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Endocrinology Vol. 144, No. 10 4433-4445
Copyright © 2003 by The Endocrine Society

Glucose-Dependent Insulinotropic Polypeptide Promotes ß-(INS-1) Cell Survival via Cyclic Adenosine Monophosphate-Mediated Caspase-3 Inhibition and Regulation of p38 Mitogen-Activated Protein Kinase

Jan A. Ehses, Vanbric R. Casilla, Tim Doty, J. Andrew Pospisilik, Kyle D. Winter, Hans-Ulrich Demuth, Raymond A. Pederson and Christopher H. S. McIntosh

Department of Physiology (J.A.E., V.R.C., T.D., J.A.P., K.D.W., R.A.P., C.H.S.M.), Faculty of Medicine, University of British Columbia, Vancouver, British Columbia V6T 1Z3, Canada; and Probiodrug Research (H.-U.D.), Biocenter, Weinbergweg 22, D-06120 Halle (Saale), Germany

Address all correspondence and requests for reprints to: Dr. C. H. S. McIntosh, Department of Physiology, Faculty of Medicine, University of British Columbia, 2146 Health Sciences Mall, Vancouver, British Columbia, Canada V6T 1Z3. E-mail: mcintoch{at}interchange.ubc.ca.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The incretin glucose-dependent insulinotropic polypeptide (GIP) is a major regulator of postprandial insulin secretion in mammals. Recent studies in our laboratory, and others have suggested that GIP is a potent stimulus for protein kinase activation, including the MAPK (ERK1/2) module. Based on these studies, we hypothesized that GIP could regulate cell fate and sought to examine the underlying mechanisms involved in GIP stimulation of cell survival. GIP potentiated glucose-induced ß-(INS-1)-cell growth to levels comparable with GH and GLP-1 while promoting cell survival in the face of serum and glucose-deprivation or treatment with wortmannin or streptozotocin. In the absence of GIP, 50% of cells died after 48 h of serum and glucose withdrawal, whereas 91 ± 10% of cells remained viable in the presence of GIP [n = 3, P < 0.05; EC50 of 1.24 ± 0.48 nM GIP (n = 4)]. Effects of GIP on cell survival and inhibition of caspase-3 were mimicked by forskolin, but pharmacological experiments excluded roles for MAPK kinase (Mek)1/2, phosphatidylinositol 3-kinase, protein kinase A, Epac, and Rap 1. Survival effects of GIP were ablated by the inhibitor SB202190, indicating a role for p38 MAPK. Furthermore, caspase-3 activity was also regulated by p38 MAPK, with a lesser role for Mek1/2, based on RNA interference studies. We propose that GIP is able to reverse caspase-3 activation via inhibition of long-term p38 MAPK phosphorylation in response to glucose deprivation (±wortmannin). Intriguingly, these findings contrasted with short-term phosphorylation of MKK3/6->p38 MAPK->ATF-2 by GIP. Thus, these data suggest that GIP is able to regulate INS-1 cell survival by dynamic control of p38 MAPK phosphorylation via cAMP signaling and lend further support to the notion that GIP regulation of MAPK signaling is critical for its regulation of cell fate.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GLUCOSE-DEPENDENT INSULINOTROPIC polypep-tide (GIP) is a meal regulated intestinal polypeptide, which is essential for the maintenance of glucose homeostasis (1, 2). The established physiological actions of GIP include glucose-dependent potentiation of insulin secretion, regulation of insulin gene transcription (3, 4), and adipose tissue effects (5, 6) as well as effects on other extrapancreatic tissues (7, 8). Many of these actions are also exhibited by the second incretin hormone, glucagon-like peptide-1 (GLP-1). Furthermore, it is now established that GLP-1 also plays an important role in regulating ß-cell proliferation, differentiation, and neogenesis (9, 10, 11, 12, 13). Studies on ß-cell models also imply that GIP is able to regulate cell fate (14, 15, 16).

GIP exerts its biological effects by interacting with its cognate G protein-coupled receptor (GPCR), a member of the family B GPCR superfamily, including receptors for glucagon, GLP-1, GLP-2, secretin, pituitary adenylate cyclase activating polypeptide (PACAP), and vasoactive intestinal polypeptide (VIP). All of the glucagon superfamily of peptide hormone receptors are coupled to the production of cAMP and, those tested, have also been shown to activate the MAPKs, ERK 1 and 2 (17, 18, 19, 20, 21). There exist at least five different mammalian MAPK signaling modules, including ERK 1/2, p38 MAPK, Jun N-terminal kinase/stress-activated protein kinase (JNK/SAPK), ERK 3, and ERK 5 (BMK 1; big MAPK). These cytoplasmic serine/threonine kinases are cellular regulators of numerous processes including gene transcription, cell differentiation and proliferation, and cell survival (22). GIP receptor activation has been shown to induce MAPK activity and activate ERK 1/2 in a ß-cell model (INS-1) (14, 15, 23). Recently we elucidated the mechanism by which GIP signals via cAMP to regulate MAPK activity (15), illustrating that cAMP is also able to positively regulate this module in INS-1 cells. Consistent with a role for MAPKs in mediating the proliferative effects of incretins, phosphatidylinositol-3 kinase (PI3K) and p38 MAPK were recently implicated in the growth promotive effects of GLP-1 (10, 24). Such studies therefore suggest that signaling pathways mediated by the glucagon peptide family can couple to mitogenesis and cell survival.

PACAP and GLP-2, both members of the VIP-secretin-glucagon family of peptides, have been convincingly shown to act as antiapoptotic agents in neurons and baby hamster kidney cells, respectively (25, 26, 27, 28). These actions are accomplished by cAMP-mediated inhibition of caspase activity; however, the role of protein kinase A (PKA) is somewhat controversial. The antiapoptotic actions of both peptide receptors have been linked to downstream kinase cascades that promote cell survival; MAPK kinase (Mek)1/2 in the case of PACAP, and protein kinase B (PKB) and glycogen synthase kinase-3ß (GSK-3ß) in mediating GLP-2 effects (28, 29). However, the coupling between B family receptors and antiapoptotic effects has been largely unexplored, and relatively little is known about antiapoptotic signaling in the pancreatic ß cell.

Generally, the activation of ERK MAPKs is associated with survival responses, whereas activity of SAPKs, such as p38 MAPK and JNK is correlated with cell death. However, the physiological actions of p38 MAPK are no longer confined to the generalized stress-response context. Agents that include cytokines, hormones, and osmotic and heat shock are able to regulate inflammation, development, differentiation, proliferation, and survival via p38 MAPK (30, 31). It is now becoming increasingly accepted that there exists a delicate balance between the ERK and p38/JNK pathways that determines whether an extracellular stimulus culminates in cell growth/differentiation or cell death.

The current study was designed to establish the underlying mechanisms by which GIP regulates ß-(INS-1) cell fate. Based on previous studies, we hypothesized that GIP was able to regulate cell survival via regulation of MAPK cascades. GIP was shown to promote cell survival in the presence of adverse or low glucose conditions or when cells were treated with wortmannin or streptozotocin. Our data suggest that these survival actions are due in part to GIP reversal of caspase-3 activity via cAMP and p38 MAPK regulation. We propose that GIP is able to contribute to the regulation of ß-cell fate via dynamic regulation of p38 MAPK and ERK1/2 activation. Taken together, we provide insight into the regulation of cell survival and caspase-3 activity by GIP via p38 MAPK and a novel role for GIP in the regulation of p38 MAPK phosphorylation.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell culture reagents and transfections
INS-1 cells (clone 832/13) were kindly provided by Dr. C. B. Newgard (University of Texas, Southwestern Medical Center) (32). Cells were cultured in 11 mM glucose RPMI 1640 (Sigma Laboratories, Natick, MA) supplemented with 2 mM glutamine, 50 µM ß-mercaptoethanol, 10 mM HEPES, 1 mM sodium pyruvate, and 10% fetal bovine serum (Cansera, Rexdale, Ontario, Canada). Before experiments, cells were harvested into 6-well (2 x 106 cells/well; Becton Dickinson, Lincoln Park, NJ), 24-well (5 x 105 cells/well), or 96-well (5 x 104 cells/well) plates. Cell passages 45–75 were used.

In experiments targeted at investigating a role for Rap1 signaling, INS-1 (832/13), cells were transiently transfected with plasmid DNA encoding the wild-type Rap1b small GTPase, constitutively active (G12V), dominant negative Rap1 (N17), or the empty vector (pCGN). Briefly, 80–90% confluent monolayers in 6-well culture plates (Becton Dickinson) were transfected using Lipofect2000 (Life Technologies, Inc., Burlington, Ontario, Canada) transfection reagent according to the manufacturer’s protocol. Regular growth medium was replaced 5 h after transfection. The Rap1 constructs and vector were a kind gift from Dr. D. Altschuler (University of Pittsburgh, Pittsburgh, PA). All transfections were performed concurrently with green fluorescent protein to ensure transfection and to estimate efficiency (~20–30%).

GIP receptor characterization studies: competitive binding, cAMP production, and insulin release
Synthetic porcine GIP (5 µg) was iodinated by the chloramine-T method, and the 125I-GIP was further purified by reverse phase HPLC to a specific activity of 250–300 µCi/µg. Competitive binding analyses were performed as previously described (33). For cAMP studies, cells were washed twice and then stimulated for 30 min with GIP in the presence of the phosphodiesterase inhibitor 3-isobutyl-1-methylxanthine (0.5 mM 3-isobutyl-1-methylxanthine; RBI/Sigma). Following stimulation, reactions were stopped, and cells lysed, in 70% ice-cold ethanol, cellular debris removed by centrifugation, and cAMP subsequently quantified by radioimmunoassay (Biomedical Technologies Inc., Stoughton, MA). All insulin release experiments were performed over 60 min, in the absence of 3-isobutyl-1-methylxanthine, and insulin secreted into the media was quantified by RIA as previously reported (33). GLP-1 competitive binding was performed using 125I-GLP-1, kindly provided by Novo Nordisk (Copenhagen, Denmark).

Determination of cell growth and survival
Cells were seeded into 96-well plates (5 x 104 cells/well) before experimentation. In the proliferation experiments, after establishing metabolic quiescence in the absence of serum for 24 h (3 mM glucose RPMI 1640 with 0.1% BSA), cells were cultured in RPMI 1640 media (with 0.1% BSA) with agents (glucose, glucose + GIP/GLP-1/GH) for an additional 24 h. Thereafter, cells were washed with KRBH [115 mM NaCl, 4.7 mM KCl, 1.2 mM KH2PO4, 10 mM NaHCO3, 1.28 mM CaCl2, 1.2 mM MgSO4 containing 10 mM HEPES and 0.1% BSA (pH 7.4)] and frozen at -70 C until assayed. Cells were quantified using the CYQUANT assay system (Molecular Probes, Eugene, OR) according to the manufacturer’s protocol. Fluorescence was measured using a microplate fluorescence reader (Bio-tek FL600, Winooski, VT) with excitation/emission set at 400/500 nm. Final cell numbers were always greater than the initial number plated (5 x 104 cells/well) when assessing cellular proliferation.

Cell survival was assessed in the presence of prolonged serum starvation and glucose deprivation. Cells were initially deprived of serum and glucose for 24 h (RPMI 1640 with 0.1% BSA); thereafter, GIP or forskolin were added for an additional 24 h, and cell number was quantified. Thus, cells had been deprived of serum and glucose for 48 h, resulting in 50% cell death in the absence of GIP or forskolin (see Fig. 2CGo). Final cell numbers were always less than the initial number plated (5 x 104 cells/well) in assessing cell survival.



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  		FIG. 2. GIP potentiates 11 mM glucose induced cell growth (A and B) and promotes cell survival of INS-1 (832/13) cells (C and D). In all experiments, cells were serum starved before and during the course of the experiment. During cell growth studies (A and B), final cell numbers were always greater than initial plating densities, indicative of mitogenesis; in which final cell numbers were quantified fluorometrically by CYQUANT. In C and D, cells were serum and glucose starved for 48 h, with or without the addition of GIP for the final 24 h. Values are means of five (A), four (B), three (C), and four (D) individual experiments done in triplicate, in which * represents P < 0.05 (t test or ANOVA with Dunnett’s post hoc test).

 
Caspase activity
Cells seeded into 6-well plates were serum starved for 12–18 h (3 mM glucose RPMI 1640 with 0.1% BSA) and then subjected to glucose deprivation (RPMI 1640 with 0.1% BSA) or treatment with wortmannin or 2 mM streptozotocin (STZ). In studies examining the effects of glucose deprivation on caspase activity, 100 nM GIP were added concomitantly with 0 mM glucose-containing media. In studies in which inhibitors were used, all inhibitors (H89, UO126, wortmannin, or SB202190) were added 15 min before glucose deprivation ± GIP. Concentrations of inhibitors used were based on previous studies, and H89 has been shown to inhibit insulin secretion, ERK1/2 phosphorylation and rat insulin promoter activity at 5 µM and 10 µM (15). In STZ studies, GIP and GLP-1 were added 10 min before 2 mM STZ and for 30 min during STZ. Following treatment, caspase-3 activity was determined after 2, 6, or 24 h according to the manufacturer’s protocol [standard: 7-amino-4-methylcoumarin (AMC), substrate: Ac-DEVD-AMC, inhibitor: Ac-DEVD-CHO; Molecular Probes]. Caspase-3 activity per well was assessed using a microplate fluorescence reader (Bio-tek FL600, excitation/emission at 360/460 nM), and corrected for total protein content using the BCA protein assay (Pierce, Roxford, IL). Caspase-9-like activity was determined using the standard 7-amino-4-(trifluoromethyl)coumarin, and the fluorescent substrate Ac-LEID-7-amino-4-(trifluoromethyl)coumarin (Calbiochem, San Diego, CA), with excitation/emission at about 400/500 nm. The caspase-9 substrate can also be cleaved by caspases-4 and -5; hence, the designation caspase-9-like activity. Caspase-3 activity was measured after 30 min. at room temperature (manufacturer’s instructions), whereas caspase-9-like activity was assessed after 2 h incubation at 37 C. The Epac-selective cAMP analog (8-CPT-2OMe-cAMP) was purchased from BIOLOG Life Sciences Institutes (Bremen, Germany) and used at 100 µM, a concentration shown to have effects on calcium-induced calcium release in INS-1 cells (34).

Kinetworks KAPS 1.0 Western blotting analysis of the expression of 25 different apoptosis-related proteins was performed by Kinexus Bioinformatics Corp. (Vancouver, British Columbia, Canada) with 400 µg of INS-1 cell lysate subjected to wortmannin treatment in the absence and presence of GIP. Samples were confirmed to have elevated caspase-3 activity, which was reversed by GIP, based on substrate cleavage analysis before KAPS. This analysis detects caspases 1 {alpha}/ß, 2, 3, 5, 6, 7, 8, 9, and 12, in addition to other apoptosis-related proteins.

Immunoblot analysis
For determination of protein kinase phosphorylation, immunoblot analysis was performed. Cells were harvested and plated into 6-well plates (2 x 106 cells/well) 2 d before overnight serum starvation and subsequent stimulation was performed on d 3. Cells were stimulated in RPMI 1640 media containing the indicated glucose concentration and 0.1% BSA. Following the elapsed stimulation period, proteins from both floating and adherent cells were extracted with cellular lysis buffer (0.5% Triton X-100; 60 mM ß-glycerophosphate; 20 mM 3[N-morpholino]propanesulfonic acid (pH 7.2); 5 mM EDTA; 5 mM EGTA; 1 mM Na3VO4; 20 mM NaF; 1% Trasylol; and 1 mM phenylmethylsulfonyl fluoride). Thereafter, samples were sonicated (30 sec), centrifuged (12,000 rpm for 30 min), and protein content was quantified using the BCA reagent (Pierce) to ensure equal loading of gels for subsequent Western blotting. Pharmacological inhibitors (wortmannin and SB202190, Calbiochem) were added for 15 min before agonist addition and maintained in the presence of agonists. Protein samples (50 µg protein/well) were separated on a 13% sodium dodecyl sulfate (SDS)/polyacrylamide gel and transferred onto nitrocellulose (Bio-Rad Laboratories, Mississauga, Ontario, Canada) membranes. Probing of the membranes was performed with phospho-T202, Y204-ERK1 (p-ERK 1/2), purchased from Santa Cruz Biotechnologies (Santa Cruz, CA), and phospho-T180, Y182-p38 MAPK (p-p38 MAPK), phospho-S189/207 MKK3/6 (p-MKK3/6), and phospho-T71 ATF2 (p-ATF2) obtained from Cell Signaling Technology (New England Biolabs, Beverly, MA). Total protein was assessed using ß-tubulin antibody (Santa Cruz). Immunoreactive bands were visualized by enhanced chemiluminescence (Amersham Pharmacia Biotech, Buckinghamshire, UK) using horseradish peroxidase-conjugated IgG secondary antibodies. For quantification of band density, as a measurement of phosphorylation state, films were analyzed using densitometric software (Eagle Eye, Stratagene, La Jolla, CA).

RNA interference (RNAi) experiments
In experiments targeted at knockdown of Mek1, cells were plated at 5 x 105 cells/well in 6-well plates or 3 x 106 cells/well in 10-cm plates the day before transfection of double-stranded (ds)RNA oligonucleotides in media without penicillin/streptomycin. Cells were transfected using Oligofectamine (Life Technologies, Inc.) for 5 h according to the manufacturer’s instructions. The following 21-mer oligoribonucleotide was used corresponding to nt 364–384 of the rat Mek1 gene; 5'-AAC TCC CCG TAC ATA GTG GGC-3' with a 3' overhang of 2 nt. Designed RNA oligonucleotides were blasted against the GenBank/EMBL database to ensure gene specificity. The dsRNA was purchased from Dharmcon Research Inc. (Lafayette, CO) along with the control scramble 21-mer oligonucleotide, which was not present in mammalian cells as of January 23, 2002. Thus, the scramble duplex served as a negative control in RNAi experiments.

Protein levels of Mek1/2 were assessed over 4 d following transfection by immunoblot analysis, with d 3 showing peak knockdown. Thus, cells transfected in 10-cm dishes were plated into six-well dishes the day following transfection, and caspase-3 experiments were carried out on d 3 post transfection. Antibodies used were as follows: Mek 1 and Mek 2 antibodies (BD Biosciences, Mississauga, Ontario, Canada) had previously been screened for selectivity (15); total ERK1/2 and ß-tubulin antibodies (Santa Cruz Biotechnologies) were used to correct for total protein loaded.

Data analysis
Data are expressed as means ± SEM with the number of individual experiments presented in the figure legend. All data were analyzed using the nonlinear regression analysis program PRISM (GraphPad, San Diego, CA), and significance was tested using t test and ANOVA with the Dunnett’s multiple comparison test or the Newman-Keuls post test (P < 0.05) as indicated in figure legends.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
INS-1 (832/13) cells express functional GIP receptors, which stimulate cell growth
Because GIP receptors in the INS-1 clone 832/13 cell line had not been previously characterized, binding, adenylyl cyclase stimulation, and insulin secretory responses to GIP were initially studied (Fig. 1Go). Cells expressed receptors at a density of 1571 ± 289 binding sites/cell (n = 3) with an IC50 for binding of 21.1 ± 2.49 nM (n = 3) and a dissociation constant of 531 ± 22 pM (n = 3); cAMP production was stimulated by GIP with an EC50 of 4.70 ± 1.81 nM (n = 4); 5.5 mM glucose stimulated insulin secretion was potentiated by 10 nM GIP (1.63 ± 0.18% total insulin secreted for 5.5 mM glucose vs. 2.44 ± 0.29% total insulin secreted (P < 0.05, n = 3). These data compare well with other insulinoma cell lines tested for GIP binding, cAMP production, and insulin release (Refs. 35 and 36 ; and Ehses, J. A., and C. H. S. McIntosh, unpublished observations). Furthermore, we have also confirmed the presence of functional GLP-1 receptors on INS-1 (832/13) cells [IC50 for binding of 5.37 ± 0.62 nM (n = 3) and an EC50 of 1.37 ± 0.62 nM (n = 4) for cAMP production].



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  		FIG. 1. INS-1 (832/13) cells express functional GIP receptors coupled to cAMP production and insulin secretion. Cells were plated in 24-well plates at 5 x 105 cells/well 2 d before competitive binding (A), saturation binding (B), cAMP production (C), and insulin secretion (D) experiments. Protocols were as described in Materials and Methods with values representing means of three or four experiments performed in triplicate, in which * represents P < 0.05 (B is a representative plot).

 
The INS-1 cell line has been previously established by Hugl et al. (37) and Dickson et al. (38) as a cellular model for ß-cell proliferation. Based on previous work demonstrating GIP could regulate MAPK signaling in INS-1 cells (14, 15), we wanted to establish that GIP was indeed a growth and survival factor. GIP was found to potentiate 11 mM glucose mediated ß-cell proliferation over 24 h (Fig. 2AGo) to levels comparable with those obtained with GH [GIP stimulated growth to 158 ± 16% of that in 5.5 mM glucose, and GH promoted growth to 158 ± 9% of control (n = 3–5)]. In a separate experiment (Fig. 2BGo), 100 nM GIP stimulated cell growth to 131 ± 7% of that measured in the presence of 5.5 mM glucose, similar to the proliferative responses to 100 nM GLP-1 (129 ± 4%; n = 4).

GIP promotes cell survival via p38 MAPK
While determining the glucose dependence of these growth-promotive effects, it was observed that GIP was capable of reversing the detrimental effects of 0 mM glucose media (serum free) on cellular survival. Incubation of cells in the presence of 0 mM glucose media for 48 h resulted in approximately 50% cell death (Figs. 2CGo and 3AGo). Surprisingly, 91 ± 10% of the cells plated remained viable when the medium was supplemented with 100 nM GIP after 24 h. These cell survival effects of GIP were found to be concentration-dependent with an EC50 value of 1.24 ± 0.48 nM GIP (n = 4; Fig. 2DGo). Thus, GIP can indeed act as a growth and survival factor in INS-1 cells, even in the complete absence of glucose.



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  		FIG. 3. GIP promotion of INS-1 (832/13) cell survival during glucose deprivation involves p38 MAPK. Cells were serum and glucose starved for 48 h with or without the addition of GIP for the final 24 h. Final cell numbers were always less than cells plated, indicative of cell death. Protein kinase inhibitors were added to the medium 15 min before the final 24 h culture in the absence or presence of 100 nM GIP. The PKA inhibitor, H89, was unable to reverse GIP (A) or forskolin (B) mediated cell survival. Wortmannin had deleterious effects on cell survival (C), which was partially reversed by GIP. D represents the involvement of p38 MAPK via specific inhibition with SB202190. Final cell numbers were quantified fluorometrically by CYQUANT, and data represent means of three to eight experiments done in triplicate, in which * and # represent P < 0.05 vs. respective controls (t test).

 
To establish which intracellular signaling pathways were involved in the GIP-induced cell survival, studies were performed with pharmacological inhibitors used at concentrations shown to exhibit selectivity for candidate protein kinases (Fig. 3Go) (15). Stimulation of adenylyl cyclase with forskolin mimicked the effects of GIP on cell survival, but the failure to inhibit the effect of either agent with H89 (Fig. 3Go, A and B) indicated a PKA-independent mode of action. Neither of the Mek1/2 inhibitors PD98059 (50 and 100 µM) nor U0126 (10 µM) blocked the effects of GIP on cell survival (data not shown; n = 3). The ability of GIP to promote cell survival was further supported by studies on the effect of the PI3K-PKB pathway inhibitor, wortmannin (Fig. 3CGo). Interestingly, cells were partially protected against wortmannin-induced cell loss by GIP treatment (n = 3, P < 0.05). The only compound tested that influenced GIP-mediated cell survival was the inhibitor SB202190 (Fig. 3DGo), indicating that, as with GLP-1 (10, 24), GIP can act via modulation of p38 MAPK activity.

GIP and cAMP inhibit caspase-3 activity in INS-1 cells
Caspase activation is a marker for the induction of cellular apoptosis (39). To establish whether the cell survival effects of GIP were due to antiapoptotic actions of the polypeptide, activation of the effector caspase-3 and a screen for various caspases was conducted during glucose deprivation and wortmannin treatment. Figure 4AGo illustrates that 0 mM glucose promoted caspase-3 activity by 6 h (not by 2 h; data not shown, n = 3) and that this effect was completely reversed by the concurrent addition of GIP or forskolin. Greater stimulation of caspase-3 activity was achieved by wortmannin treatment, and responses were also completely inhibited by concomitant GIP treatment (P < 0.05, n = 3; see caspase-3 data, Fig. 5DGo). In further experiments, it was shown that GIP treatment 24 h after initiation of caspase-3 activity by glucose deprivation was also able to reverse caspase activation (data not shown; P < 0.05, n = 3). The selective caspase-3 inhibitor, Ac-DEVD-CHO, and immunoblotting for caspase-3, were used to ensure that measured protease activity was in fact caused by caspase-3 (Fig. 4BGo and data not shown). In a recent study from our laboratory, evidence was obtained for a protective effect of the incretins on ß-cells in STZ-induced diabetes in rats (40). Therefore, we examined the ability of GIP to protect against STZ-induced ß-cell death. When added 10 min before, and during, a 30 min STZ exposure, GIP and GLP-1 were both able to protect completely against the pro-apoptotic (caspase-3 activating) effects of STZ (Fig. 4CGo).



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  		FIG. 4. GIP and cAMP abrogate caspase-3 activation by 0 mM glucose (A and B) and STZ (C) in INS-1 (832/13) cells. Cells were serum starved before and during the experiment, and 100 nM GIP, 10 µM forskolin, or 100 nM GLP-1 were added for 6 h in the presence and absence of glucose (3 mM) or STZ to assess affects on caspase-3 activity. Caspase-3 activity was quantified using the substrate, Z-DEVD-AMC, over 30 min. Caspase-3 activity was corrected for total protein concentration using the BCA protein assay. Activity was ensured to be specific by using the caspase-3 inhibitor Ac-DEVD-CHO (C). All experiments are representative of n = 3 (A and B) or n = 5 (C), in which * and # represent P < 0.05 vs. respective controls (ANOVA with Newman-Keuls multiple comparison post hoc test).

 


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  		FIG. 5. GIP inhibition of caspase-3 activation is independent of PKA, Mek1/2, and PI3K. Protein kinase inhibitors were added to the medium 15 min before the initiation of further caspase-3 activation by glucose deprivation. GIP was able to completely reverse caspase-3 activation by 0 mM glucose (A) independent of PKA (B), Mek1/2 (C), and PI3K (D). Caspase-3 activity was assessed fluorometrically after 30 min of substrate incubation and corrected for total protein. Data are representative of n = 3–4 for all, in which * and # represent P < 0.05 vs. respective controls (ANOVA with Newman-Keuls multiple comparison post hoc test).

 
In support of these data, we have begun to screen for various apoptotic proteins involved in GIP suppression of INS-1 cell death. Although there was evidence for caspase-9 like activity in INS-1 cells that was completely reversed by GIP and elevated in the presence of wortmannin (P < 0.05, n = 3), no caspase-9 protein was detected. However, cleavage of caspase-1 {alpha} and ß, caspase-5, and poly (ADP-ribose) polymerase were all reduced by GIP treatment (Ehses, J. A., and C. H. S. McIntosh, unpublished observations).

MAPKs (Mek1/2-ERK1/2 and p38) regulate caspase-3 activity in INS-1 cells
A pharmacological approach was also taken to identify the GIP receptor-mediated signaling pathway responsible for inhibition of caspase-3 activation, with the objective of correlating these findings to those for survival. In support of the cell survival data, there were no apparent roles for PKA (H89, 5 and 10 µM tested), Mek 1/2 (UO126), or PI3K (wortmannin) in GIP or cAMP interaction with caspase-3 activation (Fig. 5Go, B–D). The effect of inhibitors was controlled for by quantifying ERK1/2 phosphorylation (data not shown, n = 3). Thus, we propose that GIP is acting in a cAMP-dependent, but PKA-independent manner in regulating cell survival via caspase-3 inhibition. We have previously shown that GIP can signal via Rap1 the small GTPase upstream of B-Raf, and we therefore tested the effect of various Rap1 constructs (wild type, constitutively active G12V, dominant negative N17) on GIP inhibited caspase-3 activation. There was no significant effect of any of these (data not shown; n = 2). Supportive of this, and unlike forskolin, the Epac-selective analog, 8-CPT-OMe-cAMP was not found to be capable of reversing caspase-3 activity (data not shown, n = 4).

We were somewhat surprised by the lack of effect of the Mek1/2 inhibitor on caspase-3 activation (Fig. 5CGo), given the role for this kinase in regulating the ERK1/2 module and cell survival in other systems. Because the Mek1/2-ERK1/2 pathway is highly regulated by GIP (14, 15, 23), we wanted to assess the effect of long-term inhibition of this pathway on caspase-3 regulation by glucose deprivation and GIP. RNAi studies were performed and found to have maximal effects on Mek1/2 protein levels on d 2 and 3 post transfection. Figure 6AGo depicts the level of protein knockdown achieved on d 3, with the caspase-3 activity measured on the same day (6B). The dsRNA oligonucleotide targeted at rat Mek1 also influenced Mek2 expression (Fig. 6AGo). It is evident that even partial removal of Mek1/2 results in a parallel reduction in caspase-3 activity under all conditions (basal activity was decreased from 5.6 ± 0.4 to 2.6 ± 0.4 pmol substrate cleaved/30 min; and maximal activity was also decreased from 18.5 ± 3.5 to 6.4 ± 0.7 pmol substrate cleaved/30 min; Fig. 6BGo). The overall elevated caspase-3 activity in these studies was due to the effects of cell transfection, which was also found to increase caspase-3 activity in the aforementioned Rap1 experiments. However, we do not believe this detracts from the interpretation of the results, given the similar profile of activity.



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  		FIG. 6. MAPKs Mek-ERK (A and B) and p38 (C) regulate caspase-3 activity in INS-1 cells. The dsRNA oligonucleotides were introduced into cells by Oligofectamine, and protein knockdown was greatest on d 3 (A), with no effect on total ERK1/2 protein levels. Scramble represents the dsRNA negative control; see Materials and Methods for details. Caspase-3 activity (B and C) was assessed on d 3 post transfection, and elevated levels are due to cell trauma endured during transfections. Mek1/2 knockdown caused overall diminished caspase-3 activity with minimal effects on fold induction by glucose deprivation. GIP reversal of caspase-3 activity was significantly affected; 97 ± 6% inhibition for scramble and 68 ± 10% for Mek1/2 RNAi; n = 3, P < 0.05 (B). The protein kinase inhibitor, SB202190, was added to the medium 15 min before the initiation of further caspase activation by glucose deprivation (C). Blots in A are representative and data in B and C are representative of n = 3, in which * and # represent P < 0.05 vs. respective controls (ANOVA with Newman-Keuls multiple comparison post hoc test).

 
Although the fold caspase-3 activation by glucose deprivation is relatively similar (3.3 ± 0.6-fold for scramble and 2.5 ± .03 for Mek1/2 RNAi; n = 3, P > 0.05), the reversal of caspase-3 activity by GIP is slightly abrogated by partial removal of Mek (97 ± 6% for scramble and 68 ± 10% for Mek1/2 RNAi; n = 3, P < 0.05). Thus, we believe that there is a modest role for Mek1/2 in the regulation of caspase-3 activity in INS-1 cells, although its exact contribution could not be quantified because of the inability to completely knockdown the Mek isoforms.

Because p38 MAPK was implicated in the above cell survival effects of GIP (Fig. 3DGo), we next examined its role in regulating INS-1 caspase-3 activity. Treatment of cells with SB202190 mimicked the effects of GIP and cAMP (Fig. 6CGo), promoting the hypothesis that GIP may act on caspase-3 activation by inhibiting stress-induced p38 MAPK signaling. It therefore appears that p38 MAPK is the major short-term regulator of caspase-3 activity in INS-1 cells, whereas Mek1/2-ERK1/2 may play a more modest role and regulate long-term caspase-3 activity.

GIP regulation of p38 MAPK phosphorylation/dephosphorylation: an underlying mechanism for regulating cell survival and caspase-3 activity
To test the hypothesis that reversal of caspase-3 activation by GIP results from inhibition of stress-induced p38 MAPK, phosphorylation of the p38 MAPK module (MKK3/6, p38 MAPK, ATF-2) by glucose deprivation was studied. In contrast to its effects on ERK1/2, glucose withdrawal leads to an activation of the p38 MAPK module as assessed by phospho-specific antibodies (Fig. 7AGo). Because caspase-3 activation was assessed at 6 h, we examined the phosphorylation of MKK3/6, p38 MAPK, and ATF-2 by incubation in 0 mM glucose over this time period. Activation was pronounced and exhibited slow kinetics, reaching a phosphorylation maximum for p38 MAPK at 4 h, followed by a decline. Thus, the 4-h time point was chosen to assess the effects of GIP on 0 mM glucose activation of p38 MAPK. As predicted, GIP and forskolin were both capable of reversing phosphorylation of p38 MAPK induced by either glucose deprivation or wortmannin treatment (Fig. 7Go, B and C). The specificity for activation of p38 phosphorylation was assessed using the inhibitor SB202190 (Fig. 7BGo). Although total protein levels were assessed, changes in such long-term phosphorylation may be due to differences in total p38 MAPK protein. Examination of upstream MKK3/6 phosphorylation, however, revealed no consistent effects of GIP or forskolin at the 4-h time point. In fact, in the presence of wortmannin, there was no phosphorylation of MKK3/6 or ATF-2, in contrast to the marked phosphorylation of p38 MAPK (data not shown and Fig. 7BGo). These results, therefore, suggest that GIP acting via cAMP is able to reverse caspase-3 activity by promoting long-term dephosphorylation of p38 MAPK (via a hereto-unidentified kinase/phosphatase; see Fig. 9Go).



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  		FIG. 7. GIP and cAMP (B and C) reverse long-term phosphorylation of p38 MAPK by glucose deprivation (A) or wortmannin treatment (B and C). Cells were plated in 6-well plates 2 d before experiments at 2 x 106 cells/well and serum starved overnight before glucose deprivation for the times indicated in (A). Fifty-microgram protein samples were separated by SDS-PAGE and membranes blotted with antibodies against phospho-T180, Y182-p38 MAPK (p-p38 MAPK), phospho-S189/207 MKK3/6 (p-MKK3/6), and phospho-T71 ATF2 (p-ATF2). Glucose deprivation promoted phosphorylation of all three kinases, in which p38 MAPK was maximally phosphorylated at 4 h (A). Based on these data, glucose deprivation was sustained for 4 h in the absence or presence of 100 nM GIP or 10 µM forskolin (B and C). GIP and forskolin were also able to reverse the activation of p38 MAPK in response to a more potent death stimulus, wortmannin (B and C). Data are representative blots of three (A and B) and two (+ wortmannin; B and C) independent experiments. The far right blot in A is a positive control depicting phosphorylation of p38 MAPK in cells treated with anisomycin, and the specificity of p38 MAPK phosphorylation by glucose deprivation and wortmannin treatment was determined using the inhibitor SB202190 (B).

 


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  		FIG. 9. Proposed mechanism underlying GIP regulation of INS-1 cell fate. Under cellular stress induction (A), GIP is able to reverse caspase-3 activation by promoting long-term dephosphorylation of p38 MAPK. In contrast, under normal conditions (B), GIP rapidly promotes phosphorylation of the p38 MAPK and ERK1/2 module in INS-1 cells. This is thought to contribute to the growth-promotive actions of GIP. Hypothesis based on data from the present and previous study (15 ). See text for details.

 
Because the other incretin hormone, GLP-1, has been shown to activate p38 MAPK and thereby regulate cell growth (10, 20), and in light of the results shown in Fig. 3Go, we looked at the potential of GIP to activate the p38 MAPK module under 3 mM glucose conditions and short-term kinetics (Fig. 8Go). GIP affected phosphorylation of the entire MKK3/6, p38 MAPK, ATF-2 module. Activation of MKK3/6 and p38 MAPK was transient, but significant, with an ensuing reduction in phosphorylation after 1 h that is in agreement with a trend toward dephosphorylation. However, downstream ATF-2 phosphorylation was stimulated by GIP treatment over the entire 2-h time course. This resembled the phosphorylation kinetics of ERK1/2 more than p38 MAPK because of the sustained activation at 2 h. Because Thr (71) of ATF-2 has recently been identified as a phosphorylation target of ERK1/2 (41), this may explain these observations and also lend insight into the discrepant findings of ATF-2 phosphorylation noted at 4 h in the study above. Thus, in addition to reversing sustained stress-induced p38 activation by glucose deprivation or wortmannin treatment, GIP is also able to regulate p38 MAPK through rapid and transient phosphorylation events (see Fig. 9Go). We propose therefore, that GIP dynamically regulates p38 MAPK, rapidly promoting phosphorylation of MKK3/6, p38, and ATF-2 under normal conditions, but promoting dephosphorylation of p38 over long-term stress conditions (no glucose or serum). These two regulatory events are proposed to regulate INS-1 cell growth and apoptosis respectively (see Fig. 9Go).



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  		FIG. 8. GIP regulates the phosphorylation of MKK3/6, p38 MAPK, and ATF-2 in INS-1 (832/13) cells. Cells were plated in 6-well plates 2 d before experiments at 2 x 106 cells/well and serum starved overnight before experiments were conducted in 3 mM glucose RPMI 1640 + 0.1% BSA. Fifty-microgram protein samples were separated by SDS-PAGE, and probing of the membranes was performed with phospho-T180, Y182-p38 MAPK (p-p38 MAPK), phospho-S189/207 MKK3/6 (p-MKK3/6), and phospho-T71 ATF2 (p-ATF2) obtained from Cell Signaling Technology, and phospho-T202, Y204-ERK1 (p-ERK 1/2), purchased from Santa Cruz Biotechnologies. Densitometry of blots in A are depicted in B. All blots are representative of at least three independent experiments and quantification of phosphorylation is representative of n = 3, in which * represents P < 0.05 vs. respective controls (ANOVA with Dunnett’s post hoc test).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
It has been suggested that the etiology of both type 1 and 2 diabetes mellitus involves a reduction in the mass of functional pancreatic ß-cells. To maintain euglycemia, ß-cell mass must be held relatively constant through a dynamic process that involves neogenesis and/or differentiation, proliferation, and apoptosis (42, 43). Only recently have the growth factors and hormones responsible for maintaining this equilibrium been identified, and these include glucose, insulin, prolactin, GH, IGF, and most recently the incretin, GLP-1. From recent work (16) and the current studies, it is evident that GIP stimulates ß(INS-1)-cell proliferation as well as promotes cell survival. The proliferative effects of GIP on INS-1 (832/13) ß-cells were comparable with those obtained with two established growth factors for pancreatic ß-cells, GH, and GLP-1, and the cell survival effects may be a common property of the glucagon superfamily because GLP-2, PACAP, and VIP have also been shown to exhibit this property (25, 27). Although these data are limited to cell models, we have recently found a dysregulation of islet size in GIPR -/- mice (44) in addition to a protective role for both GIP and GLP-1 in STZ-induced diabetic rats (40), implying a physiological role for GIP in the regulation of cell fate.

A decrease in glucose transport has recently been shown to be an essential component of the execution pathway during cytokine- and growth factor-induced cell death (45, 46). Clearly, glucose depletion or withdrawal is sufficient to activate the apoptotic cascade on its own (45, 46, 47). Glucose deprivation is commonly studied in the context of cardiac and neuronal hypoxia/ischemia; however, in the current study, we employed this treatment as a simple method of activating the apoptotic cascade and causing cell death. Our data support the existence of similar pathways for apoptotic cell death in ß-cells; cell numbers were decreased by serum-free 0 mM glucose media (Fig. 2CGo), caspase-3, and caspase-9-like activity were activated (Fig. 4Go and data not shown), and the stress kinase module, p38 MAPK, was activated (MKK3/6 -> p38 MAPK -> ATF-2) (Fig. 7AGo). Similar activation of SAPKs and p38 MAPK occur after withdrawal of nerve growth factor (NGF) or nutrients from PC12 cells or neuronal cultures (48).

GIP signals via cAMP in ß-cells and non-ß-cells, and to date all physiological actions of GIP have been shown to be dependent on this pathway (1, 5, 35, 49). We have recently shown that increasing cAMP is able to positively regulate the Raf -> ERK1/2 -> p90 RSK pathway in INS-1 cells (15) and, similar to neuronal cAMP actions, the regulation of the ERK1/2 module by cAMP in ß-cells seems to underlie its ability to regulate cell fate. Interestingly, the mitogenic actions of cAMP (which are cell-type specific) often correlate well with its survival actions (50, 51). Our present data support this notion, inasmuch as cAMP (forskolin stimulation) was also able to promote cell survival in ß cells (Fig. 3Go) by inhibiting caspase-3/9 activation (Fig. 4Go and data not shown) and reversing prolonged p38 MAPK activation (Fig. 7BGo). The secretin/glucagon peptide family members, PACAP and GLP-2, are also both able to reverse caspase-3 activity via a postulated cAMP-dependent pathway (25, 27, 28), implying a common signaling property of these family B GPCRs.

PKA-independent actions of incretins have been the focus of recent studies (34, 52). Intriguingly, the actions of GIP and forskolin on survival and caspase-3 activity in INS-1 cells appear to be independent of both PKA and Epac/Rap1 (Figs. 3Go and 5Go, and data not shown; n = 4). We have obtained further evidence supporting this mechanism of action in the phospho-regulation of CREB family transcription factors by GIP (manuscript in preparation). Although these cAMP actions are contrary to the currently accepted paradigm, recent studies have demonstrated similar cAMP regulation independent of PKA and Epac and/or Rap1 in neutrophil apoptosis (53) and melanocyte mitogenesis (54, 55). Thus, it appears that there may exist an unidentified cAMP exchange factor that is critical for regulating cell fate in various tissues.

Our data also suggest that the survival actions of GIP are partially PI3K independent in INS-1 cells because wortmannin was unable to affect the influence of GIP on cell survival, caspase activity (capase-1 {alpha}/ß, caspase-3, caspase-5), or p38 MAPK phosphorylation (Figs. 3Go, 5Go, and 7BGo). These findings concur with GLP-2 studies (in the case of reversing cycloheximide-induced cell death), in which GLP-2 actions were also shown to be PI3K and Mek1/2 independent and being mimicked by cAMP (27). A recent study (16), published during the preparation of this manuscript, also identified a PKA- and PI3K-independent mechanism for GIP-mediated cell survival. The present findings extend these studies, however, by demonstrating that GIP is able to regulate cell fate through regulation of stress-induced p38 MAPK activity. Cyclic AMP has also been reported to directly regulate both PKB and GSK-3ß, independently of PI3K activity (56, 57). Interestingly, in recent studies of GLP-2 actions it was suggested that cAMP can act on GSK-3 downstream of PKB (28). Because both PKB and GSK-3ß have also been shown to be regulated by GIP in INS-1 cells (14), a common property of the secretin/glucagon receptor family may be to promote cell survival via cAMP-dependent regulation of PKB and GSK-3ß. Nevertheless, the current study indicates that regulation of p38 MAPK is central to the antiapoptotic actions of GIP and cAMP in INS-1 cells.

It is generally accepted that the balance between ERK and p38/JNK pathways determines whether an extracellular stimulus promotes (ß) cell growth or acts in a detrimental manner (apoptotic) (58, 59). Prolonged p38 MAPK activation targets cells to apoptosis, whereas selective activation of ERK generally prevents apoptosis and ensures cell survival. Thus, GIP presents itself as a prime candidate for the enhancement of cell viability by inhibiting long term p38 MAPK activation, while concurrently activating ERK1/2 rapidly and for a sustained period (Figs. 7BGo and 8Go) (14). Furthermore, caspase-3 activity is functionally regulated by both these kinases in ß-cells; GIP regulates caspase-3 activity primarily by p38 MAPK (Figs. 6CGo and 8Go) and also through Mek1/2 kinases (ERK1/2 activators; Figs. 6BGo and 8Go). The effects of GIP on ERK1/2, however, are insufficient to account for the survival and caspase-3 inhibition facilitated by GIP in ß-cells (based on inhibition with PD98059 and U0126, and RNAi studies). This contrasts with the recent proposal that the ERK1/2 pathway is central to the antiapoptotic actions of GIP (16). However, these studies were conducted in elevated glucose conditions (7.5 mM), making it impossible to distinguish actions of GIP from the autocrine insulin actions, which were absent in the present study.

Interestingly, there are now reports that rapid, transient activation of p38 and/or JNK are correlated with cellular proliferation, whereas prolonged activation of these stress pathways results in cellular execution (59, 60, 61). In neuronal cells, NGF is able to transiently activate p38 MAPK and thereby regulate neuronal differentiation in PC12 cells (62, 63). Furthermore, transient and persistent activations of ERK1/2 also lead to different cell fates because transient activation of ERK by epidermal growth factor stimulates cell growth in PC12 cells, and sustained activation is also implicated in neuronal differentiation by NGF (64). Our present study, in addition to previous work, illustrates that GIP is able to activate MKK3/6 -> p38 MAPK transiently while inhibiting its long-term activation by stress and concurrently activating ERK1/2 (and ATF-2) both rapidly and persistently. Thus, based partly on GLP-1 studies (10), we propose that GIP is able to regulate cell survival via rapid/transient activation of p38 MAPK (proliferative stimulus) and long-term inhibition in response to stress (antiapoptotic actions; see schematic Fig. 9Go).

Although much research is now being conducted on mapping the signal transduction networks of immune-mediated ß cell apoptosis (58, 65), there is a paucity of information regarding prosurvival hormones and their regulation of these networks. In the present study, insight is provided into the role of MKK3/6, p38 MAPK, ATF-2, and Mek1/2 in the survival actions of GIP on INS-1 (832/13) cells. GIP is able to reverse caspase activation in a cAMP-dependent manner through the dynamic regulation of p38 MAPK phosphorylation; furthermore, we propose that GIP is able to regulate both cell growth and cell death via dynamic control of the p38 MAPK module. It is possible that these two pathways are regulated by different isoforms of p38 MAPK. Indeed, recent data suggest that in cardiomyocytes, Jurkat and HeLa cells, p38{alpha} is proapoptotic, whereas p38ß promotes survival (66). This may explain some of the discrepancies in the present study involving the inhibitor SB202190, which is known to inhibit both of these p38 isoforms (67). A modest role for the Mek1/2 (and hence ERK1/2) MAPK pathway in the regulation of caspase-3 activity by GIP was also identified, suggesting that the regulation of ß-cell fate by GIP includes the dynamic regulation (transient vs. sustained phosphorylation) of both p38 and ERK1/2 MAPK modules. The regulation of stress kinase pathways by GIP is intriguing because they are also used in immune-mediated ß-cell attack. Our study therefore further supports the notion that GIP may be therapeutically useful for the modulation of ß-cell growth and survival and suggests that an additional contributing factor to type 2 diabetes, in which GIP actions are blunted (68), may be a lack of GIP-mediated proliferative/survival signals at the level of the ß-cell.


   Acknowledgments
 
We are grateful for the technical assistance of Cuilan Nian and the insight of S. L. Pelech.


   Footnotes
 
This work was supported by the Canadian Institutes of Health Research (Grant MOP-13192, to R.A.P. and C.H.S.M.) and the Department of Science and Technology of Sachsen Anhalt (HUD 9704/00116). J.A.E. and J.A.P. are funded by the Canadian Institutes of Health Research and the Michael Smith Foundation.

Abbreviations: AMC, 7-Amino-4-methylcoumarin; ds, double stranded; GIP, glucose-dependent insulinotropic polypeptide; GLP, glucagon-like peptide; GPCR, G protein-coupled receptor; GSK, glycogen synthase kinase; JNK, c-jun N-terminal kinase; Mek, MAPK kinase; NGF, nerve growth factor; PACAP, pituitary adenylate cyclase activating polypeptide; PI3K, phosphatidylinositol-3 kinase; PKA, protein kinase A; PKB, protein kinase B; RNAi, RNA interference; SAPK, stress-activated protein kinase; SDS, sodium dodecyl sulfate; STZ, streptozotocin; VIP, vasoactive intestinal polypeptide.

Received November 21, 2002.

Accepted for publication June 10, 2003.


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 Introduction
 Materials and Methods
 Results
 Discussion
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