PI3K p110α/Akt Signaling Negatively Regulates Secretion of the Intestinal Peptide Neurotensin Through Interference of Granule Transport

Abstract

Neurotensin (NT), an intestinal peptide secreted from N cells in the small bowel, regulates a variety of physiological functions of the gastrointestinal tract, including secretion, gut motility, and intestinal growth. The class IA phosphatidylinositol 3-kinase (PI3K) family, which comprised of p110 catalytic (α, β and δ) and p85 regulatory subunits, has been implicated in the regulation of hormone secretion from endocrine cells. However, the underlying mechanisms remain poorly understood. In particular, the role of PI3K in intestinal peptide secretion is not known. Here, we show that PI3K catalytic subunit, p110α, negatively regulates NT secretion in vitro and in vivo. We demonstrate that inhibition of p110α, but not p110β, induces NT release in BON, a human endocrine cell line, which expresses NT mRNA and produces NT peptide in a manner analogous to N cells, and QGP-1, a pancreatic endocrine cell line that produces NT peptide. In contrast, overexpression of p110α decreases NT secretion. Consistently, p110α-inhibition increases plasma NT levels in mice. To further delineate the mechanisms contributing to this effect, we demonstrate that inhibition of p110α increases NT granule trafficking by up-regulating α-tubulin acetylation; NT secretion is prevented by overexpression of HDAC6, an α-tubulin deacetylase. Moreover, ras-related protein Rab27A (a small G protein) and kinase D-interacting substrate of 220 kDa (Kidins220), which are associated with NT granules, play a negative and positive role, respectively, in p110α-inhibition-induced NT secretion. Our findings identify the critical role and novel mechanisms for the PI3K signaling pathway in the control of intestinal hormone granule transport and release.

The phosphatidylinositol 3-kinases (PI3K) are a family of intracellular lipid kinases implicated in cell proliferation, survival, metabolism, cytoskeleton reorganization, and vesicle trafficking (1). The class IA PI3K family is comprised of p110 catalytic (α, β, and δ) and p85 regulatory subunits. p110α and p110β are widely distributed in mammalian tissues, in contrast to p110δ, which shows more restricted distribution and is mainly found in leukocytes (1). Two isoforms of p85, p85α and p85β, are derived from distinct genes, Pik3r1 and Pik3r2, respectively (1). The p85 regulatory subunit binds with p110 and recruits the p85–p110 heterodimer to its substrate phosphatidylinositol 4,5-bisphosphate at the plasma membrane, which results in activation of v-Akt murine thymoma viral oncogene homolog (Akt), a well-known downstream effector of PI3K signaling (2). Another downstream effector of PI3K/Akt signaling is the mammalian target of rapamycin (mTOR), which plays a critical role in the regulation of protein synthesis and cell growth (3). mTOR signaling network contains two functionally distinct mTOR complexes (mTORC), mTORC1 and mTORC2 (4, 5). PI3K signaling contributes to vesicle trafficking and secretory function in many cell types (6–14), especially in pancreatic β-cells (10, 15–23). However, it has been controversial whether PI3K plays a positive or negative role in the regulation of insulin secretion.

Microtubules, highly dynamic polymers formed by α- and β-tubulin heterodimers, form an intracellular cytoskeletal network essential for vesicle trafficking (24). Kapeller et al. (25, 26) first found that PI3K localizes to microtubules by direct association of p85. This localization suggests that PI3K may play a role in cellular trafficking processes. In support of these findings, PI3K controls microtubule dynamics, thereby facilitating vesicle transport in neuron growth cones (27). α-Tubulin and β-tubulin are subject to numerous posttranslational modifications, including acetylation, which occurs on lysine-40 of α-tubulin (28–30). Studies have suggested that α-tubulin plays a positive role in vesicular and organelle transport (31–34). Microtubule-dependent transport of cargo is mediated by kinesin-1, a member of the kinesin superfamily that carries cargoes along the microtubule (32, 33, 35–38). Insulin secretion requires the microtubule-dependent recruitment of granules from a “reserve pool” to the cell surface (39); kinesin-1 associates with and is responsible for the transport of insulin granules during insulin secretion (40–44).

The small GTP-binding proteins ras-related protein Rab27, including Rab27A and Rab27B, is present on granules in a wide variety of secretory cells, including nonendocrine (45–51) and endocrine cells (52–55). Rab27A and its effectors associate with insulin granules and regulate the exocytosis in β-cells (54–57). The interaction between kinesin-1 and Kidins220 has been demonstrated (58). The Kinase D-interacting substrate of 220 kDa (Kidins220), originally identified as a substrate of protein kinase D (59), is a vesicle-associated protein mainly expressed in brain and neuroendocrine cells. Kidins220 interacts with tubulin and has been identified as one of the kinesin-1 cargo proteins (60).

Neurotensin (NT), a gut peptide secreted from N cells in the small bowel, has numerous physiologic functions in the gastrointestinal tract, including effects on gastrointestinal motility, facilitation of fatty acid translocation, stimulation of pancreatic secretion, and intestinal growth (61, 62). Although the PI3K pathway plays important roles in the regulation of vesicle transport (6–13, 14) and insulin secretion (10, 15–23), it remains unclear whether PI3K signaling regulates release of NT or other gut peptides. The purpose of this study was to examine the role of PI3K/Akt signaling in the regulation of NT secretion and to study the mechanisms involved. Here, we demonstrate that p110α, but not p110β, is a negative regulator in NT secretion in endocrine cell lines as well as in mice. Importantly, we demonstrate that p110α mediates NT secretion by inhibiting NT granule trafficking through mechanisms involving α-tubulin acetylation and NT granule-associated proteins, Rab27A and Kidins220.

Results

p110α, but not p110β, negatively regulates NT secretion

BON and QGP-1 cells express high levels of p110α and p110β; in contrast, p110δ and p110γ-expression is not detected (Supplemental Fig. 1A, published on The Endocrine Society's Journals Online web site at http://mend.endojournals.org). To determine whether p110α or p110β is involved in the regulation of NT secretion, we tested p110α-inhibitor (PIK-75) and p110β-inhibitor (TGX-221). PIK-75 selectively inhibits p110α and blocks activation of Akt and mTORC1 (63, 64). TGX-221 is a potent and specific TGX-221 (65, 66).

NT secretion was increased in BON cells treated with different concentrations of PIK-75 (Fig. 1A, top panel) in a dose-dependent fashion. Phosphorylation of Akt and p70S6K, an mTORC1 downstream effector, was inhibited, whereas phosphorylation of ERK1/2 was induced, by PIK-75 (Fig. 1A, lower panel). TGX-221 treatment, however, did not affect either NT release or the phosphorylation of Akt, p70S6K, and ERK1/2 in BON cells (Supplemental Fig. 1B). Consistently, NT secretion was decreased in BON cells overexpressing wild-type (WT) p110α and constitutively active forms of E545K and H1047R (Fig. 1B, top panel); the overexpression of p110α and Akt phosphorylation was monitored by Western blotting as shown in Fig. 1B, lower panel. In addition, NT secretion was also increased in QGP-1 cells treated with PIK-75 in a dose-dependent fashion (Fig. 1C, top panel). Inhibition of Akt and p70S6K phosphorylation and induction of ERK1/2 phosphorylation was also detected in QGP-1 cells treated with PIK-75 (Fig. 1C, lower panel). Again, TGX-221 treatment did not affect either NT release or the phosphorylation of Akt, p70S6K, and ERK1/2 in QGP-1 cells (Supplemental Fig. 1C).

Fig. 1.

p110α, but not p110β, negatively regulates NT secretion. A, BON cells were treated with PIK-75 at various concentrations for 1 h; medium was collected for NT measurements (top panels) (n = 6) (*, P < 0.05 vs. DMSO); signaling of Akt, p70S6K, and ERK1/2 was examined by Western blotting (lower panels). B, Medium was collected from stable BON cell lines overexpressing WT and constitutively active p110α E545E and H1047R as well as the control vector (pBabe-puro) and NT secretion measured (top panel) (n = 6) (*, P < 0.05 vs. control vector). Overexpression of p110α-plasmids and Akt phosphorylation were monitored by Western blotting (lower panel). C, QGP-1 cells were treated with PIK-75 at different concentrations for 1 h; NT EIA (top panel) (n = 6) (*, P < 0.05 vs. DMSO) and Western blotting (lower panel) were performed. D, Mice were treated with vehicle (0.5% CMC) and PIK-75 (10 mg/kg for 6 d); plasma (50 μl) was used for NT EIA (n = 6) (P < 0.05 vs. vehicle).

To confirm the findings obtained from in vitro assays, we tested the effect of PIK-75 in mice. Food intake and body weight were monitored daily and were not changed in the vehicle or PIK-75 treatment groups (data not shown). Blood glucose was measured before killing; no difference between the two groups was noted (data not shown). Plasma NT, however, was significantly increased in mice given PIK-75 (Fig. 1D). To determine whether PIK-75 treatment changed expression of NT mRNA and protein content, ileums were collected and total RNA and protein purified. Results from real-time PCR did not show changes in NT mRNA expression between PIK-75 and vehicle groups (data not shown). Similarly, NT protein content was not altered by PIK-75 treatment as determined by NT enzyme immunoassay (EIA) (data not shown). Together, these results, using both in vitro and in vivo models, demonstrate that p110α plays a negative role in the regulation of NT secretion.

Akt plays a functional role downstream of p110α in the regulation of NT secretion

Akt is a downstream effector of PI3K signaling. Akt inhibitor VIII potently and selectively inhibits Akt1, Akt2, and Akt3 activity (IC₅₀ = 58, 210, and 2.12 μm, respectively) (67–69). Akt inhibitor VIII is dependent on the presence of the pleckstrin homology domain and does not exhibit any inhibitory effect against other closely related AGC family kinases, such as cAMP-dependent protein kinase, protein kinase c, or serum and glucocorticoid-inducible kinase. Akt inhibitor VIII blocks phosphorylation of Akt Thr³⁰⁸ and Akt Ser⁴⁷³; Akt inhibitor XII is an Akt inhibitor VIII-derived allosteric inhibitor with much improved Akt2 selectivity (70). To further confirm the involvement of p110α/Akt signaling pathway in the regulation of NT secretion, we treated BON cells with Akt inhibitors VIII and XII at various doses. NT release was increased in BON cells treated with Akt VIII (Fig. 2A, top panel) and Akt XII (Fig. 2B, top panel) in a dose-dependent fashion. Phosphorylation of Akt and p70S6K was decreased by both Akt VIII (Fig. 2A, lower panel) and Akt XII (Fig. 2B, lower panel). In support of these findings, NT secretion was decreased by the overexpression of the WT Akt1 but dramatically increased by the kinase dead Akt1 (Fig. 2C, top panel). Phosphorylation and overexpression of Akt were monitored by Western blotting (Fig. 2C, lower panel). In addition, NT secretion was decreased in cells overexpressing either the WT or the myristoylated Akt2 (Fig. 2D, top panel). Western blotting confirms the overexpression of Akt2 and marked Akt induction noted in cells expressing the myristoylated Akt2 (Fig. 2D, lower panel). These results further support our findings that the PI3K/Akt signaling pathway negatively regulates NT release.

Fig. 2.

Akt1 and Akt2 negatively regulate NT secretion. A and B, BON cells were treated with Akt inhibitor VIII (A) and XII (B) at different doses for 1 h; medium was collected and NT EIA performed (top panels) (n = 6) (*, P < 0.05 vs. DMSO), and Western blotting was performed (lower panels). C and D, Medium was collected from BON cells with stable overexpression of WT and kinase dead (KD) Akt1 (C) and WT and myristoylated (myr)Akt2 (D). NT EIA was performed (top panels) (n = 6) (*, P < 0.05 vs. control vector). Overexpression of Akt1 and Akt2 and Akt phosphorylation status were examined by Western blotting (lower panels).

High levels of p85α and p110α are expressed in N cells

We analyzed the expression of p85α and p110α in the small bowel by immunofluorescent (IF) staining and confocal microscopic analysis. As shown in Fig. 3A, NT, p85α, and p110α displayed similar expression patterns in mouse ileum by single IF staining. Double staining of NT with p85α further demonstrated the colocalization of NT with p85α (Fig. 3B) and p110α (Fig. 3C), suggesting the importance of PI3K signaling in gut endocrine cells. Similar expression patterns and colocalization of p85α or p110α with NT were noted in human ileum by IF single (Fig. 3D) and double staining (Fig. 3, E and F). In some sections, NT staining was not found in cells with positive staining of p85α and/or p110α, suggesting that p85α and p110α are expressed in other types of enteroendocrine cells (data not shown).

Fig. 3.

p85α and p110α localize in N cells. A, IF and confocal microscopic analysis were performed on mouse ileum paraffin sections using anti-NT (left panel), p85α (middle panel), and p110α (right panel) antibodies. Positive cells were indicated by arrows and nuclei stained by 4′,6-diamidino-2-phenylindole (DAPI). B, Double staining with anti-p85α and anti-NT antibodies; colocalization of p85α with N cells was analyzed by confocal microscopy (arrows). C, Double staining with anti-p110α and anti-NT antibodies; colocalization of p110α with N cells was analyzed by confocal microscopy (arrows). D–F, Similar analysis as A–C was performed in human ileum.

Inhibition of p110α increased the number of NT granules docked to the plasma membrane

Previously, we reported that inhibition of mTORC1 signaling increased NT secretion via up-regulation of NT mRNA and protein content through activation of mitogen-activated protein kinase kinase/ERK/c-Jun, but not PI3K/Akt, signaling pathway (71). In the present study, we confirmed that NT mRNA and intracellular NT content were not altered by p110α and Akt inhibitors (data not shown), suggesting that p110α/Akt signaling regulates NT secretion not through the regulation of NT mRNA and protein synthesis but through regulation of the secretory processes. To prove this hypothesis, we monitored the docking status of green fluorescent protein (GFP)-tagged NT granules in live BON/GFP-NT cells by total internal reflection fluorescence (TIRF) microscopy, as described in Materials and Methods. TIRF imaging depicts the single granules docked to the plasma membrane, enabling accurate docked granule counting (72, 73). As shown in Fig. 4A, representative images at different time points showed an increase of GFP-NT granules docked to the plasma membrane after the addition of PIK-75, compared with the control images before the addition of PIK-75 (also see Supplemental video). GFP-NT granules were also monitored in the presence of dimethylsulfoxide (DMSO) (vehicle control) over the same period; no significant difference was noted after the addition of DMSO (data not shown). The docked GFP-NT granules were further quantified in cells treated with DMSO or PIK-75 by the area fraction in 31 serial images (Fig. 4B). This data suggest that the p110α/Akt signaling controls NT release at the level of NT granule trafficking or docking/fusion.

Fig. 4.

PIK-75 enhanced accumulation of NT granules on the plasma membrane. A, The glass-bottomed 35-mm dishes cultured with BON/GFP-NT cells were placed in a thermostat-controlled (37 C, 5% CO₂) humidified atmosphere chamber attached to the stage, and time serial images were taken for 30 min with an interval of 1 min. DMSO or PIK-75 (0.5 μm) was added in the interval after the first five images. Representative images are shown from PIK-75-treated cells. B, Quantification analysis was performed on 31 images from both DMSO and PIK-75 groups.

α-Tubulin acetylation is involved in p110α-mediated NT secretion

Because acetylation of α-tubulin enhances transport of vesicles (32, 33), we examined the status of α-tubulin acetylation in BON and QGP-1 cells. We detected an increase of α-tubulin acetylation induced by treatment of PIK-75 in BON and QGP-1 cells as noted by IF staining and confocal microscopic analysis using the specific antibody recognizing α-tubulin acetylation at lysine 40 (Fig. 5A). α-Tubulin acetylation was also detected by Western blotting in both cell lines treated with PIK-75 over a different time course (Fig. 5B). α-Tubulin acetylation was increased at 2 and 8 h (Fig. 5B, left panel). This reaction is rapid, as noted by a dramatic increase of α-tubulin acetylation at 5 min in both BON and QGP-1 cells (Fig. 5B, right panel). Conversely, overexpression of myristoylated p110α (Fig. 5C, left panel) and WT Akt1 and Akt2 (Fig. 5C, right panel) decreased α-tubulin acetylation compared with the cells expressing the control vector. We found that taxol, a microtubule stabilization reagent (74), but not nocodazole, a microtubule destabilization reagent (75), increased NT release in BON cells and, combined with PIK-75, further enhanced NT secretion (Fig. 5D, top panel). As shown in Fig. 5D (lower panel), α-tubulin acetylation was induced by taxol alone but was not significantly enhanced by the combination of PIK-75; nocodazole treatment decreased both basal and PIK-75-induced α-tubulin acetylation. Similar results were obtained in QGP-1 cells (Fig. 5E). Consistently, PIK-75-mediated NT secretion was further increased by overexpression of the active form of α-tubulin (K40Q) but not the inactive form (K40R) (Fig. 5F). Taken together, these results indicate that α-tubulin acetylation contributes to p110α-regulated NT release.

Fig. 5.

p110α/Akt regulated NT secretion through α-tubulin (tub) acetylation (ac). A, BON (left panel) and QGP-1 (right panel) were treated without or with PIK-75 (0.5 μm) for 1 h; IF and confocal analysis were performed. B, BON and QGP-1 cells were treated without or with PIK-75 for longer (left panel) or shorter time points (right panel) and Western blotting performed. C, Western blotting was performed on stable BON cell lines with overexpression of control vector (V), myristoylated (myr) p110α and WT Akt1 and Akt2. D and E, BON (D) and QGP-1 (E) cells were treated without or with PIK-75 (0.5 μm) in the presence or absence of taxol (1 μm) or nocodazole (Noco) (1 μm) for 1 h; medium was collected and NT EIA performed (top panels) (n = 6) (*, P < 0.05 vs. DMSO, taxol alone and Noco alone, respectively; †, P < 0.05 vs. DMSO; ‡, P < 0.05 vs. PIK-75 alone); α-tubulin acetylation was detected by Western blotting (lower panels). F, BON cells, transiently transfected with control vector, pEGFP-N1, GFP-α-tubulin K40Q, and K40R were treated without or with PIK-75 (0.5 μm) for 1 h; NT secretion was measured by NT EIA (top panel) (n = 6) (*, P < 0.05 vs. their DMSO; †, P < 0.05 vs. DMSO in control vector; ‡, P < 0.05 vs. control vector plus PIK-75); Western blotting was performed to monitor the overexpression of GFP-α-tubulin (lower panel).

HDAC6 is involved in p110α-mediated NT secretion

HDAC6, a class IIb histone deacetylase, associates with microtubules and functions to deacetylate α-tubulin (76–79). To determine whether HDAC6 is involved in NT release mediated by p110α-regulated α-tubulin acetylation, BON cells were transiently transfected with HDAC6 or the control vector pEGPF-N1 and treated with or without PIK-75. PIK-75 treatment increased NT secretion in BON cells transfected with the control vector; overexpression of HDAC6 decreased both the basal and PIK-75-enhanced NT secretion (Fig. 6, left panel). Western blot analysis showed that HDAC6 overexpression decreased both basal and PIK-75-induced α-tubulin acetylation (Fig. 6, right panel). These results demonstrate that HDAC6, an α-tubulin deacetylase, controls NT secretion by inhibiting levels of α-tubulin acetylation in BON cells. Furthermore, HDAC6-mediated α-tubulin acetylation is involved in p110α-regulated NT secretion.

Fig. 6.

HDAC6 overexpression decreased α-tubulin (tub) acetylation (ac) and NT secretion. BON cells, transiently transfected with the control vector, pEGFP-N1, and GFP-HDAC6 were treated with PIK-75 (0.5 μm) for 1 h; NT secretion was measured by NT EIA (left panel) (n = 6) (*, P < 0.05 vs. their DMSO; †, P < 0.05 vs. DMSO in control vector; ‡, P < 0.05 vs. control vector plus PIK-75); α-tubulin acetylation and GFP-HDAC6 overexpression were examined by Western blotting (right panel).

Rab27A and Kidins220 are involved in p110α-mediated NT secretion

Rab27A localizes to the membrane of insulin granules and regulates insulin granule trafficking (52, 80). We also noted the association of Rab27A with NT granules (Li, J., and B. M. Evers, unpublished data), and therefore, we suspected involvement of Rab27A in p110α-mediated NT granule transport. To test this hypothesis, BON cells were transiently transfected with Rab27A plasmids, including the WT, the active form (Q78L), and the dominant negative form (T23N). As shown in Fig. 7A, top panel, overexpression of either WT or Q78L decreased both the basal and PIK-75-stimulated NT secretion, whereas T23N had no significant effect. Interestingly, overexpression of WT and Q78L decreased α-tubulin acetylation, whereas T23N had no effect (Fig. 7B, lower panel). Previously, we showed that Kidins220 was an NT granule-associated protein that positively regulates NT secretion (81). In this study, we found that PIK-75-stimulated NT secretion was further enhanced in cells with overexpression of the active form of Kidins220-S918E (Fig. 7B). These results demonstrate that both Rab27A and Kidins220 associate with NT granules but play a negative and positive role, respectively, in p110α-mediated NT secretion.

Fig. 7.

Overexpression of Rab27A attenuated, but overexpression of Kidins220 increased, PIK-75-stimulated NT secretion. A, BON cells, transiently transfected with Flag-tagged WT, active (Q78L), and the dominant negative (T23N) Rab27A as well as the control vector pEF-Bos, were treated without or with PIK-75 (0.5 μm) for 1 h; medium was collected and NT EIA performed (top panel) (n = 6) (*, P < 0.05 vs. their DMSO; †, P < 0.05 vs. DMSO in control vector; ‡, P < 0.05 vs. control vector plus PIK-75); α-tubulin (tub) acetylation (ac) and overexpression of Flag-Rab27A were monitored by Western blotting (lower panel). B, The inducible stable cell line, BON/Kidins220–S918E, was cultured in the presence or absence of doxycyclin (Dox) (100 nm) for 48 h followed by treatment with or without PIK-75 (0.5 μm) for 1 h; NT secretion was measured by NT EIA (top panel) (n = 6) (*, P < 0.05 vs. their DMSO; †, P < 0.05 vs. PIK-75 without Dox); Western blotting was performed to detect the induction of Kidins220 expression (lower panel).

Overexpression of Rab27A blocks the release of NT granules

As shown in Fig. 8A, top panel, IF and confocal microscopic analysis showed that red fluorescent protein (RFP)-tagged Rab27A was expressed in BON cells in a similar fashion as NT and that Rab27A is colocalized with NT granules (Fig. 8A, middle panel, arrows) and Kidins220 (Fig. 8A, bottom panel, arrows). Pearson correlation coefficient (PCC) is a common way to quantify the degree of colocalization between fluorophores. Values for PCC range from −1 to 1, with −1 indicating complete inverse correlation, 0 indicating no correlation, and 1 indicating complete positive correlation for two variables (82). The colocalization of NT with Rab27A or Kidins220 was supported by the results of PCC calculations: PCC was 0.7 and 0.8, respectively, for colocalization of NT with Rab27A and Kidins220. Coimmunoprecipitation failed to detect the physical association between Rab27A and Kidins220 (data not shown), suggesting both proteins exist on NT granules but do not have direct protein-protein interactions. Interestingly, colocalization of GFP-NT and Kidins220 (PCC = 0.9) (Fig. 8B, top panel, arrows) was decreased in the presence of PIK-75 (Fig. 8B, middle panel), demonstrating that PIK-75 treatment causes an extracellular release of NT granules as well as Kidins220. However, PIK-75 induced the release of Kidins220 granules only in cells without RFP-Rab27A expression (Fig. 8B, bottom panel, arrowheads); cells expressing RFP-Rab27A showed no release of Kidins220 granules after PIK-75 treatment (Fig. 8B, bottom panel, arrows). Together, these results demonstrate that inhibition of p110α increases NT secretion through the inhibition of granule trafficking by mechanisms involving the NT granule-associated proteins, Rab27A and Kidins220.

Fig. 8.

Overexpression of Rab27A blocked the release of NT granules. A, Stable BON/RFP-Rab27A cells were stained with Alexa Fluor 488 phalloidin (top panel), NT (middle panel), and Kidins220 (bottom panel) antibodies; colocalization of RFP-Rab27A with NT or Kidins220 was analyzed by confocal microscopy (arrows), and PCC values are 0.7 and 0.8, respectively. B, BON/GFP-NT cells, in the absence (top panel) or presence (middle panel) of PIK-75 (0.5 μm, 30 min), were stained with Kidins220 antibody and confocal microscopic analysis performed (arrows); the colocalization of GFP-NT and kidins220 was analyzed, and PCC value is 0.9. BON/RFP-Rab27A cells were treated with PIK-75 (0.5 μm) for 30 min; IF and confocal microscopic analysis were performed using anti-Kidins220 antibody; Kidins220 expression was only detected in the RFP-Rab27A positive (arrows) but not in the RFP-Rab27A negative (arrow heads) cells. DAPI, 4′,6-Diamidino-2-phenylindole.

Discussion

Despite the important role of PI3K signaling in the regulation of β-cell function and insulin secretion, little is known regarding the biological effect of this pathway on intestinal hormone secretion. Here, we report that the PI3K p110α/Akt pathway plays a negative role in NT release in human endocrine cell lines and mice in vivo. The mechanisms underlying these effects include the interaction of PI3K/Akt signaling with α-tubulin acetylation and NT-granule associated proteins, Rab27A and Kidins220.

Importantly, we not only determined that NT secretion is negatively controlled by PI3K/Akt signaling, but we identified the specific involvement of the p110α-subunit in this effect. Although the role of PI3K signaling in the regulation of β-cell function and insulin secretion has been extensively studied, the specific subunits participating in the regulation of insulin secretion remain unknown. Moreover, the conclusions regarding whether PI3K signaling plays a positive or negative role are controversial. Some studies have shown that stimulated insulin secretion in β-cells was further potentiated by wortmannin or LY294002 (16, 21, 23, 83), two well-known general PI3K inhibitors (84, 85). Using human fetal-derived pancreatic cells, which grow in vitro as islet-like cell clusters, as a model for endocrine differentiation, Ptasznik et al. (86) reported that wortmannin and LY294002 induced morphological and functional endocrine differentiation associated with an increase in mRNA levels of insulin, glucagon, and somatostatin, as well as an increase in the insulin protein content and secretion. Zawalich and Zawalich (23) also reported that wortmannin and LY294002 augmented glucose-induced insulin secretion from rat islets. p85α-Deficient β-cells exhibited a marked increase in insulin secretion in response to higher concentrations of glucose (83). In contrast to these findings, an earlier study demonstrated that wortmannin selectively inhibited pituitary adenylate cyclase-activating polypeptide and vasoactive intestinal peptide-stimulated insulin release but not release induced by forskolin, a reagent that elevates cAMP and therefore activates PKA activity, or glucose (20). Mice with β-cell-specific insulin receptor knockout or IGF-I receptor knockout developed severe diabetes and a defect in insulin secretion (87–90), suggesting that PI3K plays a positive role in the regulation of insulin secretion. In support of these findings, Kaneko et al. (15) recently reported that inhibition of class IA PI3K, using a mouse model lacking the pik3r1 gene specifically in β-cells and the pik3r2 gene systemically, resulted in reduced insulin secretion in response to glucose. Our findings further highlight a differential effect of PI3K signaling on intestinal and pancreatic peptide secretion, indicating that these physiological events appear to be cell type dependent.

Although well established in neurons, little is known about whether α-tubulin acetylation regulates granule trafficking in endocrine cells. In the present study, we provide evidence demonstrating that p110α regulates NT secretion through α-tubulin acetylation in two endocrine cell lines, BON and QGP-1. HDAC6 is a well-known deacetylatase of α-tubulin in cells (76) and has been implicated in α-tubulin acetylation and kinesin-1 dependent transport (31, 91). In addition, HDAC inhibitors or HDAC6 small interfering RNA inhibits Akt phosphorylation, which parallels effects on α-tubulin acetylation (92). Consistent with these results, we also found that overexpression of HDAC6 decreased α-tubulin acetylation and, concurrently, attenuated PIK-75-enhanced NT secretion. However, whether HDAC6 is located upstream, downstream, or parallel to PI3K/Akt signaling is not known. Surprisingly, we also noted that overexpression of Rab27A decreased α-tubulin acetylation and PIK-75-enhanced NT secretion. The direct evidence showing the effect of Rab27 proteins on α-tubulin acetylation has not been reported. However, Rab27A has been demonstrated to interact with kinesin-1 through synaptotagmin-like protein 1, a Rab27 effector that is involved in the regulation of anterograde transport in axons (51). α-Tubulin acetylation influences the binding and motility of kinesin-1, and cargo transport mediated by kinesin-1 (33). We have previously reported that Kidins220 positively regulates 12-myristate 13-acetate-stimulated NT secretion downstream of protein kinase D (81). Kidins220 has been implicated as a cargo protein, and the transport of Kidins220 carriers is kinesin-1 dependent (58). Kidins220 associates with microtubule-regulating proteins that actively control the microtubule network (60). In addition, coimmunoprecipitation showed that Kidins220 interacted with acetylated α-tubulin (60). Our current findings suggest that Rab27 might mediate α-tubulin acetylation through Kidins220 and kinesin-1.

We showed that overexpression of both the WT and the active Rab27A decreased basal and PIK-75-enhanced NT secretion. The Rab27 family regulates secretory pathways in neurons and endocrine and exocrine cells (57, 93, 94). In addition, Akt interacts with and phosphorylates the Rab27 effector, synaptotagmin-like protein 1 (95), indicating the interaction of PI3K signaling and Rab27; this interaction may have implications on Rab27A-containing vesicle secretion. In contrast to our results, overexpression of WT Rab27A and its active form significantly enhanced high K⁺-induced insulin secretion (54, 55, 96–98). The differential effects might be controlled by multiple effectors of Rab27A expressed in cells. For example, overexpression of granuphilin, one of the Rab27A effectors, in endocrine cells attenuates secretion (99, 100), whereas other members of the granuphilins promote secretion (99, 100). In addition, insulin secretion is elevated in granuphilin-deficient β-cells (101). Overexpression of granuphilin significantly inhibits high K⁺-induced insulin secretion (56). Furthermore, nutrient starvation decreases insulin secretion (19, 102, 103) but increased NT secretion in our previous studies (71). Therefore, our findings suggest that different mechanisms are involved in the regulation of insulin and NT secretion. Rab27A acts positively or negatively on secretion depending on the cellular context.

In summary, our current findings identify an important and previously unrecognized role of the PI3K catalytic subunit, p110α, in the regulation of NT release and provide compelling evidence that PI3K signaling is actively involved in intestinal peptide secretion. Moreover, the interaction of PI3K with α-tubulin acetylation and granule-associated proteins provides a more in-depth understanding of the effect of this pathway in physiologic intestinal functions.

Materials and Methods

Materials

All the antibodies used in this study, except for the antibodies mentioned below, were from Cell Signaling Technology (Danvers, MA). The hemagglutinin antibody was from Covance (Princeton, NJ). Rabbit NT, p85α, Kidins220 (phospho-S919), and Kidins220 antibodies were from Abcam (Cambridge, MA). Goat NT antibody was from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Flag (clone M2), acetylated α-tubulin (clone 6–11B-1), GFP, and β-actin antibodies were from Sigma-Aldrich (St. Louis, MO). The α-tubulin antibody (12G10) was from Developmental Studies Hybridoma Bank (Iowa City, IA). TGX-221, Akt inhibitor VIII, and Akt inhibitor XII were from EMD Chemicals (Rockland, MA). PIK-75 was from Cayman Chemical (Ann Arbor, MI). Taxol and nocodazole were from Sigma-Aldrich. The NT EIA kit was from Phoenix Pharmaceuticals (Belmont, CA). The Alexa Fluor 488 phalloidin antibody, Alexa Fluor-conjugated secondary antibodies, and the NuPAGE BisTris gels for Western blotting were from Invitrogen (Carlsbad, CA). The enhanced chemiluminescence detection system was from GE Healthcare (Piscataway, NJ).

Constructs

The pBabe-puro plasmids, including p110α (WT, E545K, H1047R, and myristoylated), Akt1 (WT and kinase dead), and Akt2 (WT and myristoylated) were from Addgene (Cambridge, MA). The Flag-tagged Rab27A plasmids (WT, active form Q78L, and dominant negative T23N) and mRFP-27A were from Mitsunori Fukuda (Tohoku University, Sendai, Japan). The GFP-tagged α-tubulin K40R (inactive form) and K40Q (active form) were provided by Tso-Pang Yao (Duke University, Durham, NC). The GFP-tagged HDAC6 was from Chunming Liu (University of Kentucky). To generate GFP-tagged NT plasmid, NT cDNA was amplified by RT-PCR using template mRNA obtained from BON cells and cloned into pEGFP-N1 vector (CLONTECH, Mountain View, CA). Kidins220-S918E was generated by PCR using the Flag-tagged rat Kidins220 plasmid provided by Moses V. Chao (New York University, New York, NY) as template by the QuikChange Site-Directed Mutagenesis kit (Agilent Technologies, Santa Clara, CA) and cloned into the pcDNA4/TO expression vector (Invitrogen). This placed the Kidins220S918E under the translational control of the phCMV/TetO₂ promoter containing the tetracycline-responsive element.

Cell culture and transfection

The BON cell line was derived from a human pancreatic carcinoid tumor and characterized in our laboratory (104, 105). BON cells were maintained in a 1:1 mixture of DMEM and nutrient mixture, F12K, supplemented with 5% fetal bovine serum (FBS) (growth medium) in 5% CO₂ at 37 C. QGP-1, a pancreatic endocrine cell line purchased from Japan Health Sciences Foundation (Osaka, Japan) (106), was maintained in American Type Culture Collection-formulated RPMI 1640 medium with 10% FBS. The 293FT packaging cells (Invitrogen) were cultured in DMEM complemented with 10% FBS, 2 mm l-glutamine, and 0.1 mm MEM nonessential amino acids (growth medium). Transfections were performed by Lipofectamine 2000 (Invitrogen).

Cell treatment and NT EIA

For inhibitor treatments, both BON and QGP-1 cells were plated in 24-well plates at a density of 10 × 10⁴/cm² and grown for 48 h; cells were treated with inhibitors for 1 h in growth medium (n = 6); media were collected and stored in −80 C. NT peptide secreted into the medium was measured by NT EIA as described previously (81, 107). Data obtained from NT EIA were normalized by protein concentration from parallel cell lysates.

Generation of stable cell lines

To establish the GFP-NT and RFP-Rab27A stable cell lines, BON cells were transfected with GFP-NT or RFP-Rab27A and selected in the presence of G418 (800 μg/ml). The positive cells were further enriched by flow cytometry cell sorting. To generate the inducible stable cell line, BON/Kidins220-S918E, the pcDNA4/TO-Kidins220-S918E plasmid was transfected into BON cells with stable expression of pcDNA6/TR (Invitrogen) and selected in the presence of zeocin (10 μg/ml; Invitrogen). Single clones of BON/Kidins220-S918E, in the presence or absence of doxycycline (100 ng/ml), were assessed by Western blotting. To produce retrovirus, 293FT packaging cells cultured in 60-mm dishes were cotransfected with pBabe-puro plasmid (1 μg) and Ampho packaging plasmid by Lipofectamine 2000 and incubated in growth medium overnight. Subsequently, the cells were cultured in complete medium (growth medium plus 1 mm MEM sodium pyruvate) for 24 h; the supernatant containing the retrovirus was collected, filtered through a 0.45-μm Surfactant Free-Cellulose Acetate sterile syringe filter, and used to infect target cells. BON cells in six-well plates (5 × 10⁵ cells/well) were incubated with the viral supernatant for 24 h; cells were then incubated with growth medium for an additional 24 h. The infected cells were subcultured in 100-mm dishes in fresh medium containing puromycin (2.5 μg/ml). Puromycin-resistant cell pools were collected and the overexpression levels were monitored by Western blotting.

In vivo studies

All procedures were carried out in an animal facility according to the protocols approved by the Institutional Animal Care and Use Committee. Male C57BL/6J mice (8 wk of age, weighing 20–24 g) were obtained from Taconic (Hudson, NY). A group of seven mice was given daily oral administration of 10 mg/kg PIK-75 suspension in 0.5% carboxymethylcellulose (CMC) (Sigma Aldrich) by gavage. As a control (vehicle control), a group of seven mice was given 0.5% CMC. Food intake and body weight were monitored every day. All mice were killed after treatment for 6 d. Blood was collected from the tail vein of mice, and glucose was measured using OneTouch (LifeScan, Milpitas, CA) before killing. Mice were then anesthetized with isoflurane inhalation; blood was collected from the inferior vena cava using a heparin coated syringe and the plasma obtained by centrifuging the blood at 10,000 rpm for 10 min at 4 C. Plasma was aliquoted and stored in −80 C. Plasma (50 μl) was used for NT measurement by NT EIA. The small intestine was isolated and fixed in 10% formalin for immunohistofluorescence analysis. Sections of human ileum were obtained from University of Kentucky Hospital.

IF staining and confocal microscopy

IF staining was performed as described previously (108). Briefly, cells were grown on glass coverslips (no. 1) in 24-well plates for 48 h; after treatment, cells were fixed with 4% paraformaldehyde/PBS and permeabilized with 0.3% Triton X-100/PBS. For IF staining of small intestine, paraffin sections were deparafinized, hydrated, retrieved, and blocked with normal serum. The cells or paraffin sections were incubated with primary antibody for 1 h followed by Alexa Fluor-conjugated secondary antibody for 30 min, and nuclei were counterstained with 4′,6-diamidino-2-phenylindole. Images were observed under an FV1000 Olympus confocal microscope with a ×60, 1.35-numerical aperture oil objective (Olympus, Tokyo, Japan). Images were analyzed with Olympus FV10-ASW2.1 software. Colocalization of green and red fluorophores was quantified using PCC (82).

TIRF microscopy and image analysis

The BON/GFP-NT cells were plated on the no. 1 glass-bottom culture dishes (Mat Tek, Ashland, MA) and grown for 48 h. The dishes were then transferred into a thermostat-controlled (37 C and 5% CO₂) humidified atmosphere chamber attached to the stage. The Nikon Eclipse Ti-E inverted TIRF microscope with Perfect Focus System was used to monitor the GFP-NT granule movement (Nikon, Melville, NY). To observe GFP, a 488-nm laser line for excitation and a 515-nm long-pass filter for the barrier was used. Type 37 immersion oil (Cargille Laboratories, Cedar Grove, NJ) was used to make contact between the objective lens (×60) and the coverslip. A 30-min imaging session with images taken every minute for a total of 31 images was collected. For analysis of the TIRF images, Nikon NIS-Elements Advanced Research Software was used. Regions of interest where cells were present were identified in the images. In these regions of interest, the software performed area fraction analysis with positive fluorescent values selected in each image. Once selected, the software identified any identical values and marked them as positive. This process continued until all positive fluorescent signals in the image were identified. Once this set of values was identified, it was applied to all images in run, and changes over time were determined and graphed.

Protein preparation and Western blotting

Protein preparation and Western blotting were performed as described previously (107, 109). In brief, the cells were lysed with lysis buffer (Cell Signaling Technology), and equal amounts of protein were resolved on 4–12% NuPAGE BisTris gels and electrophoretically transferred to polyvinylidene difluoride 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 enhanced chemiluminescence detection system.

Statistical analysis

Descriptive statistics, including mean ± sd, were calculated to summarize NT secretion. Bar graphs were generated to represent mean (±sd) NT levels in different cell culture conditions and mice groups, such as different inhibitors, treatment, and dose concentrations. Within each experiment, comparisons across groups were accomplished using one-way ANOVA models, and pairwise comparisons were subsequently performed using contrast statements. Adjustment in P values due to several pairwise testing within each experiment was performed using the Holm's procedure. Trend tests for dose comparisons were likewise performed. Normality assumptions of the parametric tests for each outcome were assessed. Adjusted P values of less than 0.05 were considered statistically significant.