to AKAP79 (M90359.1)(Surdo N.C et al., 2017); a fusion of CUTie to PLM (NM_001278718.1)(Surdo N.C et al.,2017) and, for more general detection of cAMP dynamics at the whole plasmalemma, a fusion of CUTie to a short sequence (MP) which is myristoylated/palmitoylated and targets the sensor to plasma membrane raft domains (Terrin A et al., 2006). Expression of the AKAP79-CUTie, PLM-CUTie and MP-CUTie showed the expected localization both in adult rat ventricular myocytes (ARVMs) (Fig. 1B) and in neonatal rat ventricular myocytes (NRVMs) (supplementary Fig. 1A). Integration of AKAP79-CUTie into a complex including AC5/6 was previously demonstrated (Surdo et al 2017). Integration of PLM-CUTie within the expected macromolecular complex was confirmed by immunoprecipitation of the sensor and detection by western blot analysis of NKA in the immunoprecipitate, both in ARVMs (Fig. 1C) and in NRVMs (Supplementary Fig. 1B).

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Fig. 1: A) Schematic illustration of the targeted CUTie reporters used in this study. B) Wide field

fluorescence images showing the localization of AKAP79-CUTie, PLM-CUTie and MP-CUTie at the plasmalemma of ARVMs. Scale bar is 10μm. C) Western blot analysis showing co-immunoprecipitation of NKA with PLM-CUTie in ARVMs. Input is total protein. CTRL is the pulldown with beads without the GFP-Trap_A.

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THE cAMP response to β-adrenergic stimulation is attenuated at the PLM/NKA complex

We first investigated the amplitude and kinetics of the cAMP response detected by the three plasmalemma-targeted sensors in response to β-adrenergic stimulation. Application of 1nM Isoproterenol (Iso) to NRVMs expressing AKAP79-CUTie, PLM-CUTie and MP-CUTie resulted in significantly different FRET change at the three sites with, notably, no detectable response at PLM-CUTie (Fig. 2A-B). Similar results were obtained in ARVMs expressing the three sensors and treated with 5nM Iso (Fig. 2C and supplementary Fig. 2). Saturating cAMP, generated by application of the adenylyl cyclase activator forskolin and the non-selective PDE inhibitor IBMX resulted in similar amplitudes of FRET change at all sites, confirming that the three sensors respond with similar FRET change at saturation.

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Fig. 2. cAMP signals at distinct sub-plasmalemma domains on β-adrenergic stimulation. A)

Representative kinetics of FRET change on application of 1nM Isoproterenol (Iso) in NRVMs expressing MP-CUTie (blue), AKAP79-CUTie (green) or PLM-CUTie (red). Summary of FRET change recorded in NRVMs (B) and ARVMs (C) expressing MP-CUTie, AKAP79-CUTie or PLM-CUTie on application of 1nM Isoproterenol or 5nM Isoproterenol, respectively. SAT indicates application of forskolin 25μM + IBMX 100μM. Bars indicate means ± s.e.m. One-way Anova statistical analysis with Bonferonni’s post hoc correction was applied. N ≥ 9 from at least 4 biological replicates. * P≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001.

The Blunted cAMP response at PLM is specific to β-adrenergic receptor activation

We next sought to establish whether the blunted cAMP response at PLM compared to other sub-plasmalemma domains was specific for β-adrenergic

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stimulation. For this we applied a non-saturating concentration of forskolin which activates, in a non-selective manner, all cellular ACs. In NRVMs expressing the cytosolic cAMP sensor Epac1-camps (Nikolaev V.O et al., 2004), 5μM forskolin generates a global increase of cAMP in the cytosol similar to the cAMP increase elicited globally in the cytosol by 1nM Isoproterenol (Fig. 3A-C). When we applied 5μM forskolin to NRVM expressing the three targeted CUTie reporters we found that global stimulation of adenyl cyclases generates a clearly detectable increase in cAMP at all three sites (Fig. 4D) albeit the cAMP signal is higher at AKAP79 than at PLM and other membrane raft domains. These results show that cAMP levels can increase at the PLM/NKA complex to a similar extent as at other sub-plasmalemma domains and that it is the cAMP response to β-adrenergic activation that is specifically blunted at this site.

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Fig. 3. cAMP response at distinct sub-plasmalemma domains on general activation of ACs.

A) Wide field fluorescence image showing the homogeneous distribution of the cAMP Epac1-camps sensor in the cytosol. Scale bar is 10μm. B) Representative kinetics of FRET change on stimulation of NRVM expressing Epac1-camps with 5μM forskolin (F) and 1nM Isoproterenol (Iso). C) Summary of experiments performed as shown in B. D) Representative kinetics of FRET change (left panel) and summary of experiments (right panel) on application of 5μM forskolin to NRVMs expressing MP-CUTie, PLM-CUTie or AKA79-CUTie. SAT is forskolin 25μM + IBMX 100μM. N≥6. Bars indicate means ± SEM. One-way ANOVA with Bonferroni post-hoc correction was used for statistical analysis *P≤0.05, **P≤0.01, ***P≤0.001.

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PDEs attenuate the cAMP response at PLM

As PDEs are recognized as key determinants of cAMP compartmentalization, we next set out to assess whether the lack of detectable cAMP signal at the PLM/NKA complex on β-adrenergic stimulation may be due to enhanced degradation of cAMP by PDEs. FRET imaging of myocytes expressing the three targeted sensors and treated with the non-selective PDE inhibitor IBMX showed a detectable cAMP rise at all three sites, indicating that PDEs are active in unstimulated cells and contribute to regulation of the basal activity of ACs at the three locations. In addition, 0.5nM Isoproterenol in NRVMs (Fig 4A, B) and 1nM Isoproterenol in ARVMs (Fig. 4C and supplementary Fig 3) elicited a clearly detectable cAMP increase at all three sites when the cells were pre-treated with IBMX. These findings confirm that cAMP at PLM is under tight control of PDEs which degrade cAMP generated on β-adrenergic stimulation and hence attenuate the level of second messenger at this site compared to the level of cAMP at the entire plasma membrane and at AKAP79. These results indicate PLM/NKA as a unique cAMP β-adrenergic nanodomain which is regulated by a local cAMP pool that is different from that at AKAP79.

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Fig. 4. Effect of PDE inhibition on cAMP levels at different sub-plasmalemma sites. A)

Representative kinetics of FRET change recorded in NRVMs expressing MP-CUTie, PLM CUTie or AKAP79-CUTie on inhibition of PDEs with IBMX 100μM and subsequent application of 0.5nM Isoproterenol (Iso). B) Summary of FRET change in experiments performed as in A. C) Summary of FRET Change recorded in ARVM treated with IBMX 100μM and subsequent application of 1nM Isoproterenol (Iso). SAT indicates sensor saturation on application of FRSK 25μM+IBMX 100μM. One-way Anova with Bonferonni’s post hoc correction was applied. Bars indicate means ± s.e.m. N ≥ 9 from at least 6 biological replicates. * P≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001.

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Multiple PDEs contribute to the differential regulation of cAMP levels at PLM/NKA and AKAP79

To identify which PDE enzymes are involved in the differential regulation of cAMP levels at the PLM/NKA and AKAP79 complexes, we measured the FRET change reported by the three targeted CUTie reporters on application of PDE family-selective inhibitors. In all cases the PDE inhibitor was applied before the addition of Isoproterenol. We focused on PDE2, PDE4 and PDE8, three cAMP degrading enzymes that have been previously reported to be associated with regulation of cAMP signaling at the plasmalemma (Kokkonen and Kass et al., 2017)

As shown in Fig. 5A, application of the PDE2 selective blocker Bay 60-7550 (100nM) resulted in a larger cAMP increase at membrane rafts and at PLM/NKA than at AKAP79. Application of 1nM Isoproterenol to ARVM pretreated with Bay 60-7550 elicited a robust response at PLM as well as at membrane rafts that was significantly larger than the response at AKAP79.

Fig. 5B shows that application of the PDE4 selective blocker rolipram (10 μM) to ARVM expressing the targeted reporters resulted in an increase in cAMP that wass not significantly different at the three sites. Application of 1nM Isoproterenol to ARVM pretreated with rolipram also showed a robust cAMP response at PLM that was not significantly different from that at the other sites. Selective inhibition of PDE8 with PF-04957325 (PF, 100nM) resulted in a clear cAMP increase at the entire plasma membrane and at PLM/NKA but had no detectable effect at AKAP79 (Fig 5C). These results combined indicate that PDE2, 4 and 8 play a role in the regulation of cAMP at PLM/NKA whereas at AKAP79 PDE4 has the predominant role.

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Fig. 5: Effect of selective PDE inhibition on cAMP levels at sub-plasmalemma compartments.

A) Summary of FRET change on inhibition of PDE2 with Bay60-7550 (100nM) followed by β-AR stimulation with 1nM Isoproterenol (Iso) in ARVMs expressing the plasmalemma-targeted reporters. B) Summary of FRET change on inhibition of PDE4 with Rolipram (10μM) followed by β-AR stimulation with 1nM Iso in ARVMs expressing targeted reporters. C) Summary of FRET change on inhibition of PDE8 with PF-09457325 (PF)(100nM) followed by β-AR stimulation with 1nM Iso in ARVMs. SAT indicates saturating stimulus (forskolin 25μM+IBMX 100μM). One-way Anova with Bonferonni’s post hoc correction was applied. Bars indicate means ± s.e.m. N ≥ 4 from at least 3 biological replicates. * P≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001.

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β-adrenergic stimulation results in increased phosphorylation of PLM at S68

Residue S68 of PLM is reported to be one of the main targets of PKA mediated phosphorylation in cardiac myocytes (Silvermann B.Z et al.,2005; Pavlovic D et al., 2007; Han F et al., 2006; Fuller W et al., 2009). It was surprising therefore to observe undetectable increase in cAMP at this site. Indeed, western blotting analysis of ARVM treated with Iso showed a concentration-dependent increased in PLM phosphorylation at S68 (Fig 6). Of note however, and in agreement with previous findings (Han F et al., 2006; Fuller W et al.,2009), selective inhibition of PKC with bisindolylmaleimide I resulted in significant reduction of PLM phosphorylation at S68, suggesting the possibility that increased phosphorylation of this site on Iso stimulation may be indirect and rely on PKC activation and therefore be independent of a local increase in cAMP.

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Fig. 6: Phosphorylation of PLM at S68 on β-adrenergic receptor stimulation by Isoproterenol.

A) Western blot analysis of endogenous PLM in ARVMs on application of H89 30μM for PKA inhibition, Bisindolylmaleimide I (BisIndol) 1μM for PKC inhibition, 0.1nM Isoproterenol (Iso), 1nM Isoproterenol, 1μM Isoproterenol, and IBMX and Saturating stimulus (SAT) (forskolin 25μM + IBMX 100μM). B) Summary of phosphorylation of PLM on increasing Isoproterenol concentrations. One-way Anova with Bonferonni’s post hoc correction was applied. Bars indicate means ± s.e.m. N ≥ 4 biological replicates. Statistical significance was expressed as * P≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001. Significance was measured against DMSO for each respective treatment.

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Expression of targeted AKAR4 sensor in rat ventricular myocytes

To directly investigate whether the increase in phosphorylation at PLM observed on Iso application is PKA-mediated and to compare the local PKA activity at PLM/NKA with the activity at AKAP79, we targeted the FRET-based PKA activity reporter AKAR4 (Depry C et al.,2010) at the two sub-plasmalemma sites via fusion of the AKAR4 sensor to PLM or AKAP79. Expression of AKAP79-AKAR4 and PLM-AKAP79-AKAR4 in both NRVMs (supplementary Fig. 5A) and ARVMs (Fig. 7A) showed the expected localization at the plasmalemma, although PLM-AKAR4 often showed aggregate fluorescence in the sarcoplasm (Fig 7A). To exclude any artifactual signal from these aggregates, regions of interest for the analysis of FRET change were drawn selectively around the plasmalemma. To confirm incorporation of the targeted reporters in the expected multiprotein complex, western blot analysis was performed. As shown in Fig 7A and supplementary Fig 5A, PLM AKAR4 co-immunoprecipitates with endogenous NKA in both ARVMs and NRVMs, respectively. Similarly, AKAP79-AKAR4 coimmunoprecipitates with endogenous AC5/6 in ARVMs (Fig 7A).

Next, ARVMs expressing AKAP79-AKAR4 or PLM-AKAR4 were treated with Iso followed by saturation with forskolin 25μM + IBMX 100μM. The FRET change recorded on saturation was similar for PLM-AKAR4 and AKAP79-AKAR4, allowing direct comparison of the FRET changes at the two locations (Fig 7B - C). Surprisingly, despite no detectable cAMP signal was recorded at PLM when the NRVMs were treated with 1nM Iso, at this agonist concentration the FRET signal from both PLM-AKAR4 and from AKAP79-AKAR4 was close to saturation (supplementary Fig.4B - C). A very robust PKA activity that was not significantly different from the PKA activity at AKAP79, was detected at PLM also when the ARVMs were challenged with 0.5nM and 0.1 nM

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Isoproterenol (Fig 7B - C). Similar results were obtained in NRVM (supplementary Fig. 5B - C). These findings indicate that despite generating different cAMP levels at PLM and AKAP79, β-adrenergic stimulation induces a comparable level of PKA activity at those two sites.

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Figure 7. Effect of β-adrenergic stimulation on local PKA activity at PLM and AKAP79. A)

Left panels: wide field fluorescence image showing the appropriate localization of PLM-AKAR4 and AKAP79-AKAR4 at the plasmalemma in ARVMs. Scale bar = 10μm. Right panels: western blot analysis showing co-immunoprecipitation of NKA with PLM-AKAR4 and of AC5/6 with AKAP79-AKAR4 respectively from ARVMs lysates. B) Representative kinetics of FRET change on β-adrenergic stimulation of ARVMs expressing either PLM-AKAR4 (red) or AKAP79-AKAR4 (green) with 0.1nM or 0.5nM Isoproterenol (Iso), as indicated. C) Summary of FRET change in experiments performed as shown in B. Solid bars represent peak response and striped bars represent plateau amplitudes of PKA activity at each site. The FRET change at saturation (forskolin 25μM + IBMX 100μM) after each stimulus is also shown (SAT). One-way Anova with Bonferroni’s post hoc correction was applied. Bars indicate means ± s.e.m. N ≥ 6 from at least 3 biological replicates. * P≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001.

Phosphatases attenuate PKA activity at the AKAP79 complex.

A possible explanation for the apparent inconsistency between local cAMP signal and local PKA activity at PLM and at AKAP79 is that a higher phosphatase activity may be present at AKAP79 such that enhanced PKA activation is counterbalanced by more robust dephosphorylation at AKAP79 as compared to PLM, resulting in a similar phosphorylation level at the two sites. To test this hypothesis, rat ventricular myocytes expressing the PLM-AKAR4 and AKAP79-AKAR4 were treated with a combination of the phosphatase inhibitors calyculin (30nM), a non-selective phosphatase inhibitor, and cyclosporin (100nM) which inhibits calcineurin (protein phosphatase 2B) in the presence of 0.1nM Isoproterenol. Inhibition of phosphatases resulted in a significantly higher increase in phosphorylation of AKAP79-AKAR4 than at PLM-AKAR4, both in ARVM (Fig. 8A – B) and in NRVMs (Fig 8C), confirming a more prominent role of phosphatases at AKAP79 than at PLM.

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Fig. 8: Effect of phosphatases inhibition on PKA-dependent phosphorylation at AKAP79 and PLM.

A) Representative kinetics of FRET change recorded in ARVMs expressing PLM-AKAR4 and AKAP79-AKAR4 on application of the phosphatase inhibitors (PPI) calyculin 30nM + cyclosporin 100nM after β-adrenergic stimulation with 0.1nM Isoproterenol (Iso).B) Summary of FRET change of experiments performed in ARVMs as shown in A. C) Summary of FRET change of experiments

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performed in NRVMs in the same conditions as in A. One-way Anova with Bonferroni’s post hoc correction was applied. Bars indicate means ± s.e.m. N ≥ 3 from at least 2 biological replicates. Statistical significance if achieved is expressed as * P≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001.

To further assess whether the phosphatase activity at AKAP79 is higher than at PLM/NKA, maximal activation of PKA was achieved in cardiac myocytes expressing PLM-AKAR4 or AKAP79-AKAR4 before addition of the PKA inhibitor H89 (30μM). On inhibition of PKA the FRET signal is expected to decrease as the sensor is dephosphorylated by phosphatases. As shown in Fig 9A - B for NRVM and in supplementary Fig 6 for ARVM, dephosphorylation of AKAP79-AKAR4 was significantly more robust than dephosphorylation of PLM-AKAR4, both in terms of extent and in terms of rate of dephosphorylation. These findings further confirmed that the phosphatase activity at AKAP79 is significantly higher than at PLM. Interestingly, in 35% of the cells expressing PLM-AKAR4 and 23% of the cells expressing AKAP79-AKAR4, the dephosphorylation of the sensor appeared to follow a ‘two-steps’ kinetics, with the second step bringing the signal significantly below baseline (Fig 9C - D). When the two dephosphorylation steps were analyzed separately, we found that for both AKAP79-AKAR4 and PLM-AKAR4, the first dephosphorylation step shows characteristics that are similar, both in terms of amplitude and kinetics, to the ‘single–step’ dephosphorylation detected with the same sensors, with the amplitude and speed of dephosphorylation being more pronounced at AKAP79 than at PLM. In contrast, the second step of dephosphorylation showed similar features for the two sensors and showed slower kinetics at both sites (Fig 9C - D).

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Figure 9: Effect of local phosphatase activity at AKAP79 and PLM as revealed by PKA inhibition. A) ‘Single-step’ dephosphorylation. Representative kinetics of FRET change on

application of H89 30μM after saturation (SAT = 25 μM forskolin + 100 μM IBMX) in NRVMs expressing PLM-AKAR4 (red) or AKAP79-AKAR4 (green): B) Summary of FRET change under conditions as in A. C) ‘Two-steps’ dephosphorylation. Representative kinetics of FRET change on application of H89 30μM after saturation (SAT = 25 μM forskolin + 100 μM IBMX) in NRVMs expressing PLM-AKAR4 (red) or AKAP79-AKAR4 (green). D) Summary of FRET change for cells showing ‘two-steps’ dephosphorylation kinetics, as in C. One-way Anova with Bonferonni’s post hoc correction was applied. Bars indicate means ± s.e.m. N ≥ 10 from at least 6 biological replicates. * P≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001.

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DISCUSSION

PKA is a promiscuous enzyme that phosphorylates a multitude of protein targets within the cardiac myocyte. This versatility results in PKA being involved in the regulation of multiple functional effects, in some cases with opposing outcome. For example, on adrenergic stimulation, PKA-mediated phosphorylation of LTCC and PLB leads to increased [Ca2+_]i_{and positive inotropy, while} PKA-mediated phosphorylation of TPNI reduces the affinity of the myofilament for Ca2+_{, seemingly nullifying the effect of increased [Ca}2+_{]i. A similarly conflicting} outcome is achieved via PKA phosphorylation of PLM at Ser68, which results in enhanced NKA activity. The stimulated NKA pump moves Na+_{outside the cell} which in turn favors extrusion of Ca2+ _{via the NCX, resulting in reduction of} [Ca2+_{]i (Bers D.M and Despa S., 2009; Despa S. et al.,2008). How the myocyte} coordinates these apparently divergent effects of adrenergic stimulation on [Ca2+_]i is unclear. We recently demonstrated that catecholamines generate a cAMP response that is significantly smaller and delayed at the myofilament compared to the response at the LTCC/AKAP79 complex. When the difference in local cAMP levels is abrogated, a less efficient contraction ensues (Surdo N.C et al., 2017), suggesting that differential control of cAMP/PKA signals at different adrenergic targets is required to optimally coordinate regulation of contraction and relaxation. Here, we hypothesized that a similar distinct handling of cAMP/PKA signals may underpin the regulation of LTCC and PLM/NKA at the plasmalemma.

By using selectively targeted FRET-based reporters for cAMP levels and PKA activity we demonstrate both in adult and neonatal rat cardiac myocytes that a low concentration of Iso that generates a clear cAMP increase at the AKAP79/ LTCC complex fails to generate a detectable cAMP rise at the PLM/NKA

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complex. The low cAMP response at PLM/NKA does not depend on inefficient coupling of this site with ACs, as global activation of all cellular ACs results in a clear increase in cAMP at PLM/NKA, although the amplitude of the cAMP signal is not as high as at AKAP79/LTCC. The larger increase in cAMP detected at AKAP79-CUTie could be due to the sensor forming a complex with AC5/6 and therefore has closer proximity to the source of cAMP compared to PLM-CUTie.

We found that the heterogeneity in the cAMP response relies on differential local regulation of cAMP levels by PDEs. By degrading cAMP, PDEs appear to selectively contain the increase in the second messenger levels within a domain immediately surrounding PLM/sNKA. We found that PDE2 and PDE8 play a major role in selectively shielding the PLM/NKA complex from cAMP generated on activation of the β-adrenergic receptor, whereas PDE4 appears to regulate more generally cAMP levels at both AKAP79/LTCC and PLM/NKA. The exclusive increase in cAMP levels at the PLM/NKA complex, but not at the AKAP79/LTCC complex, on inhibition of PDE2 and PDE8 opens the possibility to target these enzymes to achieve selective manipulation of NKA activity for therapeutic purposes.

Despite the difference in the amplitude of the cAMP response to catecholamines at the AKAP79/LTCC and PLM/NKA complexes, we found that the PKA-dependent phosphorylation of local targets is similar at the two sites. Based on our data this apparent discrepancy is explained by a more robust phosphatase activity at AKAP79 compared to PLM. PP1 and PP2A are considered to be primarily responsible for counterbalancing the effects of PKA-mediated phosphorylation in the heart. The activity of phosphatases is known to depend on their subcellular localization and/or binding to accessory subunits (Virshup and

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Shenolikar., 2009) and there is evidence suggesting that their activity may differ substantially in different subcellular cardiac myocyte microdomains (Marx S.O et al., 2000, Yano M et al., 2005). Although further studies will be required to define the identity of the phosphatases involved in differential regulation of the AKAP79/LTCC and PLM/NKA complexes, PP1 has been shown to dephosphorylate PLM at the PKA site S68 and decrease NKA activity (El-Armouche et al., 2011) whereas AKAP79 interacts with PP2B as well as PP1 (Le A.V et al., 2011). One unexpected feature we uncovered in our experiments is a ‘two-steps’ kinetics of dephosphorylation of the PKA targets at both sites. A number of mechanisms may underpin this behavior. One possibility is the involvement of phosphatase inhibitor peptides. PP1 is specifically and potently inhibited by inhibitor peptide 1 (I-1). I-1 requires PKA-mediated phosphorylation at Thr35 (Endo S et al., 1996) and, on adrenergic stimulation, up to 70% of I-1 can be in its phosphorylated state (Foulkes J.G and Cohen P., 1979). Phosphorylation of I-1 at Thr35 is reversed by both PP2A and PP2B (El-Armouche A et al., 2006). Thus, on PKA inhibition, a second wave of PP1 mediated dephosphorylation may ensue when PP2A and PP2B have achieved dephosphorylation of I-1, releasing its inhibition on PP1. Other mechanisms, however, are equally possible, including local recruitment of additional phosphatases or phosphatase trans-activation via dephosphorylation, a mechanism that has been shown to operate in neurons where PP1 locally dephosphorylates other PP1 molecules at the synapse, providing a feed-forward mechanism for rapid activation of PP1 (Hou H et al., 2011). Additional studies will be required to establish the mechanism involved in the biphasic dephosphorylation of PKA targets and its functional significance in the regulation of PKA signaling at the AKAP79/LTCC and PLM/NKA complexes.

Overall, the finding presented here support a model whereby cAMP/PKA signaling is uniquely regulated at AKAP79/LTCC and PLM/NKA. Similar sensitivity to adrenergic stimulation is achieved at these two sites by differential

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local modulation of cAMP levels and phosphatase activity. Such an organization may be required to avoid excessive PKA activity at AKAP79/LTCC, a site that is in close proximity to the source of cAMP, while allowing at the same time PKA activation of NKA without a large amount of cAMP being required to escape the AKAP79/LTCC/β-AR/AC5-6 domain. Interestingly, there is evidence to support localization of the NKA-α2and NKA-α3 isoforms to the T-tubules, at the junction with the SR, where they could regulate local NCX and [Ca2+_{]i (Juhaszova M and} Blautein M.P., 1997; Despa S et al., 2012). The proximity of PLM/NCX to AKAP79/LTCC/β-AR/AC5-6 complexes at the T tubules combined with the balanced regulation of cAMP levels and phosphatase activity at these two complexes may be required to secure efficient adrenergic control of NKA activity without excessive spill-over of cAMP. Such arrangement would maintain signal compartmentalization and avoid inappropriate activation of PKA at more remote sites. Notably, we find similar results in adult and neonatal cardiac myocytes. As neonatal myocytes are largely devoid of T tubules, it appears that the local regulation of cAMP and phosphatase activity at AKAP79/LTCC and PLM/NKA relies on the clustering of relevant molecular components within defined subcellular domains which is independent of the complex geometry of the fully mature cardiac myocyte.

Disruption of the balanced regulation of local cAMP and phosphates activity may affect cardiac function and contribute to pathogenic mechanisms. In healthy hearts, the positive chronotropic effect of adrenergic stimulation results in increased [Na+_{]i which is normally balanced by PKA-dependent activation of} NKA and increased extrusion of Na+_{from the cell. In pathological conditions,} such as heart failure (HF), reduced expression of NKA and elevated [Na+_{]i are} well documented, both in humans and animal models (Despa S et al., 2002, Pieske B et al., 2002, Baartsheer A et al., 2003, Louch W.E et al., 2010, Schillinger W et al., 2006.), leading to imbalanced Na+_/Ca2+_{exchange at the mitochondria,} reduced ATP production and increased H2O2 (Liu T and O’Rourke B., 2008;

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Despa S and Bers D.M., 2013). In addition, elevation of [Na+_{]i favors Ca}2+_influx via NCX reversal, Ca2+_{overload and arrhythmia (Despa S et al., 2008; Park K.C.,} 2018). Both PDE2(Hua R et al., 2012; Aye T.T., 2012) and PP1(Neumann et al., 1997) have been shown to be upregulated both in human and experimental HF. The data we present here suggest that in HF further attenuation of the cAMP response at PLM/NKA by increased PDE2 activity and enhanced PLM dephosphorylation by PP1 may concur to reduce the adrenergic regulation of NKA, potentially worsening elevated [Na+_{]i and aggravating the negative effects} on cardiac metabolism and oxidative stress that associate with this condition. A detailed understanding of the local regulation of cAMP/PKA signaling in myocytes may provide much needed novel avenues for therapeutic intervention.

ACKNOWLEDGEMENTS:

This work was supported by the British Heart Foundation (PG/10/75/28537 and RG/17/6/32944), the BHF Centre of Research Excellence, Oxford (RE/13/1/30181) and Groningen Research Institute of Pharmacy, The Netherlands. The PDE8 inhibitor PF-09457325 was kindly provided by Pfizer. Some of the Artwork was kindly edited by Augustin Boisleux. Also special thanks to Stefania Monterisi and Mark Richards and Chela Nunez Alonso for technical support.

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