Search tips
Search criteria 


Logo of jbcThe Journal of Biological Chemistry
J Biol Chem. 2010 July 30; 285(31): 24248–24259.
Published online 2010 June 1. doi:  10.1074/jbc.M109.094581
PMCID: PMC2911347

Differential Roles of Epac in Regulating Cell Death in Neuronal and Myocardial Cells*An external file that holds a picture, illustration, etc.
Object name is sbox.jpg


Cell survival and death play critical roles in tissues composed of post-mitotic cells. Cyclic AMP (cAMP) has been known to exert a distinct effect on cell susceptibility to apoptosis, protecting neuronal cells and deteriorating myocardial cells. These effects are primarily studied using protein kinase A activation. In this study we show the differential roles of Epac, an exchange protein activated by cAMP and a new effector molecule of cAMP signaling, in regulating apoptosis in these cell types. Both stimulation of Epac by 8-p-methoxyphenylthon-2′-O-methyl-cAMP and overexpression of Epac significantly increased DNA fragmentation and TUNEL (terminal deoxynucleotidyltransferase-mediated biotin nick end-labeling)-positive cell counts in mouse cortical neurons but not in cardiac myocytes. In contrast, stimulation of protein kinase A increased apoptosis in cardiac myocytes but not in neuronal cells. In cortical neurons the expression of the Bcl-2 interacting member protein (Bim) was increased by stimulation of Epac at the transcriptional level and was decreased in mice with genetic disruption of Epac1. Epac-induced neuronal apoptosis was attenuated by the silencing of Bim. Furthermore, Epac1 disruption in vivo abolished the 3-nitropropionic acid-induced neuronal apoptosis that occurs in wild-type mice. These results suggest that Epac induces neuron-specific apoptosis through increasing Bim expression. Because the disruption of Epac exerted a protective effect on neuronal apoptosis in vivo, the inhibition of Epac may be a consideration in designing a therapeutic strategy for the treatment of neurodegenerative diseases.

Keywords: Apoptosis, Cell Death, Cyclic AMP (cAMP), Mitochondrial Apoptosis, Neuron, Protein Kinase A (PKA), Signal Transduction, Exchange Protein Activated by Cyclic AMP (Epac), Myocyte


Induction of apoptosis in post-mitotic cells, such as neurons and cardiac myocytes, has been thought to be responsible for such irreversible disorders as Alzheimer and Huntington diseases as well as stroke and heart failure (1). The effect on cell death of cyclic AMP (cAMP), a major second messenger, has been extensively studied. In neuronal cells it is well known that activation of cAMP signals reduces the rate of neuronal cell death under a variety of stresses (i.e. β-amyloid protein, sialoglycopeptide, low potassium-induced neurotoxicity) (2,4), although there have been several reports that dopamine or prostanoid receptor-mediated cAMP production induces neurotoxicity (5, 6). β-Adrenergic receptor signaling, on the other hand, promotes apoptosis in cardiac myocytes, resulting in heart failure (7, 8). Therefore, the model proposing that cAMP signaling plays a protective role in neuronal cells but a deteriorative role in myocardial cells is well accepted.

Most studies that have demonstrated the effect of cAMP signaling on apoptosis have focused primarily on protein kinase A (PKA),2 a classic target molecule of cAMP. Recent studies involving cAMP signaling have focused instead on Epac, an exchange protein activated by cAMP that has been identified as a new target of cAMP, independent of PKA (9). Epac has been found to regulate a variety of cellular processes, including cell proliferation, migration, secretion, and differentiation (10). It has been demonstrated that Epac either alone or with PKA plays a protective role in immune cells against apoptosis (11, 12). In post-mitotic cells such as neurons and cardiac myocytes, however, the role of Epac in apoptosis has not been reported.

To date two isoforms of Epac have been identified, Epac1 and Epac2 (9); they differ in that Epac2 contains a second binding site for cAMP. It has recently been reported that there is an up-regulation of Epac1 mRNA in Alzheimer disease (13) and an up-regulation of Epac1 protein expression in rats with inflamed neurons (14), implicating that cAMP signaling may not always play a protective role in neurons. The change in the Epac1 expression pattern has also been demonstrated in other cell types (i.e. heart, vasculature, kidney, and lung) (15,18). The stoichiometry of Epac, especially of Epac1, and that of PKA might be changed in several diseases, including neuronal and cardiac disorders; this could lead to the various effects of cAMP signaling on cell death.

Through experiments using Epac- or PKA-selective cAMP analogs and overexpression of Epac1 and the PKA catalytic subunit and Epac1-deficient mice, the present study demonstrates that cAMP signaling no longer increases neuronal cell viability when Epac is selectively activated: instead, cAMP signaling induces apoptosis through increasing Bcl-2 interacting member protein (Bim) expression. Our findings also suggest that the selective inhibition of Epac signaling may become a therapeutic strategy in the treatment of neurodegenerative diseases.


Antibodies and Reagents

8-p-Methoxyphenylthon-2′-O-methyl-cAMP (pMe-cAMP) and N6-benzoyladenosine-cAMP (Bnz-cAMP) were purchased from BioLog Life Science Institute (Bremen, Germany) and Sigma, respectively. Antibodies to Epac1, Epac2, and a PKA α catalytic subunit were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). An antibody to Bim was purchased from Stressgen Biotechnologies (Victoria, BC, Canada). Antibodies to Bim and cleaved caspase 3 were purchased from Cell Signaling Technology (Danvers, MA). An antibody to Bcl-2 was purchased from BD Biosciences.

Generation of Epac1 Knock-out Mice

Epac1 knock-out mice (Epac1 KO; accession number CDB0542K (LARGE (Laboratory for Animal Resources and Genetic Engineering)) were generated by means of homologous recombination (19). Briefly, the targeting vector was constructed by inserting loxP/PGK-Neo-pA/loxP (LARGE) into exon 1 and exon 2 of the genomic Epac1 locus (see Fig. 7A). The targeting vector was introduced into TT2 embryonic stem cells, and homologous recombinant clones were first identified by PCR, then confirmed by Southern blot analysis (see Fig. 7B). The targeted embryonic stem cell clones were injected into CD-1 8-cell stage embryos, and the resultant male chimeras were mated with C57BL/6 females to establish germ line transmission. All experiments were performed on C57BL/6 and CBA mixed-background 3–5-month-old male homozygous Epac1 KO and wild type (WT) littermates from F1 heterozygote crosses. Mice were genotyped by PCR using a mixture of three primers (F1, TGA GAA GAG CCC CAT CGT TGT G; B1, GCC TGG CAC ATG GAA GTG AT; NeoF1, TGA ATG GAA GGA TTG GAG CTA CG) as indicated in Fig. 7A. The PCR conditions consisted of 95 °C for 5 min, 35 cycles of 95 °C for 30 s each, 60 °C for 30 s, and 72 °C for 30 s followed by 72 °C for 7 min (Fig. 7C).

Generation of Epac1 gene-targeted mice. A, targeted disruption of the Epac1 gene is shown. The partial structure of the Epac1 gene (WT) and the resultant mutated allele (Epac1 KO) are shown. The positions of the phosphoglycerate kinase promoter neo cassette ...

All experiments were performed on 3–5-month-old homozygous Epac1 KO mice and WT littermates. This study was approved by the Animal Care and Use Committee at Yokohama City University School of Medicine.

Primary Culture of Fetal Mouse Cortical Neurons

Primary cortical neurons were isolated from the cortices of embryonic day 15–17 C57BL/6 or Epac1 KO mice, as previously described (20) with some modifications. Briefly, the cortex was incubated with 0.3% trypsin (Invitrogen) by titration; then cells were plated onto a 12-mm glass coverslip precoated with 6 mg/ml poly-l-lysine (Wako Pure Chemical Industries, Osaka, Japan) at a density of 1 × 105 cells/glass. The cells were incubated at 37 °C with 5% CO2, 95% atmospheric air in a neurobasal medium containing 1× GlutaMAX-1, B-27 supplement (Invitrogen), 100 μg/ml penicillin, and 100 μg/ml streptomycin. Cells were used in experiments 4–7 days later.

Primary Culture of Neonatal Mouse Cardiac Myocytes

Cardiac myocytes were isolated from the hearts of 1-day-old mice as previously described (21) with some modifications. Briefly, myocytes were chopped into small pieces and digested with 0.1% collagenase type II and 0.04% pancreatin 3 times at 7-min intervals. To remove the non-myocyte fraction, the cells were plated onto culture dishes in minimum essential medium (Invitrogen) with 10% fetal bovine serum containing 100 μg/ml penicillin and 100 μg/ml streptomycin for 45 min, after which the non-attached myocyte-rich fraction was plated onto a 12-mm glass coverslip precoated with 20 mg/ml laminin (Sigma) at a density of 1 × 105 cells/glass in the same medium. Twenty-four hours after plating, the culture medium was changed to minimum essential medium with an insulin-transferrin-selenium-A supplement (ITS-A, Invitrogen) containing 100 μg/ml penicillin and 100 μg/ml streptomycin. The cells were maintained in a humidified 5% CO2, 95% atmospheric air incubator at 37 °C.

Primary Culture of Mouse Renal Epithelial Cells

Primary culture of mouse renal epithelial cells was performed as previously described (22).

Quantitative Reverse Transcription (RT)-PCR

Total RNA was extracted from cortical neurons using TRIzol (Invitrogen) according to the manufacturer's instructions. Both the generation of cDNA and the RT-PCR analysis were performed as previously described (17, 23). Real-time PCR was executed using a MyiQ Single-Color Real-Time PCR Detection System (Bio-Rad) and an SYBR Green kit (Takara Bio, Shiga, Japan). Primers for amplification were designed based on Bim (5′-CCCGGAGATACGGATTGCAC-3′ and 5′- GCCTCGCGGTAATCATTTGC-3′) and 18 S ribosomal RNA. The forward and reverse primer set was designed between multiple exons. Abundance of mRNA was determined relative to that of 18 S ribosomal RNA.

Northern Blotting

Partial fragments of mouse Epac1 and Epac2 cDNA clones were obtained by PCR. A mouse glyceraldehyde-3-phosphate dehydrogenase probe was used as an internal control. Northern blotting was performed as previously described (24).

Western Blot Analysis

Western blot analysis of cortical neurons and cardiac myocytes was performed as previously described (25) with some modifications. Briefly, cells in 35-mm plastic dishes were lysed and collected with a lysis buffer (25 mm Tris-HCl (pH 8.0), 10 mm EGTA, 10 mm EDTA, 10 mm Na4P2O7, 100 mm NaF, 10 mm Na3VO4, 20 μg/ml 1-chloro-3-tosylamido-7-amino-2-heptanone or Nα-p-tosyl-l-lysine chloromethyl ketone, 10 μg/ml leupeptin, 1 mm phenylmethylsulfonyl fluoride, 50 units of erythrina trypsin inhibitor, 2 μg/ml aprotinin, and 1% Nonidet P-40). After protein concentrations were determined using the RC DC protein assay kit (Bio-Rad), SDS-PAGE and Western blotting were performed followed by densitometric analysis using LAS3000 and Science Lab Multi Gauge Version 3.0 software (Fujifilm, Tokyo, Japan).


Lysates from cells treated with pMe-cAMP or Bnz-cAMP for 24 h were incubated with 2 μg of anti-Bcl-2 or anti-Bim antibody overnight. Immune complexes were captured with protein G-Sepharose 4 Fast Flow (GE Healthcare). Beads were washed 3 times in the lysis buffer and boiled in an SDS sample buffer. Samples were subjected to SDS-PAGE and blotted onto a polyvinylidene difluoride membrane (Immobilon-P; Millipore, Billerica, MA).

Adenovirus Construction

For construction of adenoviral vectors, full-length cDNA-encoding human Epac1, Epac2, and PKA α catalytic subunits were cloned into an adenoviral vector using an AdenoX adenovirus construction kit (Clontech, Mountain View, CA) (17). Human Epac1 and Epac2 cDNAs were kindly provided by Dr. J. L. Bos of University Medical Center, Utrecht, The Netherlands. Adenovirus-mediated transfection was performed using LacZ control. The cells were infected with adenoviruses at the indicated multiplicities of infection and used for their corresponding assays 24 h later.

Transfection of siRNA in Cortical Neurons

Silencing of Bim was carried out using Accell SMARTpool siRNAs (Dharmacon Inc., Lafayette, CO), each of which contains four siRNAs designed for use with the Bim gene (5′-CUGGCUUCCUUUACGUUUU-3′, 5′-CUAUGAAUUGUAGAAGUAU-3′, 5′-CGCUUAUUUAAAUGUCUUA-3′, and 5′-UCAUAAUUAAGGAUUUGUA-3′) according to the manufacturer's instructions. Briefly, 48 h after plating, cortical neurons were transfected with 1 μm siRNA and subsequently cultured in a neurobasal medium. The efficiency of the knockdown of the Bim protein was evaluated 72 h after transfection by Western blot analysis. Scrambled siRNAs for Bim-targeted siRNA (Dharmacon Inc.) were used as a negative control.

Terminal Deoxynucleotidyltransferase-mediated Biotin Nick End-labeling (TUNEL) Assay

In situ labeling of fragmented DNA in cultured cortical neurons and cardiac myocytes was performed using the DeadEndTM fluorometric TUNEL system (Promega, Madison, WI) according to the manufacturer's instructions. Cells were incubated with the presence or absence of pMe-cAMP or Bnz-cAMP for 48 h, fixed with 4% paraformaldehyde for 25 min, and then incubated with 0.2% Triton X-100 for 5 min. The cells were equilibrated with a buffer consisting of 200 mm potassium cacodylate (pH 6.6), 25 mm Tris-HCl (pH 8.0), 0.2 mm dithiothreitol, 0.25 mg/ml bovine serum albumin, and 2.5 mm cobalt chloride at room temperature for 10 min followed by 60 min of incubation with a terminal deoxynucleotidyltransferase reaction buffer containing 100 μm dATP, 5 μm fluorescein-12-dUTP, 10 mm Tris-HCl (pH 7.6), 1 mm EDTA, and 40 μm terminal deoxynucleotidyltransferase enzyme at 37 °C. DNAs were stained with DAPI (4′, 6-diamidino-2 phenylindole). The percentage of the total cells that were TUNEL-positive was determined in a blinded manner. Approximately 2000–3000 cells in 10 randomly selected fields from each sample were counted. For detection of apoptosis in brain tissues from WT or Epac1 KO mice, deparaffinized tissue sections were treated with 20 μg/ml proteinase K and 50 mm EDTA in 100 mm Tris-HCl (pH 8.0). The sections were fixed with 4% paraformaldehyde for 15 min at room temperature and then subjected to the equilibration step in the procedures described above.

Analysis of DNA Fragmentation by Enzyme-linked Immunosorbent Assay

Histone-associated DNA fragments were quantified using the Cell Death Detection enzyme-linked immunosorbent assay kit (Roche Diagnostics) according to the manufacturer's instructions. After cortical neurons and cardiac myocytes were incubated in the presence or absence of pMe-cAMP or Bnz-cAMP for 48 h, they were gently washed with phosphate-buffered saline and incubated with a lysis buffer (phosphate-buffered saline containing 10 mm EDTA (pH 7.2) and 0.1% Triton-X) for 1 h at 37 °C followed by vigorous shaking for 30 s. The cell lysates containing cytoplasmic histone-associated DNA fragments were applied to a streptavidin-coated microtiter plate. Subsequently, a mixture of biotin-labeled anti-histone antibody and peroxidase-conjugated anti-DNA antibody was added, and the resulting mixture was incubated with moderate shaking for 2 h. After unbound antibodies were removed by washing, the amount of nucleosomes was quantified based on the peroxidase retained in the immune complex. The activity of the peroxidase was determined photometrically using 2,2-azino-di-[3-ethylbenzthiazoline-sulfonate] as a substrate. The values from triplicate absorbance (at 405 nm) measurements were then averaged.

Mitochondrial Membrane Potential Analysis

Mitochondrial membrane potential of cortical neurons was quantified using a MitocaptureTM Mitochondrial Apoptosis Detection kit (BioVision Inc., Mountain View, CA) according to the manufacturer's instructions. Cortical neurons were incubated on a 12-mm glass coverslip in the presence or absence of pMe-cAMP or Bnz-cAMP for 48 h, then stained with Mitocapture reagent and incubated in DAPI to allow the visualization of all nuclei. The images were obtained using an inverted microscope (TE2000-E, Nikon, Japan). The red emission of the dye detected at 543 nm is due to a potential-dependent aggregation in the mitochondria reflecting normal membrane potential. Green fluorescence detected at 488 nm reflects the monomeric form of MitocaptureTM, appearing in the cytosol after mitochondrial membrane depolarization. The percentage of the total cells representing apoptotic cells was determined in a blinded manner by counting ~1000–3000 cells in 10 randomly selected fields from each sample.

Rap1 Activation Assay

Rap1 activity was measured using the EZ-Detect RAP1 activation kit (Pierce) according to the manufacturer's instructions. Primary renal epithelial cells from WT and Epac1 KO mice were lysed 15 min after stimulation with pMe-cAMP (50 μm). Cell lysates were incubated with the Rap binding domain RalGDS-RBD fused to a glutathione S-transferase disk. After cells were washed several times, bound GTP-Rap1 was removed from the disk through boiling in an SDS sample buffer and analyzed by Western blotting using an anti-Rap1 antibody.

In Vivo Experiment and Tissue Preparation

3-Propionic acid (3-NP, Sigma) was prepared and administered as previously described (26). 3-NP was dissolved in saline, and the resulting solution was adjusted to pH 7.3–7.4 with 5 n NaOH. 3-NP (140 mg/kg/day) was the injected intraperitoneally into the animals once per day for 2 days. Twenty-four hours after the second injection, the mice were anesthetized with pentobarbital and transcardially injected with 10% paraformaldehyde/phosphate-buffered saline (pH 7.4). Brains were fixed in the same fixative solution overnight, immersed in 70% ethanol for 24 h, then embedded in paraffin. Sections 4 μm in thickness were subjected to TUNEL staining.

Statistical Analysis

All data are reported as the mean ± S.E. Comparisons between two groups were analyzed using Student's t test. For multiple groups, one-way analysis of variance was used with a Bonferroni post-hoc test. p < 0.05 was considered to indicate significance.


Differential Effects of Epac on Apoptosis in Mouse Cortical Neurons and Myocytes

We first investigated whether the stimulation of Epac has similar effects on apoptosis in various kinds of post-mitotic cells using a primary culture of mouse cortical neurons and cardiac myocytes. Apoptosis was detected by means of TUNEL staining 48 h after treatment with pMe-cAMP or Bnz-cAMP, the Epac- and PKA-selective cAMP analogs, respectively (27). We observed strong TUNEL labeling of apoptotic bodies in cortical neurons treated with pMe-cAMP (Fig. 1A, upper panels, inset). In contrast, pMe-cAMP did not induce apoptosis in cardiac myocytes (Fig. 1A, lower panels). The number of TUNEL-positive cells was quantified, showing that stimulation of Epac significantly increased apoptosis in cortical neurons but not in cardiac myocytes (Fig. 1, B and C). Because a high Bax/Bcl-2 ratio is associated with greater vulnerability to apoptotic activation (28, 29), we quantified the protein expression of Bax/Bcl-2 to confirm our findings. Activation of Epac by pMe-cAMP increased the Bax/Bcl-2 ratio in cortical neurons but not in cardiac myocytes (supplemental Fig. 1, A–D).

Effects of activation of Epac and PKA on apoptosis in cortical neurons and cardiac myocytes. A, apoptotic cells (green) in cortical neurons and myocytes were examined by means of TUNEL staining 48 h after treatment with pMe-cAMP (50 μm) or Bnz-cAMP ...

It is already known that activation of cAMP/PKA signaling plays a protective role in neuronal cells but a deteriorative role in myocardial cells (30, 31). In accordance with previous reports, the PKA-selective cAMP analog Bnz-cAMP induced apoptosis in cardiac myocytes but not in cortical neurons, even at 100 μm (Fig. 1, A–C). These results suggest that the activation of Epac has different effects on apoptosis in neuronal cells and cardiac myocytes.

Effects of Overexpression of Epac on Apoptosis in Cortical Neurons and Cardiac Myocytes

To confirm that Epac was involved in neuronal cell death, we performed adenovirus-mediated gene transfer of Epac1 and Epac2 (as both isoforms of Epac (9) are known to be activated by pMe-cAMP), a PKA α catalytic subunit, or a LacZ control. Overexpression of Epac1, Epac2, PKA α catalytic subunit proteins, or PKA α regulatory subunit proteins in cortical neurons and cardiac myocytes 12 h after infection with each adenovirus is shown in Fig. 2, A and B. Endogenous protein expression of Epac1, Epac2, and PKA subunits was not significantly affected by any of the adenoviruses. We also confirmed that overexpression of PKA α catalytic subunit significantly increased PKA activity in cortical neurons (supplemental Fig. 2). Overexpression of Epac1 or Epac2, unlike LacZ, significantly increased the incidence of TUNEL-positive apoptotic cells in cortical neurons but not in cardiac myocytes (Fig. 2, C and D). In contrast, overexpression of PKA increased the number of TUNEL-positive cells in cardiac myocytes but not in cortical neurons. An enzyme-linked immunosorbent assay yielded results similar to those of TUNEL staining (Fig. 2, E and F), indicating that overexpression of Epac1 or Epac2 increased DNA fragmentation in cortical neurons but not in cardiac myocytes. Importantly, these results are different from those obtained through PKA overexpression, suggesting that, at least in part, Epac promotes neuronal, but not myocardial, apoptosis.

Effects of overexpression of Epac and PKA on apoptosis in cortical neurons and cardiac myocytes. A, shown are the representative immunoblots of cortical neurons or myocytes transfected with Epac1, Epac2, PKA α catalytic subunit, PKA regulatory ...

Epac Activation Increases Bim Expression

The Bcl-2 interacting member (Bim) is a sensor of apoptotic stress located upstream of the Bcl-2 family. Bim regulates Bcl-2 in the mitochondrial membrane, resulting in apoptosis (32, 33). Bim protein was highly expressed in brain tissue and cortical neurons (Fig. 3, A and B) but was expressed either slightly or not at all in heart tissue and cardiac myocytes, as previously described (34, 35). A previous paper suggested the involvement of cAMP/PKA signaling in regulating Bim expression in lymphoid cells (55). We, therefore, examined the change in the expression levels of Bim mRNA and protein in cortical neurons using pMe-cAMP. We found that pMe-cAMP increased expression of Bim mRNA and protein in a time-dependent manner (Fig. 3C and see Fig. 5, C and F). Furthermore, pMe-cAMP, but not Bnz-cAMP, significantly increased Bim protein in cortical neurons 24 h after treatment (Fig. 3,D and E). These results suggest that the stimulation of Epac increases Bim expression by enhancing transcription in cortical neurons, although the stimulation of PKA does not.

Activation of Epac increased the expression of Bim mRNA and protein in cortical neurons. A and B, shown is expression of endogenous Bim protein in brain and heart tissues (A) and cultured cortical cells and cardiac myocytes (B). C, the expression of Bim ...
Activation of Epac promoted interaction between Bcl-2 and Bim protein and decreased mitochondrial transmembrane potential in cortical neurons. A–C, a representative immunoblot (IB) shows the interaction between Bcl-2 and Bim protein was increased ...

Inhibition of p38 MAPK Attenuates Epac-induced Bim Expression and Apoptosis

Three signal pathways have been implicated in regulating Bim protein expression: the JNK/c-Jun, cell cycle (Cdk4/E2F/Myb), and p38MAPK/FoxO pathways (36, 37). We next sought to determine which of these pathways plays the most important role. We found that an Epac-selective cAMP analog increased phosphorylation of p38 MAPK in cortical neurons, although a PKA-selective cAMP analog did not (Fig. 4, A and B). In contrast, there was no significant difference between Epac and PKA stimulation in terms of their effects on the phosphorylation of p44/42 MAPK, JNK1, or JNK2/3 (data not shown). Furthermore, SB203580, a p38 MAPK inhibitor, attenuated Epac-induced Bim expression and apoptosis (Fig. 4, C and D). These results suggest that Epac-induced neuronal apoptosis is mediated by the elevation of Bim expression via p38 MAPK.

Effect of p38 MAPK on Epac-induced Bim expression and apoptosis in cortical neurons. A and B, shown are representative images and quantification of phosphorylation/total p38 protein in mouse cortical neurons treated with pMe-cAMP (50 μm) or Bnz-cAMP ...

Epac Activation Increases Interaction of Bim with Bcl-2

Bim is thought to exert its pro-apoptotic activity by binding to Bcl-2, thereby blocking the anti-apoptotic function of Bcl-2 (38). After demonstrating that stimulation of Epac increased Bim expression, we needed to confirm that the binding of Bim to Bcl-2 was likewise increased. We conducted pulldown assays using anti-Bcl-2 and anti-Bim antibodies after treatment with pMe-cAMP or Bnz-cAMP and found that the activation of Epac by pMe-cAMP significantly increased the amount of Bim associated with Bcl-2 in cortical neurons 24 h after the treatment (Fig. 5, A and D). Association of Bim with Bcl2 was also increased 10 h after Epac activation (Fig. 5, B and E), suggesting that association of Bim with Bcl2 was increased in accordance with increased Bim expression. Activation of PKA by Bnz-cAMP did not promote binding Bcl-2 with Bim even 24 h after treatment (Fig. 5, H and I).

Because it is already known that Bcl-2 regulates the mitochondrial pathway of apoptosis, we next explored whether pMe-cAMP induced apoptosis through the mitochondrial pathway in cortical neurons. Disruption of mitochondrial transmembrane potential is one of the earliest intracellular events, and such disruption occurs after induction of apoptosis via mitochondria (1). In apoptotic cells, the mitochondrial membrane potential is dissipated, and thus, the Mitocapture dye is dispersed in the cell as green fluorescent monomers detected at 488 nm. We found that Epac activation with pMe-cAMP (50 μm) promoted the disruption of mitochondrial transmembrane potential based on 488-nm-positive (green fluorescence) intensity in cortical neurons (Fig. 5, J and K). Taken together, these data suggest that stimulation of Epac promotes the binding of Bim to Bcl-2, leading to neuronal apoptosis via the mitochondrial pathway.

Epac-induced Neuronal Apoptosis Is Mediated by Bim

To further confirm the contribution of Bim to Epac-induced apoptosis is in cortical neurons, we used Bim-targeted siRNA. Changes in Bim protein expression caused by the siRNAs are shown in Fig. 6A. When Bim was silenced, the effect of pMe-cAMP on the number of TUNEL-positive cells in cortical neurons became significantly smaller (Fig. 6, B and C), although Epac-induced apoptosis was not completely abolished. The evidence suggests that enhanced Bim expression via p38 MAPK appears to play an important role in Epac-induced apoptosis in neuronal cells.

Silencing of Bim attenuated Epac-induced apoptosis in cortical neurons. A, shown is a representative immunoblot of Bim 72 h after transfection of Bim-targeted siRNA or negative siRNA control in cortical neurons. B, cortical neurons were transfected with ...

Effects of Apoptotic Stimuli on Cortical Neurons of Epac1 KO Mice

Two isoforms of Epac, Epac1 and Epac2, have been previously identified (9) and are known to be activated by pMe-cAMP. In the present study we focused on the role of Epac1, because changes in Epac1 expression have been demonstrated in Alzheimer disease and in neuronal cells (13,15). It has, therefore, been tentatively proposed that Epac inactivation might play a protective role against neuronal apoptosis.

To test this theory, we generated Epac1 KO mice (see “Experimental Procedures” and Fig. 7A), which lacked Epac1 expression in neuronal cells as shown by Northern blot analysis (Fig. 7D). pMe-cAMP-induced Rap1 activation in Epac1 KO mice was significantly decreased in renal epithelial cells (Fig. 7E). Because renal epithelial cells do not express Epac2, this decrease in Rap1 activation most likely mirrors the impact of Epac1 deletion.

Induction of neuronal apoptosis has been well demonstrated in cortical neuronal cells using 3-NP (26, 39) or hydroxyl peroxide (40, 41). Using these pharmacological stressors, we examined whether the induction of apoptosis could be altered in cultured neuronal cells obtained from Epac1 KO mice. Apoptosis induced by hydroxyl peroxide or 3-NP, an irreversible inhibitor of mitochondria complex II, and detected through TUNEL staining was significantly decreased in neuronal cells from Epac1 KO mice (Fig. 8, A and B). Furthermore, both mRNA and protein expression levels of Bim remained significantly lower in cells from Epac1 KO mice than in those from WT mice (Fig. 8, C–E), suggesting that Epac1 deletion plays a protective role against neuronal stresses.

The effect of 3-NP and hydroperoxide on apoptosis in cortical neurons from WT and Epac1 KO mice. A and B, apoptosis was evaluated by means of TUNEL staining 48 h after the addition of the indicated reagents in cortical neurons from WT and Epac1 KO mice. ...

Deletion of Epac1 Attenuates 3-NP-induced Neuronal Apoptosis in Vivo

To examine the effect of Epac1 deletion in vivo, we administered 3-NP systemically to intact mice; this is a chemical and pathological way to induce mitochondrial and degenerative disorders in vivo (26, 39). We determined the number of apoptotic cells in cortical and striatal regions of Epac1 KO and WT mice through TUNEL staining. We found that 3-NP-induced apoptosis was significantly decreased in both the cortices and the striata of Epac1 KO mice in vivo (Fig. 9, A, B, D, and E). We confirmed that TUNEL-positive cells were stained with NeuN, a neuron-specific marker (supplemental Fig. 3). Furthermore, the number of cleaved caspase 3-positive cells was significantly increased in WT mice treated with 3-NP and was attenuated in Epac1 KO mice treated with 3-NP (Fig. 9, A, C, D, and F).

Deletion of Epac1 suppressed 3-NP-induced brain cell apoptosis in vivo. A and D, representative images of TUNEL staining and immunohistochemistry of cleaved caspase 3 of cortical and striatal sections from WT or Epac1 KO mice 24 h after injection of 3-NP. ...

Taken together, these results reveal that Epac plays an important role in inducing neuronal, but not myocardial, apoptosis. More importantly, its role in this process is different from that of PKA. We found that neuronal apoptosis was, at least partially, mediated by Epac-Bim signaling and that Epac silencing had a protective role against apoptosis in vivo. Inhibition of Epac might be considered as a therapeutic strategy for the treatment of neurodegenerative diseases.


It is well known that cAMP signaling increases neuronal cell survival and decreases myocardial cell survival. We have demonstrated here that the activation of cAMP signaling does not protect neuronal cells when Epac is selectively activated. Rather, cAMP signaling increased apoptosis in neuronal cells when Epac1 was activated. In myocardial cells, however, Epac activation does not promote apoptosis. To our knowledge this is the first demonstration of the differential role of Epac in apoptosis in neuronal and myocardial cells, both of which are typical post-mitotic cells. The present study suggests that neuronal apoptosis is partly mediated by Epac through increased Bim expression and that the inhibition of Epac signaling plays a protective role in neuronal apoptosis in vivo.

The Roles of Epac and PKA in Apoptosis

The effect of cAMP signaling on cell death has been explored in multiple cell types, although most of these studies were conducted before Epac was identified. In neuronal cells activation of cAMP/PKA signaling inhibited apoptosis induced by KCl in cerebella granule neurons (42) or by human immunodeficiency virus protein gp120 in the brain (43), promoting survival pathways in multiple neuronal cells (44, 45); these findings are in agreement with ours (supplemental Fig. 4). In cardiac myocytes, on the other hand, activation of cAMP signaling through such triggers as β-adrenergic receptor stimulation increased apoptosis (7, 8). In these studies the role of cAMP has been described primarily in terms of the activation of PKA.

Recently, several studies have suggested a contribution of either Epac alone or both Epac and PKA to apoptosis in restricted cell types including B-cell chronic lymphocytic leukemia (12), human leukocytes (11), immature B lymphoma cells (46), RINm5F β-cells (47), and H9c2 cells (48), showing that Epac and PKA play a protective role in apoptosis either alone and/or in concert in immune cells. However, the role of Epac in neuronal and myocardial apoptosis remains unknown despite the importance of cell death in tissues composed of post-mitotic cells. Our results show that stimulation and overexpression of Epac induces apoptosis in neurons but not in cardiac myocytes, implying that there are cell type-based differences in the effect of Epac activation on cell survival.

Epac-induced Apoptosis through Increased Bim Expression in Neuronal Cells

Our study demonstrated that Epac-induced apoptosis is mediated through the regulation of Bim, which acts on mitochondria as a pro-apoptotic factor, leading to disruption of the mitochondrial membrane potential. Bim binds to Bcl-2 and neutralizes its pro-survival function, resulting in apoptosis in multiple cell types (38, 49, 50). Bim is known to be expressed in neurons, hematopoietic cells, germ cells, lymphoid tissues, myeloid cells, and epithelial cells but not in cardiac myocytes, skeletal muscle, or neural-supporting cells, including glial, astrocytes, and oligodendrocytes (35). In agreement with these reports, our results show that Bim protein was highly expressed in primary culture of mouse cortical neurons but not in mouse cardiac myocytes. In cortical neurons we found that an Epac-selective cAMP analog increased Bim protein at the transcriptional level. When Bim was silenced, Epac-induced apoptosis was attenuated in neuronal cells. It should be noted that we were not able to exclude the possibility of off-target effects of the siRNAs because the rescue experiment that might exclude them is technically difficult. However, our results together with other data indicating that the suppression of the p38 MAPK pathway inhibits the elevation of Bim mRNA expression and Epac-induced apoptosis suggest that Epac-induced apoptosis is at least partly mediated by increased Bim expression. In fact, gene transfer of Bim to cardiac myocytes, which do not express Bim protein, induced apoptosis (supplemental Fig. 5). Taken together, the evidence strongly suggests that the expression of Bim is responsible for Epac-triggered apoptosis in neuronal cells, whereas Epac does not induce apoptosis in cardiac myocytes due to a lack of endogenous Bim expression. Further investigation is needed to identify the precise mechanism of Epac-induced transcriptional regulation of Bim in neuronal cells.

Changes in Epac Expression in Pathological/Physiological Conditions

Recent studies have indicated that the expression profile of Epac is altered during chronic degenerative inflammatory diseases. Epac1 mRNA, but not Epac2 mRNA, was increased in a mouse vascular injury model and was decreased in cardiac fibroblasts activated by transforming growth factor β (17, 23). Studies have reported the up-regulation of Epac1 mRNA and down-regulation of Epac2 mRNA in Alzheimer disease (13) and the up-regulation of Epac1 protein expression in inflamed rat neurons (14). These studies indicate that the stoichiometry of Epac and especially that of Epac1 can be changed and selectively activated in disease conditions including neurodegenerative disorders.

Approximately half of all neurons in the nervous system undergo apoptosis during embryonic and early postnatal development (51), a period when Epac1 is highly expressed in the brain (15). Our results indicate that Epac1-induced neuronal apoptosis may be involved in the mechanisms underlying neuronal development. Nevertheless, Epac1 KO mice showed normal development up to at least 12 months of age, although no detailed assessment of their behavior, cognition, or learning memory has been made. Further studies using Epac2 KO mice and Epac1 and Epac2 double KO mice will need to be conducted given our observation that overexpression of Epac2 induced neuronal apoptosis in vitro.

The Effect of Epac1 Deletion on Apoptosis in Vivo

The mechanisms of neurological disorders such as Alzheimer disease, Huntington disease, and Parkinson disease are thought to stem from mitochondrial dysfunction (52). 3-NP, an irreversible inhibitor of the mitochondrial enzyme succinate dehydrogenase, is often administered systemically to treat these conditions and is considered to possess unique chemical and pharmacological traits that are accordingly considered in the generation of models of mitochondrial disorders and degenerative disorders (26, 39). The mechanisms of 3-NP toxicity are also thought to involve enhanced production of reactive oxygen species, including hydrogen peroxide, which can cause oxidative damage to DNA, lipids, and proteins (53). In the present study both 3-NP and hydrogen peroxide failed to induce apoptosis in cultured cortical neurons from Epac1 KO mice, and 3-NP-induced neuronal apoptosis was abolished in Epac1 KO mice in vivo. In contrast, there was no difference between Epac1 KO and WT mice in terms of 3-NP-induced apoptosis. Cardiac myocytes from Epac1 KO mice did not differ in their sensitivity to cAMP analogs (supplemental Figs. 6 and 7). Although this and our previous studies show that the inhibition of Epac1 protects 3-NP-mediated neuronal apoptosis in vivo and in vitro, the relevance of Epac2 to this phenomenon needs to be examined in future studies. A recent study has demonstrated that Epac is involved in the secretion of an amyloid precursor protein, which has been known to induce apoptosis leading to Alzheimer disease (54). Together with our data, this indicates that selective inhibition of the Epac signal may prove useful as a therapeutic strategy in treating neurodegenerative diseases.

In conclusion, Epac induces neuronal apoptosis through increased Bim expression. Because disruption of Epac1 exerts a protective effect on neuronal apoptosis in vivo, inhibition of Epac may be a useful tactic in the treatment of neurodegenerative diseases.

Supplementary Material

Supplemental Data:

*This work was supported by a grant-in-aid for Scientific Research (KAKENHI) (to U. Y. and S. S.) and by grants from the Ministry of Health, Labor, and Welfare (to Y. I.), the Ministry of Education, Culture, Sports, Science, and Technology of Japan (to Y. I.), the Yokohama Foundation for Advanced Medical Science (to U. Y.), the Kanae Foundation for the Promotion for Medical Science (to U. Y.), the Miyata Cardiology Research Promotion Funds (to U. Y.), the Takeda Science Foundation (to U. Y.), the Sumitomo Foundation (to U. Y.), the Japan Heart Foundation Research Grant (to U. Y.), the Kowa Life Science Foundation (to U. Y.), the Cosmetology Research Foundation (to Y. I.), the Uehara Memorial Foundation (to U. Y.), the Kitsuen Research Foundation (to Y. I.), and the Japan Space Forum (to Y. I.).

An external file that holds a picture, illustration, etc.
Object name is sbox.jpgThe on-line version of this article (available at contains supplemental Methods and Figs. 1–7.

2The abbreviations used are:

protein kinase A
Bcl-2 interacting member protein
reverse transcription
terminal deoxynucleotidyltransferase-mediated biotin nick end-labeling
3-propionic acid
small interfering RNA
c-Jun N-terminal kinase
mitogen-activated protein kinase


1. Mattson M. P., Kroemer G. (2003) Trends Mol. Med. 9, 196–205 [PubMed]
2. D'Mello S. R., Galli C., Ciotti T., Calissano P. (1993) Proc. Natl. Acad. Sci. U.S.A. 90, 10989–10993 [PubMed]
3. Kobayashi Y., Shinozawa T. (1997) Brain Res. 778, 309–317 [PubMed]
4. Parvathenani L. K., Calandra V., Roberts S. B., Posmantur R. (2000) Neuroreport 11, 2293–2297 [PubMed]
5. Jonakait G. M., Ni L. (2009) Brain Res. 1285, 30–41 [PubMed]
6. Porat S., Simantov R. (1999) Ann. N.Y. Acad. Sci. 893, 372–375 [PubMed]
7. Hirotani S., Otsu K., Nishida K., Higuchi Y., Morita T., Nakayama H., Yamaguchi O., Mano T., Matsumura Y., Ueno H., Tada M., Hori M. (2002) Circulation 105, 509–515 [PubMed]
8. Iwatsubo K., Minamisawa S., Tsunematsu T., Nakagome M., Toya Y., Tomlinson J. E., Umemura S., Scarborough R. M., Levy D. E., Ishikawa Y. (2004) J. Biol. Chem. 279, 40938–40945 [PubMed]
9. Bos J. L. (2003) Nat. Rev. Mol. Cell Biol. 4, 733–738 [PubMed]
10. Roscioni S. S., Elzinga C. R., Schmidt M. (2008) Naunyn. Schmiedebergs Arch. Pharmacol. 377, 345–357 [PubMed]
11. Grandoch M., Bujok V., Fleckenstein D., Schmidt M., Fischer J. W., Weber A. A. (2009) J. Leukoc. Biol. 86, 847–849 [PubMed]
12. Tiwari S., Felekkis K., Moon E. Y., Flies A., Sherr D. H., Lerner A. (2004) Blood 103, 2661–2667 [PubMed]
13. McPhee I., Gibson L. C., Kewney J., Darroch C., Stevens P. A., Spinks D., Cooreman A., MacKenzie S. J. (2005) Biochem. Soc. Trans. 33, 1330–1332 [PubMed]
14. Wang C., Gu Y., Li G. W., Huang L. Y. (2007) J. Physiol. 584, 191–203 [PubMed]
15. Murray A. J., Shewan D. A. (2008) Mol. Cell. Neurosci. 38, 578–588 [PubMed]
16. Ulucan C., Wang X., Baljinnyam E., Bai Y., Okumura S., Sato M., Minamisawa S., Hirotani S., Ishikawa Y. (2007) Am. J. Physiol. Heart Circ. Physiol. 293, H1662–H1672 [PubMed]
17. Yokoyama U., Patel H. H., Lai N. C., Aroonsakool N., Roth D. M., Insel P. A. (2008) Proc. Natl. Acad. Sci. U.S.A. 105, 6386–6391 [PubMed]
18. Yokoyama U., Minamisawa S., Quan H., Akaike T., Jin M., Otsu K., Ulucan C., Wang X., Baljinnyam E., Takaoka M., Sata M., Ishikawa Y. (2008) Am. J. Physiol. Heart Circ. Physiol. 295, H1547–H1555 [PubMed]
19. Okumura S., Kawabe J., Yatani A., Takagi G., Lee M. C., Hong C., Liu J., Takagi I., Sadoshima J., Vatner D. E., Vatner S. F., Ishikawa Y. (2003) Circ. Res. 93, 364–371 [PubMed]
20. Yamashita N., Uchida Y., Ohshima T., Hirai S., Nakamura F., Taniguchi M., Mikoshiba K., Honnorat J., Kolattukudy P., Thomasset N., Takei K., Takahashi T., Goshima Y. (2006) J. Neurosci. 26, 13357–13362 [PubMed]
21. Geng Y. J., Ishikawa Y., Vatner D. E., Wagner T. E., Bishop S. P., Vatner S. F., Homcy C. J. (1999) Circ. Res. 84, 34–42 [PubMed]
22. Biju M. P., Akai Y., Shrimanker N., Haase V. H. (2005) Am. J. Physiol. Renal Physiol. 289, F1217–F1226 [PubMed]
23. Yokoyama U., Minamisawa S., Adachi-Akahane S., Akaike T., Naguro I., Funakoshi K., Iwamoto M., Nakagome M., Uemura N., Hori H., Yokota S., Ishikawa Y. (2006) Am. J. Physiol. Heart Circ. Physiol. 290, H1660–H1670 [PubMed]
24. Ishikawa N., Shimada N., Munakata Y., Watanabe K., Kimura N. (1992) J. Biol. Chem. 267, 14366–14372 [PubMed]
25. Iwatsubo K., Suzuki S., Li C., Tsunematsu T., Nakamura F., Okumura S., Sato M., Minamisawa S., Toya Y., Umemura S., Ishikawa Y. (2007) Am. J. Physiol. Cell Physiol 293, C1498–C1508 [PubMed]
26. Brouillet E., Jacquard C., Bizat N., Blum D. (2005) J. Neurochem. 95, 1521–1540 [PubMed]
27. Christensen A. E., Selheim F., de Rooij J., Dremier S., Schwede F., Dao K. K., Martinez A., Maenhaut C., Bos J. L., Genieser H. G., Døskeland S. O. (2003) J. Biol. Chem. 278, 35394–35402 [PubMed]
28. Chae I. H., Park K. W., Kim H. S., Oh B. H. (2004) Clin. Chim. Acta 341, 83–91 [PubMed]
29. Dong J. W., Zhu H. F., Zhu W. Z., Ding H. L., Ma T. M., Zhou Z. N. (2003) Cell Res. 13, 385–391 [PubMed]
30. Communal C., Colucci W. S. (2005) Arch. Mal. Coeur. Vaiss. 98, 236–241 [PubMed]
31. Wang X., Tang X., Li M., Marshall J., Mao Z. (2005) J. Biol. Chem. 280, 16705–16713 [PubMed]
32. Puthalakath H., Strasser A. (2002) Cell Death Differ. 9, 505–512 [PubMed]
33. Putcha G. V., Moulder K. L., Golden J. P., Bouillet P., Adams J. A., Strasser A., Johnson E. M. (2001) Neuron 29, 615–628 [PubMed]
34. Guo J., Gertsberg Z., Ozgen N., Steinberg S. F. (2009) Circ. Res. 104, 660–669 [PMC free article] [PubMed]
35. O'Reilly L. A., Cullen L., Visvader J., Lindeman G. J., Print C., Bath M. L., Huang D. C., Strasser A. (2000) Am. J. Pathol. 157, 449–461 [PubMed]
36. Biswas S. C., Shi Y., Sproul A., Greene L. A. (2007) J. Biol. Chem. 282, 29368–29374 [PubMed]
37. Cai B., Xia Z. (2008) Apoptosis 13, 803–810 [PMC free article] [PubMed]
38. Adams J. M., Cory S. (1998) Science 281, 1322–1326 [PubMed]
39. Pandey M., Varghese M., Sindhu K. M., Sreetama S., Navneet A. K., Mohanakumar K. P., Usha R. (2008) J. Neurochem. 104, 420–434 [PubMed]
40. Hoyt K. R., Gallagher A. J., Hastings T. G., Reynolds I. J. (1997) Neurochem. Res. 22, 333–340 [PubMed]
41. Whittemore E. R., Loo D. T., Watt J. A., Cotman C. W. (1995) Neuroscience 67, 921–932 [PubMed]
42. Li M., Wang X., Meintzer M. K., Laessig T., Birnbaum M. J., Heidenreich K. A. (2000) Mol. Cell. Biol. 20, 9356–9363 [PMC free article] [PubMed]
43. Chun H., Hao W., Honghai Z., Ning L., Yasong W., Chen D. (2009) Brain Res. 1257, 75–88 [PubMed]
44. O'Driscoll C., Wallace D., Cotter T. G. (2007) J. Neurochem. 103, 860–870 [PubMed]
45. Pugh P. C., Margiotta J. F. (2006) Mol. Cell. Neurosci. 31, 586–595 [PubMed]
46. Grandoch M., López de Jesús M., Oude Weernink P. A., Weber A. A., Jakobs K. H., Schmidt M. (2009) Cell Signal 21, 609–621 [PubMed]
47. Kwon G., Pappan K. L., Marshall C. A., Schaffer J. E., McDaniel M. L. (2004) J. Biol. Chem. 279, 8938–8945 [PubMed]
48. Kwak H. J., Park K. M., Choi H. E., Chung K. S., Lim H. J., Park H. Y. (2008) Cell. Signal. 20, 803–814 [PubMed]
49. Kaufmann T., Jost P. J., Pellegrini M., Puthalakath H., Gugasyan R., Gerondakis S., Cretney E., Smyth M. J., Silke J., Hakem R., Bouillet P., Mak T. W., Dixit V. M., Strasser A. (2009) Immunity 30, 56–66 [PMC free article] [PubMed]
50. Puthalakath H., O'Reilly L. A., Gunn P., Lee L., Kelly P. N., Huntington N. D., Hughes P. D., Michalak E. M., McKimm-Breschkin J., Motoyama N., Gotoh T., Akira S., Bouillet P., Strasser A. (2007) Cell 129, 1337–1349 [PubMed]
51. Oppenheim R. W. (1991) Annu. Rev. Neurosci. 14, 453–501 [PubMed]
52. Wallace D. C. (1999) Science 283, 1482–1488 [PubMed]
53. Mandavilli B. S., Boldogh I., Van Houten B. (2005) Brain Res. Mol. Brain Res. 133, 215–223 [PubMed]
54. Müller T., Meyer H. E., Egensperger R., Marcus K. (2008) Prog. Neurobiol. 85, 393–406 [PubMed]
55. Zhang L., Insel P. A. (2004) J. Biol. Chem 279, 20858–20865 [PubMed]

Articles from The Journal of Biological Chemistry are provided here courtesy of American Society for Biochemistry and Molecular Biology