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Cyclic nucleotide phosphodiesterases (PDEs): coincidence detectors acting to spatially and temporally integrate cyclic nucleotide and non-cyclic nucleotide signals Donald H. Maurice*†1 , Lindsay S. Wilson†2 , Sarah N. Rampersad†, Fabien Hubert*, Tammy Truong†, Milosz Kaczmarek*, Paulina Brzezinska*, Silja I. Freitag*, M. Bibiana Umana* and Alie Wudwud†
Biochemical Society Transactions
www.biochemsoctrans.org
*Department of Biomedical and Molecular Sciences, Queen’s University, Kingston, Ontario, Canada, K7L 3N6 †Department of Pathology and Molecular Medicine, Queen’s University, Kingston, Ontario, Canada, K7P 3N6
Abstract The cyclic nucleotide second messengers cAMP and cGMP each affect virtually all cellular processes. Although these hydrophilic small molecules readily diffuse throughout cells, it is remarkable that their ability to activate their multiple intracellular effectors is spatially and temporally selective. Studies have identified a critical role for compartmentation of the enzymes which hydrolyse and metabolically inactivate these second messengers, the PDEs (cyclic nucleotide phosphodiesterases), in this specificity. In the present article, we describe several examples from our work in which compartmentation of selected cAMP- or cGMP-hydrolysing PDEs co-ordinate selective activation of cyclic nucleotide effectors, and, as a result, selectively affect cellular functions. It is our belief that therapeutic strategies aimed at targeting PDEs within these compartments will allow greater selectivity than those directed at inhibiting these enzymes throughout the cells.
Introduction Cellular homoeostasis requires efficient integration of signals encoded by myriad primary messengers, including numerous hormones and transmitters. Inefficient integration of these signals destabilizes critical cellular processes, and this instability can promote disease-initiating maladaptive cellular responses [1]. Many cellular systems have evolved to sense the presence of primary messengers, to decode their signals and, subsequently, to operationalize their intent in cells. These systems are collectively referred to as signal transduction systems. One of the earliest signal transduction systems described allows cells to respond to primary messengerencoded signals by altering the intracellular concentration of a second messenger, cAMP [2]. Subsequently, a cGMP signalling system was also identified and described [2,3]. Both the cAMP- and the cGMP-signalling systems regulate a large number of important physiological processes, including visual transduction, inflammation, apoptosis, cell proliferation and differentiation, as well as critical metabolic pathways such as steroidogenesis, insulin secretion and glycogen synthesis, as well as glycogenolysis, lipogenesis and lipolysis [1–3]. Key words: cAMP, cGMP, cyclic nucleotide phosphodiesterase (PDE), endothelium, platelet. Abbreviations: AC, adenylate cyclase; AJ, adherens junction; AKAP, A-kinase-anchoring protein; ECM, extracellular matrix; Epac, exchange protein directly activated by cAMP; ER, endoplasmic reticulum; HAEC, human arterial endothelial cell; HMVEC, human microvascular endothelial cell; IP3R, InsP3 receptor; IRAG, IP3R-associated cGMP kinase substrate; PDE, cyclic nucleotide phosphodiesterase; PI3K, phosphoinositide 3-kinase; PKA, protein kinase A; PKG, protein kinase G; VEC, vascular endothelial cell; VE-cadherin, vascular endothelial cadherin. 1 2
To whom correspondence should be addressed (email [email protected]). Present address: Pfizer Neuroscience, 610 Main Street, Cambridge, MA 02139, U.S.A.
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Synthesis of cellular cAMP or cGMP is catalysed by both plasma-membrane-associated and cytosolic nucleotidyl cyclases. Indeed, nine distinct plasma membrane ACs (adenylate cyclases) and one cytosolic AC catalyse the synthesis of cAMP from ATP [4]. Similarly, several particulate and cytosolic GCs (guanylate cyclases) convert GTP into cGMP in cells [3]. Since diffusion of ‘free’ cyclic nucleotides in cells is very fast (∼400 μm2 ·s − 1 ) [5,6], significant efforts have focused on delineating mechanisms which allow for spatiotemporal resolution of the cellular actions of cAMP and cGMP. Considerable evidence supports the notion that cyclic nucleotide hydrolysis catalysed by the members of a large multigene family of enzymes called PDEs (cyclic nucleotide phosphodiesterases) actively participate in this spatiotemporal resolution. Informed by data obtained during the sequencing of the human genome, it is now clear that there are 11 different PDE families, that most families have several genes, and that these genes can be differentially expressed and processed in cells [5,7]. In addition to differences in structure, distinct PDEs also have distinct kinetic properties and modes of regulation and can be expressed selectively by cells and differentially localized within cells. Of therapeutic importance, selective therapeutic agents for many of the PDE families are known and are being used in the treatment of many diseases and conditions. If not hydrolysed, cAMP and cGMP can affect cell functions through actions at multiple effectors, including cyclicnucleotide-activated protein kinases [PKA (protein kinase A) and PKG (protein kinase G)], Epacs (exchange proteins directly activated by cAMP) and cyclic-nucleotide-gated Biochem. Soc. Trans. (2014) 42, 250–256; doi:10.1042/BST20130268
Targeting cAMP Signalling to Combat Cardiovascular Diseases
cationic channels [5,7]. The idea that interactions between individual components of the cAMP-signalling system [GPCRs (G-protein-coupled receptors), G-proteins, ACs, PKAs/Epacs/cyclic-nucleotide-gated channels and PDEs] allow the formation of multiple distinct intracellular cAMPsignalling complexes (hereinafter ‘cAMP-signalosomes’) is gaining acceptance [5,6,8]. Although less well developed, a similar concept is proving useful in assessing how cGMP effects in cells are co-ordinated. In our laboratory, we are largely interested in elucidating how compartment-specific cAMP, or cGMP, signalling selectively regulates actions of cells in the cardiovascular system [5,9–15] and build on excellent earlier work in multiple cell types in which cAMPsignalosomes were shown to contain cAMP effectors (PKA or Epac1, or both) and individual PDEs, including PDE3A, PDE3B, PDE4D or PDE7A [7,16]. Continuing advances in our understanding of mechanisms whereby biological and/or therapeutic agents affect cellular functions by altering levels of these cyclic nucleotides are enabling the identification and development of therapeutic agents for use in numerous human diseases [4,6,17–19].
A PDE4D/Epac1/Rap1-containing cAMP-signalosome co-ordinates cAMP effects on VEC (vascular endothelial cell) intercellular contact adhesions and paracellular permeability Numerous cAMP-elevating agents have been reported to reduce VEC permeability to both solutes and cells by strengthening intercellular AJ (adherens junction)-based structures and by promoting the formation of large rings of cortical actin. Studies have shown that activation of either PKA or Epac can co-ordinate these cAMP-dependent effects [20–26]. Since PDE-mediated hydrolysis of cAMP controls the magnitude, duration, and subcellular compartmentation of cAMP [16,19], we and others have addressed their roles in these actions of cAMP [14,21]. Overall, these studies have identified a dominant role for PDE4 over PDE3 and PDE2, the other VEC PDEs. Four genes (PDE4A–PDE4D) yield 20 PDE4 variants, and these can be stratified into long or short forms [5]. Whereas long PDE4s contain N-terminal UCRs (upstream conserved regions) (UCR1/2) and unique targeting domains, short PDE4s do not. Individual PDE4s interact with binding partners, and this often allows their localization within discrete subcellular domains in cells, including VECs [5,14,27]. Although PDE4A, PDE4B and PDE4D variants are each expressed in VECs, most data identify PDE4D as the dominant PDE4 family members involved in controlling these effects of cAMP [11,14,28]. PDE4 inhibitors reduce VEC migration, proliferation and permeability [11,14,28–31]. In early work, we showed that macrovascular and microvascular human VECs could form numerous distinct cAMP-signalosomes, that these cAMP-signalosomes contained either PKA or Epac1, either PDE3B or PDE4D, and that they probably represented the tools through which
cAMP could regulate selectively multiple distinct events in these cells [11,14,28,32]. These data were consistent with an earlier suggestion [33] that cAMP-regulated functions in VECs were co-ordinated within distinct ‘pools’ or compartments. Previously, we uncovered a role for compartmentation of a PDE4D/Epac1-based cAMP-signalosome in regulating HAEC (human arterial endothelial cell) permeability [14] (Figure 1). Moreover, we showed that these actions were coordinated through the integration of this cAMP-signalosome into the VE-cadherin (vascular endothelial cadherin)-based structure known to form VEC AJs. Thus we observed that PDE4 inhibition with rolipram, but not PDE3 inhibition with cilostamide, reduced HAEC permeability. Consistent with a dominant role for Epac1 over PKA in mediating this effect, siRNA-based knockdown of Epac1, but not of PKACα (PKA catalytic subunit α), markedly increased HAEC permeability. Knockdown of individual PDE4 gene products identified PDE4D, but not PDE4A or PDE4B, as dominant in controlling cAMP-mediated activation of Epac1 and in the control of its permeability-reducing effects. Most interestingly, our data defined a role for PDE4D as the link that allowed Epac1 integration into VE-cadherin-based structures. Indeed, although PDE4 inhibition with rolipram decreased VEC permeability, PDE4D knockdown increased permeability and caused a marked reduction in VE-cadherinassociated Epac1. Since PDE4 inhibitors had decreased VEC permeability, we had hypothesized that PDE4D knockdown would similarly reduce HAEC permeability. In contrast, knockdown of PDE4A or PDE4B, or of the dominant HAEC PDE3, namely PDE3B, had little or no effect on VEC permeability. Interestingly, although HAEC VE-cadherinbased isolates also contain PKA, in these cells PDE4D did not locally regulate integration of this cAMP effector or its ability to affect AJ-based adhesions of permeability. How our findings relate to other work of Beavo and colleagues in which cGMP-elevating agents were shown to alter the effect of cAMP-elevating agents on VEC permeability [34] is currently unclear. Although PDE2A, PDE3A and PDE3B are expressed in many human, bovine and porcine arterial or venous VECs used in studies of VEC permeability [21,29,34– 38], these enzymes were not found to integrate into the Epac1/PDE4D-signalosome [14] and as such are unlikely to affect VEC permeability through local events within AJassociated cAMP-signalosomes. Although our data identified Rap1 as the likely downstream actor in promoting actions of this cAMP-signalosome, elucidating the involvement of other agents known to affect VEC permeability such as Cdc42, Rac, PAK (p21-activated kinase), Krit1 (Krev1-interaction trapped gene) or of other PDE4D-interacting proteins, including RACK1 (receptor for activated C-kinase 1), AKAPs (Akinase-anchoring proteins) and β-arrestins or spectrin will require further work [39]. Of potential therapeutic importance, we identified the sequences within Epac1 that promote a direct interaction with PDE4D [14]. We also demonstrated that permeability of HAECs was perturbed when a cell-permeant peptide designed to antagonize this interaction was added to cells. C The
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Figure 1 Examples of cAMP- and cGMP-signalosomes present in cells of the cardiovascular system Proteins identified in the distinct macromolecular complexes are defined in the text.
Thus an Epac1-based peptide designed to interfere with Epac1–PDE4D binding markedly reduced the ability of PDE4D to control Epac1-based effects at the border. These data strongly suggest that small molecules designed to compete for binding between PDE4D and Epac1 might represent powerful agents with which to selectively control VEC barrier functions (Figure 1).
A PDE3B/Epac1/R-Ras signalosome co-ordinates cAMP-mediated VEC adhesions to ECM proteins, migration and tubule formation Many cAMP-elevating agents promote integrin-dependent VEC adhesions to ECM (extracellular matrix) proteins and through this effect can modulate VEC migration [28,29,32]. The relative dependence of this effect on PKA or Epac1 activation is currently unclear. Although PKA activation can promote integrin engagement and stimulate VEC adhesions, PKA activity itself is reported to be either increased or decreased in response to integrin–ECM engagement, and PKA inhibitors do not antagonize fully these cAMPstimulated adhesive events [40,41]. We have concluded that these somewhat contradictory data are most likely reflective of the fact that, whereas each PKA and Epac can regulate integrin engagement, the activity of these effectors are in C The
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turn affected by events downstream and upstream of integrin signalling [32]. Also, the species or vascular structures from which the VECs were derived, the identity of the ECM protein chosen for study and the choice of cAMP-elevating agent each significantly affected the relative involvement of PKA or of Epac in these events. In this context, we reported previously that cAMP-promoted increases in HAECs (macro-VECs) or HMVECs (human microvascular endothelial cells) (micro-VECs) adhesions to distinct ECM proteins were not equally dependent on PKA or Epac1 [32]. Briefly, we showed that, whereas cAMP-induced VEC adhesion was more PKA-dependent in HAECs than in HMVECs, the magnitude of the difference was ECMprotein-dependent. Although most cAMP-elevating agents tested (isoprenaline, forskolin, rolipram or cilostamide) could promote HAEC adhesion when PKA could be activated, in the presence of PKA inhibitors these agents only promoted adhesion of HMVECs, not HAECs. These data identified a functional link between Epac1 and PDE3-catalysed, or PDE4-catalysed, cAMP hydrolysis that was more dominant in HMVECs than in HAECs and suggest that integrin-based cAMP-signalosomes may underpin these differences [32]. Because initiation of angiogenesis involves VEC adhesion, and subsequent migration and tubule formation by these cells, and because cAMP-elevating agents affect each of these events, the ability of certain cAMP-elevating agents to
Targeting cAMP Signalling to Combat Cardiovascular Diseases
affect angiogenesis has been investigated [42–45]. Although PKA, or Epac1, probably co-ordinates these events through numerous separate cAMP-signalosomes, only recently have these events been studied comprehensively. In this context, we identified an Epac1/PDE3B/R-Ras-based signalosome that regulates integrin-based adhesions, migration and tubule formation by VECs in the context of angiogenesis [11] (Figure 1). In this signalosome, PDE3B and Epac1 reciprocally regulated one another’s activities as well as their association with a membrane compartment. We showed that PDE3B bound Epac1 directly via protein–protein interactions and tethered this cAMP effector to cellular membranes. In addition, we showed that PDE3B-catalysed cAMP hydrolysis acted to antagonize Epac1 binding of cAMP within this signalosome, and consequently inhibited Epac1 activation by cAMP. Interestingly, PDE3B association with Epac1 increased PDE3B activity through an as yet unknown mechanism. As described previously, cAMPmediated activation of Epac1 in cells promotes accumulation of GTP-bound/activated Rap1/2 and/or R-Ras proteins [11]. In turn, activated Rap or R-Ras can signal by activating one or many of a group of effectors including PLCε (phospholipase Cε), PLD (phospholipase D), Raf-1, p38MAPK (p38 mitogenactivated protein kinase) and PI3K (phosphoinositide 3kinase) [46–49]. Since a previous study had shown that PDE3B could be isolated with one of the two known PI3Kγ regulatory subunits, namely p84 [50], and a cardiac phenotype was associated with the loss of PI3Kγ catalytic domain (p110γ ) in mice had been attributed to increases in cAMP signalling [51], we investigated whether PI3Kγ was also tethered via PDE3B in VECs and whether this interaction was related to the PDE3B–Epac1 complex. Consistent with the idea that these three activities were integrated into a common cAMP-signalosome, PDE3B was found to interact directly with p84 through a protein–protein interaction distinct from the one that co-ordinated PDE3B–Epac1 binding. Since p84 and Epac1 bound PDE3B via distinct sites, we surmised that PDE3B could simultaneously tether and anchor both Epac1 and the p84-regulated form of PI3Kγ in cells. Consistent with this, PDE3 inhibition with cilostamide and selective PDE3B knockdown in HAECs each promoted Epac1dependent activation of Rap1 and R-Ras, and of PI3Kγ and its downstream activated kinases, in these cells. Indeed, our data clearly showed that this PDE3B-based scaffold allowed for a PDE3B-regulated pool of cAMP to directly promote Epac1dependent activation of PI3Kγ , PKB (protein kinase B) and ERK1/2 (extracellular-signal-regulated kinase 1/2) [11]. At a cell functional level, experiments confirmed that the PDE3B/Epac1/R-Ras/p84-regulated PI3Kγ -signalosome controlled cAMP-mediated HAEC adhesion, migration and tubule formation. Thus, whereas PDE3 inhibition with cilostamide and PDE3B knockdown each promoted HAEC adhesion to ECM proteins, and promoted tubule formation, knockdown of Epac1 and of p84 each abolished tubule formation behaviour in these cells. Most strikingly, all of these events occurred without addition of activators of ACs and, as such, occurred without changing global cAMP levels.
In a recent study, the catalytic domain of PI3Kγ (p110γ ) was identified as a potential AKAP that could tether PKA and PDE3B in murine cardiac myocytes and in cells expressing these proteins heterologously [52]. Since p110γ was not integrated into the PDE3B-based cAMP-signalosome which we identified in VECs when p84 was knocked down, it is currently unclear how this recent finding relates to the system that we describe in the present article. Of potential translational importance, we identified the sequence of PDE3B and two sequences in Epac1 that allowed these proteins to interact directly. On the basis of this information, we designed a cell-permeant PDE3Bbased peptide with which to antagonize binding between these proteins in cells. Consistent with our predictions, the PDE3B-based Epac1-displacing peptide effectively competed the binding between PDE3B and Epac1, promoted Epac1/RRas-dependent activation of PI3Kγ in these cells and stimulated the rate at which tubules formed on MatrigelTM . Most strikingly, the internal consistency of these data to our model was observed when the displacing peptide rendered cells insensitive to the effects of PDE3B inhibition. Together, the observations detailed in the two previous sections validate the concept that selective regulation of individual cAMP-sensitive cellular events can be achieved with reagents, or drugs, able to selectively disturb the formation of individual cAMP-signalosomes in cells.
PDE5A is resident in an ER (endoplasmic reticulum)-based cGMP-signalling complex On a return flight from a conference in Glasgow, during which my head was full of ideas concerning PDE-based signalling compartments (and whisky!), it struck me that compartments might not be needed in cells that expressed only one PDE, or only one effector. After all, if signalosomes were designed to allow enzyme-based selectivity, surely they would be redundant in a cell in which there was no choice. In contrast, if compartments were critical in allowing selective cell responses, independent of the enzyme involved, one would predict that they would be found in cells irrespective of the number of tools to which that cell had access. Because I was unaware of a cell type in which only one cAMPPDE, or one cAMP effector, was expressed, Lindsay Wilson and I turned to the platelet to investigate these ideas. On the basis of my previous work with the late Dr Richard Haslam, I ‘knew’ that PDE5A accounted for >90% of cGMP-PDE activity in platelets and that only PKG served as a cGMP effector in these cells. As a result of Lindsay’s observation that sildenafil could inhibit thrombin-induced Ca2 + transients without increasing global intraplatelet cGMP, we tested the hypothesis that PDE5 might be acting locally at the platelet ER [15]. More precisely, we determined whether PDE5 formed a functional part of the IP3R1 (InsP3 receptor 1)–IRAG (IP3R-associated cGMP kinase substrate)–PKG1β signalling complex which had been reported previously to allow cGMPdependent control of thrombin-induced Ca2 + transients in C The
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platelets [53]. Differential centrifugation of platelet lysates identified PKG1β (∼25%), PDE5 (∼45%) and IP3R1 (∼100%) in particulate cellular fractions, and selective immunoprecipitation of PKG1β, PDE5 or IP3R1 allowed their co-immunoprecipitation, a finding consistent with the idea that PDE5 was integral to the IP3R1–IRAG–PKG1β intraplatelet complex. Interestingly, this complex was only influenced by cGMP, as PKA inhibition did not affect the sildenafil-induced inhibition of Ca2 + transients and PKA and the cAMP PDE (PDE3A) were not co-immunoprecipitated with the cGMP signalling proteins. In addition to being able to co-immunoprecipitate these proteins, we also discovered that PDE5A could be differentially regulated depending on its cellular localization. We began to investigate the two pools of PDE5A present in platelets, one particulate and PKGassociated pool of PDE5 and another cytosolic or ‘bulk’ PDE5A which was not associated with PKG. Interestingly, we discovered that these two pools of PDE5A had different intrinsic activities. Whereas the non-PKG-associated pool had a higher specific activity, the PKG-associated PDE5A had a very low specific activity. Since previous work had indicated that PDE5 could be activated upon either cGMP binding to a PDE5 GAF-A domain or PKG phosphorylation or PDE5 at Ser102 [54–56], we investigated these two pools further to determine whether they could be differentially regulated. Uniquely, only the PKG1B-associated PDE5A could be selectively phosphorylated by PKG at Ser102 , whereas phosphorylation of the non-PKG-associated pool was not observed. In addition, this phosphorylation resulted in a significant activation of PDE5A, again, a result not observed in the cytosolic and non-PKG-associated pool of enzyme. We also observed that the PKG-associated pool of PDE5 could not be activated by cGMP alone in the absence of ATP, adding to our findings that these two pools were regulated differently. Together, these studies demonstrate that a single PDE isoform, namely PDE5A, can be differentially regulated depending on its binding partners and localization in the cell. Although previous work indicated that PDE5 was activated upon either cGMP binding to a PDE5 GAFA domain, or PKG phosphorylation of PDE5 at Ser102 [54–56], these earlier studies were silent on the relative importance of these mechanisms in cells. To address this issue, we compared the phosphorylation and activation of the PKG-associated and non-PKG-associated forms of PDE5 in 8-bromo-cGMP (1 mM, 15 min)-treated platelets. 8-Bromo-cGMP treatment of platelets markedly increased the Ser102 phosphorylation status and activity of the PKGassociated form of PDE5, but not that which was not associated with the kinase. Consistent with the idea that the phosphorylated PDE5 was resident within the IP3R1– IRAG–PKG1β complex, IP3R1 was recovered in the antiPKG immune complexes, but not in those representing bulk PDE5. Similarly, when anti-IP3R1 immune complexes were obtained from control or 8-bromo-cGMP-treated platelets, only the IP3R1-associated PDE5 was activated by 8-bromo-cGMP. An identical pattern of PDE5 activation C The
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was obtained when PDE5 was isolated using the method used originally to isolate and characterize the IP3R1–IRAG– PKG1β complex in platelets, namely cGMP–agarose [53]. Taken together, these data were consistent with the idea that only PKG-associated PDE5 was subject to PKG-catalysed phosphorylation and activation in cells treated with 8-bromocGMP. In addition to being selectively activated by PKG in cells, our data indicate that the IP3R1–IRAG–PKG1βassociated PDE5 had a lower specific activity than that isolated from the bulk cytosol. Indeed, the IP3R1–IRAG– PKG1β-associated PDE5 was only approximately 14% as active as that isolated in anti-PDE5 immunoprecipitates. Interestingly, our data are inconsistent with the idea that PDE5 and PKG form a stable complex in platelet cellular fractions devoid of IP3R1. Indeed, although PKG and PDE5 were each efficiently individually immunoprecipitated from platelet cytosolic fractions, these proteins never coimmunoprecipitated from this fraction. Because PDE5, PKG or IP3R1 was not enriched in IP3R1-containing subcellular fractions in cells incubated with a cGMP analogue, or with an NO (nitric oxide) donor, our data were also inconsistent with the idea that cGMP elevation triggers movement of these proteins between these fractions. Taken together, we believe that these results are consistent with the idea that PDEs and cyclic nucleotide effectors are compartmented in cells, independent of the number of such proteins available, resulting in efficient regulation and integration of intracellular signals. The examples highlighted in the present review support the notion that PDE-containing signalosomes are formed to regulate selected cellular processes and do not simply exist to isolate similar enzymes from each other. In the future, and of therapeutic importance, investigation into signalosomes and protein interaction partners will help us in identifying potential new drug targets. PDEs are favourable targets, and, although multiple family-specific inhibitors exist, the ability to distinguish between individual isoforms of PDEs is challenging. Targeting individual PDEs contained in signalosomes provides an opportunity to achieve a desired signal while avoiding another pool of this enzyme where potential side effects could interfere. Determining the protein interaction partners and protein-interface residues can only inform on the differences between pools of PDEs and allow us to use this information to make small molecules that can inhibit interactions or target specific nodes in signalling pathways. Since small-molecule compounds targeting global signalling pathways have the potential for unwanted toxicity and side effects, finer more precise therapeutics that target intracellular pools have the potential to be the next class of drugs to potentially benefit multiple disease areas.
Funding Funding for the studies described were provided by the Canadian Institutes of Health Sciences (CIHR).
Targeting cAMP Signalling to Combat Cardiovascular Diseases
References 1 Dennis, E.A. and Bradshaw, R.A. (2011) Signaling in development and disease. In Intracellular Signaling in Development and Disease (Dennis, E.A. and Bradshaw, R.A., eds), pp. 3–8, Academic Press, San Diego 2 Beavo, J. and Brunton, L. (2002) Cyclic nucleotide research: still expanding after half a century. Nat. Rev. Mol. Cell Biol. 3, 710–718 3 Kots, A., Martin, E., Sharina, I. and Murad, F. (2009) A short history of cGMP, guanylyl cyclases, and cGMP-dependent protein kinases. Handb. Exp. Pharmacol. 191, 1–14 4 Pierre, S., Eschenhagen, T., Geisslinger, G. and Scholich, K. (2009) Capturing adenylyl cyclases as potential drug targets. Nat. Rev. Drug Discov. 8, 321–335 5 Houslay, M.D., Baillie, G.S. and Maurice, D.H. (2007) cAMP-specific phosphodiesterase-4 enzymes in the cardiovascular system: a molecular toolbox for generating compartmentalized cAMP signaling. Circ. Res. 100, 950–966 6 Lee, L.C.Y., Maurice, D.H. and Baillie, G.S. (2013) Targeting protein–protein interactions within the cyclic AMP signaling system as a therapeutic strategy for cardiovascular disease. Future Med. Chem. 5, 451–464 7 Bender, A. and Beavo, J. (2006) Cyclic nucleotide phosphodiesterases: molecular regulation to clinical use. Pharmacol. Rev. 58, 488–520 8 Pawson, C.T. and Scott, J.D. (2010) Signal integration through blending, bolstering and bifurcating of intracellular information. Nat. Struct. Mol. Biol. 17, 653–658 9 Kerfant, B.-G., Zhao, D., Lorenzen-Schmidt, I., Wilson, L.S., Cai, S., Chen, S.R.W., Maurice, D.H. and Backx, P.H. (2007) PI3Kγ is required for PDE4, not PDE3, activity in subcellular microdomains containing the sarcoplasmic reticular calcium ATPase in cardiomyocytes. Circ. Res. 101, 400–408 10 Palmer, D., Jimmo, S.L., Raymond, D.R., Wilson, L.S., Carter, R.L. and Maurice, D.H. (2007) Protein kinase A phosphorylation of human phosphodiesterase 3B promotes 14-3-3 protein binding and inhibits phosphatase-catalyzed inactivation. J. Biol. Chem. 282, 9411–9419 11 Wilson, L.S., Baillie, G.S., Pritchard, L.M., Umana, B., Terrin, A., Zaccolo, M., Houslay, M.D. and Maurice, D.H. (2011) A phosphodiesterase 3B-based signaling complex integrates exchange protein activated by cAMP 1 and phosphatidylinositol 3-kinase signals in human arterial endothelial cells. J. Biol. Chem. 286, 16285–16296 12 Raymond, D.R., Carter, R.L., Ward, C.A. and Maurice, D.H. (2009) Distinct phosphodiesterase-4D variants integrate into protein kinase A-based signaling complexes in cardiac and vascular myocytes. Am. J. Physiol. Heart Circ. Physiol. 296, H263–H271 13 Beca, S., Helli, P.B., Simpson, J.A., Zhao, D., Farman, G.P., Jones, P.P., Tian, X., Wilson, L.S., Ahmad, F., Chen, S.R.W. et al. (2011) Phosphodiesterase 4D regulates baseline sarcoplasmic reticulum Ca2 + release and cardiac contractility, independently of L-type Ca2 + current. Circ. Res. 109, 1024–1030 14 Rampersad, S.N., Ovens, J.D., Huston, E., Umana, M.B., Wilson, L.S., Netherton, S.J., Lynch, M.J., Baillie, G.S., Houslay, M.D. and Maurice, D.H. (2010) Cyclic AMP phosphodiesterase 4D (PDE4D) tethers EPAC1 in a vascular endothelial cadherin (VE-Cad)-based signaling complex and controls cAMP-mediated vascular permeability. J. Biol. Chem. 285, 33614–33622 15 Wilson, L., Elbatarny, H., Crawley, S., Bennett, B. and Maurice, D.H. (2008) Compartmentation and compartment-specific regulation of PDE5 by protein kinase G allows selective cGMP-mediated regulation of platelet functions. Proc. Natl. Acad. Sci. U.S.A. 105, 13650–13655 16 Omori, K. and Kotera, J. (2007) Overview of PDEs and their regulation. Circ. Res. 100, 309–327 17 Potter, L.R. (2011) Regulation and therapeutic targeting of peptide-activated receptor guanylyl cyclases. Pharmacol. Ther. 130, 71–82 18 Derbyshire, E. and Marletta, M. (2012) Structure and regulation of soluble guanylate cyclase. Annu. Rev. Biochem. 81, 533–559 19 Francis, S., Blount, M. and Corbin, J. (2011) Mammalian cyclic nucleotide phosphodiesterases: molecular mechanisms and physiological functions. Physiol. Rev. 91, 651–690 20 Yamamizu, K. and Yamashita, J. (2011) Roles of cyclic adenosine monophosphate signaling in endothelial cell differentiation and arterial–venous specification during vascular development. Circ. J. 75, 253–260 21 Sayner, S. (2011) Emerging themes of cAMP regulation of the pulmonary endothelial barrier. Am. J. Physiol. Lung Cell. Mol. Physiol. 300, L667–L678
22 Fantidis, P. (2010) The role of intracellular 3 5 -cyclic adenosine monophosphate (cAMP) in atherosclerosis. Curr. Vasc. Pharmacol. 8, 464–472 23 Lorenowicz, M., Fernandez-Borja, M., Kooistra, M., Bos, J. and Hordijk, P. (2008) PKA and Epac1 regulate endothelial integrity and migration through parallel and independent pathways. Eur. J. Cell Biol. 87, 779–792 24 Cullere, X., Shaw, S., Andersson, L., Hirahashi, J., Luscinskas, F. and Mayadas, T. (2005) Regulation of vascular endothelial barrier function by Epac, a cAMP-activated exchange factor for Rap GTPase. Blood 105, 1950–1955 25 Kooistra, M.R., Corada, M., Dejana, E. and Bos, J.L. (2005) Epac1 regulates integrity of endothelial cell junctions through VE-cadherin. FEBS Lett. 579, 4966–4972 26 Birukova, A., Zagranichnaya, T., Fu, P., Alekseeva, E., Chen, W., Jacobson, J. and Birukov, K. (2007) Prostaglandins PGE2 and PGI2 promote endothelial barrier enhancement via PKA- and Epac1/Rap1-dependent Rac activation. Exp. Cell Res. 313, 2504–2520 27 Houslay, M.D. (2010) Underpinning compartmentalised cAMP signalling through targeted cAMP breakdown. Trends Biochem. Sci. 35, 91–100 28 Netherton, S. and Maurice, D. (2005) Vascular endothelial cell cyclic nucleotide phosphodiesterase and regulated cell migration: implication in angiogenesis. Mol. Pharmacol. 67, 263–272 29 Favot, L., Keravis, T., Holl, V., Le Bec, A. and Lugnier, C. (2003) VEGF-induced HUVEC migration and proliferation are decreased by PDE2 and PDE4 inhibitors. Thromb. Haemost. 90, 334–343 30 Zhu, B., Kelly, J., Vemavarapu, L., Thompson, W. and Strada, S. (2004) Activation and induction of cyclic AMP phosphodiesterase (PDE4) in rat pulmonary microvascular endothelial cells. Biochem. Pharmacol. 68, 479–491 31 Thompson, W., Ashikaga, T., Kelly, J., Liu, L., Zhu, B., Vemavarapu, L. and Strada, S. (2002) Regulation of cyclic AMP in rat pulmonary microvascular endothelial cells by rolipram-sensitive cyclic AMP phosphodiesterase (PDE4). Biochem. Pharmacol. 63, 797–807 32 Netherton, S.J., Sutton, J.A., Wilson, L.S., Carter, R.L. and Maurice, D.H. (2007) Both protein kinase A and exchange protein activated by cAMP coordinate adhesion of human vascular endothelial cells. Circ. Res. 101, 768–776 33 Sayner, S., Alexeyev, M., Dessauer, C. and Stevens, T. (2006) Soluble adenylyl cyclase reveals the significance of cAMP compartmentation on pulmonary microvascular endothelial cell barrier. Circ. Res. 98, 675–681 34 Surapisitchat, J., Jeon, K., Yan, C. and Beavo, J. (2007) Differential regulation of endothelial cell permeability by cGMP via phosphodiesterases 2 and 3. Circ. Res. 101, 811–818 35 Sadhu, K., Hensley, K., Florio, V. and Wolda, S. (1999) Differential expression of the cyclic GMP-stimulated phosphodiesterase PDE2A in human venous and capillary endothelial cells. J. Histochem. Cytochem. 47, 895–906 36 Keravis, T., Komas, N. and Lugnier, C. (2000) Cyclic nucleotide hydrolysis in bovine aortic endothelial cells in culture: differential regulation in cobblestone and spindle phenotypes. J. Vasc. Res. 37, 235–249 37 Lugnier, C. and Schini, V. (1990) Characterization of cyclic nucleotide phosphodiesterases from cultured bovine aortic endothelial cells. Biochem. Pharmacol. 39, 75–84 38 Suttorp, N., Ehreiser, P., Hippenstiel, S., Fuhrmann, M., Krull, ¨ M., Tenor, H. and Schudt, C. (1996) Hyperpermeability of pulmonary endothelial monolayer: protective role of phosphodiesterase isoenzymes 3 and 4. Lung 174, 181–194 39 Gloerich, M. and Bos, J.L. (2010) Epac: defining a new mechanism for cAMP action. Annu. Rev. Pharmacol. Toxicol. 50, 355–375 40 Howe, A., Baldor, L. and Hogan, B. (2005) Spatial regulation of the cAMP-dependent protein kinase during chemotactic cell migration. Proc. Natl. Acad. Sci. U.S.A. 102, 14320–14325 41 Howe, A. and Juliano, R. (2000) Regulation of anchorage-dependent signal transduction by protein kinase A and p21-activated kinase. Nat. Cell Biol. 2, 593–600 42 Ruegg, ¨ C., Dormond, O. and Mariotti, A. (2004) Endothelial cell integrins and COX-2: mediators and therapeutic targets of tumor angiogenesis. Biochim. Biophys. Acta 1654, 51–67 43 Sakurai, T., Suzuki, K., Yoshie, M., Hashimoto, K., Tachikawa, E. and Tamura, K. (2011) Stimulation of tube formation mediated through the prostaglandin EP2 receptor in rat luteal endothelial cells. J. Endocrinol. 209, 33–43 44 Jin, H., Garmy-Susini, B., Avraamides, C., Stoletov, K., Klemke, R. and Varner, J. (2010) A PKA–Csk–pp60Src signaling pathway regulates the switch between endothelial cell invasion and cell–cell adhesion during vascular sprouting. Blood 116, 5773–5783 C The
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45 Namkoong, S., Kim, C., Cho, Y., Kim, J., Lee, H., Ha, K., Choe, J., Kim, P., Won, M., Kwon, Y., Shim, E. and Kim, Y. (2009) Forskolin increases angiogenesis through the coordinated cross-talk of PKA-dependent VEGF expression and Epac-mediated PI3K/Akt/eNOS signaling. Cell. Signal. 21, 906–915 46 Breckler, M., Berthouze, M., Laurent, A., Crozatier, B., Morel, E. and Lezoualc’h, F. (2011) Rap-linked cAMP signaling Epac proteins: compartmentation, functioning and disease implications. Cell. Signal. 23, 1257–1266 47 Jyaraj, S., Unger, N. and Chotani, M. (2011) Rap1 GTPases: an emerging role in the cardiovasculature. Life Sci. 88, 645–652 48 Chrzanowska-Wodnicka, M. (2010) Regulation of angiogenesis by a small GTPase Rap1. Vasc. Pharmacol. 53, 1–10 49 Frische, E. and Zwartkruis, F. (2010) Rap1, a mercenary among the Ras-like GTPases. Dev. Biol. 340, 1–9 50 Voigt, P., Dorner, M. and Schaefer, M. (2006) Characterization of p87PIKAP , a novel regulatory subunit of phosphoinositide 3-kinase γ that is highly expressed in heart and interacts with PDE3B. J. Biol. Chem. 281, 9977–9986 51 Patrucco, E., Notte, A., Barberis, L., Selvetella, G., Maffei, A., Brancaccio, M., Marengo, S., Russo, G., Azzolino, O., Rybalkin, S. et al. (2004) PI3Kγ modulates the cardiac response to chronic pressure overload by distinct kinase-dependent and -independent effects. Cell 118, 375–387
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52 Perino, A., Ghigo, A., Ferrero, E., Morello, F., Santulli, G., Baillie, G.S., Damilano, F., Dunlop, A.J., Pawson, C., Walser, R. et al. (2011) Integrating cardiac PIP3 and cAMP signaling through a PKA anchoring function of p110γ . Mol. Cell 42, 84–95 53 Antl, M., von Bruhl, ¨ M.-L., Eiglsperger, C., Werner, M., Konrad, I., Kocher, T., Wilm, M., Hofmann, F., Massberg, S. and Schlossmann, J. (2007) IRAG mediates NO/cGMP-dependent inhibition of platelet aggregation and thrombus formation. Blood 109, 552–559 54 Corbin, J.D., Turko, I.V., Beasley, A. and Francis, S.H. (2000) Phosphorylation of phosphodiesterase-5 by cyclic nucleotide-dependent protein kinase alters its catalytic and allosteric cGMP-binding activities. Eur. J. Biochem. 267, 2760–2767 55 Rybalkin, S.D., Rybalkina, I.G., Feil, R., Hofmann, F. and Beavo, J.A. (2002) Regulation of cGMP-specific phosphodiesterase (PDE5) phosphorylation in smooth muscle cells. J. Biol. Chem. 277, 3310–3317 56 Rybalkin, S.D., Rybalkina, I.G., Shimizu-Albergine, M., Tang, X.-B. and Beavo, J.A. (2003) PDE5 is converted to an activated state upon cGMP binding to the GAF A domain. EMBO J. 22, 469–478
Received 5 December 2013 doi:10.1042/BST20130268
Biochemical Society Transactions (2014) Volume 42, part 2
Cyclic nucleotide phosphodiesterases (PDEs): coincidence detectors acting to spatially and temporally integrate cyclic nucleotide and non-cyclic nucleotide signals Donald H. Maurice*†1 , Lindsay S. Wilson†2 , Sarah N. Rampersad†, Fabien Hubert*, Tammy Truong†, Milosz Kaczmarek*, Paulina Brzezinska*, Silja I. Freitag*, M. Bibiana Umana* and Alie Wudwud†
Biochemical Society Transactions
www.biochemsoctrans.org
*Department of Biomedical and Molecular Sciences, Queen’s University, Kingston, Ontario, Canada, K7L 3N6 †Department of Pathology and Molecular Medicine, Queen’s University, Kingston, Ontario, Canada, K7P 3N6
Abstract The cyclic nucleotide second messengers cAMP and cGMP each affect virtually all cellular processes. Although these hydrophilic small molecules readily diffuse throughout cells, it is remarkable that their ability to activate their multiple intracellular effectors is spatially and temporally selective. Studies have identified a critical role for compartmentation of the enzymes which hydrolyse and metabolically inactivate these second messengers, the PDEs (cyclic nucleotide phosphodiesterases), in this specificity. In the present article, we describe several examples from our work in which compartmentation of selected cAMP- or cGMP-hydrolysing PDEs co-ordinate selective activation of cyclic nucleotide effectors, and, as a result, selectively affect cellular functions. It is our belief that therapeutic strategies aimed at targeting PDEs within these compartments will allow greater selectivity than those directed at inhibiting these enzymes throughout the cells.
Introduction Cellular homoeostasis requires efficient integration of signals encoded by myriad primary messengers, including numerous hormones and transmitters. Inefficient integration of these signals destabilizes critical cellular processes, and this instability can promote disease-initiating maladaptive cellular responses [1]. Many cellular systems have evolved to sense the presence of primary messengers, to decode their signals and, subsequently, to operationalize their intent in cells. These systems are collectively referred to as signal transduction systems. One of the earliest signal transduction systems described allows cells to respond to primary messengerencoded signals by altering the intracellular concentration of a second messenger, cAMP [2]. Subsequently, a cGMP signalling system was also identified and described [2,3]. Both the cAMP- and the cGMP-signalling systems regulate a large number of important physiological processes, including visual transduction, inflammation, apoptosis, cell proliferation and differentiation, as well as critical metabolic pathways such as steroidogenesis, insulin secretion and glycogen synthesis, as well as glycogenolysis, lipogenesis and lipolysis [1–3]. Key words: cAMP, cGMP, cyclic nucleotide phosphodiesterase (PDE), endothelium, platelet. Abbreviations: AC, adenylate cyclase; AJ, adherens junction; AKAP, A-kinase-anchoring protein; ECM, extracellular matrix; Epac, exchange protein directly activated by cAMP; ER, endoplasmic reticulum; HAEC, human arterial endothelial cell; HMVEC, human microvascular endothelial cell; IP3R, InsP3 receptor; IRAG, IP3R-associated cGMP kinase substrate; PDE, cyclic nucleotide phosphodiesterase; PI3K, phosphoinositide 3-kinase; PKA, protein kinase A; PKG, protein kinase G; VEC, vascular endothelial cell; VE-cadherin, vascular endothelial cadherin. 1 2
To whom correspondence should be addressed (email [email protected]). Present address: Pfizer Neuroscience, 610 Main Street, Cambridge, MA 02139, U.S.A.
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Synthesis of cellular cAMP or cGMP is catalysed by both plasma-membrane-associated and cytosolic nucleotidyl cyclases. Indeed, nine distinct plasma membrane ACs (adenylate cyclases) and one cytosolic AC catalyse the synthesis of cAMP from ATP [4]. Similarly, several particulate and cytosolic GCs (guanylate cyclases) convert GTP into cGMP in cells [3]. Since diffusion of ‘free’ cyclic nucleotides in cells is very fast (∼400 μm2 ·s − 1 ) [5,6], significant efforts have focused on delineating mechanisms which allow for spatiotemporal resolution of the cellular actions of cAMP and cGMP. Considerable evidence supports the notion that cyclic nucleotide hydrolysis catalysed by the members of a large multigene family of enzymes called PDEs (cyclic nucleotide phosphodiesterases) actively participate in this spatiotemporal resolution. Informed by data obtained during the sequencing of the human genome, it is now clear that there are 11 different PDE families, that most families have several genes, and that these genes can be differentially expressed and processed in cells [5,7]. In addition to differences in structure, distinct PDEs also have distinct kinetic properties and modes of regulation and can be expressed selectively by cells and differentially localized within cells. Of therapeutic importance, selective therapeutic agents for many of the PDE families are known and are being used in the treatment of many diseases and conditions. If not hydrolysed, cAMP and cGMP can affect cell functions through actions at multiple effectors, including cyclicnucleotide-activated protein kinases [PKA (protein kinase A) and PKG (protein kinase G)], Epacs (exchange proteins directly activated by cAMP) and cyclic-nucleotide-gated Biochem. Soc. Trans. (2014) 42, 250–256; doi:10.1042/BST20130268
Targeting cAMP Signalling to Combat Cardiovascular Diseases
cationic channels [5,7]. The idea that interactions between individual components of the cAMP-signalling system [GPCRs (G-protein-coupled receptors), G-proteins, ACs, PKAs/Epacs/cyclic-nucleotide-gated channels and PDEs] allow the formation of multiple distinct intracellular cAMPsignalling complexes (hereinafter ‘cAMP-signalosomes’) is gaining acceptance [5,6,8]. Although less well developed, a similar concept is proving useful in assessing how cGMP effects in cells are co-ordinated. In our laboratory, we are largely interested in elucidating how compartment-specific cAMP, or cGMP, signalling selectively regulates actions of cells in the cardiovascular system [5,9–15] and build on excellent earlier work in multiple cell types in which cAMPsignalosomes were shown to contain cAMP effectors (PKA or Epac1, or both) and individual PDEs, including PDE3A, PDE3B, PDE4D or PDE7A [7,16]. Continuing advances in our understanding of mechanisms whereby biological and/or therapeutic agents affect cellular functions by altering levels of these cyclic nucleotides are enabling the identification and development of therapeutic agents for use in numerous human diseases [4,6,17–19].
A PDE4D/Epac1/Rap1-containing cAMP-signalosome co-ordinates cAMP effects on VEC (vascular endothelial cell) intercellular contact adhesions and paracellular permeability Numerous cAMP-elevating agents have been reported to reduce VEC permeability to both solutes and cells by strengthening intercellular AJ (adherens junction)-based structures and by promoting the formation of large rings of cortical actin. Studies have shown that activation of either PKA or Epac can co-ordinate these cAMP-dependent effects [20–26]. Since PDE-mediated hydrolysis of cAMP controls the magnitude, duration, and subcellular compartmentation of cAMP [16,19], we and others have addressed their roles in these actions of cAMP [14,21]. Overall, these studies have identified a dominant role for PDE4 over PDE3 and PDE2, the other VEC PDEs. Four genes (PDE4A–PDE4D) yield 20 PDE4 variants, and these can be stratified into long or short forms [5]. Whereas long PDE4s contain N-terminal UCRs (upstream conserved regions) (UCR1/2) and unique targeting domains, short PDE4s do not. Individual PDE4s interact with binding partners, and this often allows their localization within discrete subcellular domains in cells, including VECs [5,14,27]. Although PDE4A, PDE4B and PDE4D variants are each expressed in VECs, most data identify PDE4D as the dominant PDE4 family members involved in controlling these effects of cAMP [11,14,28]. PDE4 inhibitors reduce VEC migration, proliferation and permeability [11,14,28–31]. In early work, we showed that macrovascular and microvascular human VECs could form numerous distinct cAMP-signalosomes, that these cAMP-signalosomes contained either PKA or Epac1, either PDE3B or PDE4D, and that they probably represented the tools through which
cAMP could regulate selectively multiple distinct events in these cells [11,14,28,32]. These data were consistent with an earlier suggestion [33] that cAMP-regulated functions in VECs were co-ordinated within distinct ‘pools’ or compartments. Previously, we uncovered a role for compartmentation of a PDE4D/Epac1-based cAMP-signalosome in regulating HAEC (human arterial endothelial cell) permeability [14] (Figure 1). Moreover, we showed that these actions were coordinated through the integration of this cAMP-signalosome into the VE-cadherin (vascular endothelial cadherin)-based structure known to form VEC AJs. Thus we observed that PDE4 inhibition with rolipram, but not PDE3 inhibition with cilostamide, reduced HAEC permeability. Consistent with a dominant role for Epac1 over PKA in mediating this effect, siRNA-based knockdown of Epac1, but not of PKACα (PKA catalytic subunit α), markedly increased HAEC permeability. Knockdown of individual PDE4 gene products identified PDE4D, but not PDE4A or PDE4B, as dominant in controlling cAMP-mediated activation of Epac1 and in the control of its permeability-reducing effects. Most interestingly, our data defined a role for PDE4D as the link that allowed Epac1 integration into VE-cadherin-based structures. Indeed, although PDE4 inhibition with rolipram decreased VEC permeability, PDE4D knockdown increased permeability and caused a marked reduction in VE-cadherinassociated Epac1. Since PDE4 inhibitors had decreased VEC permeability, we had hypothesized that PDE4D knockdown would similarly reduce HAEC permeability. In contrast, knockdown of PDE4A or PDE4B, or of the dominant HAEC PDE3, namely PDE3B, had little or no effect on VEC permeability. Interestingly, although HAEC VE-cadherinbased isolates also contain PKA, in these cells PDE4D did not locally regulate integration of this cAMP effector or its ability to affect AJ-based adhesions of permeability. How our findings relate to other work of Beavo and colleagues in which cGMP-elevating agents were shown to alter the effect of cAMP-elevating agents on VEC permeability [34] is currently unclear. Although PDE2A, PDE3A and PDE3B are expressed in many human, bovine and porcine arterial or venous VECs used in studies of VEC permeability [21,29,34– 38], these enzymes were not found to integrate into the Epac1/PDE4D-signalosome [14] and as such are unlikely to affect VEC permeability through local events within AJassociated cAMP-signalosomes. Although our data identified Rap1 as the likely downstream actor in promoting actions of this cAMP-signalosome, elucidating the involvement of other agents known to affect VEC permeability such as Cdc42, Rac, PAK (p21-activated kinase), Krit1 (Krev1-interaction trapped gene) or of other PDE4D-interacting proteins, including RACK1 (receptor for activated C-kinase 1), AKAPs (Akinase-anchoring proteins) and β-arrestins or spectrin will require further work [39]. Of potential therapeutic importance, we identified the sequences within Epac1 that promote a direct interaction with PDE4D [14]. We also demonstrated that permeability of HAECs was perturbed when a cell-permeant peptide designed to antagonize this interaction was added to cells. C The
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Figure 1 Examples of cAMP- and cGMP-signalosomes present in cells of the cardiovascular system Proteins identified in the distinct macromolecular complexes are defined in the text.
Thus an Epac1-based peptide designed to interfere with Epac1–PDE4D binding markedly reduced the ability of PDE4D to control Epac1-based effects at the border. These data strongly suggest that small molecules designed to compete for binding between PDE4D and Epac1 might represent powerful agents with which to selectively control VEC barrier functions (Figure 1).
A PDE3B/Epac1/R-Ras signalosome co-ordinates cAMP-mediated VEC adhesions to ECM proteins, migration and tubule formation Many cAMP-elevating agents promote integrin-dependent VEC adhesions to ECM (extracellular matrix) proteins and through this effect can modulate VEC migration [28,29,32]. The relative dependence of this effect on PKA or Epac1 activation is currently unclear. Although PKA activation can promote integrin engagement and stimulate VEC adhesions, PKA activity itself is reported to be either increased or decreased in response to integrin–ECM engagement, and PKA inhibitors do not antagonize fully these cAMPstimulated adhesive events [40,41]. We have concluded that these somewhat contradictory data are most likely reflective of the fact that, whereas each PKA and Epac can regulate integrin engagement, the activity of these effectors are in C The
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turn affected by events downstream and upstream of integrin signalling [32]. Also, the species or vascular structures from which the VECs were derived, the identity of the ECM protein chosen for study and the choice of cAMP-elevating agent each significantly affected the relative involvement of PKA or of Epac in these events. In this context, we reported previously that cAMP-promoted increases in HAECs (macro-VECs) or HMVECs (human microvascular endothelial cells) (micro-VECs) adhesions to distinct ECM proteins were not equally dependent on PKA or Epac1 [32]. Briefly, we showed that, whereas cAMP-induced VEC adhesion was more PKA-dependent in HAECs than in HMVECs, the magnitude of the difference was ECMprotein-dependent. Although most cAMP-elevating agents tested (isoprenaline, forskolin, rolipram or cilostamide) could promote HAEC adhesion when PKA could be activated, in the presence of PKA inhibitors these agents only promoted adhesion of HMVECs, not HAECs. These data identified a functional link between Epac1 and PDE3-catalysed, or PDE4-catalysed, cAMP hydrolysis that was more dominant in HMVECs than in HAECs and suggest that integrin-based cAMP-signalosomes may underpin these differences [32]. Because initiation of angiogenesis involves VEC adhesion, and subsequent migration and tubule formation by these cells, and because cAMP-elevating agents affect each of these events, the ability of certain cAMP-elevating agents to
Targeting cAMP Signalling to Combat Cardiovascular Diseases
affect angiogenesis has been investigated [42–45]. Although PKA, or Epac1, probably co-ordinates these events through numerous separate cAMP-signalosomes, only recently have these events been studied comprehensively. In this context, we identified an Epac1/PDE3B/R-Ras-based signalosome that regulates integrin-based adhesions, migration and tubule formation by VECs in the context of angiogenesis [11] (Figure 1). In this signalosome, PDE3B and Epac1 reciprocally regulated one another’s activities as well as their association with a membrane compartment. We showed that PDE3B bound Epac1 directly via protein–protein interactions and tethered this cAMP effector to cellular membranes. In addition, we showed that PDE3B-catalysed cAMP hydrolysis acted to antagonize Epac1 binding of cAMP within this signalosome, and consequently inhibited Epac1 activation by cAMP. Interestingly, PDE3B association with Epac1 increased PDE3B activity through an as yet unknown mechanism. As described previously, cAMPmediated activation of Epac1 in cells promotes accumulation of GTP-bound/activated Rap1/2 and/or R-Ras proteins [11]. In turn, activated Rap or R-Ras can signal by activating one or many of a group of effectors including PLCε (phospholipase Cε), PLD (phospholipase D), Raf-1, p38MAPK (p38 mitogenactivated protein kinase) and PI3K (phosphoinositide 3kinase) [46–49]. Since a previous study had shown that PDE3B could be isolated with one of the two known PI3Kγ regulatory subunits, namely p84 [50], and a cardiac phenotype was associated with the loss of PI3Kγ catalytic domain (p110γ ) in mice had been attributed to increases in cAMP signalling [51], we investigated whether PI3Kγ was also tethered via PDE3B in VECs and whether this interaction was related to the PDE3B–Epac1 complex. Consistent with the idea that these three activities were integrated into a common cAMP-signalosome, PDE3B was found to interact directly with p84 through a protein–protein interaction distinct from the one that co-ordinated PDE3B–Epac1 binding. Since p84 and Epac1 bound PDE3B via distinct sites, we surmised that PDE3B could simultaneously tether and anchor both Epac1 and the p84-regulated form of PI3Kγ in cells. Consistent with this, PDE3 inhibition with cilostamide and selective PDE3B knockdown in HAECs each promoted Epac1dependent activation of Rap1 and R-Ras, and of PI3Kγ and its downstream activated kinases, in these cells. Indeed, our data clearly showed that this PDE3B-based scaffold allowed for a PDE3B-regulated pool of cAMP to directly promote Epac1dependent activation of PI3Kγ , PKB (protein kinase B) and ERK1/2 (extracellular-signal-regulated kinase 1/2) [11]. At a cell functional level, experiments confirmed that the PDE3B/Epac1/R-Ras/p84-regulated PI3Kγ -signalosome controlled cAMP-mediated HAEC adhesion, migration and tubule formation. Thus, whereas PDE3 inhibition with cilostamide and PDE3B knockdown each promoted HAEC adhesion to ECM proteins, and promoted tubule formation, knockdown of Epac1 and of p84 each abolished tubule formation behaviour in these cells. Most strikingly, all of these events occurred without addition of activators of ACs and, as such, occurred without changing global cAMP levels.
In a recent study, the catalytic domain of PI3Kγ (p110γ ) was identified as a potential AKAP that could tether PKA and PDE3B in murine cardiac myocytes and in cells expressing these proteins heterologously [52]. Since p110γ was not integrated into the PDE3B-based cAMP-signalosome which we identified in VECs when p84 was knocked down, it is currently unclear how this recent finding relates to the system that we describe in the present article. Of potential translational importance, we identified the sequence of PDE3B and two sequences in Epac1 that allowed these proteins to interact directly. On the basis of this information, we designed a cell-permeant PDE3Bbased peptide with which to antagonize binding between these proteins in cells. Consistent with our predictions, the PDE3B-based Epac1-displacing peptide effectively competed the binding between PDE3B and Epac1, promoted Epac1/RRas-dependent activation of PI3Kγ in these cells and stimulated the rate at which tubules formed on MatrigelTM . Most strikingly, the internal consistency of these data to our model was observed when the displacing peptide rendered cells insensitive to the effects of PDE3B inhibition. Together, the observations detailed in the two previous sections validate the concept that selective regulation of individual cAMP-sensitive cellular events can be achieved with reagents, or drugs, able to selectively disturb the formation of individual cAMP-signalosomes in cells.
PDE5A is resident in an ER (endoplasmic reticulum)-based cGMP-signalling complex On a return flight from a conference in Glasgow, during which my head was full of ideas concerning PDE-based signalling compartments (and whisky!), it struck me that compartments might not be needed in cells that expressed only one PDE, or only one effector. After all, if signalosomes were designed to allow enzyme-based selectivity, surely they would be redundant in a cell in which there was no choice. In contrast, if compartments were critical in allowing selective cell responses, independent of the enzyme involved, one would predict that they would be found in cells irrespective of the number of tools to which that cell had access. Because I was unaware of a cell type in which only one cAMPPDE, or one cAMP effector, was expressed, Lindsay Wilson and I turned to the platelet to investigate these ideas. On the basis of my previous work with the late Dr Richard Haslam, I ‘knew’ that PDE5A accounted for >90% of cGMP-PDE activity in platelets and that only PKG served as a cGMP effector in these cells. As a result of Lindsay’s observation that sildenafil could inhibit thrombin-induced Ca2 + transients without increasing global intraplatelet cGMP, we tested the hypothesis that PDE5 might be acting locally at the platelet ER [15]. More precisely, we determined whether PDE5 formed a functional part of the IP3R1 (InsP3 receptor 1)–IRAG (IP3R-associated cGMP kinase substrate)–PKG1β signalling complex which had been reported previously to allow cGMPdependent control of thrombin-induced Ca2 + transients in C The
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platelets [53]. Differential centrifugation of platelet lysates identified PKG1β (∼25%), PDE5 (∼45%) and IP3R1 (∼100%) in particulate cellular fractions, and selective immunoprecipitation of PKG1β, PDE5 or IP3R1 allowed their co-immunoprecipitation, a finding consistent with the idea that PDE5 was integral to the IP3R1–IRAG–PKG1β intraplatelet complex. Interestingly, this complex was only influenced by cGMP, as PKA inhibition did not affect the sildenafil-induced inhibition of Ca2 + transients and PKA and the cAMP PDE (PDE3A) were not co-immunoprecipitated with the cGMP signalling proteins. In addition to being able to co-immunoprecipitate these proteins, we also discovered that PDE5A could be differentially regulated depending on its cellular localization. We began to investigate the two pools of PDE5A present in platelets, one particulate and PKGassociated pool of PDE5 and another cytosolic or ‘bulk’ PDE5A which was not associated with PKG. Interestingly, we discovered that these two pools of PDE5A had different intrinsic activities. Whereas the non-PKG-associated pool had a higher specific activity, the PKG-associated PDE5A had a very low specific activity. Since previous work had indicated that PDE5 could be activated upon either cGMP binding to a PDE5 GAF-A domain or PKG phosphorylation or PDE5 at Ser102 [54–56], we investigated these two pools further to determine whether they could be differentially regulated. Uniquely, only the PKG1B-associated PDE5A could be selectively phosphorylated by PKG at Ser102 , whereas phosphorylation of the non-PKG-associated pool was not observed. In addition, this phosphorylation resulted in a significant activation of PDE5A, again, a result not observed in the cytosolic and non-PKG-associated pool of enzyme. We also observed that the PKG-associated pool of PDE5 could not be activated by cGMP alone in the absence of ATP, adding to our findings that these two pools were regulated differently. Together, these studies demonstrate that a single PDE isoform, namely PDE5A, can be differentially regulated depending on its binding partners and localization in the cell. Although previous work indicated that PDE5 was activated upon either cGMP binding to a PDE5 GAFA domain, or PKG phosphorylation of PDE5 at Ser102 [54–56], these earlier studies were silent on the relative importance of these mechanisms in cells. To address this issue, we compared the phosphorylation and activation of the PKG-associated and non-PKG-associated forms of PDE5 in 8-bromo-cGMP (1 mM, 15 min)-treated platelets. 8-Bromo-cGMP treatment of platelets markedly increased the Ser102 phosphorylation status and activity of the PKGassociated form of PDE5, but not that which was not associated with the kinase. Consistent with the idea that the phosphorylated PDE5 was resident within the IP3R1– IRAG–PKG1β complex, IP3R1 was recovered in the antiPKG immune complexes, but not in those representing bulk PDE5. Similarly, when anti-IP3R1 immune complexes were obtained from control or 8-bromo-cGMP-treated platelets, only the IP3R1-associated PDE5 was activated by 8-bromo-cGMP. An identical pattern of PDE5 activation C The
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was obtained when PDE5 was isolated using the method used originally to isolate and characterize the IP3R1–IRAG– PKG1β complex in platelets, namely cGMP–agarose [53]. Taken together, these data were consistent with the idea that only PKG-associated PDE5 was subject to PKG-catalysed phosphorylation and activation in cells treated with 8-bromocGMP. In addition to being selectively activated by PKG in cells, our data indicate that the IP3R1–IRAG–PKG1βassociated PDE5 had a lower specific activity than that isolated from the bulk cytosol. Indeed, the IP3R1–IRAG– PKG1β-associated PDE5 was only approximately 14% as active as that isolated in anti-PDE5 immunoprecipitates. Interestingly, our data are inconsistent with the idea that PDE5 and PKG form a stable complex in platelet cellular fractions devoid of IP3R1. Indeed, although PKG and PDE5 were each efficiently individually immunoprecipitated from platelet cytosolic fractions, these proteins never coimmunoprecipitated from this fraction. Because PDE5, PKG or IP3R1 was not enriched in IP3R1-containing subcellular fractions in cells incubated with a cGMP analogue, or with an NO (nitric oxide) donor, our data were also inconsistent with the idea that cGMP elevation triggers movement of these proteins between these fractions. Taken together, we believe that these results are consistent with the idea that PDEs and cyclic nucleotide effectors are compartmented in cells, independent of the number of such proteins available, resulting in efficient regulation and integration of intracellular signals. The examples highlighted in the present review support the notion that PDE-containing signalosomes are formed to regulate selected cellular processes and do not simply exist to isolate similar enzymes from each other. In the future, and of therapeutic importance, investigation into signalosomes and protein interaction partners will help us in identifying potential new drug targets. PDEs are favourable targets, and, although multiple family-specific inhibitors exist, the ability to distinguish between individual isoforms of PDEs is challenging. Targeting individual PDEs contained in signalosomes provides an opportunity to achieve a desired signal while avoiding another pool of this enzyme where potential side effects could interfere. Determining the protein interaction partners and protein-interface residues can only inform on the differences between pools of PDEs and allow us to use this information to make small molecules that can inhibit interactions or target specific nodes in signalling pathways. Since small-molecule compounds targeting global signalling pathways have the potential for unwanted toxicity and side effects, finer more precise therapeutics that target intracellular pools have the potential to be the next class of drugs to potentially benefit multiple disease areas.
Funding Funding for the studies described were provided by the Canadian Institutes of Health Sciences (CIHR).
Targeting cAMP Signalling to Combat Cardiovascular Diseases
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Received 5 December 2013 doi:10.1042/BST20130268
