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Molecular Endocrinology, doi:10.1210/me.2006-0340
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Molecular Endocrinology 21 (1): 159-171
Copyright © 2007 by The Endocrine Society

Cyclic Adenosine 5'-Monophosphate-Stimulated Neurotensin Secretion Is Mediated through Rap1 Downstream of both Epac and Protein Kinase A Signaling Pathways

Jing Li, Kathleen L. O’Connor, Xiaodong Cheng, Fang C. Mei, Tatsuo Uchida, Courtney M. Townsend, Jr and B. Mark Evers

Department of Surgery (J.L., K.L.O., C.M.T., B.M.E.), Sealy Center for Cancer Cell Biology (J.L., K.L.O., X.C., B.M.E.), Department of Pharmacology and Toxicology (X.C., F.C.M.), and Office of Biostatistics (T.U.), The University of Texas Medical Branch, Galveston, Texas 77555

Address all correspondence and requests for reprints to: B. Mark Evers, M.D., Department of Surgery, The University of Texas Medical Branch, 301 University Boulevard, Galveston, Texas 77555-0536. E-mail: mevers{at}utmb.edu.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Neurotensin (NT), a gut peptide, plays important roles in gastrointestinal secretion, inflammation, and growth of normal and neoplastic tissues. cAMP regulates the secretion of hormones via its effector proteins protein kinase A (PKA) or Epac (exchange protein directly activated by cAMP). The small GTPase Rap1 can be activated by both PKA and Epac; however, the role of Rap1 in hormone secretion is unknown. Here, using the BON human endocrine cell line, we found that forskolin (FSK)-stimulated NT secretion was reduced by inhibition of Rap1 expression and activity. FSK-stimulated NT secretion was enhanced by overexpression of either wild-type or constitutively active Rap1. Epac activators and wild-type Epac enhanced NT release and Rap1 activity. In contrast, overexpression of a cAMP binding mutant, EpacR279E, decreased NT release and Rap1 activity. PKA activation increased NT release and Rap1 activity. FSK-stimulated NT release was reduced by PKA inhibition and the dominant negative Rap1N17. NT secretion, stimulated by Epac activation, was reduced by PKA inhibition; NT release, stimulated by PKA activation, was enhanced by wild-type Epac but reduced by the mutant EpacR279E. Finally, prostaglandin E2 (PGE2), a physiological agent that increases cAMP, stimulated NT secretion via cAMP/PKA/Rap1. Importantly, we demonstrate that PKA and Epac mediate the cAMP-induced NT secretion synergistically by converging at the common downstream target protein Rap1. Moreover, PGE2, a potent mediator of inflammation and associated with colorectal carcinogenesis, stimulates NT release suggesting a possible link between PGE2 and NT on intestinal inflammatory disorders and colorectal cancers.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
NEUROTENSIN (NT), A GASTROINTESTINAL hormone, is localized to enteroendocrine cells (N cells) found predominantly in the distal small bowel and is released by intraluminal fats (1, 2). NT facilitates fatty acid translocation (3), affects gut motility (4), and stimulates growth of normal gut mucosa (5, 6) as well as certain pancreatic, colonic, and prostatic cancers bearing NT receptors (NTR) (7, 8). NT, acting through the high-affinity NTR (NTR1), is a known proinflammatory peptide in the colon with increased expression of NT and NTR1 noted in the inflamed mucosa of rats after exposure to Clostridium difficile toxin A; treatment with the NTR1 receptor antagonist decreased inflammation and prostaglandin E2 (PGE2) secretion (9, 10, 11).

Our laboratory is focused on delineating the signaling mechanisms regulating NT secretion. In this regard, we utilize a novel human endocrine cell line, BON, which expresses the NT/neuromedin N gene, synthesizes and secretes NT peptide, and processes the NT/neuromedin N peptide in a manner analogous to that of N cells in the small bowel (12). Using the BON cell line, we have shown that protein kinase C (PKC), particularly PKC isoforms-{alpha} and -{delta}, plays a role in the release of NT mediated by the phorbol ester, phorbol 12-myristate 13-acetate (PMA) (13, 14). Furthermore, we found that protein kinase D, a serine/threonine protein kinase that is structurally distinct from the PKC family members, and the Rho/Rho kinase are involved in PMA-mediated NT secretion as well as NT secretion mediated by bombesin, the amphibian equivalent of gastrin-releasing peptide, which is a physiological stimulant of NT release in vivo (15). We have previously reported that forskolin (FSK), an adenylyl cyclase activator that increases intracellular cAMP levels, stimulated NT secretion from BON cells (16); however, the precise signaling molecules involved in this process are not known.

cAMP has been implicated in the secretion of certain hormones (17, 18); protein kinase A (PKA) was thought to be the principal effector for cAMP in mammalian cells (19, 20) and in cAMP-regulated exocytosis in secretory cells (21). Recently, Epac (exchange protein directly activated by cAMP), a guanine nucleotide-exchange factor activated by cAMP, has been identified as a direct cAMP target (22, 23, 24). The Epac family of proteins, including Epac1 and Epac2, contains a functional cAMP-binding domain and a guanine nucleotide-exchange factor domain that activates a small GTPase Rap1 in response to cAMP (22, 23). Epac regulates several important biological functions, such as integrin-dependent cell adhesion, insulin secretion, and calcium release (25). By utilizing the cAMP analog 8-pCPT-2'-O-Me-cAMP, Epac was first implicated in the regulation of insulin secretion in ß-cells (26). Furthermore, the dominant negative Epac2 blocks stimulatory actions of glucagon-like peptide-1 and 8-pCPT-2'-O-Me-cAMP on Ca2+ signaling and exocytosis in ß-cells (27, 28, 29). Using either antisense oligonucleotides directed against Epac2 mRNA or expression of a mutant Epac2 protein defective for cAMP binding, Epac2 was shown to be responsible for the cAMP effect in the incretin response (30).

The Rap family of small GTPases are involved in cell adhesion, proliferation, and differentiation (31). In humans, four Rap isoforms, Rap1A, Rap1B, Rap2A, and Rap2B, have been identified. Rap1 cycles between a GDP-bound inactive and a GTP-bound active form; this switching is regulated by specific guanine nucleotide exchange factors and GTPase activating proteins. Rap1 is rapidly activated by a multitude of stimuli, including growth factors, hormones, neurotransmitters, and cytokines via common second messengers such as calcium, diacylglycerol, and cAMP/PKA (25, 31, 32). Epac catalyzes the exchange of GTP for GDP and activates Rap1 (33, 34), which is independent of PKA. In addition, PKA can phosphorylate Rap1 and is necessary for Rap1 activation in certain cell types (35, 36, 37, 38). However, Rap1-dependent signaling and function are not well understood.

The purpose of our present study was to determine whether cAMP/PKA or cAMP/Epac pathways are involved in NT secretion and, whether Rap1 is downstream of these pathways. Here, we demonstrate that Rap1 promotes NT secretion from the human endocrine cell line BON and serves as the common effector protein, downstream of both PKA and Epac. Moreover, we demonstrate a synergistic effect of PKA and Epac in cAMP-regulated NT secretion. We also show that PGE2, a G protein-coupled receptor (GPCR) agonist, potentiates NT secretion through the cAMP/PKA/Rap1 pathway, suggesting that cAMP is a physiological regulator of NT secretion in the intestine.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Inhibition of Rap1 Decreases cAMP-Stimulated NT Secretion
Rap1 is a downstream signaling molecule that may be activated by either Epac (33, 34) or PKA (37, 38, 39). We demonstrated that FSK stimulated NT secretion from BON cells (16). BON cells express high levels of Rap1 (data not shown); therefore, we sought to determine whether Rap1 is involved in cAMP-mediated NT release (Fig. 1Go). For this purpose, Rap1 GTPase-activating protein II (Rap1GAPII), which specifically catalyzes the hydrolysis of GTP to GDP and inactivates Rap1 activity (40), was transiently overexpressed in BON cells. The overexpression of Rap1GAPII dramatically abrogated FSK-stimulated NT secretion compared with the control vector (pMT2) (Fig. 1AGo, top panel). Furthermore, FSK-stimulated NT secretion in BON cells, transfected with Rap1A small interfering RNA (siRNA), was significantly decreased compared with BON cells transfected with the nontargeting control siRNA (Fig. 1BGo, top panel). Rap1 protein expression was suppressed by Rap1A siRNA, as determined by Western blotting (Fig. 1BGo, bottom panel).


Figure 1
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  		Fig. 1. Inhibition of Rap1 Decreases FSK-Stimulated NT Secretion

A, BON cells in 12-well plates were transiently transfected with an HA-tagged Rap1GAPII plasmid or the empty vector (pMT2) and cultured for 48 h. Cells were treated with FSK for 30 min, and NT secretion was measured (top). Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. empty vector; {dagger}, P < 0.05 vs. empty vector plus FSK. The cells were lysed, and overexpression of Rap1GAPII was detected using anti-HA antibody (bottom). B, BON cells were transfected with either 50 nM of control or Rap1A siRNA by electroporation. The cells were trypsinized the next day and plated in 24-well plates. Three days after transfection, cells were treated with FSK for 30 min, and NT secretion was measured (top). Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control siRNA; {dagger}, P < 0.05 vs. control siRNA plus FSK. The cells were lysed, and expression of Rap1 was detected using anti-Rap1 antibody (bottom).

 
Wild-Type and Constitutively Active Rap1A Enhance but Dominant Negative Rap1A Inhibits FSK-Stimulated NT Release
To further confirm the role of Rap1 in cAMP-mediated NT secretion, BON cells stably expressing wild-type Rap1A (Rap1A WT), constitutively active Rap1 (Rap1A V12), and dominant negative Rap1 (Rap1A N17) were established. Overexpression of either Rap1A WT (Fig. 2AGo, left) or Rap1A V12 (Fig. 2AGo, middle) dramatically enhanced FSK-stimulated NT release. In contrast, FSK-stimulated NT secretion was markedly inhibited in Rap1A N17 cells (Fig. 2AGo, right). Next, the expression of green fluorescent protein (GFP)-tagged Rap1A in BON cells was analyzed by confocal microscopy (Fig. 2BGo). Rap1A WT localized in the membrane as well as in the cytosol (Fig. 2BGo, left). The constitutively active Rap1A V12 demonstrated a similar expression pattern as Rap1A WT (Fig. 2BGo, middle). In contrast, dominant negative Rap1A N17 was not localized to the membrane but entirely localized to the cytosol and the nucleus (Fig. 2BGo, right). Taken together, these results further provide evidence for the involvement of Rap1 in cAMP-mediated NT secretion.


Figure 2
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  		Fig. 2. Overexpression of the Rap1A WT and Constitutively Active Rap1A V12 Enhances, but the Dominant Negative Rap1A N17 Decreases FSK-Stimulated NT Release

A, BON cells, stably overexpressing Rap1A WT (left, top), were cultured in 24-well plates for 24 h. BON cells, stably overexpressing Rap1A V12 (middle, top) and Rap1A N17 (right, top), were cultured in 12-well and 24-well plates, respectively, for 48 h and treated with FSK for 30 min, and NT secretion was measured. Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. vector; {dagger}, P < 0.05 vs. vector plus FSK. The cells were lysed, and Western blotting was performed to monitor the overexpression of Rap1 using anti-GFP antibody (bottom). B, BON cells, stably overexpressing Rap1A WT (left), Rap1A V12 (middle), and Rap1A N17 (right) were plated on coverslips in 24-well plates and cultured for 48 h. GFP was detected by confocal microscopy.

 
Epac Stimulates cAMP-Mediated NT Secretion via Rap1
Epac activates Rap1 and is a downstream target of cAMP, independent of PKA (33, 34). Based on the involvement of Rap1 in cAMP-mediated NT secretion and the fact that BON cells express endogenous Epac1 mRNA and protein (data not shown), we next examined whether Epac regulates NT secretion and activates Rap1. To test whether the activation of endogenous Epac is sufficient to stimulate NT secretion, we used a cAMP analog, 8-pMeOPT-cAMP, which preferentially activates Epac without affecting PKA activity (41). 8-pMeOPT-cAMP significantly increased NT secretion in a dose-dependent manner (Fig. 3AGo, left). This effect was also confirmed using another Epac activator, 8-Br-O-cAMP (Fig. 3AGo, middle). Furthermore, BON cells were treated with the Epac activators 8-pMeOPT-cAMP and 8-Br-O-cAMP, and Rap1 pull-down assays were performed. Similar to FSK, the Epac activators induced Rap1 activation in BON cells (Fig. 3AGo, right). To elucidate further the involvement of Epac on cAMP-mediated NT release, we next determined whether Epac is involved in FSK-stimulated NT secretion in BON cells (Fig. 3BGo, left). FSK-stimulated NT secretion was significantly enhanced in BON cells transiently transfected with Epac wild-type (Epac WT) compared with BON cells transfected with empty vector. In addition, we performed Rap1 pull-down assays using cell lysates from BON cells stably expressing Epac WT treated with FSK; FSK-stimulated Rap1 activity was further enhanced in Epac WT cells (Fig. 3BGo, right). Moreover, cAMP-mediated NT secretion was inhibited in BON cells transiently transfected with the mutant EpacR279E (Fig. 3CGo, left), and, consistently, Rap1 activity was also decreased in EpacR279E cells (Fig. 3CGo, right). Collectively, these results demonstrate regulation of NT secretion by Epac via Rap1 activation in BON cells.


Figure 3
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  		Fig. 3. Epac Regulates NT Secretion via Rap1

A, BON cells, plated in 12-well plates for 24 h, were treated with either pMeOPT-cAMP (left) or 8-Br-O-cAMP (middle) for 1 h, and NT secretion was measured. Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control. Forty-eight hours after plating in 60-mm dishes, BON cells were treated with pMeOPT-cAMP (right, top) or 8-Br-O-cAMP (right, bottom) for 15 min, and Rap1 pull-down assay was performed. GDP and GTP{gamma}S were used as negative and positive controls, respectively. Protein (15 µl) from each lysate was used in the Western blotting to monitor for equal protein amount. B, BON cells, transiently transfected with Epac WT, were cultured in 12-well plates for 48 h and treated with FSK for 30 min, and NT secretion was measured (left, top). Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. vector; {dagger}, P < 0.05 vs. vector plus FSK. The cells were lysed, and Western blotting was performed to monitor the overexpression of Epac using anti-GFP antibody (left, bottom). BON cells, stably overexpressing the Epac WT, were cultured for 48 h in 60-mm dishes and treated with FSK for 15 min, and Rap1 pull-down assay was performed (right, top). The anti-Rap1 and -GFP antibodies were used to monitor for equal protein amount (right, middle) and overexpression of Epac (right, bottom), respectively. C, BON cells, transiently transfected with EpacR279E, were cultured for 48 h in 12-well plates and treated with FSK for 30 min, and NT secretion was measured (left, top). Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. vector; {dagger}, P < 0.05 vs. vector plus FSK. The cells were lysed, and Western blotting was performed to monitor the overexpression of the mutant Epac using anti-Flag antibody (left, bottom). BON cells, stably overexpressing EpacR279E, were cultured for 48 h and treated with FSK for 15 min, and Rap1 pull-down assay was performed (right, top). The anti-Rap1 and -Flag antibodies were used to monitor for equal protein amount (right, middle) and overexpression of Epac (right, bottom), respectively.

 
PKA Stimulates NT Secretion via Rap1
PKA is a well-characterized downstream effector of cAMP and is thought to be involved in cAMP-enhanced secretion (21). Rap1 is phosphorylated and activated by PKA in neutrophils and platelets (37, 38, 39). Thus, we first investigated the involvement of the cAMP/PKA pathway in the regulation of NT release from BON cells stimulated by FSK (Fig. 4Go). We used the potent PKA activators, N6-phenyladenosine-3', 5'-cyclic monophosphate (6-Phe-cAMP) and N6-benzoyladenosine-3', 5'-cyclic monophosphate (6-Bnz-cAMP), to activate BON cells to determine whether PKA plays a role in NT secretion. Both 6-Phe-cAMP (Fig. 4AGo, left) and 6-Bnz-cAMP (Fig. 4AGo, right) increased NT secretion in a dose-dependent manner. Furthermore, we determined whether FSK- or 6-Phe-cAMP-stimulated NT release is altered by pretreatment with a selective PKA chemical inhibitor, H89 (42). As shown in Fig. 4BGo (left), FSK- and 6-Phe-cAMP-induced NT secretion was significantly inhibited by H89 in BON cells. To verify the specific role of H89, BON cells were pretreated with the peptide protein kinase inhibitor (PKI) (14, 15, 16, 17, 18, 19, 20, 21, 22), a myristoylated PKA inhibitor, and then treated with FSK. Similar to results using H89, FSK-induced NT secretion was significantly inhibited by PKI treatment (Fig. 4BGo, right). Next, we examined whether PKA regulated NT secretion via Rap1. For this purpose, we first tested whether PKA also induced Rap1 activation using Rap1 activity assays in vitro. BON cells were treated with the potent PKA activator 6-Phe-cAMP, and Rap1 pull-down assays were performed. 6-Phe-cAMP, at concentrations which significantly stimulated NT secretion, activated Rap1 activity (Fig. 4CGo, left). Furthermore, we treated BON cells stably transfected with Rap1A N17 using 6-Phe-cAMP (Fig. 4CGo, right). 6-Phe-cAMP-induced NT secretion was significantly inhibited in cells expressing Rap1A N17. Taken together, these results suggest a cAMP/PKA/Rap1-dependent signaling pathway that is involved in cAMP-induced NT secretion in BON cells.


Figure 4
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  		Fig. 4. PKA Regulates FSK-Stimulated NT Secretion via Rap1

A, BON cells, plated in 12-well plates for 24 h, were treated with either 6-Phe-cAMP (left) or 6-Bnz-cAMP (right) for 1 h. NT secretion was measured. Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control. B, Forty-eight hours after plating in 12-well plates, BON cells were pretreated with either H89 (left) or PKI (right) for 30 min, followed by combination treatment of either FSK or 6-Phe-cAMP and inhibitors for another 1 h. NT secretion was measured. Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control; {dagger}, P < 0.05 vs. FSK or 6-Phe-cAMP alone. C, Forty-eight hours after plating, BON cells were treated with 6-Phe-cAMP for 15 min, and Rap1 pull-down assay was performed (left, top); total Rap1 was examined as a control for equal protein amount (left, bottom). Twenty-four hours after plating in 24-well plates, BON cells, stably overexpressing Rap1A N17, were treated with 6-Phe-cAMP for 1 h, and NT release was measured (right panel). Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. vector; {dagger}, P < 0.05 vs. vector plus 6-Phe-cAMP.

 
Interaction of PKA and Epac Pathways
PKA and Epac pathways coexist and function independently (30, 43). To test whether stimulation of the PKA and Epac pathways can synergistically enhance NT secretion, BON cells were treated with either the PKA activator 6-Phe-cAMP or the Epac activator pMeOPT-cAMP, alone or in combination, and NT secretion was measured (Fig. 5AGo). Treatment with either activator alone significantly stimulated NT release; the PKA activator 6-Phe-cAMP appeared to be a more potent stimulator of NT release compared with the Epac activator pMeOPT-cAMP. Combination treatment with both activators produced a synergistic effect on NT secretion. To determine further whether PKA and Epac pathways act dependently or independently, BON cells were pretreated with H89 (10 µM) for 30 min, followed by combination treatment with H89 and the Epac activator pMeOPT-cAMP for another 1 h; NT release was measured. Consistent with the results shown in Fig. 4BGo (left), FSK-stimulated NT secretion was significantly decreased by H89 treatment (Fig. 5BGo, left). Importantly, NT release induced by the Epac activator was also significantly inhibited by H89 treatment. To confirm further the inhibitory effects of H89 on Epac activator-stimulated NT secretion, BON cells were pretreated with PKI for 30 min, followed by combination treatment of PKI and pMeOPT-cAMP (Fig. 5BGo, right). Consistent with the results using H89, PKI also decreased Epac activator-stimulated NT release.


Figure 5
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  		Fig. 5. Interaction of PKA and Epac Pathways

A, BON cells, plated in 12-well plates for 48 h, were treated with either pMeOPT-cAMP or 6-Phe-cAMP alone or with the combination of both for 1 h. NT release was measured. Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control; {dagger}, P < 0.05 vs. either pMeOPT-cAMP or 6-Phe-cAMP alone. B, Forty-eight hours after plating in 24-well plates, BON cells were pretreated with either H89 (left) or PKI (right) for 30 min, followed by combination treatment of H89 or PKI with either FSK or pMeOPT-cAMP for another 1 h. NT release was measured. Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control; {dagger}, P < 0.05 vs. either FSK or pMeOPT-cAMP alone. C, BON cells, transiently transfected with Epac WT, were cultured for 48 h in 12-well plates and treated with 6-Phe-cAMP for 1 h. NT secretion was measured (top). Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. vector; {dagger}, P < 0.05 vs. vector plus 6-Phe-cAMP. The cells were lysed, and Western blotting was performed using anti-GFP antibody to monitor the Epac overexpression (bottom). D, BON cells, transiently transfected with EpacR279E, were cultured for 48 h in 12-well plates and treated with pMeOPT-cAMP or 6-Phe-cAMP for 1 h. NT secretion was measured (top). Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. vector; {dagger}, P < 0.05 vs. vector or vector plus pMeOPT-cAMP or 6-Phe-cAMP. The cells were lysed, and Western blotting was performed using anti-Flag antibody to monitor Epac overexpression (bottom).

 
Next, BON cells were transiently transfected with Epac WT and treated with the PKA activator 6-Phe-cAMP (Fig. 5CGo). Compared with the control vector, 6-Phe-cAMP-stimulated NT secretion was further enhanced in cells overexpressing Epac WT, demonstrating the interaction of PKA and Epac. Furthermore, NT secretion was significantly reduced in BON cells transfected with the mutant EpacR279E and treated with either 6-Phe-cAMP or pMeOPT-cAMP compared with BON cells transfected with the control vector (Fig. 5DGo). These results demonstrate that Epac is partially dependent on PKA in cAMP-stimulated NT secretion in BON cells.

PGE2 Stimulates NT Secretion via a PKA/Rap1 Pathway
The BON cell line is similar in many respects to the terminally differentiated N cell of the small bowel. In this regard, we used PGE2, a physiological agent that increases cAMP levels in certain cells including intestinal cells (44), to confirm further the role of cAMP on NT release. We treated BON cells with PGE2 and measured NT release. As shown in Fig. 6AGo (left), PGE2 significantly enhanced NT secretion in a dose-dependent manner. PGE2 binds to the E prostanoid (EP) receptors, including EP1, EP2, EP3, and EP4. BON cells express all four types of EP receptors as determined by RT-PCR (data not shown). We further discriminated which of the PGE2 receptors actually regulate NT release. For this purpose, BON cells were treated with various PGE2 receptor agonists, and NT secretion was measured. Compared with the control cells, sulprostone, an EP3 activator, did not induce NT secretion, suggesting that EP3 is not involved in PGE2-mediated NT release. PGE1 alcohol (EP3 and EP4 receptor agonist), 17-phenyl trinor PGE2 (EP1 and EP3 agonist), and 11-deoxy-16,16-dimethyl PGE2 (EP2 and EP3 agonist) potently stimulated NT release (Fig. 6AGo, right), suggesting the involvement of EP1, EP2, and EP4 receptors. To examine whether the effect of PGE2 is through a cAMP/PKA-dependent pathway, downstream of EP2 and EP4 receptors, BON cells were pretreated with either H89 or PKI and then treated with PGE2. PGE2-stimulated NT secretion was significantly inhibited by both H89 (Fig. 6BGo, left) and PKI (Fig. 6BGo, right). To confirm induction of cAMP by PGE2, BON cells were treated with various concentrations of PGE2, and cAMP levels were measured; levels of cAMP were increased with PGE2 treatment in a dose-dependent fashion (data not shown). To determine whether the increase in NT release by PGE2 occurs through Rap1 activation, we first assessed Rap1 activity in BON cells treated with PGE2; PGE2 strongly stimulated Rap1 activity (Fig. 6CGo, left). Moreover, BON cells were transfected with Rap1A siRNA, and NT secretion was measured after PGE2 treatment. PGE2-stimulated NT secretion was significantly decreased by Rap1A siRNA (Fig. 6CGo, right).


Figure 6
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  		Fig. 6. PGE2 Stimulates NT Secretion via cAMP/PKA/Rap1 Pathway

A, BON cells, plated in 24-well plates for 48 h, were treated with PGE2 at various concentrations (left) for 1 h or in 12-well plates, BON cells were treated with PGE2 receptor agonists (right) for 1 h, and NT secretion was measured. Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control. B, Twenty-four hours after plating in 12-well plates, BON cells were pretreated with either H89 (left) or PKI (right) for 30 min, followed by combination treatment of PGE2 and inhibitors for 1 h. NT secretion was measured. Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control; {dagger}, P < 0.05 vs. PGE2 only. C, Forty-eight hours after plating in 60-mm dishes, BON cells were treated with PGE2 as well as FSK for 15 min, and Rap1 pull-down assay was performed (left, top); total Rap1 was detected to monitor for equal protein amount (left, bottom). BON cells were transfected with 50 nM of Rap1A siRNA as well as the control siRNA. Cells were trypsinized and plated in 24-well plates for 72 h. Cells were treated with PGE2 for 1 h. NT secretion was measured (right, top). Results are expressed as mean ± SE (n = 6). *, P < 0.05 vs. control siRNA; {dagger}, P < 0.05 vs. control siRNA plus PGE2. Western blotting was performed to monitor the expression of Rap1 using an anti-Rap1 antibody; ß-actin was probed as a loading control (right, bottom).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
The cAMP signaling pathway, acting through its downstream effector proteins PKA and Epac, regulates a wide array of cellular processes, including proliferation, differentiation, and secretion (21, 25, 45, 46, 47). In our current study, we show that both cAMP/PKA and cAMP/Epac pathways are involved in FSK-mediated NT secretion in BON endocrine cells. We demonstrate the inhibition of NT secretion using the PKA inhibitors, H89 and PKI, and the increase of NT release with specific PKA activators. We also demonstrate that Epac plays a positive role in the regulation of cAMP-stimulated NT secretion. More importantly, we have shown that combination treatment of both PKA and Epac activators synergistically enhanced NT secretion. NT release, stimulated by the Epac activator, was reduced by H89, a potent and selective inhibitor of PKA (42), and PKI, a highly specific inhibitor of PKA (48, 49). Additionally, PKA activator-induced NT release was decreased in BON cells overexpressing the cAMP binding mutant EpacR279E. Therefore, using the human endocrine cell line as a model, we demonstrate the presence of both functional cAMP/PKA and cAMP/Epac pathways that stimulate NT release in a synergistic fashion.

Both PKA and Epac have been implicated in cAMP-stimulated secretion of glucagon in pancreatic {alpha}-cells (50) and insulin from ß-cells (18, 26). According to previous studies (51), both the cAMP/PKA and cAMP/Epac pathways coexist in cells but function independently. For example, estrogen increases hormonal output in mouse melanotrophs from fresh pituitary tissue slices through both cAMP/PKA- and cAMP/Epac-dependent mechanisms that are independent of each other (51). The intestinal hormone glucagon-like peptide-1 independently stimulates both PKA and Epac, resulting in insulin secretion (52). In nonendocrine cells, such as vascular endothelial cells, both the cAMP/PKA and cAMP/Epac signaling pathways independently regulate barrier function (53, 54). Epac2 was shown to mediate PKA-independent insulin exocytosis in mouse pancreatic ß-cells (30, 43). The proposed explanation is that PKA is more targeted to the plasma membrane and exerts a stimulatory effect on docked secretory granules (55, 56), whereas Epac associates with secretory granule membranes and mediates stimulatory effects of cAMP on granule priming (29). In addition, PKA binds cAMP with a greater affinity than Epac, suggesting that Epac can further enhance cAMP signaling when PKA is already saturated (25).

Rap1, located on secretory granule membranes (57), is expressed in ß-cells in which Epac regulates glucose-dependent insulin secretion (58). Yang et al. (59) first suggested that Rap1 might be involved in norepinephrine release from rat pheochromocytoma PC12 cells. In a pancreatic ß-cell line (ß-TC), Rap1 translocated from the cytosol to particulate fractions in response to GTP{gamma}S (58). Therefore, these findings, albeit indirect, suggest that Rap1 plays a role in exocytosis downstream of Epac or PKA in the regulation of hormone secretion. Other studies, in nonendocrine secretory cells, showed that Epac acted through Rap1 to regulate cAMP-dependent processing of amyloid precursor protein, a key protein in Alzheimer’s disease, and secretion of the nonamyloidogenic soluble form of amyloid precursor protein (60). D’Silva et al. (57) showed the localization and high concentration of Rap1 on secretory granule membranes in rat parotid acinar cells. In addition, previous studies have shown that Rap1 is activated by cAMP/PKA (37, 38, 39). PKA phosphorylation of Src was required for Rap1 activation in PC12 cells (35). The PKA-dependent effects of cAMP on growth inhibition were mediated through Rap1 (36). In platelets, both Rap1A and Rap1B are phosphorylated by PKA and are necessary for Rap1 activation (38). Phosphorylation of Rap1B by PKA did not affect its basal GDP/GTP exchange reaction or the GAP-stimulated GTPase activity (38, 61, 62). However, there are limited studies that actually link cAMP, Epac, and Rap1 to protein secretion.

In our study, we inhibited endogenous Rap1 activity and expression by Rap1GAPII, a Rap1 GTPase-activating protein that inactivates Rap1, and Rap1 siRNA, respectively, and found that cAMP-mediated NT secretion was significantly attenuated. FSK-stimulated NT secretion was enhanced in BON cells stably overexpressing either the Rap1A WT or the constitutively active Rap1A V12 but inhibited in BON cells stably overexpressing the dominant negative Rap1A N17. Importantly, we found that both PKA and Epac enhanced Rap1 activity and PKA activator-stimulated NT secretion was reduced in BON cells overexpressing the dominant negative Rap1A N17. Therefore, we provide strong evidence supporting the role of Rap1 in hormone secretion, downstream of PKA and Epac pathways. Rap1, as a common effector, converges these two pathways to facilitate release of NT.

PGE2, a product of arachidonate metabolism, is released locally and acts in an autocrine or paracrine fashion (63) to affect a variety of physiological and pathological processes in the gastrointestinal tract (64). For example, PGE2 is a potent mediator of inflammation and has been clearly linked to colorectal carcinogenesis. In addition, PGE2 is involved in the regulation of cAMP-mediated chloride secretion in intestinal epithelial cells (65). PGE2 exerts its biological effects by binding to its membrane receptors; four subtypes of EP receptors have been described, EP1, EP2, EP3, and EP4, encoded by different genes. Each subtype uses different intracellular signaling mechanisms. Binding to EP1 receptor increases intracellular Ca2+ and activates PKC. EP2 and EP4 activate Gs, which stimulates cAMP generation, followed by the activation of PKA. Conversely, EP3 acts via Gi to inhibit cAMP generation (66). We found that PGE2 stimulates NT secretion through the cAMP/PKA pathway, and most likely involving the EP2/EP4 receptors. We found that Rap1 activity was significantly increased by PGE2, suggesting that the effect of PGE2 was mediated via cAMP, PKA, and Rap1. Consistent with our present study, it has been reported that cAMP, produced by Gs-linked GPCR, can activate Rap proteins to regulate certain cell functions, such as proliferation (37, 67, 68, 69, 70). Because both PGE2 and NT are mediators of intestinal inflammation (9, 10, 11, 64) and have been shown to promote colon cancer cell growth (71), our findings suggest that PGE2-regulated NT release may play a role in intestinal inflammation and carcinogenesis.

In conclusion, we found that both the cAMP/PKA and cAMP/Epac pathways are involved in cAMP-mediated NT release in the BON endocrine cell line (summarized in Fig. 7Go). We also show that Rap1 plays a critical role as a common effector protein for both the cAMP/Epac and cAMP/PKA pathways and converges the two pathways in mediating hormone release. Previously, we showed that PKC-{alpha} and PKC-{delta} regulated PMA-stimulated NT secretion through protein kinase D1 and myristoylated alanine-rich C kinase substrate, respectively (13, 14, 15). In our current study, we identify an important role for cAMP/PKA/Rap1 and cAMP/Epac/Rap1 pathways in NT release. It is clear that multiple pathways contribute to intestinal hormone release with opportunities for cross talk between the various pathways. This system allows for altered cellular responses depending upon the stimuli for hormone release. Finally, our findings of PGE2-mediated NT secretion may have important clinical ramifications related to the pathogenesis of intestinal inflammatory disorders and colorectal cancers.


Figure 7
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  		Fig. 7. cAMP and Its Effectors Regulate NT Secretion

This diagram summarizes the results in our study demonstrating stimulation of NT release through both PKA and Epac acting via the common effector protein, Rap1. Gs, G proteins stimulate the formation of cAMP; AC, adenylate cyclase).

 

   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Materials
GFP-tagged Rap1A WT, the constitutively active Rap1A V12, and the dominant negative Rap1A N17 plasmids were provided by Dr. Mark R. Philips (New York University School of Medicine) (72). Rap1GAPII plasmid and the control vector pMT2 were provided by Dr. Johannes L. Bose (University Medical Center, Utrecht, The Netherlands) (73). H89 and PKI (14, 15, 16, 17, 18, 19, 20, 21, 22) were from BIOMOL Research Laboratories Inc. (Plymouth Meeting, PA). FSK was from Calbiochem (La Jolla, CA). 8-Br-2'-O-Me-cAMP, 8-pMeOPT-2'-O-Me-cAMP, 6-Phe-cAMP, and 6-Bnz-cAMP were from BIOLOG (Bremen, Germany). Sulprostone, 17-phenyl trinor PGE2 (17-phenyl), PGE1 alcohol, and 11-deoxy-16,16-dimethyl PGE2 (11-deoxy) were from Cayman Chemical Company (Ann Arbor, MI). PGE2, mouse monoclonal anti-GFP antibody (clone GFP-20), anti-Flag M2 monoclonal antibody, and anti-ß-actin antibody were from Sigma Chemical Co. (St. Louis, MO). The anti-hemagglutinin (HA) antibody (clone 3F10) was from Roche (Nutley, NJ). The SmartPool Rap1A siRNA and nontargeting siRNA #1 (control siRNA) were from Dharmacon, Inc. (Lafayette, CO). The antisecondary antibodies for Western blotting were from Pierce (Rockford, IL). The enhanced chemiluminescence (ECL) system for Western immunoblot analysis was from Amersham (Buckinghamshire, UK). The concentrated protein assay dye reagent was from Bio-Rad (Hercules, CA). Tissue culture media and reagents were from GIBCO-BRL (Grand Island, NY). All other reagents were of molecular biology grade and were purchased from Sigma.

Cells and Transfection
The BON cell line was derived from a human pancreatic carcinoid tumor and characterized in our laboratory (74). BON cells are maintained in a 1:1 mixture of DMEM and nutrient mixture, F12K, supplemented with 5% fetal bovine serum in 5% CO2 at 37 C. BON cells stably expressing the GFP-tagged Rap1A WT, the constitutively active Rap1A V12, and the dominant negative Rap1A N17 were established by culturing cells in the growth medium containing G418 (800 µg/ml); GFP-positive cells were sorted by flow cytometry. Stable BON cell lines overexpressing the Epac WT tagged with GFP, the mutant EpacR297E (defective in cAMP binding) tagged with Flag (75), and their control vectors pEGFP-N1 and pcDNA3, respectively, were established and cultured in the growth medium and supplemented with G418 (400 µg/ml). For some experiments, BON cells were transiently transfected with Epac WT, EpacR279E, Rap1GAPII, or the control vector pMT2 using Lipofectamine 2000 (Invitrogen, Carlsbad, CA). For experiments utilizing siRNA and for the establishment of stable cell lines, BON cells were transfected by electroporation (400V, 500 µF) using GenePulser XCell (Bio-Rad).

Cell Treatments and NT Enzyme Immunoassay
All experiments were performed 24–72 h after seeding cells as described in the figure legends. Before each experiment, BON cells were starved in serum-free medium for 5 h. For NT release experiments, BON cells were plated in 12- or 24-well plates and treated with the agonists in 0.5 or 0.2 ml, respectively, of serum-free medium per well for 30 or 60 min. For inhibitor experiments, cells were pretreated with inhibitors for 30 min, followed by the combination of inhibitors with the agonists for another 30 or 60 min. Medium was collected and stored at –80 C, and NT level in the medium was measured by NT enzyme immunoassay kit (Phoenix Pharmaceuticals, Inc., Belmont, CA).

Protein Preparation and Western Blotting
Protein preparation and Western blotting were performed as described previously (13). In brief, equal amounts of protein were resolved on NuPAGE Bis-Tris gels (Invitrogen) and electrophoretically transferred to polyvinylidene diflouride membranes; the membranes were incubated with primary antibodies overnight at 4 C, followed by secondary antibodies conjugated with horseradish peroxidase. Membranes were developed using the ECL detection system.

Confocal Microscopy
Cells were grown on glass coverslips in 24-well plates. The GFP fluorescence was observed under an LSM 510 META confocal system configured with an Axiovert 200 M inverted microscope (Zeiss, Jena, Germany). Images were acquired using a plan-apochromat 63x, 1.4 NA oil immersion objectives. The image acquisition and processing were carried out using the Zeiss LSM510 workstation (v. 3.0) and the Zeiss Image Browser (v. 3.1) software.

cAMP and Rap1-GTP Pull-Down Assays
The intracellular cAMP levels were measured by cAMP direct immunoassay kit (BioVision, Mountain View, CA). Rap1 activity was detected by a Rap1 activation assay kit (Upstate Biotechnology, Inc., Lake Placid, NY) following the modified method described by the manufacturer. Briefly, 5 x 104/cm2 cells were seeded on 60 mm-culture dishes and grown for 24 h. Cells starved in serum-free medium overnight were treated with various compounds. Cells were lysed with 500 µl of lysis buffer. For loading controls, a 15-µl aliquot of lysate was removed before bead addition. A bead volume of 30 µl was added. The beads were washed with PBS three times, and bound proteins were eluted in 30 µl of 2X Tris-glycine sodium dodecyl sulfate sample buffer. The samples were loaded onto a NuPAGE Novex 4–12% Bis-Tris gel electrophoresis and transferred to polyvinylidene diflouride membranes. The membranes were gently washed once with PBS for 5 min after primary or secondary antibody. The active Rap1 signals were detected using ECL Western Blotting Detection System.

Statistical Analysis
One- and two-factor experiments were used to analyze the data in our current study. The one-factor experiments were used for the dose studies (pMeOPT-cAMP, 8-Br-O-cAMP, 6-Phe-cAMP, 6-Bnz-cAMP, and PGE2) and PGE2 receptor agonist study. Due to heterogeneous variability among treatment groups, data were analyzed using the Kruskal-Wallis test. The two-factor experiments were used to analyze effects of combinations, especially significant synergistic effects of two factors. The two factor experiments were Rap1GAPII, Rap1A siRNA, Rap1A WT, V12 and N17, Epac WT, EpacR279E or H89 or PKI (present or control) and FSK (present or absent). Other combinations of the two factors were pMeOPT-cAMP, Rap1A N17 or Epac WT (present or control) and 6-Phe-cAMP (present or absent), and Rap1A siRNA and PGE2. Due to heterogeneous variability among treatment groups, data were transformed using logarithm to the base 10. The log-transformed data were analyzed using analysis of variance for a two-factor experiment. The effects were assessed at the 0.05 level of significance. Because synergistic effects were expected, interactions were assessed at the 0.15 level of significance. Multiple comparisons were assessed using Fisher’s least significant difference procedure with Bonferroni adjustment for the number of comparisons. All statistical computations were conducted using the SAS system, release 9.1 (SAS Institute Inc., Cary, NC) (76).


   ACKNOWLEDGMENTS
 
The authors thank Emily E. Bayne for technical assistance and Karen Martin for manuscript preparation.


   FOOTNOTES
 
This paper was presented, in part, at the annual meeting of the American Gastroenterological Association (May 14–19, 2005, Chicago, IL, and May 20–25, 2006, Los Angeles, CA).

This work was supported by National Institutes of Health Grants 2R37 AG10885, RO1 DK48489, PO1 DK35608, R21 CA10212, and RO1 GM66170.

Disclosure Statement: The authors have nothing to disclose.

First Published Online October 26, 2006

Abbreviations: 6-Bnz-cAMP, N6-Benzoyladenosine-3', 5'-cyclic monophosphate; ECL, enhanced chemiluminescence; EP, E prostanoid; Epac, exchange protein directly activated by cAMP; FSK, forskolin; GFP, green fluorescent protein; GPCR, G protein-coupled receptor; HA, hemagglutinin; NT, neurotensin; NTR, NT receptors; NTR1, high-affinity NTR; PGE2, prostaglandin E2; 6-Phe-cAMP, N6-phenyladenosine-3', 5'-cyclic monophosphate; PKA, protein kinase A; PKC, protein kinase C; PKI, protein kinase inhibitor; PMA, phorbol 12-myristate 13-acetate; Rap1GAPII, Rap1 GTPase-activating protein II; Rap1A N17, dominant negative Rap1; Rap1A V12, constitutively active Rap1; siRNA, small interfering RNA; WT, wild type.

Received for publication August 16, 2006. Accepted for publication October 13, 2006.


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