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Departments of Medicine and Physiology University of Toronto Toronto, Ontario, Canada M5S 1A8
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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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.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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and Table 1). 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. 1, A and B, and Table 1). 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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, A and B). GLP-1 (10
nM), in the presence of 5 mM glucose, elicited
a large cAMP response (Fig. 2C) 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. 2D). 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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) 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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, 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. 4B) 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. 4C) 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. 4, 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. 4D) 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. 4D). These studies appear to indicate that
GLP-1 has more profound effects on insulin secretion than
on biosynthesis.
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). 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. 6B, 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. 6). As shown in Fig. 6, 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. 6C). 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. 7). 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. 8). 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. 8C;
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. 8C). 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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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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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. 2A
) nor were the levels of basal insulin secretion (Fig. 2B). These
studies were repeated in growth (Fig. 4A) 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. 2 and 4)
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. 1). 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. 2C) in cells expressing WT-GLP-1R that was
accompanied by a concomitant increase in insulin secretion (Fig. 2D).
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. 6).
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. 6). 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. 7). 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. 8).
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. 8). 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. 8, 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. 3) 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. 4C) 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. 4D) 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.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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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
. 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. 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).
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FOOTNOTES |
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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.
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