Departments of 1Metabolic Diseases and 2Pharmacology, Graduate School of Medicine, University of Tokyo, Tokyo 113-8655; 3CREST of Japan Science and Technology Corporation, Saitama 332-0012; 4Institute for Diabetes Care and Research, Asahi Life Foundation, Tokyo 100-0005, Japan; and 5Department of Biochemistry, School of Medical Sciences, University of Bristol, Bristol BS8 1TD, United Kingdom
Submitted 13 December 2002 ; accepted in final form 10 March 2003
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ABSTRACT |
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glutamate metabolism; alkalinization of vesicular pH; regulation of insulin secretion
Recently, two important findings were reported regarding a relationship
between glutamate metabolism and insulin secretion. A new form of persistent
hyperinsulinemia with hypoglycemia of the infant (PHHI) was demonstrated to be
caused by an excessive activity of glutamate dehydrogenase, which catalyzes a
conversion between glutamate and a TCA cycle substrate, -ketoglutarate
(26,
49,
50). Second, glutamate
produced via
-ketoglutarate from glucose was reported to enhance
insulin secretion under conditions of clamped cytosolic
[Ca2+] and [ATP] at high levels
(29). The insulin secretion
stimulated by glutamate was blocked by an inhibitor of vacuolar type
H+-ATPase or by an inhibitor of vesicular glutamate transporter.
Although the direction of metabolic flux between glutamate and
-ketoglutarate upon stimulation with glucose has been controversial in
-cells (18,
25), these results raised a
novel postulation that glutamate might play a role in transducing secretory
signals from glucose metabolism to secretory vesicles and that this pathway
might involve a modulation of secretory vesicle pH, the acidity of which is
thought to be generated mainly by vacuolar-type H+-ATPase
(9,
21,
22,
33).
Here, we studied the roles of glucose and glutamate in the regulation of vesicular pH and glucose-stimulated insulin secretion. Our results indicated that glucose was capable of acutely alkalinizing secretory vesicle pH and that this effect was dependent on glucose metabolism but not on cytosolic [Ca2+] elevation. Glutamate dimethyl ester (GME), a cell-permeable analog of glutamate (10, 46, 55), potentiated glucose-stimulated insulin secretion without changing cellular ATP content or cytosolic [Ca2+]. Application of GME reproduced the alkalinizing effect of glucose on vesicular pH at the basal glucose concentration. These results suggested that glucose metabolism increased secretory vesicle pH, at least in part, through generation of glutamate, which may be related to an alternative pathway of insulinotropic effect of glucose to the KATP channel-dependent pathway.
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MATERIALS AND METHODS |
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Plasmid construction. Plasmid phogrin-Ds red was generated by subcloning an AgeI-NotI fragment bearing Ds Red cDNA from plasmid prepro-atrial natriuretic factor (kindly provided by Dr. David K. Apps, University of Edinburgh, Scotland, UK) into phogrin-EGFP, previously digested to remove the EGFP-encoding region.
Preparation of islets. Islets were isolated by collagenase digestion and manual picking from the pancreata of C57BL6J mice at 34 mo old (54). These islets were used for experiments immediately after isolation.
Insulin secretion from islets. Insulin secretion from pancreatic islets was measured in static incubation or perifusion incubation with Krebs-Ringer bicarbonate (KRB) buffer composed of (in mM) 129 NaCl, 4.8 KCl, 1.2 MgSO4, 1.2 KH2PO4, 2.5 CaCl2, 5 NaHCO3, 10 HEPES (pH 7.4), and 0.2% bovine serum albumin (15). In static incubation, batches of 10 islets were preincubated at 37°C for 30 min in KRB buffer containing 2.8 mM glucose. The preincubation solutions were replaced with KRB buffer containing test agents, and batches of islets were incubated at 37°C for 60 min. Insulin released into supernatants was measured by radioimmunoassay. In perifusion incubation, 30 islets were suspended in 500 µl of Bio-Gel G-10 beads in each perifusion chamber and perifused with KRB buffer at 37°C at a flow rate of 0.6 ml/min. Islets were perifused for 30 min in the presence of 2.8 mM glucose before stimulation. Effluent fractions were periodically collected, and insulin in these samples was measured by radioimmunoassay.
Fluorescence study. For monitoring intraluminal pH of
insulin-secretory vesicles, islets were loaded with 2 µM DND-189 for 30 min
at 37°C in Sol AII buffer composed of (in mM) 150 NaCl, 5 KCl, 1
MgCl2, 2 CaCl2, and 10 HEPES (pH 7.4)
(3,
20). After completion of the
dye loading, a single islet on a glass coverslip in a perifusion chamber was
placed on the stage of an inverted fluorescence microscope IX70 (Olympus,
Osaka, Japan) equipped with a charge-coupled device camera (Photometrics). The
islet was viewed under a water immersion objective LUMPlanFL lens (Olympus)
with an excitation wavelength of 373 nm. Images of 256 x 256 pixels (170
x 170 µm) were obtained every 20 s with an exposure time of 100 ms.
Image analysis was carried out using the IPLab program (Signal Analysis). A
standard curve for DND-189 fluorescence was obtained by dissolving DND-189 in
the intracellular solution (in mM: 136 KCl, 4 NaCl, 5 MgCl2, 5
glucose, and 20 HEPES) with varying pH values from 5.0 to 6.0. For monitoring
cytosolic pH, islets were loaded with 1 µM BCECF for 30 min at 37°C and
excited with alternate wavelengths of 440 and 490 nm
(23,
44,
47,
48). The emission intensity
from islets when excited at 490 nm compared with that at 440 nm of BCECF's
isosbestic point was measured and plotted on a standard curve made with the
intracellular solution with varying pH values from 6.0 to 8.0. For monitoring
cytosolic [Ca2+], islets were loaded with 15 µM fura
2-acetoxymethyl ester at 37°C for 60 min in Sol AII buffer and excited
with alternate wavelengths of 355 and 380 nm
(4,
17). Cytosolic
[Ca2+] was expressed as ratios of the emission
intensities excited at 355/380 nm. For double staining of -cells with
DND-189 and phogrin-Ds red, a confocal microscopic examination was performed
with the FLUOVIEW system and an oil-immersion objective UPlanApo lens
(Olympus). Dispersed single
-cells on a coverslip were transfected with
0.4 µg of plasmid encoding cDNA for phogrin-Ds red by Effectene
transfection reagent (Qiagen, Hilden, Germany)
(37,
38). After 48 h, islets were
loaded with 1 µM DND-189 for 30 min at 37°C. The fluorescence of
DND-189 was excited at 488 nm, and that of phogrin-Ds red was excited at 568
nm. The image analysis was performed with FLUOVIEW software (version 2.0.32,
Olympus).
Glucose oxidation. Oxidation of [U-14C]glucose in islets was measured by generation of 14CO2 (54). Batches of 10 islets were incubated at 37°C for 90 min in KRB buffer containing the isotope. The 14CO2 generated in the buffer was made volatile by an addition of HCl, captured in NaOH, and measured with liquid scintillation counting.
ATP content. To determine ATP content in islets, batches of 10 islets were incubated at 37°C for 60 min in KRB buffer. The incubation was stopped by addition of ice-cold HClO4, and islets were homogenized by sonication. The homogenates were neutralized by addition of NaOH. ATP content in the supernatants was measured with an ATP bioluminescence assay kit, using known amounts of ATP as internal controls (54).
Statistical analysis. Statistical analysis was performed using Student's t-test for unpaired comparisons and analysis of variance. Values were presented as means ± SE.
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RESULTS |
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Next, islets loaded with DND-189 were examined under perifusion conditions.
When the glucose concentration of the perifusate was raised from 2.8 to 22.2
mM, DND-189 fluorescence was decreased by 0.66 ± 0.10% at 148.9
± 16.0 s (Fig.
1C). Because the pH of secretory vesicles has been
reported to be 5.5 (21),
we estimated that the fluorescence decrease corresponded to an increase in pH
by 0.016 ± 0.002 unit, using a standard curve in
Fig. 1B, in which a
decrease in DND-189 fluorescence by 0.41% indicated an increase in pH by 0.010
unit at
pH 5.5. For comparison, the cytosolic pH of islet cells was
monitored with a pH-sensitive fluorescent probe, BCECF
(23,
44,
47,
48). In response to 22.2 mM
glucose stimulation, cytosolic pH immediately and transiently rose by 0.02
± 0.01 unit (Fig.
1D). After the pH returned to baseline level, it was
further decreased below the level by 0.04 ± 0.01 unit but was finally
restored to the baseline level. Thus the profile of cytosolic pH change in
response to glucose was distinct from that of secretory vesicles in their
phasic pattern and time course.
Glucose-induced vesicular alkalinization is dependent on glucose metabolism. To study the mechanism of glucose-induced alkalinization of secretory vesicles, we examined effects of glucose metabolism and cytosolic [Ca2+] on the phenomenon. The nonmetabolizable glucose analog 3-O-methylglucose, at 19.4 mM, was unable to induce a decrease in DND-189 fluorescence (Fig. 2A) (30). Inhibitor of glycolysis monoiodoacetic acid at 1 mM (data not shown) or inhibitor of mitochondrial ATP synthase oligomycin at 10 µg/ml (Fig. 2B) completely blocked the effect of glucose on secretory vesicle pH (16, 24, 57). These results indicated that the vesicular alkalinization was dependent on glucose-metabolizing steps finally leading to ATP generation in mitochondria. We next studied a relationship between elevating cytosolic [Ca2+] and vesicular alkalinization. Diazoxide is known to suppress glucose-induced Ca2+ influx into the cytosol by opening the KATP channels on the plasma membranes (19, 45). Indeed, the drug at 250 µM completely inhibited glucose-stimulated insulin secretion (data not shown), indicating that Ca2+ influx into the cytosol failed to occur. The alkalinizing change of vesicles in response to 22.2 mM glucose was preserved in the presence of 250 µM diazoxide (Fig. 2C). When islets were stimulated with 50 mM KCl in the presence of 2.8 mM glucose, no changes in DND-189 fluorescence were observed (Fig. 2D). Glibenclamide at 5 µM, which closed the KATP channels and thereby activated Ca2+ influx into the cytosol, was also unable to induce the fluorescence change at 2.8 mM glucose (data not shown). These results indicated that cytosolic [Ca2+] elevation was neither necessary nor sufficient for the glucose-induced vesicular alkalinization and that the alkalinizing effect of glucose was mediated through its metabolic processes but independently of the closure of KATP channels and subsequent elevation of cytosolic [Ca2+].
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GME enhances glucose-stimulated insulin secretion without modulating
the KATP channel-dependent pathway. In the
next series of experiments, we examined the possibility that the effect of
glucose on secretory vesicle alkalinization might be relevant to intracellular
generation of glutamate. It has been reported that cellular concentrations of
glutamate were increased after stimulation of -cells with glucose
(29), probably via breakdown
of glucose molecules through glycolysis and the TCA cycle, generation of
-ketoglutarate in the cycle, and its conversion to glutamate by the
enzyme glutamate dehydrogenase. Insulin secretion in response to 22.2 mM
glucose was potentiated by GME (Fig.
3A), a cell-permeable analog of glutamate
(10,
46,
55). The effect of GME was
dose dependent until 1 mM, but the higher dose of 5 mM was employed thereafter
in the present study. This was because the dose makes it easier to compare our
results with the previous ones
(29,
46). GME at 5 mM increased the
secretion by 123 ± 29%. This potentiating effect of GME at 5 mM was not
observed at 2.8 or 5.6 mM glucose but was evident at higher concentrations of
glucose, indicating that GME was not an initiator of insulin secretion but a
potentiator of glucose stimulation (Fig.
3B). In a perifusion study, 5 mM GME potentiated both the
first-phase and the second-phase secretions in response to 22.2 mM glucose
(Fig. 3C).
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Next, we examined whether this effect of GME was mediated through activated
oxidation of glucose, which should lead to an increase in ATP content of
islets. [U-14C]glucose oxidation was not affected by 5 mM GME at
2.8 mM glucose (Fig.
4A). However, GME decreased [U-14C]glucose
oxidation by 28 ± 8% at 22.2 mM glucose
(Fig. 4A). These
results suggested that application of GME might decrease glucose-stimulated
generation of ATP in islets. Thus we next measured ATP content of islets
(Fig. 4B)
(54). Stimulation of islets
with 22.2 mM glucose significantly increased ATP content from 10.2 ±
0.4 to 13.9 ± 1.2 pmol/islet. In the presence of 5 mM GME, ATP content
was 13.7 ± 0.3 pmol/islet at 22.2 mM glucose, and this value was
comparable to that in the absence of GME. GME at 0.52.0 mM also failed to
significantly change ATP content of islets stimulated with 22.2 mM glucose
(Fig. 4B). Thus,
although application of GME decreased glucose oxidation, we assumed that
overall ATP generation from oxidation of both glucose and GME-derived
glutamate remained largely unchanged. ATP is known to play a crucial role in
inhibiting the KATP channels, thereby depolarizing the plasma
membranes and elevating cytosolic [Ca2+]
(1,
5). Consistent with the
unchanged ATP content of islets, elevation of cytosolic
[Ca2+] in response to 22.2 mM glucose (from 0.37
± 0.04 to 0.55 ± 0.06; Fig.
4C, top) was not changed by a simultaneous
application of 5 mM GME (from 0.36 ± 0.03 to 0.53 ± 0.05;
Fig. 4C,
bottom). These results suggested that GME's potentiating effect on
glucose-stimulated insulin secretion was exerted not through augmenting ATP
generation or elevating cytosolic [Ca2+] but mainly
through nonoxidative utilization of glutamate in the
-cells.
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GME mimics the alkalinizing effect of a high concentration of glucose. We next examined whether GME modified the pH of secretory vesicles in perifusion experiments. At 2.8 mM glucose, application of 5 mM GME decreased DND-189 fluorescence by 0.83 ± 0.19% (an increase in pH by 0.020 ± 0.005 unit) at 37.9 ± 2.6 s (Fig. 5), which was comparable in magnitude to that of 22.2 mM glucose stimulation (Fig. 1C). However, the time required for attaining the level was significantly less (P < 0.01) than that of 22.2 mM glucose stimulation (Fig. 1C).
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DISCUSSION |
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The alkalinizing change of vesicular pH in response to a glucose challenge was monotonous and lasted more than 10 min, and thus it was totally distinct from the pH change of the cytosolic space in response to glucose (Fig. 1D), which has been reported to have a biphasic pattern (44, 48). Upon stimulation with glucose, cytosolic pH quickly moved to an initial transient alkalinization. This was explained by mitochondrial consumption of H+ ions in the course of glucose metabolism. Next, the pH shifted to acidification, which was thought to be coupled with Ca2+ influx from the extracellular space and the action of HCO3-/Cl- exchanger. These results further supported that the fluorescence of DND-189 originated from secretory vesicles but was not contaminated by the fluorescence of the cytosolic space, as was also demonstrated by the copresence of the DND-189 fluorescence with that of vesicular membrane-bound phogrin-Ds red (Fig. 1A) (37, 38).
We assumed glutamate to be a candidate for molecular links between glucose
metabolism and acute vesicular alkalinization. Intracellular concentration of
glutamate was reported to be increased in -cells exposed to glucose
(29), probably via generation
of
-ketoglutarate through a series of steps in glycolysis and the TCA
cycle, and finally via its reductive amination reaction into glutamate
catalyzed by glutamate dehydrogenase. In fact, glutamate, which was derived
from GME by intracellular removal of dimethyl moiety
(10,
46,
55), mimicked the alkalinizing
effect of glucose on the vesicular pH (Fig.
5). However, the times required to attain these fluorescence
changes were quite different. The longer time required for the effect of
glucose to occur probably corresponded to a time for metabolizing glucose
until it becomes glutamate through the intermediate substrate
-ketoglutarate. It was known that it took
2 min for the
mitochondrial electrochemical membrane potential to be fully formed after
glucose stimulation (16),
which well reflected the activity of the TCA cycle and thus generation of
-ketoglutarate in the cycle.
We observed that GME at 5 mM was effective in potentiating insulin
secretion in response to 22.2 mM glucose in mouse islets. Previous reports
indicated that the potentiating effect of GME was exhibited at low-to-moderate
concentrations of glucose but disappeared at higher concentrations (16.7 to 25
mM) in rat islets and rat -cell-derived INS-1 cells
(29,
46). One possible
interpretation of these discrepancies is the difference in efficiencies of
intrinsic glutamate generation from glucose. Thus rat
-cells might be
active in generating glutamate from glucose and an additional application of
external glutamate, as a form of GME, failed to show the potentiating effect.
In contrast, the effect was manifested in our study with mouse
-cells
because the intrinsic generation of glutamate from glucose might not reach the
threshold required for the potentiation. It is well known that the amplitude
of the second-phase insulin secretion to glucose is much higher in rat
-cells than in mouse
-cells. A recent report showed that the
expression of glutamate dehydrogenase was indeed lower in mouse islets than in
rat islets and supported our data because GME induced the secretory response
even at 16.7 mM glucose in the mouse islets
(27).
At present, a causal relationship between the vesicular alkalinization and glucose-stimulated insulin secretion or glutamate-induced potentiation of the insulin secretion is unclear. In fact, addition of GME did not further alkalinize the pH of secretory vesicles attained by glucose treatment (data not shown), indicating that the potentiating effect of GME on glucose-stimulated insulin secretion was, at least in part, not directly linked to the vesicular alkalinization. However, the following observations prompted us to consider that the alkalinization would not be merely an event that coincided with those increases in insulin secretion but would be, at least in part, causally related to them. Maechler et al. (29) reported that a direct application of glutamate to the vesicles evoked increases in insulin secretion and cytosolic [ATP] and that these increases were blocked by an inhibitor of the vesicular glutamate transporters. They argued that the vesicles themselves were the ATP source of the increase in cytosolic [ATP], probably through a reverse reaction of the vacuolar-type H+-ATPase. In fact, similar effects were observed with an inhibitor of vacuolar-type H+-ATPase (29). If this were the case, an outward flux of protons across vesicular membranes would reduce the proton gradient, and the luminal pH of the vesicles would tend to be alkalinized, which is what we observed in the present study. Generation of ATP in this manner from the vesicles was assumed to elevate the cytosolic [ATP] immediately near the exocytotic machinery composed of soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) complex and thus to be favorable to the priming and final fusion processes of the vesicles (11, 52). Another merit of the vesicular alkalinization during exocytosis might be prompting the crystals of Zn2+-insulin hexamers stored inside the vesicles to dissolve for faster diffusion into the bloodstream. Interestingly, Aspinwall et al. (6) demonstrated with amperometric detection of insulin molecules that inhibiting the vacuolar-type H+-ATPase and thereby increasing the vesicular pH caused insulin to be more rapidly extruded from the vesicles. Thus alkalinizing the vesicular pH could be a form of priming process of the vesicles for exocytosis, in which the insulin crystals were "primed" for the forthcoming dissolution, although the extent of the alkalinization by glutamate was small.
Lately, it has been found that glutamate is taken up into synaptic vesicles
by brain-type Na+-dependent inorganic phosphate transporter (BNPI)
(7,
53) to prepare for its later
release in excitatory neuronal cells. Another Na+-dependent
phosphate transporter, differentiation-associated Na+-dependent
inorganic phosphate transporter (DNPI), which has a close resemblance in amino
acid sequence to BNPI, was also cloned in neural tissues
(2). We screened for these two
transporters in a cDNA library made from insulin-producing INS-1 cells,
resulting in isolation of both the types of transporters in the -cells
(Yamashita T, Eto K, and Kadowaki T, unpublished observations). It was thus
assumed that
-cells also exhibited glutamate uptake activities via these
transporters. It is suggested that H+ may play a role as a counter
ion during the glutamate uptake by those transporters
(36), where the outward efflux
of H+ should contribute to alkalinization of the intravesicular pH,
although precise expression and function of BNPI and DNPI in
-cell
secretory vesicles remain to be addressed.
In summary, -cell secretory vesicle alkalinization occurred acutely
after a glucose challenge depending on glucose metabolism but independently of
cytosolic [Ca2+] elevation or the change of cytosolic
pH. Our results suggest the possibility that one of the mediators of this
glucose effect was glucose-derived glutamate, because the cell-permeable
glutamate analog GME reproduced such alkalinization at a basal concentration
of glucose. GME potentiated glucose-stimulated insulin secretion without
affecting the KATP channel-dependent pathway of the secretion. Thus
it is likely that glutamate exhibited such a potentiating effect on the
secretion by acting directly on secretory vesicles, which may be coupled to
changes of proton transport across the vesicle membrane and their acute
alkalinization. It was reported that insulin-secretory vesicles were
organelles actively importing and releasing Ca2+ via
certain types of Ca2+-ATPases and ryanodine receptors,
respectively, in response to glucose
(32). Further elucidation of
acute-phase regulation of the ionic milieu in the secretory vesicles will be
needed for a comprehensive understanding of the exocytotic mechanism of
glucose-stimulated insulin secretion in a normal state and in pathological
states such as insulin-deficient type 2 diabetes and PHHI caused by mutations
of glutamate dehydrogenase.
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DISCLOSURES |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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