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Spatial Control of Epac2 Activity by cAMP and Ca2+-Mediated Activation of Ras in Pancreatic β Cells

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Science Signaling  30 Apr 2013:
Vol. 6, Issue 273, pp. ra29
DOI: 10.1126/scisignal.2003932


The cAMP (adenosine 3′,5′-monophosphate)–activated guanine nucleotide exchange factor (GEF) Epac2 is an important mediator of cAMP-dependent processes in multiple cell types. We used real-time confocal and total internal reflection fluorescence microscopy to examine the spatiotemporal regulation of Epac2, which is a GEF for the guanosine triphosphatase (GTPase) Rap. We demonstrated that increases in the concentration of cAMP triggered the translocation of Epac2 from the cytoplasm to the plasma membrane in insulin-secreting β cells. Glucose-induced oscillations of the submembrane concentration of cAMP were associated with cyclic translocation of Epac2, and this translocation could be amplified by increases in the cytoplasmic Ca2+ concentration. Analyses of Epac2 mutants identified the high-affinity cAMP-binding and the Ras association domains as crucial for the translocation. Expression of a dominant-negative Ras mutant reduced Epac2 translocation, and Ca2+-dependent oscillations in Ras activity synchronized with Epac2 translocation in single β cells. The cyclic translocation of Epac2 was accompanied by oscillations of Rap GTPase activity at the plasma membrane, and expression of an inactive Rap1B mutant decreased insulin secretion. Thus, Epac2 localization is dynamically controlled by cAMP as well as by Ca2+-mediated activation of Ras. These results help to explain how oscillating signals can produce pulses of insulin release from pancreatic β cells.


The exchange proteins directly activated by cAMP (adenosine 3′,5′-monophosphate), Epac1 and Epac2, also known as the cAMP-activated guanine nucleotide exchange factors cAMP-GEFI and cAMP-GEFII, are mediators of cAMP-regulated cellular processes (1). These proteins regulate neurotransmitter and hormone release (26), cell adhesion and junction formation (710), cell proliferation (11), gene transcription (12), and intracellular Ca2+ handling (13). Epac1 and Epac2 are multidomain proteins with an N-terminal regulatory region and a C-terminal catalytic region with GEF activity toward the small Ras family guanosine triphosphatases (GTPases) Rap1 and Rap2 (1). The regulatory region consists of a DEP (disheveled, Egl-10, pleckstrin) domain and one or two cyclic nucleotide binding (CNB) domains. The catalytic region contains a Ras exchange motif (REM), a Ras association (RA) domain, and a CDC25 homology domain (1). In the absence of cAMP, the Epac proteins have a compact structure in which the regulatory region blocks the catalytic region, keeping the protein in an autoinhibited state. Binding of cAMP to the CNB domain releases the autoinhibition (1416). Whereas Epac1 has one CNB domain, Epac2 has two with low and high affinities for cAMP. The high-affinity domain binds micromolar cAMP, a considerably higher concentration range than that necessary to activate cAMP-dependent protein kinase (PKA) (17, 18). The function of the low-affinity CNB domain is unclear, but it may be required for appropriate subcellular targeting of Epac2 (19).

The Epac targets Rap1 and Rap2 localize to various membrane compartments, including the Golgi network, the nuclear membrane, the vesicular membranes, and the plasma membrane (2022). Activation of a specific pool of these proteins might be achieved by subcellular targeting of their upstream GEFs. Epac1 and Epac2 have been found both at the plasma membrane and at various organelle membranes (12, 2327). The mechanisms regulating the subcellular distribution of Epac proteins are not well understood. It has been reported that increases in the cAMP concentration trigger Epac1 translocation to the plasma membrane in a process that requires cAMP binding as well as an intact DEP domain (27), which mediates binding to phosphatidic acid (28). In addition, the cAMP-induced plasma membrane translocation of Epac1 is increased by binding of the N terminus of Epac1 to activated ezrin-radixin-moesin (ERM) proteins (29). Epac2 differs from Epac1 in that its RA domain can bind to Ras–GTP (guanosine 5′-triphosphate). Overexpression of Ras in H1299 lung carcinoma cells thus targets Epac2 to the plasma membrane (25), and the membrane association of Epac2 is lost if the RA domain is deleted or mutated (25, 26). Whether binding of cAMP is required for membrane targeting of Epac2 is a matter of debate. The conformational change induced by cAMP binding is not necessary for exposing the RA domain (16), and although Epac2 has been suggested to target to membranes by cAMP-independent interaction with Ras-GTP (26), others have failed to observe membrane association of Epac2 in the absence of cAMP (25).

Whereas Epac1 is present in most cells, Epac2 is primarily present in neuronal tissue, adrenal glands (30), and pancreatic β cells (31). In β cells, glucose triggers oscillations of the sub–plasma membrane Ca2+ and cAMP concentrations that synergize to generate pulsatile insulin secretion (32). Epac2 mediates PKA-independent amplification of insulin secretion by cAMP during stimulation with glucose (33) and the incretin hormone glucagon-like peptide-1 (GLP-1) (3). Using quantitative real-time imaging with confocal and total internal reflection fluorescence (TIRF) microscopy, we have investigated how the subcellular distribution of Epac2 is controlled by dynamic changes of the sub–plasma membrane concentrations of cAMP and Ca2+ as well as of endogenous Ras activity. We found that Epac2 cyclically translocated to and from the plasma membrane in response to cAMP and Ca2+ signals. The Ca2+-dependent translocation was mediated by increased Ras activity and required a concomitant increase in the cAMP concentration. The periodic spatial redistribution of Epac2 was translated into oscillations of Rap GTPase activity in the plasma membrane compartment.


cAMP induces translocation of Epac2 to the plasma membrane

Confocal imaging of MIN6 β cells immunostained for Epac2 showed that an increase in cAMP concentration induced by the phosphodiesterase inhibitor IBMX (3-isobutyl-1-methylxanthine) caused increased localization of endogenous Epac2 at the plasma membrane (Fig. 1A). To study the kinetics of this translocation, we transfected MIN6 β cells with Epac2 with an N-terminal GFP (green fluorescent protein) fusion (GFP-Epac2). Under basal conditions, GFP-Epac2 distributed evenly in the cytoplasm, but an IBMX-induced increase in the cAMP concentration caused reduced cytoplasmic fluorescence due to increased localization of the construct to the plasma membrane (Fig. 1A). This translocation was more conspicuous when analyzed with TIRF microscopy, which selectively visualizes fluorescent molecules near the plasma membrane (Fig. 1, A and B). Similar translocations were observed when the cells were exposed to the membrane-permeable cAMP analog 8-Br-cAMP or the Epac-selective cAMP analog 8-pCPT-2′-O-Me-cAMP-AM (“007-AM”; Fig. 1B). Inhibition of PKA with Rp-8-CPT-cAMPS did not affect the response to IBMX (fig. S1A). The translocation was readily reversible, and repeated increases of the cAMP concentration through pulsatile application of IBMX caused oscillatory redistribution of GFP-Epac2 to and from the plasma membrane (Fig. 1C). To minimize the potential influence of fluorescence changes that may occur as a result of alterations in cell adhesion or cell volume, we expressed a membrane-targeted red fluorescent protein (td2-CAAX) together with GFP-Epac2 and used the GFP/td2 (tdimer2) fluorescence ratio as readout for Epac2 localization (Fig. 1C). The ability of cAMP to trigger Epac2 translocation was verified in α-toxin–permeabilized cells, where the nucleotide induced dose-dependent plasma membrane association of GFP-Epac2 (Fig. 1, D and E). Also, 007 caused rapid and reversible membrane translocation of Epac2 in this preparation (Fig. 1F).

Fig. 1 cAMP-induced translocation of Epac2 to the plasma membrane.

(A) Confocal microscopy images showing endogenous Epac2 (left column) and GFP-Epac2 (middle column) distribution in a MIN6 β cell under basal conditions and after exposure to IBMX. Translocation results in 14 ± 2% decrease of cytoplasmic GFP-Epac2 fluorescence (fluor.) (P < 0.001; n = 5 cells). Right column shows TIRF microscopy images of GFP-Epac2. Images are representative of 36 and 10 (upper and lower left, respectively), 5 (middle), and 45 cells (right column). Scale bars, 10 μm. (B) Real-time TIRF intensity recordings of GFP-Epac2 from single MIN6 β cells exposed to IBMX (black trace, representative of 45 cells; average fluorescence increase: 26 ± 1%, P < 0.001), 8-Br-cAMP (magenta trace, representative of 41 cells; fluorescence increase: 10 ± 1%, P < 0.001), or 007-AM (blue trace, representative of 42 cells; fluorescence increase: 13 ± 1%, P < 0.001). (C) TIRF intensity recording of GFP-Epac2 (blue trace) and td2-CAAX (magenta trace) as well as the GFP/td2 fluorescence ratio (black trace) during intermittent IBMX application (I, shaded areas). Representative of six cells. (D) TIRF microscopy recording of GFP-Epac2 fluorescence in an α-toxin–permeabilized MIN6 β cell exposed to cAMP. Representative of 41 cells exposed to increasing concentrations of cAMP. (E) Dose dependence of cAMP-induced GFP-Epac2 membrane translocation in an α-toxin–permeabilized MIN6 β cells as shown in (D). Means ± SE for 8 to 41 cells at each concentration. Half-maximal and maximal effects were obtained at 28 and 200 μM, respectively. (F) TIRF recording from an α-toxin–permeabilized MIN6 β cell showing mCherry-Epac2 membrane translocation after addition of 007. Representative of five cells.

Epac2 translocates to the membrane in an oscillatory manner in glucose-stimulated β cells

We next tested if physiological stimulation influences the localization of Epac2. Glucose is the main stimulus for insulin secretion. Exocytosis of insulin granules from β cells is triggered by glucose-induced increase in the cytoplasmic Ca2+ concentration and further amplified by increases in the cAMP concentration (32, 33). An increase in the glucose concentration from 3 to 11 mM triggered membrane translocation of GFP-Epac2 after a 2- to 4-min delay (Fig. 2A). The translocation pattern varied between cells. Most of the cells responded with oscillations with frequencies of <0.3 min−1 (Fig. 2A), but some cells showed transient plasma membrane association (fig. S1B), stable association (fig. S1C), or faster oscillations (>0.3 min−1; fig. S1D). In most cells, IBMX amplified the translocation induced by glucose, indicating that the glucose response was not saturated (fig. S1, B and C). Similarly, augmenting the cAMP concentration by the addition of GLP-1 to glucose-stimulated β cells triggered further GFP-Epac2 translocation, often associated with pronounced oscillations (fig. S1E). Confocal imaging of immunostained cells showed that the cAMP-increasing stimuli also triggered translocation of endogenous Epac2 (fig. S1F). The glucose-induced translocation of fluorescent protein–tagged Epac2 was not limited to the clonal MIN6 β cells, but was also observed in primary mouse β cells within intact pancreatic islets (Fig. 2B). Parallel measurements of the sub–plasma membrane cAMP concentration with a translocation biosensor (34) in MIN6 β cells showed that glucose-induced Epac2 translocation was preceded by an increase in the cAMP concentration (Fig. 2C). Blocking cAMP formation with 2′,5′-dideoxyadenosine (DDA) or SQ 22,536 reduced the glucose-induced Epac2 translocation (Fig. 2, D and E).

Fig. 2 Glucose induces oscillatory Epac2 membrane translocation that depends on cAMP and Ca2+.

(A) TIRF microscopy recording of the GFP-Epac2/td2-CAAX fluorescence ratio from an individual MIN6 β cell during glucose stimulation. Average ratio increase, 15 ± 1% (P < 0.001). Representative of 35 of 60 cells with glucose-induced Epac2 translocation. (B) Epac2 translocation in a primary cell within an intact islet stimulated by an increase in glucose concentration. Representative of 15 of 21 cells from five islets. (C) Simultaneous recordings of mCherry-Epac2 (black trace) and sub–plasma membrane cAMP concentration (blue trace) from a MIN6 β cell during glucose stimulation. The cAMP trace is presented to show concentration increases as upward deflections. cAMP increases 33 ± 8 s (P < 0.001) before Epac2 starts to translocate. Representative of 29 cells. (D) Inhibition of glucose-induced Epac2 translocation by DDA. Representative of 16 cells. (E) Means ± SE for the effect of DDA, SQ 22,536, and Ca2+-deficient medium on the time-average GFP-Epac2 fluorescence (average intensity during exposure to test condition) normalized to control (11 mM glucose; n = 16 to 23 cells). *P < 0.05, **P < 0.01 compared to control (Student’s paired t test). (F) Simultaneous recordings of GFP-Epac2 (black trace) and sub–plasma membrane Ca2+ concentration (magenta trace) from an individual MIN6 β cell during glucose stimulation. The Ca2+ trace is presented to show concentration increases as upward deflections. Ca2+ increases precede Epac2 translocation by 11 ± 2 s (P < 0.001). Representative of 40 cells. (G) Inhibition of glucose-induced Epac2 translocation by omission of extracellular Ca2+. Representative of 23 cells.

Ca2+ regulates cAMP-induced Epac2 translocation

Because glucose induces oscillations in the cytoplasmic Ca2+ concentration in β cells, we investigated the relationship between Ca2+ and Epac2 localization. Simultaneous TIRF microscopy recordings of the sub–plasma membrane Ca2+ concentration and GFP-Epac2 revealed that the glucose-induced GFP-Epac2 translocation was preceded by an increase in the Ca2+ concentration (Fig. 2F). Removal of extracellular Ca2+ and addition of EGTA decreased the GFP-Epac2 association with the plasma membrane (Fig. 2, E and G). Although Ca2+ removal did not completely prevent the glucose-induced GFP-Epac2 translocation, it always transformed oscillatory translocations into stable membrane association. These data demonstrate that membrane association of Epac2 is determined by both cAMP and Ca2+.

To clarify how Ca2+ influences Epac2 translocation, we depolarized MIN6 cells with a high concentration of K+, which resulted in immediate translocation of Epac2 to the plasma membrane (fig. S2A). However, the response was suppressed after adenylyl cyclase inhibition with DDA (fig. S2, A to C). Thus, an increase in the Ca2+ concentration without concomitant formation of cAMP is a poor stimulator of Epac2 translocation.

We next investigated cAMP-dependent Epac2 translocation under conditions in which increases in the cytoplasmic Ca2+ concentration were prevented by depleting Ca2+ from the intracellular stores with the sarco-endoplasmic reticulum Ca2+ adenosine triphosphatase (SERCA) inhibitor cyclopiazonic acid (CPA) and by maintaining the cells in Ca2+-deficient medium supplemented with EGTA. Treatment with a combination of IBMX and the adenylyl cyclase activator forskolin caused distinct instances of GFP-Epac2 translocation in Ca2+-deficient medium, but the amplitude and rate of increase were suppressed compared to control (Fig. 3, A and B). The reduced translocation cannot be explained by a lower intracellular cAMP concentration because 8-Br-cAMP–induced translocation of GFP-Epac2 was also suppressed in Ca2+-deficient medium (fig. S2, D and E). We directly evaluated the ability of IBMX or 8-Br-cAMP to increase the intracellular cAMP concentration in MIN6 β cells transfected with the cAMP translocation biosensor. Although the rate of IBMX-induced increase of cAMP concentration was slightly reduced in the absence of Ca2+, there was no difference in the steady-state sub–plasma membrane concentration of cAMP (fig. S2F). After depletion of intracellular Ca2+ stores with CPA, the rate of the IBMX-induced increase in the cAMP concentration was further decreased, but the steady-state concentration of cAMP beneath the membrane induced by IBMX or 8-Br-cAMP was not reduced (fig. S2, F to I). Together, these results indicate that Ca2+ has a permissive effect on cAMP-induced Epac2 translocation.

Fig. 3 Ca2+ affects cAMP-induced Epac2 membrane association.

(A) TIRF microscopy of Epac2 translocation (GFP/td2 ratio) after an increase in the intracellular cAMP concentration with a combination of IBMX and forskolin (Fsk) under control conditions and when increases of cytoplasmic Ca2+ are prevented by removal of extracellular Ca2+ and addition of EGTA and by depletion of intracellular Ca2+ stores with CPA. Representative of 26 cells. (B) Means ± SE of the peak GFP/td2 ratio and the time to half-maximal translocation in the experiments in (A). n = 26 cells; **P < 0.01, ***P < 0.001 for the differences from control (Student’s paired t test). (C) Effect of membrane depolarization on GFP-Epac2 membrane association triggered by 8-Br-cAMP. Representative of 28 of 37 cells. Average response, 49 ± 14% increase in fluorescence (P < 0.01); n = 37 cells. (D) Ca2+ potentiates cAMP-induced GFP-Epac2 plasma membrane translocation in an α-toxin–permeabilized MIN6 β cell. (E) Means ± SE for the effect of Ca2+ on cAMP-induced GFP-Epac2 plasma membrane translocation in permeabilized MIN6 β cells as shown in (D). Data are expressed in relation to the translocation observed at 0.1 μM Ca2+ (n = 19 cells); **P < 0.01 for difference from the condition with 0.1 μM Ca2+ (Student’s unpaired t test). (F) Ca2+ amplification of cAMP-induced GFP-Epac2 translocation in an α-toxin–permeabilized cell depends on the presence of GTP. (G) Means ± SE for the effect of Ca2+ on cAMP-induced GFP-Epac2 translocation in α-toxin–permeabilized cells in the presence or absence of 250 μM GTP as shown in (F). n = 11 cells; **P < 0.01 for difference between groups (Student’s unpaired t test).

In most cells exposed to 8-Br-cAMP, depolarization with K+ caused an increase in GFP-Epac2 plasma membrane fluorescence (Fig. 3C). The remaining cells instead showed a reduction in fluorescence, but the average response from all cells was positive. Similar results were obtained with cells depolarized in the presence of forskolin and IBMX (fig. S3A). Moreover, a stepwise increase in the intracellular buffer Ca2+ concentration in permeabilized cells dose-dependently amplified the cAMP-induced plasma membrane translocation of Epac2 (Fig. 3, D and E). This amplification required the presence of GTP in the buffer (Fig. 3, F and G). Like in intact cells, a fraction of the permeabilized cells showed dissociation of Epac2 from the membrane when exposed to a sudden increase in Ca2+ concentration (fig. S3B). These data suggest that increases in the cytoplasmic Ca2+ concentration may exert dual effects: predominantly an amplification of Epac2 translocation with a less pronounced inhibition of the Epac2 membrane interaction.

Epac2 membrane translocation requires binding to cAMP and Ras

Previous studies suggest that binding of Epac2 to the plasma membrane requires intact CNB (25) and RA (25, 26) domains. To determine their roles for the translocation of Epac2, we used mutants lacking the entire CNB region (GFP-Epac2-ΔCNB) or containing single amino acid substitutions in either the low-affinity cAMP-binding domain (mCherry-Epac2G114E), the high-affinity cAMP-binding domain (mCherry-Epac2G422D), or both (mCherry-Epac2G114E/G422D) (2), as well as in the Ras binding site (mCherry-Epac2K684E) (26). Although the absence of CNB domains resulted in enhanced association with the membrane (Fig. 4A), the cAMP-binding– and Ras-binding–deficient mutants were predominantly localized in the cytoplasm even after an IBMX-induced increase in the cAMP concentration (Fig. 4, A to F), except for the mCherry-Epac2G114E mutant, which showed translocation similar to control (Fig. 4, C and F). We verified the unaltered cAMP-induced membrane translocation of Epac2G114E and the impaired translocation of the other Epac2 mutants in α-toxin–permeabilized cells (Fig. 4, G to K). Glucose induced membrane translocation of the mCherry-Epac2G114E/G422D mutant in intact MIN6 β cells, but to a substantially reduced extent compared to wild-type control (Fig. 4L). Similarly, the glucose-induced membrane translocation of the Ras-binding–deficient mCherry-Epac2K684E mutant in intact cells was suppressed compared to wild type and did not oscillate (Fig. 4M). Consistent with the requirement for Ras binding, overexpression of a dominant-negative Ras mutant (RasS17N) suppressed the glucose-induced plasma membrane translocation of wild-type GFP-Epac2 (Fig. 4N).

Fig. 4 Dynamic Epac2 localization requires intact high-affinity CNB and RA domains.

(A) Confocal images of MIN6 β cells showing the subcellular distribution of different fluorescent protein–tagged Epac2 mutants. WT, GFP-tagged wild-type Epac2; ΔRA, GFP-Epac2 lacking the RA domain; ΔCNB, GFP-Epac2 lacking both CNB domains and the intervening sequence; G114E/G422D, mCherry-tagged Epac2 with G114E/G422D substitutions in the cAMP-binding domains; K684E, mCherry-Epac2 with a K684E mutation in the RA domain. Each image is representative of 27 to 42 cells from three independent experiments. (B to E) TIRF microscopy recordings of the IBMX-induced translocation of wild-type GFP-Epac2 (black traces) and coexpressed mutated versions of mCherry-Epac2 (colored traces) in single MIN6 β cells. (F) Means ± SE of the IBMX-induced translocation of wild-type and mutant Epac2 as shown in (B) to (E). n = 11 to 32 cells in each group. ***P < 0.001 for difference from wild type (Student’s paired t test). (G to J) TIRF microscopy recordings from individual α-toxin–permeabilized MIN6 β cells coexpressing wild-type GFP-Epac2 (black traces) and mCherry-tagged Epac2 mutants (colored traces) and exposed to increasing concentrations of cAMP. (K) Dose dependence of cAMP-induced translocation of wild-type and mutated Epac2 in α-toxin–permeabilized MIN6 β cells from experiments in (G) to (J). n = 28 to 79 cells at each concentration for WT, 9 cells for G114E, 18 cells for G422D, 11 cells for G114E/G422D, and 6 to 14 cells for K684E. (L) Means ± SE of the glucose-induced translocation of GFP-tagged wild-type Epac2 (black trace) and mCherry-tagged Epac2 with both cAMP-binding sites mutated (G114E/G422D; green trace) recorded from 68 individual MIN6 β cells. (M) TIRF microscopy recordings from an intact MIN6 β cell expressing wild-type GFP-Epac2 (black trace) and mCherry-Epac2K684E (orange trace) during glucose stimulation. Representative of 12 cells. (N) Means ± SE of glucose-induced GFP-Epac2 plasma membrane translocation in control cells and cells expressing RasS17N. n = 32 to 33; **P < 0.01 for difference from control (Student’s paired t test).

Ca2+-mediated Ras activation triggers translocation of Epac2 to the plasma membrane

To further investigate the role of Ras in Epac2 translocation, we monitored active (GTP-loaded) Ras using TIRF microscopy and a biosensor consisting of the Ras-binding domain from Raf1 fused to cyan fluorescent protein (CFP-Raf1RBD) (35). An increase in the glucose concentration induced translocation of CFP-Raf1RBD to the plasma membrane without changing the td2-CAAX reference signal, indicating activation of Ras in the plasma membrane compartment (Fig. 5A). Simultaneous measurements revealed that the oscillations of Epac2 translocation induced by glucose or a combination of glucose and GLP-1 were paralleled by oscillations of CFP-Raf1RBD without a significant time lag (Fig. 5, B and C). These effects on Ras activity are probably driven by changes of the sub–plasma membrane Ca2+ concentration. Accordingly, increases in the Ca2+ concentration generated by repeated K+ depolarizations in MIN6 β cells exposed to 007-AM consistently induced synchronized membrane recruitment of CFP-Raf1RBD and mCherry-Epac2 that depended on the presence of Ca2+ in the extracellular medium (Fig. 5, D and E). Moreover, after treatment with 007-AM and the Ca2+ ionophore ionomycin, both mCherry-Epac2 and CFP-Raf1RBD translocated to the plasma membrane in response to increasing concentrations of Ca2+ (Fig. 5, F and G). Together, these findings indicate that Epac2 membrane translocation is controlled by Ca2+-mediated activation of Ras.

Fig. 5 Ca2+-stimulated Ras activity controls Epac2 plasma membrane dynamics.

(A) TIRF microscopy recording from a single MIN6 β cell expressing the Ras activity reporter CFP-Raf1RBD and td2-CAAX during glucose stimulation. Average CFP-Raf1RBD fluorescence increase, 28 ± 2% (P < 0.001); representative of 47 cells. The lower black trace shows the CFP/td2 fluorescence ratio. (B) Simultaneous TIRF microcopy recording of CFP-Raf1RBD (blue trace) and mCherry-Epac2 (black trace) fluorescence upon glucose stimulation of a MIN6 β cell. Representative of 40 cells. Time difference between Epac2 and CFP-Raf1RBD translocation: 2 ± 3 s. (C) Simultaneous TIRF microscopy recordings of CFP-Raf1RBD (blue trace) and mCherry-Epac2 (black trace) during stimulation with a combination of glucose and GLP-1. Representative of 25 cells. (D) TIRF microscopy recording of mCherry-Epac2 (black trace) and CFP-Raf1RBD (blue trace) fluorescence from a single MIN6 β cell during periodic increases of the sub–plasma membrane Ca2+ concentration induced by intermittent K+ depolarizations when Epac2 is maximally activated by 007-AM. Representative of 31 cells. (E) Simultaneous TIRF microscopy recordings of CFP-Raf1RBD (blue trace) and mCherry-Epac2 (black trace) after K+ depolarization and temporary removal of extracellular Ca2+ from the medium. Representative of 12 cells. (F) Simultaneous TIRF microscopy recordings of CFP-Raf1RBD (blue trace) and mCherry-Epac2 (black trace) in an individual MIN6 β cell exposed to 007-AM and ionomycin in buffer containing increasing Ca2+ concentrations. Representative of 20 cells. (G) Ca2+ concentration dependence of the translocation of CFP-Raf1RBD (blue) and mCherry-Epac2 (black) from experiments as in (F). Means ± SE for the translocation expressed in percentage of that observed at 10 μM Ca2+. n = 20 cells at each concentration.

Epac2 translocation activates Rap1 at the plasma membrane

To investigate if Epac2 translocation was associated with activation of effector proteins at the plasma membrane, we monitored the activity of Rap GTPases using a fluorescent protein–tagged Rap-binding domain from RalGDS, which shows a high selectivity for GTP-bound Rap over the closely related Ras (36). An increase of the glucose concentration induced oscillatory translocation of the GFP-RalGDSRBD in both MIN6 and primary mouse β cells, indicating periodic activation of Rap GTPases at the plasma membrane (Fig. 6, A and B). Activation of Epac with 007-AM or a combination of forskolin and IBMX induced stable translocation of the Rap reporter, although with smaller magnitude compared to glucose (Fig. 6, C and D). Subsequent depolarization with K+, which caused translocation of Epac2 to the membrane (Fig. 5, D and E), induced further GFP-RalGDSRBD translocation, indicating that Rap activity is indeed controlled by the subcellular localization of Epac2 (Fig. 6, C and E). The results from the optical assay were confirmed in a traditional Rap-GTP pull-down assay (Fig. 6F). Consistent with the involvement of Ras in Epac2 activation, expression of the dominant-negative RasS17N mutant reduced glucose-induced Rap activity as determined with GFP-RalGDSRBD (Fig. 6G). To evaluate the involvement of Rap in insulin secretion, we monitored insulin secretion kinetics from individual MIN6 β cells using an assay based on detection of phosphatidylinositol-3,4,5-trisphosphate (PIP3) after autocrine activation of insulin receptors (32, 37). Control MIN6 β cells expressing CFP and a yellow fluorescent protein (YFP)–tagged PIP3-binding PH domain from Akt responded to glucose stimulation or K+ depolarization with translocation of the biosensor to the plasma membrane, which reflects insulin secretion. The magnitude of this translocation was significantly reduced in cells expressing a CFP-tagged dominant-negative mutant of Rap1 (Rap1BS17N) (Fig. 6, H to J).

Fig. 6 Epac2 translocation leads to Rap activation at the plasma membrane.

(A and B) TIRF microscopy recording showing glucose stimulation causes oscillatory translocation of GFP-RalGDSRBD to and from the plasma membrane in a single MIN6 β cell (A) and a cell within an intact mouse pancreatic islet (B). Representative of 16 cells (A) and 29 of 46 cells in nine islets (B). (C) Means ± SE for the GFP-RalGDSRBD translocation in MIN6 β cells stimulated with glucose (n = 44 cells), 007-AM (n = 26 cells), forskolin with IBMX (n = 50 cells), or forskolin/IBMX combined with K+ depolarization (n = 16 cells). (D) TIRF microscopy recording of GFP-RalGDSRBD (green trace) and td2-CAAX (magenta trace) fluorescence upon stimulation by 007-AM. The black trace shows the GFP/td2 fluorescence ratio. Representative of 31 cells. (E) TIRF microscopy recording of GFP-RalGDSRBD fluorescence from a single MIN6 β cell during periodic increases of the sub–plasma membrane Ca2+ concentration induced by intermittent increases in K+ under conditions of maximal Epac2 activation by forskolin and IBMX. Representative of 23 cells. (F) Western blot showing GTP-loaded and total amount of Rap1 in MIN6 β cell lysates. The bar diagram shows means ± SE of the densitometric quantification of the data. n = 4 experiments. (G). Means ± SE for the glucose-induced Rap activity in cells expressing or not a dominant-negative RasS17N mutant. *P < 0.05 for difference from control (Student’s unpaired t test.). n = 44 cells for control and 47 for RasS17N. (H) Single-cell recordings of insulin secretion reflected as plasma membrane PIP3 concentration in cells expressing the PHAkt-YFP biosensor combined with a dominant-negative CFP-tagged Rap1BS17N mutant (magenta trace) or CFP alone (black trace) as control. Representative of 52 and 30 cells, respectively. (I) Means ± SE for the glucose-induced insulin secretion quantified as the time-average PHAkt-YFP fluorescence in control cells (n = 31) and cells expressing the Rap1B mutant S17N (n = 57) as shown in (H). **P < 0.01 for difference from control (Student’s unpaired t test). (J) Means ± SE for the K+-induced insulin secretion quantified as the amplitude of the PHAkt-YFP fluorescence in control cells (n = 30) and in cells expressing Rap1BS17N (n = 52). *P < 0.05 for difference from control (Student’s unpaired t test).


Spatial compartmentalization of signaling is an important mechanism for the specific regulation of distinct cellular functions. The present study demonstrates that the subcellular localization of Epac2 is dynamically controlled by cAMP as well as by Ca2+-mediated Ras activity. Epac2 undergoes glucose- and hormone-induced oscillatory translocation between the cytoplasm and the plasma membrane in insulin-secreting β cells, and this translocation is associated with activation of Rap in the plasma membrane compartment (Fig. 7). In addition, both the high-affinity CNB domain and the RA domain are required for the dynamic redistribution of the protein.

Fig. 7 Proposed model of the regulation of Epac2 localization by cAMP and Ca2+.

cAMP generated by adenylyl cyclases (AC) binds to one of the two cAMP binding domains of Epac2. The protein interacts with GTP-loaded Ras through its RA domain at the plasma membrane. This translocation of Epac2 is positively regulated by Ca2+, probably by activation of Ras through Ca2+-dependent activation of Ras-GEF proteins. By an unknown mechanism, Ca2+ also prevents Epac2 membrane association. At the plasma membrane, Epac2 stimulates GTP/GDP (guanosine diphosphate) exchange of the Rap family of small GTPases.

The intracellular concentration of cAMP is the most important determinant of Epac2 translocation. The effects of agents increasing the cAMP concentration were mimicked by an Epac-selective agonist and were resistant to PKA inhibition, indicating that the translocation is directly driven by the activation of Epac and not by other cAMP-regulated processes. Half-maximal translocation was observed at 28 μM cAMP in permeabilized cells, which is similar to the 46 μM reported for the activation of Epac protein in vitro (14). Studies of the Epac2 structure indicate that cAMP activates the protein by releasing autoinhibition of catalytic activity exerted by the regulatory region (15, 16). However, the mechanism by which Epac2 localizes to the plasma membrane is unclear. Various observations indicate that the membrane association of Epac2 relies on the interaction between its RA domain and Ras (25, 26), but consensus is lacking regarding the involvement of cAMP in this process. Whereas overexpression of an active Ras mutant was sufficient for Epac2 membrane localization in one study (26), other investigators failed to observe membrane binding in the absence of cAMP (25). It has been proposed that the N-terminal, low-affinity CNB domain is crucial for membrane association by a mechanism that does not depend on its cAMP-binding capacity (19). The present data indicate that Epac2 resembles Epac1 in its dynamic translocation in response to cAMP signals. In addition to cAMP binding, the translocation required a functional RA domain. The deletion mutant lacking the CNB domains was localized to the membrane, indicating that the N-terminal region of the protein is not required for membrane interaction. On the basis of the observation that Epac2 with mutations that prevent either cAMP binding or interaction with Ras showed cytoplasmic distribution with little tendency to translocate, we conclude both that cAMP-induced translocation involves interaction with Ras and that Epac2 interaction with endogenous Ras protein cannot occur in the absence of cAMP. Because Epac2 does not activate Ras (30, 38), cAMP promotes the interaction between Epac2 and Ras that is active under basal conditions.

In addition to the control by cAMP, Epac2 translocation was regulated in a complex manner by Ca2+. The ability of cAMP to induce Epac2 translocation was suppressed in Ca2+-deficient media. Although the rate of cAMP formation was lower under these conditions, inhibition of Epac2 translocation could not be explained by a reduced sub–plasma membrane concentration of cAMP. The cAMP-induced translocation of Epac2 was similarly suppressed when Ca2+ was omitted from the intracellular-like buffer in permeabilized MIN6 cells. Increases in the cytoplasmic Ca2+ concentration typically promoted Epac2 translocation, but in some cells, the ion instead seemed to counteract the Epac2-membrane interaction. Epac2 does not have a Ca2+ binding domain, and the ion probably affects translocation by indirect mechanisms. Because Epac2 membrane association depended on Ras, whose activity is regulated by Ca2+-sensitive GEFs and GTPase-activating proteins (3941), we propose that Ca2+ controls Epac2 localization by changes in Ras activity. The observation that Ca2+ did not affect Epac2 localization in permeabilized cells unless GTP was present in the intracellular medium supports the involvement of a G protein (heterotrimeric guanine nucleotide–binding protein). Moreover, using a Raf kinase–based activity reporter for Ras (36), we found that glucose- and depolarization-induced Ca2+ increases lead to activation of Ras at the plasma membrane with kinetics similar to the translocation of Epac2. It can therefore be concluded that at least the stimulatory effect of Ca2+ on Epac2 translocation is mediated by Ras. It is unclear why Ca2+ sometimes promoted dissociation of Epac2 from the membrane. This effect probably involves mechanisms other than modulation of Ras because Ras activity was invariably increased by K+ depolarization.

Glucose triggers pulsatile insulin secretion by generating coordinated oscillations of the sub–plasma membrane concentrations of Ca2+ and cAMP in β cells (32). These oscillations can explain the periodic Epac2 translocation observed in glucose- and GLP-1–stimulated cells. Epac2 is important for mediating the PKA-independent stimulatory actions of cAMP on insulin secretion (2, 42). The mechanisms involve alterations of the membrane potential, Ca2+ signaling, secretory granule priming, and exocytosis through activation of Rap1 (6) and its effectors, including phospholipase C-ε (4345), as well as through interactions with the SUR1 subunit of the KATP channel (42) and proteins involved in the exocytosis machinery, such as Rim2, Piccolo (46), and SNAP25 (47). Because these are membrane-associated proteins, it seems appropriate that Epac2 translocates to the plasma membrane upon activation. Ras and Rap are closely related GTPases. Whereas Rap1 has been implicated in cell adhesion, junction formation, and exocytosis (6, 48), Ras promotes cell proliferation and survival (4850) and has not been previously implicated in the regulation of insulin secretion. The present findings indicate that activation of Ras in glucose- and hormone-stimulated β cells is involved in mediating the effect of cAMP on insulin secretion by recruiting Epac2 to the plasma membrane. Epac2 subsequently activates Rap1 and other potential effectors at sites of exocytosis.

Protein translocation is an efficient means to achieve spatial control of signaling processes (51), and oscillations in signaling systems, in particular of Ca2+, contribute to the efficiency and specificity in the regulation of downstream events (52). cAMP oscillations have previously been suggested to contribute to specificity by spatially restricting PKA activity (34), and cAMP oscillations are translated into oscillations of PKA activity with the oscillatory frequency positively correlating with nuclear PKA activity and CREB (cAMP response element–binding protein) phosphorylation (53). We now demonstrate that cAMP oscillations can be decoded also by Epac2. Our present findings show that Epac2 signaling is under dynamic spatiotemporal control of both cAMP and Ca2+. Coordinated oscillations of the two messengers provide a distinct signal for precise temporal regulation of the plasma membrane pool of Epac2 effectors.

Materials and Methods


Reagents of analytical grade and deionized water were used. IBMX, EGTA, forskolin, DDA, 9-(tetrahydro-2-furanyl)-9H-purin-6-amine (also known as SQ 22,536), and ionomycin were from Sigma. The Epac agonist 8-pCPT-2′-O-Me-cAMP (007) and its acetoxymethyl ester derivative 007-AM as well as the protein kinase A inhibitor Rp-8-CPT-cAMPS were from Biolog Life Science Institute. The acetoxymethyl ester of the Ca2+ indicators Fura Red and Fluo-4, Dulbecco’s modified Eagle’s medium, Opti-MEM, and Lipofectamine 2000 were obtained from Invitrogen. A plasmid encoding mouse Epac2 fused to GFP (GFP-Epac2) was obtained from D. Altschuler (University of Pennsylvania, Philadelphia, PA). mCherry-Epac2 was generated by polymerase chain reaction amplification of mCherry with Age I and BsrG I restriction sites. Point mutations were generated by site-directed mutagenesis (QuikChange II XL Site-Directed Mutagenesis Kit, Agilent Technologies) according to the manufacturer’s instructions. Epac2 lacking both CNB domains and the intervening sequence (GFP-Epac2ΔCNB), and Epac2 lacking the RA domain (GFP-Epac2ΔRA) were gifts from L. Quilliam (Indiana University School of Medicine, Indianapolis, IN). The red fluorescent protein td2 was provided by R. Y. Tsien (University of California, San Diego, CA), and a version anchored to the plasma membrane (td2-CAAX) was used as an independent reference signal in ratiometric recordings. CFP-Raf1RBD, which was used to detect the localization of Ras-GTP, was provided by M. Fivaz (Duke Graduate Medical School Singapore, Singapore), and GFP-RalGDSRBD for localization of GTP-bound Rap as well as the pGST-RalGDS-RBD plasmid for expression of GST-RalGDSRBD in bacteria were gifts from H. Rehmann and J. L. Bos (University Medical Centre Utrecht, Netherlands). Adenoviral vectors expressing mCherry-Epac2, GFP-RalGDSRBD, and RasS17N were generated by Vector BioLabs.

Cell culture and transfection

MIN6 β cells (54) of passages 17 to 30 were maintained in Dulbecco’s modified Eagle’s medium containing glucose (4.5 g/liter) and supplemented with 15% fetal calf serum, 2 mM l-glutamine, 50 μM β-mercaptoethanol, penicillin (100 U/ml), and streptomycin (100 μg/ml), at 37°C in a 5% CO2 humidified atmosphere. For transfection, ~2 × 105 cells were suspended in 100 μl of Opti-MEM medium containing 0.5 to 1 μg of Lipofectamine 2000 (Invitrogen) and 0.15 to 0.4 μg of plasmid DNA and plated on 25-mm poly-l-lysine–coated coverslips. Where indicated, the plasmid DNA transfection was followed by infection of the cells with 10–30 plaque-forming units per cell of adenovirus encoding RasS17N. After 3 to 5 hours, when the cells were firmly attached, the transfection was terminated by the addition of 3 ml of complete cell culture medium, in which cells were maintained for an additional 18 to 36 hours.

Pancreatic islets were isolated with collagenase from the pancreases of 5- to 7-month-old female C57Bl/6J mice. The Uppsala ethical committee approved all animal experimental procedures. The islets were transferred to RPMI 1640 culture medium containing 5.5 mM glucose and supplemented with 10% fetal calf serum, penicillin (100 U/ml), and streptomycin (100 μg/ml) for culture for 1 to 2 days at 37°C in a 5% CO2 humidified air atmosphere. The islets were infected with mCherry-Epac2– or GFP-RalGDSRBD–expressing adenoviruses at a concentration of 107 plaque-forming units per islet in 250 to 500 μl for 2 hours, followed either by washing with normal RPMI 1640 medium and further culture for 16 to 20 hours before use (GFP-RalGDSRBD) or by addition of 2.5 ml of medium (without washing) and overnight culture (mCherry-Epac2).

Buffers and permeabilization

Before imaging experiments, the cells or islets were transferred to a buffer containing 125 mM NaCl, 4.8 mM KCl, 1.3 mM CaCl2, 1.2 mM MgCl2, 3 mM glucose, and 25 mM Hepes (pH adjusted to 7.40 with NaOH), and incubated for 40 to 60 min at 37°C. Continued experiments were made with the coverslips mounted in an open 50-μl chamber with buffer superfused at a rate of 0.3 ml/min. If not otherwise stated, test substances were used at the following concentrations: 11 or 20 mM glucose, 10 nM GLP-1, 10 μM forskolin, 100 μM IBMX (50 μM when combined with forskolin), 30 mM K+ (total concentration; additions made with equimolar reduction of Na+ to maintain osmolarity), 100 μM CPA, 50 to 100 μM DDA, 2 mM EGTA, 0.25 mM GTP, 1 μM ionomycin, 400 μM SQ 22,536, 100 μM Rp-8-CPT-cAMPS, 2.5 mM 8-Br-cAMP, 100 μM cAMP, 1 μM 007-AM, and 100 μM 007 sodium salt. Where indicated, the MIN6 cells were permeabilized with α-toxin from Staphylococcus aureus (PhPlate). The cells were then superfused with an intracellular-like medium containing 140 mM KCl, 6 mM NaCl, 1 mM MgCl2, 0.465 mM CaCl2, 2 mM EGTA, 2 mM N-(2-hydroxyethyl)ethylenediamine-N,N′,N′-triacetic acid (HEDTA), 1 mM Mg-ATP, 0.25 mM Na-GTP, 100 μM IBMX, and 10 mM Hepes (pH adjusted to 7.00 with KOH). For permeabilization with α-toxin, superfusion was temporarily interrupted, and 5 μl of α-toxin (0.46 mg/ml) was added directly into the 50-μl superfusion chamber. After 2 to 5 min, cells were washed and 0.1 μM to 1 mM cAMP were added to the medium, whereas the concentrations of Mg2+ and Ca2+ were maintained at 1 mM and 0.1 μM, respectively. In some experiments, the free Ca2+ concentration was increased to 0.5–10 μM by addition of appropriate amounts of CaCl2. Calculations of the Mg2+ and Ca2+ concentrations were done with MAXCHELATOR software (

Measurements of the cytoplasmic Ca2+ concentration beneath the plasma membrane

The cells were loaded with the Ca2+ indicators Fluo-4 or Fura Red by 30 to 60 min of incubation at 37°C in experimental buffer supplemented with 5 μM of respective acetoxymethyl ester. After rinsing the cells in indicator-free medium, measurements of the sub–plasma membrane Ca2+ concentration were performed with a TIRF microscope setup described below.

Measurements of cAMP concentration beneath the plasma membrane

The sub–plasma membrane concentration of cAMP was measured with a fluorescent translocation biosensor as previously described (34). The biosensor consists of a membrane-anchored and truncated form of the PKA regulatory RIIβ subunit and an YFP-labeled PKA catalytic Cα subunit (Cα-YFP). Upon rise of the cAMP concentration beneath the plasma membrane, Cα-YFP dissociates from the regulatory subunit, and this translocation is detected with TIRF microscopy as loss of YFP fluorescence. In some experiments, a CFP-tagged RIIβ subunit was used and the cAMP concentration was instead recorded as changes of the CFP/YFP ratio.

Single-cell recordings of insulin secretion

The dynamics of insulin secretion from individual cells was determined by monitoring the phosphoinositide 3-kinase–mediated formation of PIP3 in the plasma membrane, which follows autocrine activation of insulin receptors. The pleckstrin homology domain from Akt fused to YFP (PHAkt-YFP) was used as a translocation biosensor for PIP3 as previously described (32).

Pull-down assay for Rap activity

The amount of GTP-Rap1 was determined essentially as previously reported (38). In brief, MIN6 β cells were seeded in 100-mm dishes (107 cells per dish) and cultured for 2 days. The cells were subsequently washed with cold phosphate-buffered saline on ice and lysed in cell lysis buffer [1% NP-40; 10% glycerol, 50 mM tris-HCl (pH 7.4), 200 mM NaCl, 2.5 mM MgCl2, 1 mM phenylmethylsulfonyl fluoride, 2 mM Na3VO4, and Halt protease inhibitor cocktail (Thermo Scientific)]. Lysates were clarified and incubated with an excess of GST-RalGDS-RBD immobilized on glutathione beads (Sigma) to pull out active Rap1. The amount of active Rap1 and total Rap1 was determined by Western blotting with a monoclonal anti-Rap1 antibody (BD Biosciences) that recognizes both Rap1A and Rap1B. A goat anti-mouse secondary antibody conjugated to horseradish peroxidase (GE Healthcare) was applied, and the chemiluminescence reaction was carried out with Immobilon Western Chemiluminescent HRP Substrate (Millipore). Quantification of Rap1 protein was performed by densitometric analysis of Western blots with ImageJ (National Institutes of Health).


Cells grown on chamber slides were fixed with 4% paraformaldehyde for 15 min at room temperature followed by 40 min of washing and permeabilization in 0.1% Triton X-100. The cells were then incubated with an Epac2 mouse monoclonal antibody (1:50 dilution, Sc-28326; Santa Cruz Biotechnology) at 4°C for 19 hours, followed by overnight incubation with a goat anti-mouse Alexa Fluor 568 (AF-568)–conjugated secondary antibody (1:300; Invitrogen). The chamber slides were finally mounted in ProLong Gold mounting medium (Invitrogen), and the samples were imaged with a confocal microscope (see below).

Fluorescence microscopy

The subcellular distribution of fluorescence-tagged proteins was assessed with a spinning disc confocal unit (Yokogawa CSU-10) attached to a Nikon TE2000-U microscope. Light of 488 nm for excitation of GFP and of 561 nm for excitation of AF-568 and mCherry were provided by a multiline argon laser (ALC 60X, Creative Laser Production) and a diode-pumped solid-state laser (Jive, Cobolt AB), respectively. An acousto-optic tunable filter (AA Optoelectronics) was used for selection of excitation wavelength and to block the beam between image captures. Fluorescence from the cells was collected with a 60× 1.4 NA (numerical aperture) oil immersion objective (Nikon) and detected through a 530/50 nm half-bandwidth interference filter (GFP) or a 625LP filter (AF-568, mCherry) with a back-illuminated EMCCD (electron-multiplying charge-coupled device) camera (DU-888, Andor Technology, Belfast) controlled by MetaFluor software (Molecular Devices Corp.).

Fluorescence in the sub–plasma membrane compartment was recorded with a TIRF microscopy setup based on an E600FN upright microscope (Nikon) contained in an acrylic glass box thermostated at 37°C by an air stream incubator. A helium-cadmium laser (Kimmon) provided 442-nm light for excitation of CFP, and an argon laser (ALC 60X) provided 488- and 514-nm light for excitation of GFP and YFP, respectively. Another Jive diode-pumped solid-state laser (Cobolt AB) provided 561-nm light for td2 and mCherry excitation. The laser beams were merged with dichroic mirrors (Chroma Technology), homogenized, and expanded by a rotating Light Shaping Diffuser (Physical Optics Corp.) before being refocused through a modified quartz dove prism (Axicon) with 70° angle to achieve total internal reflection. Laser lines were selected with interference filters (Semrock) in a motorized filter wheel equipped with a shutter (Sutter Instruments) blocking the beam between image captures. The coverslips with attached cells were used as exchangeable bottoms of an open 200-μl superfusion chamber. The chamber was mounted on the custom-built stage of the microscope such that the coverslip was maintained in contact with the dove prism by a thin layer of immersion oil. Fluorescence from the cells was collected through a 40× 0.8 NA water immersion objective (Nikon) and detected at 530/50 nm for GFP, 485/25 nm for CFP, and 542/27 nm for YFP (Semrock interference filters) or 625LP for td2/mCherry/Fura Red (Melles Griot glass filter) with a CCD camera (ORCA-AG) under MetaFluor software control. Images were acquired every 1 to 5 s with exposure times in the 100- to 400-ms range. Some experiments, such as those involving intact pancreatic islets or α-toxin–permeabilized cells, were performed at an alternative TIRF imaging setup based on an inverted Eclipse Ti microscope (Nikon) equipped with a 60× 1.45 NA objective as previously described (55).

Image analysis and statistics

Images were analyzed with MetaFluor. Changes of fluorescence intensity over time were recorded from manually defined regions of interest corresponding to individual cells. The data were normalized by expressing the fluorescence or fluorescence ratio relative to the prestimulatory intensity or ratio after background subtraction. All experiments were repeated a minimum of three times, and the data are expressed as means ± SE. A two-tailed Student’s t test was used to assess statistical differences, and P values <0.05 were considered statistically significant.

Supplementary Materials

Fig. S1. Glucose- and cAMP-induced Epac2 translocation.

Fig. S2. Permissive effect of Ca2+ on cAMP-induced translocation of Epac2.

Fig. S3. Dual effects of Ca2+ on cAMP-induced translocation of Epac2.

References and Notes

Acknowledgments: We are indebted to D. Altschuler (University of Pennsylvania, Philadelphia, PA), L. Quilliam (Indiana University School of Medicine, Indianapolis, IN), R. Y. Tsien (University of California, San Diego, CA), M. Fivaz (Duke Graduate Medical School, National University of Singapore, Singapore), and H. Rehmann and J. L. Bos (University Medical Centre Utrecht, Netherlands) for gifts of plasmids. I.-M. Mörsare and P. Ahooghalandari are acknowledged for their skilful technical assistance. Funding: This study was supported by grants to A.T. from the European Foundation for the Study of Diabetes/Merck Sharp and Dohme, the Family Ernfors Foundation, Novo Nordisk Foundation, the Swedish Diabetes Association, and the Swedish Research Council. Y.X. was supported by a postdoctoral fellowship from the Swedish national strategic grant consortium EXODIAB (Excellence of Diabetes Research in Sweden). Author contributions: O.I.-H. performed most of the Epac2, Raf1RBD, and RalGDSRBD translocation experiments, analyzed the data, and contributed to the writing of the manuscript. I.J. performed Epac2 and RalGDSRBD translocation experiments in primary mouse β cells, constructed Epac2 point mutants, investigated their translocation, and analyzed the data. Y.X. performed the pull-down experiments and the PIP3 measurements. A.T. conceived the study, analyzed the data, and wrote the manuscript. All authors critically revised the manuscript and approved the final version. Competing interests: The authors declare that they have no competing interests.
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