Expression of Epac1 and Epac2
Epac1 (cAMP-GEF-1) and Epac2 (cAMP-GEF-II) are expressed in both mature and developing tissues. Recent studies have indicated that alterations in the cellular microenvironment are present in chronic degenerative inflammatory diseases and seem to affect the expression profile of Epac1 and Epac2. Initially, Epac1 mRNA expression was reported to be most abundant in the heart and kidney, although subsequently, it was found to be expressed in all human tissues being analysed (de Rooij et al., 1998). Additionally, expression of Epac1 has been found in monocytes, macrophages, B and T lymphocytes, eosinophils, neutrophils, platelets, and in CD34-positive haematopoietic cells (Tiwari et al., 2004; Bryn et al., 2006; Gerlo et al., 2006; Lorenowicz et al., 2006). Recently, developmental changes of Epac1 mRNA expression have been reported in the heart, vasculature, brain, kidneys and lungs (Ulucan et al., 2007; Yokoyama et al. 2008a,b; Murray and Shewan, 2008). Epac2 mRNA expression has been found to be prominent in the brain and adrenal glands (Kawasaki et al., 1998), whereas Epac2 was undetectable in all hematopoietic cell types being studied (Tiwari et al., 2004). It has also been reported that Epac2 mRNA expression in the heart, vasculature, brain, kidneys and lungs is subject to developmental changes (Ulucan et al., 2007; Murray and Shewan, 2008; Yokoyama et al., 2008a,b). Due to the recent identification of a novel splice variant of Epac2, designated Epac2B, the previously identified Epac2 has been renamed Epac2A; it has been reported that Epac2A mRNA is expressed in pancreatic islets and cerebral cortex, whereas Epac2B mRNA expression is restricted to adrenal glands (Niimura et al., 2009).
There are distinct expression patterns of Epac1 and Epac2 proteins in rat brain, spinal cord and dorsal root ganglia at embryonic, neonatal and adult stages of development. Interestingly, it has been shown that the expression of Epac1 declines in adulthood, whereas Epac2 expression is dramatically up-regulated in adults. Such developmental regulation of Epac expression seems to promote axon growth and regeneration of the nervous system (Murray and Shewan, 2008). Expression of Epac1 mRNA, but not Epac2 mRNA, was found to be transiently increased in a mouse model of vascular injury, and up-regulation of Epac1 protein expression correlated with the progression of neointimal thickening (Yokoyama et al., 2008a). Although the expression of Epac1 and Epac2 mRNAs was up-regulated in rat ductus arteriosus during the perinatal period, Epac1 activity correlated with the intimal cushion formation (Yokoyama et al., 2008b). Studies on developmental changes of Epac1 and Epac2 mRNA expression have shown alterations of Epac1 and Epac2 in the heart, brain, kidneys and lungs of mice at different stages of development from foetus into adulthood. Interestingly, relative to Epac1, Epac2 became dominant in the adult brain and heart compared with fetal organs, whereas Epac1, relative to Epac2, became dominant in the adult kidney and lung compared with fetal organs (Ulucan et al., 2007), suggesting that Epac1 and Epac2 differentially contribute to fetal and adult organ function. In mycocardial hypertrophy induced by chronic catecholamine infusion, Epac1 and Epac2 mRNA were up-regulated, whereas only Epac1 mRNA was increased in pressure overload-induced hypertrophy (Ulucan et al., 2007). At present, it is not known whether Epac is a cause of the development of cardiac hypertrophy, or if its expression is a consequence of hypertrophy. Transforming growth factor β1 (TGF-β1) has been shown to reduce the expression of Epac1 mRNA in rat cardiac, lung and skin fibroblasts, and a decrease in Epac1 mRNA expression paralleled the increase in expression of TGF-β1 in a rat myocardial infarction model. Such a process may promote the synthesis of extracellular matrix at site of injury (Yokoyama et al., 2008c). The reduction of Epac1 mRNA expression in U937 monocytic cells by TGF-β1 might protect against aberrant transendothelial migration of leukocytes during inflammation (Basoni et al., 2005). Although the underlying molecular mechanisms have yet to be addressed, attenuation of Epac1 expression by (fibrogenic) agonists may represent a mode of adaptation of cAMP responses to pathophysiological alterations present in inflammatory diseases.
Localization of Epac1 and Epac2
Epac1 is localized in the plasma membrane, cytoplasm, perinuclear regions, nuclear membrane and mitochondria in various cell types, including HEK293, N1E-115, CHO, COS1(7), HeLa, PC12, PCCL3, primary hippocampal (cortical) neurons, peritoneal macrophages, rat cholangiocytes, primary rat neonatal ventriculocytes, and different tubular segmental cells of rat and human kidney (Qiao et al., 2002; DiPilato et al., 2004; Nikolaev et al., 2004; Ponsioen et al., 2004; Dodge-Kafka et al. 2005; Morel et al., 2005; Borland et al., 2006; Hochbaum et al., 2008; Li et al., 2008; Métrich et al., 2008; Di Benedetto et al. 2008; Banales et al., 2009). In particular, studies in HEK293 and COS1(7) cells demonstrated that subcellular (re)distribution of Epac1 is subject to cell cycle- and cytoskeleton-dependent dynamics (Qiao et al., 2002; Borland et al., 2006; Huston et al., 2008). Epac2 is localized to the (sub)plasma membrane, cytosolic fractions, actin cytoskeleton, meiotic midzone region and Golgi in several cells, such as MIN6, pancreatic islets, H1299, human microvascular endothelial cells, different tubular segmental cells of rat (human) kidney, rat cholangiocytes, adult rat ventricular myocytes and mouse spermatocytes (Ozaki et al., 2000; Hong et al., 2007; Shibasaki et al., 2007; Leroy et al., 2008; Li et al., 2008; Aivatiadou et al., 2009; Banales et al., 2009; Niimura et al., 2009). Clearly, Epac1 and Epac2 are localized to a number of subcellular regions, and their cell type-specific co-localization at the plasma membrane and cytoplasm indicates that Epacs might act in concert to modulate cellular responses.
Meanwhile, recent studies have reported on spatial and temporal regulation of cellular Epac signalling by using Epac-based fluorescence resonance energy transfer cAMP sensors or green fluorescent protein (GFP)/Flag-tagged Epacs (DiPilato et al., 2004; Nikolaev et al., 2004; Ponsioen et al., 2004, 2009; Di Benedetto et al., 2008; Leroy et al., 2008; Liu et al., 2008; Shafer et al., 2008). The Epac-based cAMP sensor studies were important for the concept that the concentration of cellular cAMP rises to a level sufficient to activate Epac1 and Epac2. Initially, Nikolaev and colleagues demonstrated, by using single-chain cAMP sensors based on the cAMP-binding domains of Epac and PKA, that β-adrenoceptor-induced cAMP signals are rapidly propagated throughout the entire cell body of primary hippocampal neurons and peritoneal macrophages (Nikolaev et al., 2004). Using Epac as an indicator of cAMP (either targeted to plasma membrane, mitchondria or nucleus), DiPilato and co-workers demonstrated differential dynamics of cAMP signaling in response to β-adrenoceptor or prostanoid receptor activation in HEK293, HeLa and in PC12 cells (DiPilato et al., 2004). In contrast, Ponsioen et al. showed, by using a CFP-Epac-YFP construct, that Epac activation was not limited to membranes, but rather occurs throughout the cell (A431, HEK293, N1E-115 and MCF-7 cells (Ponsioen et al., 2004). More recently, Ponsioen and colleagues reported, by using GFP-tagged Epac1, that activation of β- adrenoceptors induced a rapid translocation of Epac1 to the plasma membrane in A431, HEK293, OVCAR-3, ACHN, RCC10, MDCK, N1E-115, HeLa, Rat-1, GE11 and H1299 cells (Ponsioen et al., 2009). Similarly, Liu et al. demonstrated, by using Flag-tagged Epac2, that Epac2 activation requires compartmentalization of Epac2 to Ras-containing membranes, a process that operates in COS, HEK293 and PC12 cells (Liu et al., 2008). Although the concept of subcellular (re)distribution of Epac as a prerequisite for their activation remains controversial, Epac1 and Epac2 clearly represent novel cAMP sensors that contribute to the highly dynamic features of cAMP signalling.
Because the binding affinity of cAMP for PKA and for Epac has been found to be very similar (kd∼2.9 µM), it has been proposed that Epac and/or PKA are activated in response to moderate increases of cellular cAMP, and that such activation depends upon cellular compartmentalization of cAMP formation and effector protein availability (Dao et al., 2006). Indeed, studies in primary rat neonatal ventriculocytes have identified distinct intracellular cAMP signalling compartments composed of distinct PKA subtypes, Gs-coupled receptors and cAMP-hydrolyzing PDEs (Di Benedetto et al., 2008). Recently, Leroy and colleagues reported on spatio-temporal dynamics of β-adrenoceptor-induced cAMP signalling in adult rat ventricular myocytes, and speculated that PDE3 regulates a ‘constitutive’ cAMP pool linked to cardiac contractility, whereas PDE4 regulates the cAMP microdomains driven by β-adrenoceptor stimulation (Leroy et al., 2008). Intriguingly, very recent research suggests that different mechanisms contribute to the plasma membrane targeting of Epac1 and Epac2 (Liu et al., 2008; Niimura et al., 2009; Ponsioen et al., 2009). Thus, an increasing weight of experimental data infers that cellular compartmentalization of cAMP signalling affects the net outcome of biological functions. Spatio-temporal cAMP signalling is believed to involve members of the AKAP family (Wong and Scott, 2004), and indeed cAMP-responsive multiprotein complexes, including Epac1 and Epac2, have been identified in the heart and neurones (Dodge-Kafka et al., 2005; Nijholt et al., 2008) (Figure 4). In neonatal rat cardiomyocytes, Dodge-Kafka and colleagues identified a cAMP-responsive multiprotein complex composed of nuclear envelope-associated mAKAP, PKA, PDE4D3 and Epac1 (Dodge-Kafka et al., 2005). This cardiac-specific multiprotein complex was sensitive to differential cellular cAMP concentrations. Thus, at high cAMP concentrations, cardiac hypertrophy was attenuated upon Epac1-Rap1-dependent inhibition of extracellular signal-regulated kinase5 (ERK5) and subsequent activation of PDE4D3, whereas at low cAMP concentrations, cardiac hypertrophy was enhanced upon ERK5-mediated inactivation of PDE4D3 and subsequent increased PKA signalling (Dodge-Kafka et al., 2005) (Figure 4). Recent studies in our laboratory reported a novel link between neuronal plasma membrane-associated AKAP79/150, PKA, Epac2 and phosphatidylinositol 3-kinase (PI 3-kinase)-dependent protein kinase B (PKB)/Akt (Nijholt et al., 2008) (Figure 4). Direct activation of PKA or Epac2 complexed to AKAP79/150 exerted opposing effects on neuronal PKB/Akt: direct activation of PKA inhibited PKB/Akt phosphorylation, whereas direct activation of Epac2 enhanced PKB/Akt phosphorylation (Nijholt et al., 2008). At present, it is not known whether distinct PDEs are also tethered to the neuronal AKAP79/150 complex, and the input of distinct receptor-driven cAMP alterations has yet to be determined. Of particular interest, a recent study by Raymond and co-workers demonstrated the presence of distinct PKA- and Epac-based signalling complexes comprised of several PDEs and AKAPs (Raymond et al., 2007). Because co-localization of PKA and Epacs to distinct AKAPs correlates with opposing biological effects in cardiomyocytes and neurones (Figure 4), the capacity of AKAPs to interact with additional signalling components might be crucial (Wong and Scott, 2004; Beene and Scott, 2008). Clearly, further studies are required to analyze AKAP-dependent cellular cAMP compartmentalization and its effect on cAMP-dependent biological functions.
Structure of Epac1 and Epac2
Initial studies indicated that Epac1 and Epac2 consist of an auto-inhibitory N-terminal regulatory region that contains a DEP (dishevelled, Egl-10, pleckstrin) domain responsible for membrane association and a high-affinity cAMP-binding domain (cAMP-B), a C-terminal catalytic region bearing a CDC25 homology domain (CDC25HD) that exhibits GEF activity for Ras-like GTPases, a Ras-exchange motif (REM) domain believed to stabilize the GEF domain, and a Ras-associating (RA) domain present in several Ras-interacting proteins. Indeed, Epac2 has been shown to interact with GTP-bound Ras (Li et al., 2006) (Figure 1). In addition, Liu et al. demonstrated that the interaction of Epac2 with Ras via its RA domain is required to redirect Epac2 to Ras-containing membranes and to subsequently induce cAMP-dependent activation of Rap proteins (Liu et al., 2008). The authors proposed that coincident detection of both cAMP and Ras signals is required for Epac2 to activate Rap1 in a temporally and spatially controlled manner (Liu et al., 2008). Similarly, Ponsioen and colleagues recently characterized the molecular mechanisms underlying the direct spatial control of Epac1 by cAMP (Ponsioen et al., 2009). It has been reported that cAMP is required to release Epac1 from auto-inhibitory restraints, and that cAMP induces translocation of Epac1 to the plasma membrane, a process dependent on the DEP domain of Epac1 (Ponsioen et al., 2009). Until very recently, it was believed that the second low-affinity cAMP-A domain of Epac2 exerted an as-yet undetermined biological function. However, Niimura and co-workers identified a novel splice variant of Epac2, designated Epac2B, and the initially identified Epac2 has now been renamed as Epac2A (Niimura et al., 2009). Expression of Epac2A mRNA is restricted to pancreatic islets and cerebral cortex, and it has been shown that Epac2A-driven insulin secretion from pancreatic cells requires the cAMP-A domain, independent of its cAMP-binding capacity, to localize Epac2A near the plasma membrane (Niimura et al., 2009). Clearly, translocation of Epac1, Epac2A and Epac2B to the plasma membrane is driven by rather different mechanisms, and it will be of interest to analyse the impact of AKAP proteins in these processes. Recent X-ray crystallography of Epac2 and NMR spectroscopy of Epac1 have provided novel insights into the dynamic equilibrium of critical conformational switches encompassing a closed, auto-inhibited state in which the N-terminal regulatory region sterically blocks the C-terminal catalytic region to a completely different, open and catalytically active state (Rehmann et al., 2006, 2008; Mazhab-Jafari et al., 2007; Das et al., 2008; Harper et al., 2008). It is to be hoped that development of Epac-specific cAMP analogues will arise as a result of these types of studies.
Methods to validate cAMP signalling via PKA and Epac
Membrane-permeable cyclic nucleotide analogues have been synthesized, and these pharmacological tools seem to differentiate between PKA and Epac signalling (Table 1). For example, N6-benzyladenosine-3′,5′-cyclic monophosphate (6-Bnz-cAMP), 8-(4-chlorophenylthio)-adenosine-3′,5′-cyclic monophosphorothioate, Rp-isomer (Rp-8-CPT-cAMPS), 8-(4-chlorophenylthio)-2′-O-methyl-cAMP (8-pCPT-2′-O-Me-cAMP), acetoxymethyl 8-pCPT-2′-O-Me-cAMP (8-pCPT-2′-O-Me-cAMP-AM), 8-(4-chlorophenylthio)-2′-O-methyladenosine-3′,5′-cyclic monophosphorothioate, Sp-isomer (Sp-8-pCPT-2′-O-Me-cAMPS) and 8-(4-chlorophenylthio)-2′-O-methyl-cGMP (8-pCPT-2′-O-Me-cGMP) are now used to activate and/or inhibit PKA and Epac, and 8-pCPT-2′-O-Me-cGMP is used as a negative control for 8-pCPT-2″-O-Me-cAMP (Enserink et al., 2002; Christensen et al. 2003; Holz et al. 2006, 2007; Vliem et al. 2008; Haag et al. 2008) (Table 1). A 2′-O-methyl substitution on the ribose ring of cAMP of Epac-selective cAMP analogues confers specificity towards Epac, and such compounds are therefore used as pharmacological tools to analyse Epac-driven cAMP dynamics independent of PKA. However, currently available Epac activators do not differentiate between Epac1 and Epac2 (Table 1). As Epac1 and Epac2 are insensitive to inhibitors of PKA, such as Rp-8-CPT-cAMPS (Holz et al., 2007) (Table 1), PKA inhibitors are used to demonstrate that Epac-specific analogues act via Epac independently of PKA. Studies in Trypanosoma brucei indicated, however, that 8-pCPT-2′-O-Me-cAMP might act via its 5′-AMP derivative upon inhibition of PDEs (Laxman et al., 2006). In addition, recent studies in human platelets showed that various cyclic nucleotide analogues, including 6-Bnz-cAMP and 8-pCPT-2′-O-Me-cAMP might, in addition to their primary effects, also cause elevation of cAMP or cGMP upon inhibition of phosphodiesterases (Poppe et al., 2008). Altogether, these studies suggest caution when interpreting data obtained with cyclic nucleotide analogues of dubious selectivity, especially when such agents are used to dissect PKA-dependent and -independent versus Epac-dependent and -independent signalling properties. At present, highly specific pharmacological inhibitors of individual Epac isoforms, Epac1 and Epac2, are not available (Holz et al., 2007; Poppe et al. 2008). However, the successful suppression of the endogenous expression of Epac1 and Epac2 by specific siRNAs has been reported (López de Jesús et al., 2006; Haag et al. 2008, Huang et al. 2008; Yokoyama et al. 2008a,c). Recently, Seino et al. developed Epac2-deficient mice and demonstrated that Epac2/Rap1 signalling is essential in the regulation of insulin granule dynamics (Shibasaki et al., 2007). Generation of additional Epac knockout mice and/or Epac reporter mice might help to gain further insights into Epac-related signalling.
Table 1. Cyclic nucleotide compounds
|N6-benzyladenosine-3′,5′-cyclic monophosphate (6-Bnz-cAMP), selective and membrane-permeable protein kinase A (PKA) activator||Christensen, Selheim et al., 2003|
|Poppe et al., 2008|
|8-(4-chlorophenylthio)-adenosine-3′,5′-cyclic monophosphorothioate, Rp-isomer (Rp-8-CPT-cAMPS), competetive inhibitor of PKA||Christensen, Selheim et al., 2003|
|Poppe et al., 2008|
|8-(4-chlorophenylthio)-2′-O-methyl-cAMP (8-pCPT-2′-O-Me-cAMP), selective and membrane-permeable activator of Epac1 and Epac2||Enserink et al., 2002|
|Christensen, Selheim et al., 2003|
|Poppe et al., 2008|
|Acetoxymethyl 8-pCPT-2′-O-Me-cAMP (8-pCPT-2′-O-Me-cAMP-AM), see above||Vliem et al., 2008|
|8-(4-chlorophenylthio)-2′-O-methyladenosine-3′,5′-cyclic monophosphorothioate, Sp-isomer (Sp-8-pCPT-2′-O-Me-cAMPS), selective and membrane-permeable Epac activator, insensitive to phosphodiesterases||Laxman et al., 2006|
|Poppe et al., 2008|
|8-(4-chlorophenylthio)-2′-O-methyl-cGMP (8-pCPT-2′-O-Me-cGMP), negative control for 8-pCPT-2′-O-Me-cAMP ||Haag et al., 2008|
Epac effectors and biological functions
Epac1 and Epac2 were initially characterized as cAMP-activated GEFs for Rap1 and Rap2 (de Rooij et al., 1998; Kawasaki et al. 1998). Recent studies have indicated that Epac proteins also function as a molecular link between different Ras family members (Maillet et al., 2003; Krugmann et al., 2004; Morel et al., 2005; López de Jesús et al., 2006; Li et al. 2006; Métrich et al., 2008). Furthermore, our studies demonstrated that Epac1 binds to and activates R-Ras, a process that is likely to facilitate cytoskeleton dynamics and calcium handling driven by Epac (López de Jesús et al., 2006). Over the last 10 years, considerable progress has been made into the Epac-related cAMP dynamics associated with learning and memory (Gekel and Neher, 2008; Gelinas et al., 2008; Murray and Shewan, 2008; Ouyang et al., 2008), inflammation, fibrosis, and hypertrophy (Holz et al., 2006, 2007; Schmidt et al., 2007a; Roscioni et al., 2008; Schaafsma et al., 2008; Borland et al., 2009b). Epac1 and Epac2 seem to control these distinct cellular responses by signalling to a range of effectors. The number and diversity of these effectors is considerable. For example, phospholipase C-ε (Schmidt et al., 2001; Oestreich et al., 2007), phospholipase D (López de Jesús et al., 2006; Han et al., 2007) and ERK1/2 (Lin et al., 2003; Keiper et al., 2004; Kiermayer et al., 2005; Wang et al., 2006) have been implicated in the regulation of hypertrophic responses. ERKs (Ster et al., 2007, 2009; Gelinas et al., 2008; Ma et al., 2009) and PI 3-kinase-dependent PKB/Akt (Mei et al., 2002; Misra and Pizzo, 2005; Yano et al. 2007; Misra et al. 2008; Kwak et al., 2008; Nijholt et al. 2008) seem to modulate learning and memory. TGF-β1 receptor-regulated Smads are important for fibrogenic and inflammatory responses (Basoni et al., 2005; Conrotto et al., 2006; Yokoyama et al., 2008c). Finally, NF-κB (Fuld et al., 2005; Scheibner et al., 2008), the suppressor of cytokine signalling-3 (SOCS-3) (Sands et al., 2006; Yarwood et al., 2008; Borland et al., 2009a), the CCAAT/enhancer-binding protein C/EBP and glycogen synthase kinase-3 (GSK-3) (Jing et al., 2004; Xu et al., 2008) (Figure 1 and Figure 3), have been reported to regulate inflammation. Further details on the molecular mechanisms of the regulation of Epac-related effectors and their biological functions are outlined in recent excellent reviews on these topics (Cohen and Frame, 2001; Scheid and Woodgett, 2001; Kolch, 2005; Bunney and Katan, 2006; Oude Weernink et al., 2007; Perkins, 2007; Schmierer and Hill, 2007; Yoshimura et al., 2007; Borland et al., 2009b). Despite the novel insights into the molecular mechanisms linking Epac to a diversity of downstream effectors, it still remains to be determined whether co-localization of Epac to PKA-AKAP-based signalling complexes are of central importance to the net outcome of cAMP signalling.