Mutations to the Third Cytoplasmic Domain of the Glucagon-Like Peptide 1 (GLP-1) Receptor Can Functionally Uncouple GLP-1-Stimulated Insulin Secretion in HIT-T15 Cells

Anne Marie F. Salapatek, Patrick E. MacDonald, Herbert Y. Gaisano and Michael B. Wheeler

Departments of Medicine and Physiology University of Toronto Toronto, Ontario, Canada M5S 1A8


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Glucagon-like peptide-1 (GLP-1) is an insulinotropic hormone with powerful antidiabetogenic effects that are thought to be mediated by adenylyl cyclase (AC). Recently, we generated two GLP-1 receptor mutant isoforms (IC3–1 and DM-1) that displayed efficient ligand binding and the ability to promote Ca2+ mobilization from intracellular stores but lacked the ability to couple to AC. In the present study, the wild-type rat GLP-1 receptor (WT-GLP-1 R) or the IC3–1 and DM-1 mutant forms were expressed for the first time in the insulin-producing HIT-T15 cells. Only cells expressing WT-GLP-1 R displayed dramatically elevated GLP-1-induced cAMP responses and elevated insulin secretion. The increase in GLP-1-stimulated secretion in cells expressing WT-GLP-1 R, however, was not accompanied by differences in glucose-stimulated insulin release. Prolonged exposure to GLP-1 (10 nM, 17 h), not only led to an increase in insulin secretion but also increased insulin mRNA levels, but only in cells expressing the WT-GLP-1R and not the mutant isoforms. Electrophysiological analyses revealed that GLP-1 application enhanced L-type voltage-dependent Ca2+ channel (VDCC) currents > 2-fold and caused a positive shift in VDCC voltage-dependent inactivation in WT-GLP-1R cells only, not control or mutant (DM-1) cells. This action on the Ca2+ current was further enhanced by the VDCC agonist, BAYK8644, suggesting GLP-1 acts via a distinct mechanism dependent on cAMP. These studies demonstrate that the GLP-1 receptor efficiently couples to AC to stimulate insulin secretion and that receptors lacking critical residues in the proximal region of the third intracellular loop can effectively uncouple the receptor from cAMP production, VDCC activity, insulin secretion, and insulin biosynthesis.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The incretin hormone glucagon-like peptide-1 (GLP-1) is released from intestinal L cells to stimulate insulin secretion from pancreatic ß-cells in a glucose-sensitive manner. The biologically active forms of GLP-1, GLP-1(7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37), and GLP-1(7–36 amide), have been shown to be among the most potent insulinotropic agents identified to date in mammals (1). Clinical studies indicate that GLP-1 not only stimulates insulin secretion in normal subjects, but also in those with non-insulin-dependent diabetes mellitus (type 2) (2, 3, 4), supporting its therapeutic potential. A possible role for GLP-1 in the central control of feeding has also been demonstrated (5). GLP-1 administered via an intracerebroventricular injection was a powerful inhibitor of feeding in fasted rats.

The insulinotropic properties of GLP-1 are mediated through a high-affinity GLP-1 receptor on the insulin-secreting ß-cells of the pancreas (6, 7). The receptor cDNA, initially cloned from rat pancreatic islet cells, and subsequently from human pancreas (8, 9), predicts a seven-transmembrane G protein-coupled receptor (GPCR) of the glucagon/vasoactive intestinal peptide/secretin receptor subfamily (10). Work on the endogenous receptor in isolated ß-cells and ß-cell lines, and with the recombinant GLP-1 receptors expressed in cell lines, strongly suggests that the insulinotropic actions of GLP-1 are mediated by cAMP-dependent activation of protein kinase A (11). The mechanisms whereby protein kinase A (PKA) mediates GLP-1-induced insulin exocytosis appears to be multifaceted including proposed actions on ATP-sensitive K+ channels, nonselective cation channels, L-type voltage-dependent Ca2+ channels (VDCCs), and on the exocytotic machinery (12, 13, 14, 15, 16).

The majority of studies correlating GLP-1-induced insulin secretion with PKA have relied heavily on the use of cAMP analogs with varying degrees of specificity for the activity of this kinase. In the present study, we have used a novel approach to correlate the insulinotropic actions of GLP-1 with cAMP, employing a series of recombinant GLP-1 receptor isoforms that are specifically uncoupled from adenylyl cyclase (AC). Previously, we and others reported that the third intracellular loop (IC3) and the carboxyl-terminal (CT) domain were shown to contain specific amino acids required for efficient signaling of the receptor (17, 18, 19, 20). Two adjacent amino acid block deletion mutations in the predicted N-terminal portion of the IC3 domain (DM-1, lacking V331-I332-A333, and IC3–1, lacking K334-L335-K336) were shown to be required for the efficient coupling of receptor to AC (18, 19) when expressed in COS cells. In the present study these mutant receptor isoforms and the recombinant WT-GLP-1R have been examined functionally in the insulin-producing ß-cell line HIT-T15.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
GLP-1R Expression
To examine the level of expression of endogenous GLP-1Rs in HIT-T15 cells and to assess the level of expression of the wild-type (WT) GLP-1R and mutant isoforms, Northern blotting and standard competitive binding displacement assays were performed (Fig. 1Go and Table 1Go). Using the rat receptor as a probe, under moderate stringency, GLP-1R transcripts were not observed in HIT cells (data not shown); however, specific binding was detected, albeit at extremely low levels (Fig. 1Go, A and B, and Table 1Go). Cells transfected with pCDNA-3 ß-gal demonstrated high-level transient transfection efficiency (45–80%, data not shown). Transfection also resulted in the efficient expression of receptor cDNAs as determined by Northern blotting (data not shown). Binding assays revealed that transfection resulted in the functional expression of the receptor cDNAs with specific binding (Bmax) of the WT-GLP-1R, IC3–1, and DM-1 (3049 ± 478, 3347 ± 530, 2939 ± 843 cpm/106 cells, respectively) found to be similar. The affinity for binding (IC50) was not found to differ significantly among the receptor isoforms (WT-GLP-1R, 6.5 ± 1.7 nM; IC3–1, 4.6 ± 2.0 nM; and DM-1, 3.4 ± 1.8 nM, n >= 6). These data are consistent with the Bmax and IC50 values reported for these receptor isoforms expressed in COS-7 cells (18, 19).



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Figure 1. Expression of GLP-1R Constructs in HIT-T15 Cells

Binding displacement of 125I-GLP-1 in control HIT-T15 cells and cells expressing the GLP-1R isoforms. Data are expressed as specific binding in counts per min (A) and percentage B/Bo (B).

 

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Table 1. Characterization of GLP-1 R Binding in Transfected HIT-T15 Cells

 
Analysis of GLP-1-Stimulated Insulin Secretion
To analyze the effects of GLP-1 in HIT-T15 cells overexpressing the receptor isoforms, cells were prepared as described in Materials and Methods. The patterns of basal cAMP production (5 mM glucose with no peptide) and insulin secretion were not significantly different among the test groups (Fig. 2Go, A and B). GLP-1 (10 nM), in the presence of 5 mM glucose, elicited a large cAMP response (Fig. 2CGo) in cells expressing WT GLP compared with control cells (13.9 ± 2.4 vs. 4.6 ± 1.0 pmol/well, respectively), which was accompanied by a significant increase in insulin secretion compared with control cells (60 ± 9 vs. 30 ± 7 ng/well/h, Fig. 2DGo). Increases in cAMP accumulation and insulin secretion in response to GLP-1 were considered significant compared with control cells and those expressing the mutant receptor isoforms (n >= 7; P <= 0.001 and P <= 0.01 for cAMP and insulin secretion, respectively). Collectively, these studies suggest a strong correlation between cAMP accumulation induced by GLP-1 and insulin secretion. They also demonstrated that the receptor can be functionally uncoupled from cAMP and insulin secretion in a ß-cell line through a modification to the proximal portion of the third intracellular loop.



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Figure 2. Basal and GLP-1-Stimulated cAMP Accumulation and Insulin Secretion in HIT-T15 Cells

HIT-T15 cells transfected with WT-GLP-1R or receptor mutants were analyzed for total cAMP content and insulin secretion rate under basal conditions (5 mM) glucose (A and B) and in the presence of 10 nM GLP-1 (C and D) over a 2-h test period.

 
Effects on Glucose-Stimulated Insulin Secretion
GLP-1R overexpression has been shown to increase glucose responsivity in RIN cells in the absence of a GLP-1 stimulus (21). To examine this possibility in HIT-T15 cells, those expressing the WT-GLP-1R were examined for responsiveness to glucose and compared with control cells and those expressing the cAMP-defective mutant isoforms. Basal (0 mM glucose) insulin secretion (Fig. 3Go) was not found to differ among control, WT-GLP-1R, IC3–1, and DM-1 transfection groups (15.2 ± 2.0, 14.0 ± 1.2, 15.3 ± 1.1, and 13.0 ± 2.1 ng/well/h, respectively; P > 0.05, n >= 7). Ten millimolar glucose elicited approximately a 2-fold increase in insulin secretion; however, no significant difference was observed among control, WT-GLP-1R, IC3–1, and DM-1 transfectants (36.3 ± 3.0, 37.3 ± 3.9, 39.5 ± 3.9, and 38.6 ± 4.9 ng/well, respectively; P > 0.05, n >= 7).



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Figure 3. Comparison of WT-GLP-1R with Deletion Mutations on Glucose-Stimulated Insulin Secretion

Cells in each test group were treated in the absence or presence of 10 mM glucose and insulin measured over a 2-h test period.

 
Effects on Insulin Biosynthesis
In COS-7 cells expressing DM-1 or IC3–1, receptor coupling to AC was dramatically reduced (18, 19). To examine the effects of receptor isoform overexpression on cAMP accumulation and insulin-secretory function in insulin-secreting cells, the HIT-T15 cell line was transfected with either expression vector, WT-GLP-1R, or DM-1. For these studies, cAMP content, insulin secretion rate, total insulin content, and insulin mRNA abundance were examined under basal conditions and in the presence of GLP-1 (10 nM or 1 µM) during a 17-h treatment period. As shown in Fig. 4AGo, under basal conditions, cAMP content was not significantly different among control cells and the test groups (control, 5.4 ± 0.4 pmol/well; WT-GLP-1R, 6.1 ± 1.0 pmol/well; and DM-1, 6.6 ± 1.1 pmol/well; n >= 6, P > 0.77). Insulin secretion rate over the 17-h period (Fig. 4BGo) in cells expressing control, WT-GLP-1R, or DM-1 were also not found to differ significantly (23 ± 3, 28 ± 3, and 22 ± 4 ng/well/h, respectively; n = 8). Total cellular insulin content (Fig. 4CGo) was also similar in control and WT-GLP-1R and DM-1 transfected cells (1618 ± 371, 1454 ± 300, and 1601 ± 327 ng/well, respectively; n = 8, P > 0.9). GLP-1 treatment increased cAMP accumulation and insulin secretion maximally at 100 nM; however, insulin content was not affected by exposure to GLP-1 for the 17-h test period (Fig. 4Go, A–C). GLP-1 failed to significantly change these parameters in control and DM-1-transfected cells. Northern blot analysis and subsequent quantification (insulin/18 s, Fig. 4DGo) revealed a significant increase in proinsulin gene transcripts in cells expressing WT-GLP-R at the 10 nM GLP-1 concentration (n = 3, P = 0.05), with similar levels being observed in control cells and those expressing the DM-1 mutant. No significant differences in transcript levels were observed under basal conditions or in the presence of 1 µM GLP-1 (Fig. 4DGo). These studies appear to indicate that GLP-1 has more profound effects on insulin secretion than on biosynthesis.



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Figure 4. Comparison of WT-GLP-1R with Deletion Mutations on Insulin Biosynthesis under Basal Conditions and in the Presence of GLP-1 over a 17-h Test Period

cAMP accumulation in the presence of 0, 10 nM, and 1 µM GLP-1 (panel A), insulin secretion (panel B), total cellular insulin content (panel C), and insulin mRNA abundance as presented in relative density units (R.D.U.) (panel D).

 
Effects on L-Type VDCC Currents
Cells expressing the WT-GLP-1R or DM-1 receptors tagged with enhanced green fluorescent protein (EGFP) were shown to bind GLP-1. Only cells expressing WT-GLP-1R showed a GLP-1-mediated increase in cAMP (data not shown) similar to cells expressing WT-GLP-1R. Specific HIT cells expressing the receptors or control cells (cells expressing EGFP only), were identified under UV light (480 nm) (typical GFP-positive cells shown in Fig. 5Go). When studied under current clamp conditions with standard high K+ pipette solution, the mean resting membrane potentials were not significantly different for that recorded for control compared with WT-GLP-1R-expressing cells, -61.4 ± 2.2 mV vs. -59.3 ± 1.5 mV. To enhance VDCC current amplitude and reduce the rate of VDCC run-down, Ba2+ was used as the charge carrier and resulted in a mean peak steady state Ba2+ current of -206 ± 54 pA at +10 mV (Fig. 6BGo, control). Nifedipine (10 µM) or Cd2+ (50 µM) markedly suppressed the inward current (not shown). Taken together, these results indicate that the current was carried by Ba2+ through L-type VDCCs. There was no difference in VDCC current magnitude under basal, unstimulated conditions among WT-GLP-1R, DM-1, or control cells (Fig. 6Go). As shown in Fig. 6Go, B and C, GLP-1 (10-8 M) caused a leftward shift in the current-voltage curve with a marked increase in peak VDCC current magnitude over the -30 to +30 mV range in WT-GLP-1R compared with control cells (-487 ± 68 pA and -201 ± 51 pA at +10 mV, respectively; P < 0.001, n = 31 cells). GLP-1 had no effect on DM-1 cells or control cells (Fig. 6CGo). These effects on VDCC current magnitude and voltage dependence are consistent with the positive shift in the steady state inactivation curve observed after GLP-1 (10-8 M) application in WT-GLP-1R cells (Fig. 7Go). This shift was expressed by parameters calculated from the Boltzmann function (as described in Materials and Methods) in which there was a significant shift in the half-maximal voltage of inactivation (V1/2) and the slope factor (k) in the basal (untreated) state compared with treatment with GLP-1 (V1/2: -26.45 ± 2.5 mV and -16.29 ± 4.3 mV; and k: 6.6 ± 0.4 mV and 5.2 ± 0.7 mV, respectively; P < 0.01, n = 16 cells). To gain insight into the mechanism of action of GLP-1 on VDCC, the combinatory effects of the VDCC agonist BAYK8644 (BAYK) and GLP-1 were tested (Fig. 8Go). BAYK (10 µM), added alone, increased VDCC current to the same extent in WT-GLP-1R, DM-1, and control cells (an additional increase in VDCC current of 56.4 ± 7.1%, 73.5 ± 11.3%, 64.1 ± 13.5% over basal levels, respectively; see Fig. 8CGo; P < 0.01, n = 15 ). After pretreatment with BAYK (10 µM), GLP-1 (10-8) application further increased VDCC currents in WT-GLP-1R cells only (an additional 62.3 ± 17.2% over BAYK addition; P < 0.01 n = 12 cells; see Fig. 8CGo). After pretreatment with GLP-1 (10-8), BAYK (10 µM) addition caused a further increase in VDCC current in WT-GLP-1R cells only (an additional 29.8 ± 9.9% over GLP-1 addition, P < 0.01 or by a greater than 2-fold increase (171.1 ± 8.5%) over BAYK addition alone; P < 0.01, n = 10 cells; see Fig. 8B).



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Figure 5. HIT-T15 Cells Expressing WT-GLP-1R-GFP

Cells transfected with WT-GLP-1R-GFP are viewed under standard light conditions (A) or under a UV light source (480 nm) (B) approximately 48 h posttransfection.

 


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Figure 6. Effect of GLP-1 on VDCCs in GLP-1R-Expressing Cells

A, Representative sequential VDCC current traces from a single WT-GLP-1 R cell in which one voltage step protocol was applied in which cells are held at -70 mV and stepped up to +10 mV. VDCC current is increased significantly with 10-8 M GLP-1 addition. B, Current-voltage curve for WT-GLP-1R cells in the basal, unstimulated state (open circles) and after application of GLP-1 (10-8 M). C, Mean peak steady state VDCC current measured in the presence of increasing GLP-1 doses in control (open bars), DM-1 (solid bars), and WT-GLP-1R (hatched bars) cells. VDCC current was significantly altered by GLP-1 (10-8 M) in WT-GLP-1R cells alone.

 


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Figure 7. Effect of GLP-1 on WT-GLP-1R VDCC Inactivation

A, Using a standard double-pulse protocol pictured at the top, representative sequential VDCC traces are shown obtained from a single WT-GLP-1R cell under basal and GLP-stimulated (10-8 M) conditions. B, Mean steady state inactivation curves for VDCC current obtained under basal (open circles) and GLP (10-8 M)-stimulated (filled circles) conditions. The fitted curves are obtained from the Boltzmann equation as described in the text.

 


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Figure 8. Effects of GLP-1 and BAYK8644 on VDCC Currents

A, Superimposed VDCC current traces from one cell obtained using a one-step voltage protocol, illustrating the sequential, cumulative responses in the basal state (labeled 1), after GLP-1 (10-8 M) (labeled 2) and after BAYK (labeled 3) in control, WT-GLP-1R, and DM-1 cells. B, Mean peak steady state VDCC currents measured in the basal state (open bars) and after the sequential, cumulative addition of GLP-1 (10-8 M) (hatched bars) and then BAYK (solid bars). C, Mean peak steady state VDCC currents measured in the basal state (open bars) and after the sequential, cumulative addition of BAYK (solid bars) and then GLP-1 (10-8 M) (hatched bars).

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
It is postulated that G protein activation results from the formation of a high-affinity complex between a GPCR, its ligand, and the heterotrimeric G protein. Studies involving the structural alteration of two closely related members of the B class of GPCRs (GLP-1R and the glucagon receptor) provide strong evidence that the cytoplasmic (IC) domains of these receptors facilitate G protein activation. Chicchi et al. (22) using a series of deletion mutations to the glucagon receptor showed complete loss of coupling to AC with a receptor isoform lacking eight residues in IC2 domain and dramatically reduced coupling with a series of deletions directed to the IC3 domain. Employing a similar strategy to the IC2 loop of the GLP-1R, we also described decreases in GLP-1-activated cAMP production but attributed this attenuation to reduced functional receptor expression. Heller et al. (20) reported that a point mutation to R348G compromised AC activation by GLP-1, supporting a role for the IC3 loop in G protein coupling within the GLP-1R. We have also characterized the IC3 loop of the GLP-1R, between residues K334-K351, using a series of deletion and substitution mutations scanning the region (18, 19). As a logical extension of these studies, we have now examined the functional consequences of this uncoupling on GLP-1-ß-cell signal transduction.

In COS cells expressing DM-1 or IC3–1, receptor coupling to AC was dramatically reduced compared with cells expressing WT-GLP-1R, while basal production was not affected (18, 19). In HIT-T15 cells transfected with WT-GLP-1R, DM-1, or IC3–1, basal cAMP content was also not significantly different (Fig. 2AGo) nor were the levels of basal insulin secretion (Fig. 2BGo). These studies were repeated in growth (Fig. 4AGo) or serum-free media (data not shown) with similar negative results, suggesting that a biologically active GLP-1 component of the serum is negligible. The similarity in cAMP levels over 2- and 17-h test periods (Figs. 2Go and 4Go) in the absence of exogenous GLP-1 also suggests that HIT-T15 cells are not producing significant amounts of biologically active GLP-1. Interestingly, insulin-producing RIN 1046–38 cells stably expressing the rat WT-GLP-1 displayed elevated basal cAMP levels and insulin secretion, elevations that were evident in the absence and presence of glucose in the incubation medium (21). The authors suggest several possibilities for the elevated responses, including the possibility that the receptor is constitutively active when overexpressed in RIN cells or that the cells may be responding to small amounts of endogenous GLP-1 secreted by the cells. In our transient assays in HIT cells there is no evidence to suggest that the recombinant GLP-1R is appreciably activated in the basal state.

The mechanisms whereby GLP-1 exerts its insulinotropic activity have been under intense study in recent years (reviewed in Ref. 1). Collectively, these studies suggest that the actions of GLP-1 are multifaceted, with several targets for action in the ß-cell. Targets and mechanisms include the inhibition of ATP-sensitive K+ channels to facilitate cell depolarization (12, 15), excitatory effects on VDCCs to increase [Ca2+]i, including a suppression of time-dependent inactivation (23, 24), and potentiation of activation of L-type VDCC (23). Recent studies suggest that GLP-1 could also exert an effect on insulin release at a level distal to an elevation in [Ca2+]i (15, 25). Although there may be several targets for GLP-1 action in the ß-cell, pharmacological agents that inhibit the PKA pathway appear to negate the effects of GLP-1 on KATP, VDCC, or events distal to cell depolarization and Ca2+ influx. Increases in cAMP accumulation and insulin secretion in response to GLP-1 were not significant in control cells. These data are in contrast to studies by Lu et al. (13), who showed an increase in cAMP accumulation and insulin secretion in HIT-T15 cells. Although the cells used in the previous study were not examined for GLP-1R expression, it is highly likely that the level of receptors would be considerably higher than we report in the present study. Our data support previous studies, which found that functional GLP-1Rs are indeed expressed in our HIT-T15 cell line, but the level of expression is extremely low (Fig. 1Go). This low level of GLP-1R expression made this insulin-secreting cell line ideal for the present overexpression studies. Indeed, GLP-1 (10 nM) elicited a large cAMP response (Fig. 2CGo) in cells expressing WT-GLP-1R that was accompanied by a concomitant increase in insulin secretion (Fig. 2DGo). These data demonstrate a strong correlation between cAMP accumulation induced by GLP-1 and insulin secretion. Increases were not observed in cells expressing either of the mutant receptors. GLP-1 treatment had no effect on inward currents in the absence of glucose (control or WT-GLP-1R cells; see Fig. 6Go).

Since previous reports demonstrated that GLP-1 had direct stimulatory effects on voltage-clamped L-type VDCC in rat ß-cells (23, 24), we conducted a series of experiments to compare the effects of GLP-1 on voltage-clamped L-type VDCC activity in WT-GLP-1R, mutant (DM-1) GLP-1R, and control (GFP expressing only) cells. Similar to the findings of Suga et al. (23), GLP-1 had signficant stimulatory effects on L-type VDCC activity, acting to increase peak VDCC current amplitude and shift the current-voltage relationship leftward in WT-GLP-1R-expressing cells (Fig. 6Go). In fact, we observed a 142% increase in VDCC current magnitude with 10-8 M GLP-1 in WT-GLP-1R cells, an effect that was much larger than the 30% increase with 2 x 10-8 M GLP-1 reported by Suga et al. (23). This finding suggests that the increased GLP-1R expression in WT-GLP-1R cells acts to amplify the actions of GLP-1 on VDCC. In addition to reported GLP-1-stimulated changes in time-dependent inactivation, which act to slow VDCC inactivation (24), GLP-1 caused a rightward shift in voltage-dependent inactivation (Fig. 7Go). This positive shift in inactivation would contribute to increased VDCC availability and contribute to the observed GLP-1-induced changes in the voltage dependence of VDCC currents.

The lack of GLP-1 response in mutant and control cells was not a result of nonfunctional VDCC expression, since the voltage dependence or stimulatory effects of the L-type VDCC-specific agonist, BAYK8644 (BAYK), were preserved and similar in all cells (Fig. 8Go). Since BAYK is not known to increase [cAMP]i (26), this finding would support our hypothesis that AC uncoupling leads to the abolished GLP-1 response in DM-1 cells. The slight upward trend in the VDCC response to GLP-1 but its failure to reach significance in control cells suggests that low GLP-1R expression is likely responsible. This hypothesis is supported by previous reports, which also demonstrated that the actions of GLP-1 on VDCC are mediated through the AC pathway via changes in [cAMP]i (11, 23, 24). Previous studies in mouse ß-cells have demonstrated that elevation of [cAMP]i or activation of PKA by forskolin (24, 27, 29) caused little increase in VDCC current magnitude but caused a slowed time course of inactivation. We find that GLP-1 caused marked changes in VDCC current magnitude and voltage-dependent inactivation in VDCC. In fact, the GLP-1-mediated changes in VDCC activity we observed are similar to those reported for cardiac myocyte VDCCs phosphorylated by PKA and protein kinase C (28, 29).

To further explore the mechanism of action of GLP-1 on VDCC, we performed studies on the combinational effects of GLP-1 and BAYK (Fig. 8Go). BAYK is known to act at an extracellular site on VDCC to cause VDCC current potentiation. Although BAYK effects occur without elevation of [cAMP]i (26), its potentiating effects are positively modulated by cAMP-dependent phosphorylation (29). In fact, in a recent study on newborn rat cardiac myocytes, the degree to which BAYK stimulated VDCC was found to be a good indicator of the degree of VDCC phosphorylation in developing myocytes (30). In our studies, the cumulative stimulatory effect of GLP-1 and BAYK on VDCC activity was greater than either drug added alone, irrespective of their order of addition. These findings support the putative actions of GLP-1 to increase [cAMP]i and positively modulate the actions of BAYK (Fig. 8Go, A and B).

GLP-1R overexpression has been shown to increase glucose responsivity and secretion in RIN cells in the absence of a GLP-1 stimulus (21). Using reverse hemolytic plaque assays, Rafizadeh et al. (21) demonstrated that this increase in glucose-mediated secretion could be explained by an increase in the number of glucose-responding cells. In contrast, islets isolated from GLP-1R-deficient mice appeared to display normal glucose responsivity (31). To examine glucose responsivity, HIT-T15 cells expressing the WT-GLP-1R were examined and compared with control cells and those expressing the cAMP-defective mutant isoforms. Basal (0 mM glucose) insulin secretion (Fig. 3Go) was not found to differ among control, WT-GLP-1R, IC3–1, and DM-1 transfection groups, and glucose either 5 or 10 mM elicited a characteristic 1.5- to 2-fold increase in insulin secretion in all groups examined. This was supported by the fact that mean resting membrane potentials were not significantly different from that recorded for control, DM-1-, or WT- GLP-1R-expressing cells. These results suggest that in the absence of GLP-1, in HIT cells, the GLP-1R remains primarily inactive and does not influence glucose competence. They also suggest that in the case of RIN cells overexpressing the GLP-1R (21), the possibility that the receptor prefers an active conformation is supported, and that elevated cAMP accumulation renders the cells more sensitive to glucose.

Binding of GLP-1 to its ß-cell receptor stimulates not only insulin secretion but also increases insulin mRNA production (32), likely via induction of insulin gene transcription through a cAMP-dependent mechanism (33, 34). In keeping with this hypothesis, mice deficient in the GLP-1R have reduced insulin gene expression and reduced total pancreatic insulin content (35). In the present study, insulin secretion rate over a 17-h culture period was compared with total insulin content and insulin mRNA abundance, to examine the effects of overexpression of the WT-GLP-1R and the cAMP-defective mutants on insulin biosynthesis. Total insulin content (Fig. 4CGo) was similar in control and WT-GLP-1R-transfected cells in the presence or absence of a prolonged GLP-1 stimulus capable of significantly elevating cAMP and insulin secretion. It is possible that the increased secretion rate may have offset any increase in insulin synthesis resulting from a GLP-1 stimulus. Northern blot analysis and subsequent quantification (insulin/18 s, Fig. 4DGo) revealed a small but significant increase in proinsulin gene transcripts in cells expressing WT-GLP-R in the presence of GLP-1 (10 nM), consistent with a role for GLP-1 on insulin gene transcription. However, the correlation is not entirely clear, since cAMP levels are maximal at 100 nM GLP-1, when the increase in insulin mRNA did not reach statistical significance. Perhaps future studies in models that will allow a more prolonged expression of the DM-1 mutant isoform may allow a more clear correlation between insulin gene transcription and biosynthesis to be observed. Nevertheless, the present studies clearly demonstrate that the GLP-1R efficiently couples to AC to stimulate insulin secretion, actions mediated via effects on the VDCC. Furthermore, receptors lacking critical residues in the proximal region of the third intracellular loop can efficiently uncouple the receptor from cAMP production, L-type VDCC action, and insulin secretion.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Binding Assays
Synthetic human GLP-1(7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36) amide (Bachem Torrance, CA) was used in all binding studies. Radioiodination was accomplished by the chloramine-T method as previously described (19). The [125I]GLP-1(7–36 amide) product was purified by reverse phase adsorption to a C-18 Sep-pak column (Waters Associates, Milford, MA) and had a specific activity of approximately 125–250 µCi/µg. Whole-cell binding assays were performed as previously described (19). GLP-1R and GLP-1R mutations were generated as previously described (18, 19). For binding assays, 8 x 106 HIT-T15 cells (gift from Dr. Paul Robertson, Pacific Northwest Research Institute, Seattle WA) between passage 72 and 90 cultured in RPMI-1640 medium supplemented with 10% FBS, 1% Penicillin/Streptomycin, and 1% L-glutamine were seeded into 10-cm plates and transfected with 20 µg of expression plasmids (pcDNA3 vector alone (control); WT-GLP-1R; V331-I332-A333-deletion mutant (DM-1); and K334-L335-K336-deletion mutant (IC3–1)] using Pfx1 lipid reagent (cells were incubated in the transfection media for 3.5–4 h) according to the product specifications (Invitrogen, Carlsbad CA). Binding assays were carried out 48 h posttransfection. The cells were washed twice in PBS and recovered from plates with 2 mM EDTA in PBS. Cells (~1 x 106/tube) were incubated for 45 min at 37 C in binding buffer (RPMI containing 0.4% glucose, 1% BSA, pH 7.4) with radiolabeled tracer [125I]GLP-1 amide (100,000 cpm, ~270 pM) and unlabeled GLP-1 at concentrations of 10-12 to 10-6 M, in a final volume of 200 µl. Cell suspensions were centrifuged at 12,000 x g, and the cell-associated radioactivity was counted (Cobra II, Canberra Packard, Meriden, CT). Specific binding (total binding less nonspecific binding) measured in the presence of excess (1 µM GLP-1) was determined for the WT and each mutant receptor. Binding characteristics, including specific binding and IC50, were calculated from competitive binding-displacement curves generated using curve-fitting software (Prism, GraphPad Software, Inc., San Diego, CA) as we have previously reported (19).

Insulin and cAMP Assays
Twelve-well plates were seeded with 4 x 105 cells per well in a total volume of 1 ml. The cells were incubated for 48 h and then transfected with 2 µg of plasmid DNA (three wells of pcDNA3 vector alone, one well for ß-gal plasmid, four wells for WT-GLP-1R, four wells for DM-1 or IC3–1 mutant) using Pfx1 as described above. After a 48 h posttransfection incubation period, the culture media were replaced.

Insulin Assays
The following day (17 h), medium was replaced with 1 ml of Krebs-Ringer buffer (KRB) containing 0.1% BSA (RIA grade), 0.238% (10 mM) HEPES, pH 7.4, and incubated twice for 30 min, after which the cells were washed twice with 1 ml of KRB buffer and then with 2 ml buffer. The final wash was replaced with 1 ml of experimental buffer containing KRB, 0.1% BSA (RIA grade), 0.238% (10 mM) HEPES, pH 7.4, and 5 mM glucose and stimulated with 10-8 M GLP-1 for 2 h. GLP-1 was prepared from lyophilized samples on the day of assay and added from concentrated stocks. For glucose-stimulation assays, the protocol remained the same with the exception of the glucose concentration (0 mM, 5 mM, or 10 mM). For overnight secretion experiments, the growth media were replaced after 48 h with media containing 0, 10 nM, or 1 µM GLP-1 and collected for assay after the 17-h test period. In all cases, 700 µl of the experimental media from each well were transferred to a microfuge tube and spun at 3000 rpm for 2 min. The top 300 µl of supernatant were transferred to a new tube and stored at -70 C. Insulin RIAs were performed using rat insulin RIA kit from Linco (St. Charles, MO). Total cell insulin content was determined using acid extraction as previously described (36).

cAMP Assays
Immediately after media were collected for insulin RIA, the cells were washed in cold PBS, and intracellular cAMP was extracted with 80% ethanol. Lyophilized samples were reconstituted in sodium acetate buffer (pH 6.2) and cAMP production was measured by RIA (Biomedical Technologies, Stoughton, MA). ß-Galactosidase assays were performed to assess transfection efficiency on each multiwell plate using the manufacturer’s protocol (Invitrogen, San Diego, CA). If efficiencies below 40% were observed with ß-galactosidase, the cells were not used in assays.

Northern Blot Analysis
Plates (10 cm) were seeded with 8 x 106 HIT-T15 cells and transfected with 20 µg of plasmid as described for binding assays. After 48 h, the plates were washed twice with PBS, and total cellular RNA was extracted using 2.5 ml of TRIzol reagent (Life Technologies, Inc., Burlington, Ontario, Canada) according to the protocol provided. Total RNA (25 µg) from HIT-T15 cells was suspended in sample buffer (6% formaldehyde, 50% formamide, 100 µl of 1x MOPS, 10% glycerol, and bromophenol blue). The RNA was denatured and run on a 1.2% agarose-formaldehyde denaturing gel and transferred to nylon membranes (Amersham Pharmacia Biotech, Oakville, Ontario, Canada) as previously described (37, 38). cDNA probes were prepared using the random primer labeling kit from Life Technologies, Inc.. A partial hamster insulin cDNA was obtained by RT-PCR on HIT-T15 cell RNA to yield a 350-bp fragment corresponding to coding sequence. The probe used to detect GLP-1R transcripts was a Kpn-I/HincII fragment of the rat GLP-1R (kindly provided by Bernard Thorens, Institute of Pharmacology and Toxicology, Lausanne, Switzerland) The 18S probe was generated as previously described (31). The blots were hybridized at 40 C overnight as previously described (37, 38) and washed with 0.5x SSC and 0.1% SDS at 55 C for 30 min. Densitometry was performed as previously described to quantitate insulin transcripts (35). Briefly, the autoradiogram was scanned and a constant size area was used to convert the intensity of the bands to pixels.

Electrophysiological Assays
WT GLP-1R or DM-1 cDNAs lacking a stop codon were generated by PCR where the primers were designed to amplify a product where the stop codon was removed. This product was first cloned into PCR 2.1 (Invitrogen) and then into the HindIII-SmaI sites of pEBFP-N2 (CLONTECH, Palo Alto, CA). Cells transfected with a WT-GLP-1 or the DM-1 mutant receptors tagged at the C terminus with EGFP. Control cells were transfected with pEBFP-N2 alone. All cells were lightly trypsinized (0.05% trypsin), washed with extracellular solution, and allowed to equilibrate and adhere to a patch clamp study chamber that was mounted on an inverted microscope (CK-2, Olympus Corp., Lake Success, NY). Membrane currents through L-type VDCC were recorded using standard whole-cell patch clamp techniques. To block K+ flux and observe large inward currents through L-type VDCCs, which were not prone to run-down, extracellular solutions were used that contained the following: 20 mM BaCl2, 90 mM NaCl, 5 mM CsCl, 1 mM MgCl2, 10 mM glucose, and 10 mM HEPES. Intracellular solutions in which K+ was replaced with cesium were used: 75 mM Cs2-aspartate, 1 mM MgCl2, 20 mM tetraethylammonium chloride (TEA)-Cl, 5 mM EGTA, 4 mM ATP-Mg, and 20 mM HEPES. Patch pipettes were prepared from 1.5-mm thin-walled borosilicate glass using a two-state patch-pipette puller (model pp83, Nari-shige, Tokyo, Japan). Pipette tips were fire polished to resistances of 4–5 M{Omega}. The current flow between the pipette and the bath solution was compensated to achieve a zero baseline before seal formation. Standard tight-seal recording techniques for seal formation were used, and access to the interior of the cell was obtained by further suction to rupture the patch membrane.

All electrophysiological experiments were performed at 22–24 C according to Hamill et al. (39), on representative cells expressing EGFP. Currents were measured with an Axopatch-1D patch clamp amplifier (Axon Instruments, Foster City, CA), filtered with a Bessel filter (-3 decibels at 1 kHz) and recorded online by a computer (IBM PC) using pCLAMP (v.6) software (Axon Instruments). Whole-cell capacitance was routinely measured measured by the intermittent initial testing by cancellation of the capacity transient and measured on average 10.5 ± 0.6 pF. The average series resistance was 5.2 ± 1.5 M{Omega}. Neither cell capacitance nor series resistance was electronically compensated. VDCC currents were elicited by a protocol in which cells were incrementally depolarized in +20 mV steps that were held for 200 ms from -70 mV to +70 mV from a holding potential of -70 mV and peak steady state currents were measured at 250 ms. Inward, Ca2+ currents were assessed between drug additions by a single depolarizing step to +10 mV. ß-Cells studied in this manner could be routinely patch clamped for up to 1 h with no significant change in cell currents or membrane potential. Steady state inactivation curves were obtained using a two-pulse protocol. From a holding potential of -70 mV, a 15-sec depolarizing conditioning pulse to different voltages was followed by a 250-msec test pulse to the voltage at which the maximal VDCC currents were obtained (+10 mV). Conditioning and test pulses were separated by a 20-msec return to the holding potential (-70 mV). Steady state inactivation curves were normalized by dividing the current amplitude (I) during the test pulse by the maximal amplitude obtained in the absence of a conditioning pulse (Imax). The data were fitted with the Boltzmann function using pClamp6 software: I/Imax = {1 + exp[Vm-V1/2)/k}-1 where I/Imax is the relative current, Vm is the membrane potential, V1/2 the half-maximum voltage of inactivation, and k is the slope factor. V1/2 and k were determined for each current obtained from individual cells.

Stock solutions or lyophilized drugs tested were stored at -22 C with each aliquot being defrosted once and used over a 6-h study period. All drugs were added to the chamber in microliter volumes, and routine controls with the vehicles used for dissolution were done to exclude nonspecific effects of the diluent. GLP-1 was lyophilized and distilled water added. BAYK8644 (+/-, Calbiochem, La Jolla, CA) was prepared in dimethylsulfoxide to 10-3 M stock and diluted further to 10 µM.

Statistics
All values are expressed as the mean ± SEM of at least three independent observations unless stated otherwise. Statistical analysis was performed using one way ANOVA followed by Tukey’s postanalysis (InStat, GraphPad Software, Inc., San Diego, CA).


    FOOTNOTES
 
Address requests for reprints to: Michael B. Wheeler, Department of Physiology, University of Toronto, 1 Kings College Circle, Toronto, Ontario, Canada M5S 1A8.

This work was funded by grants to M.B.W. from the Medical Research Council of Canada (MT-12898) and the Canadian Diabetes Association, and to H.Y.G. and M.B.W. from the Eli Lilly & Co./Banting and Best Diabetes Centre Research Program.

Received for publication February 16, 1999. Revision received April 14, 1999. Accepted for publication April 26, 1999.


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

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