Yale University School of Nursing, New Haven, Connecticut 06536-0740
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ABSTRACT |
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The capacity of 20 mM glucose to desensitize insulin release was determined. A prior
exposure to 20 mM glucose impaired the response of rat islets to
subsequent restimulation. Compared with control islets, insulin
secretory rates measured 25-30 min after the onset of 20 mM
glucose stimulation were reduced by 75%. Restimulation of
glucose-desensitized islets with 20 mM glucose plus 500 nM forskolin
resulted in a dramatic enhancement of both phases of secretion. In
contrast to the desensitization of rat islets induced by prior 20 mM
glucose exposure, mouse islets were immune to this adverse effect of
the hexose. Prior exposure to 20 mM glucose had no adverse effect on
glucose usage rates. The activation of phospholipase C in
glucose-desensitized rat islets was compromised when compared with
control islets. The impairment could not be accounted for by a decrease
in immunoreactive content of several major phospholipase C isozymes
(1 or
1) or their partitioning between the membrane and cytosolic
compartments. In contrast to rat islets, prior exposure of mouse islets
to 20 mM glucose for 180 min had no effect on inositol phosphate
accumulation. These observations document an additional difference
between rat and mouse islets and suggest that the evolution of
desensitization is a consequence of the impaired activation of
phospholipase C in rat islets.
phospholipase C; inositol phosphates; toxicity; islets
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INTRODUCTION |
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The insulin secretory response of human islets in vivo and perfused
rat pancreatic islets in situ to sustained hyperglycemic stimulation is
biphasic, characterized by a first phase of several minutes duration
and a slowly rising and sustained second phase (16-18, 20, 32,
41). This secretory pattern has been duplicated using perifused islets
studied in vitro (7, 22, 56). In addition to the evocation of biphasic
secretion, prior short-term exposure of human or rat islets to glucose
primes them to subsequent restimulation (11, 19, 45). Also termed
sensitization or time-dependent potentiation, this response can be
induced by several agonists, linked by their capacity to activate
phospholipase C (PLC) and/or protein kinase C (31, 38, 44, 54).
An adverse action of glucose on -cell performance has also been
documented. Termed glucose-induced desensitization, time-dependent
suppression, or third-phase release, chronic exposure of islets to
stimulatory glucose impairs their capacity to respond to subsequent
restimulation (6, 9, 26, 33, 37, 55). The devolution of islet sensitivity to glucose stimulation may play a particularly important role in the etiology of type II diabetes (14, 15).
In marked contrast to human or rat islets, mouse islet responses to glucose stimulation are notable for the lack of a rising second-phase insulin secretory response and for the failure of prior short-term glucose exposure to induce time-dependent potentiation as well (3, 4, 28, 29, 42, 51, 53). An analysis of information flow in the PLC signaling cascade has revealed that mouse islet responses to glucose are significantly less than the comparable responses of rat islets to the hexose (43, 50, 53) and that the expression of several PLC isozymes is reduced as well (53).
If the ability of glucose to stimulate a rising second-phase secretory response and to induce both time-dependent potentiation and time-dependent suppression of release in rat and human islets is linked by a common signaling cascade, then the failure of glucose to elicit the same phenomena in mouse islets may be a consequence of defective or reduced signaling via the same cascade. If this is a reasonably accurate portrayal of biochemical events in the islet, then mouse islets should be immune to the desensitizing effects of sustained stimulation with glucose. The present experiments were designed to address this issue and to explore possible biochemical mechanisms involved in the induction of desensitization in rat islets.
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MATERIALS AND METHODS |
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Islet isolation. The detailed methodologies employed to assess insulin output from collagenase-isolated islets have been previously described (53). Young adult male CD-1 mice (body weights at time of study 32-45 g) or male Sprague-Dawley rats (body weights at time of study 300-450 g) were purchased from Charles River. All animals were treated in a manner that complied with the National Research Council Guide for the Care and Use of Laboratory Animals. The animals were fed ad libitum. After pentobarbital sodium anesthesia (Nembutal, 50 mg/kg; Abbott, North Chicago, IL) was induced, islets were isolated by collagenase digestion and handpicked by use of a glass loop pipette under a stereo microscope. They were free of exocrine contamination.
Perifusion studies. Groups of 14-18 rat or mouse islets were loaded onto nylon filters (Tetko, Briarcliff Manor, NY) and incubated for 180 min in a Krebs-Ringer bicarbonate (KRB) buffer supplemented with 5 or 20 mM glucose. After washing with 5 ml of KRB, these islets were then perifused for 20 min with 3 mM glucose and for an additional 30 min with 20 mM glucose with or without 500 nM forskolin. The flow rate was maintained at 1 ml/min. Perifusate solutions were gassed with 95% O2-5% CO2 and maintained at 37°C. At the termination of the perifusion, islets still on the nylon filters were placed in small vials. Hanks' solution (0.25 ml) was added, the islets were sonicated, and insulin contents were measured. Insulin was measured by radioimmunoassay (1).
Islet labeling for inositol phosphate studies. After isolation, groups of 18-26 islets were loaded onto nylon filters, placed in a small glass vial, incubated for 180 min in a KRB solution containing myo-[2-3H]inositol, and prepared as follows. myo-[2-3H]inositol (10 µCi; specific activity 16-23 Ci/mmol) was placed in a 10 × 75-mm culture tube. To this aliquot of tracer, 250 µl of warmed (to 37°C) and oxygenated KRB medium supplemented with 5 mM or 20 mM glucose were added. After mixing, 240 µl of this solution were gently added to the vial with islets. The vial was capped with a rubber stopper, gassed for 10 s with 95% O2-5% CO2, and incubated at 37°C. After 90 min, the vials were again gently oxygenated. After the labeling period, the islets still on nylon filters were washed with 5 ml of fresh KRB.
Inositol phosphate measurements. After washing, the islets on nylon filters were placed in small glass vials. Added gently to the vial were 400 µl of KRB supplemented with 10 mM LiCl, to prevent inositol phosphate (IP) degradation, and the appropriate agonists as indicated. The vials were capped and gently gassed for 5 s with 95% O2-5% CO2. After 30 min, the generation of IPs was stopped by adding 400 µl of 20% perchloric acid. Total IPs formed were then measured using Dowex columns, as described previously (5, 48).
Glucose utilization rates. In experiments with rat islets, the usage of glucose was measured by determining the rate of 3H2O formation from [5-3H]glucose by use of methods previously described (46). These islets were first incubated for 180 min with 5 mM or 20 mM glucose and then were washed with 5 ml of fresh KRB. The islets were then incubated for 60 min in 0.125 ml of 20 mM glucose supplemented with tracer [5-3H]glucose. The 3H2O formed during the 60-min incubation period was separated from the unused [3H]glucose, as described previously (46).
PLC isozyme content and distribution.
Rat islets were loaded onto nylon filters and incubated for 180 min in
either 5 mM or 20 mM glucose. The islets were retrieved and gently
sonicated in 150 µl of buffer containing 20 mM Tris, pH 7.4, 0.5 mM
EGTA, 50 µM leupeptin, 1 mM phenylmethylsulfonyl fluoride, 0.1%
2-mercaptoethanol, 10 µM pepstatin A, and aprotinin at 25 µg/ml.
The samples were then centrifuged in a Beckman Airfuge (25 psi for 10 min; 1 psi = 6.89 kPa). The supernatant was collected as the cytosolic
fraction. The pellet was resuspended in 150 µl of sonication buffer
and collected as the membrane fraction. All samples were dried for
45-50 min in a Savant speedvac. Dried samples were resuspended in
18.75 µl water plus 6.25 µl of a 4× concentrated Laemmli
solution. For the Western blots, protein samples of cytosol and
membrane fractions were separated by SDS-PAGE with a 4% stacking gel
and a 7% running gel at 12 mA and 16 mA, respectively. Gel-resolved proteins were electrotransferred onto Immobilon polyvinylidene fluoride
membranes at 15 V for 20 h. The membranes were stained with Ponceau S
solution for protein, washed, and blocked for 1 h in TBS (0.5% Tween
20 and 5% milk powder). The membranes were incubated for 1 h with
PLC-1 antibody (0.5 µg/ml dilution) or for 2.5 h with PLC-
1
antibody (1.0 µg/ml dilution), washed, incubated for 45 min with
horseradish peroxidase (HRP)-conjugated secondary antibodies for
PLC-
1 and 1.5 h for PLC-
1, and washed again. The antigen-antibody
complexes were visualized using the enhanced chemiluminescence system
(Amersham, Arlington Heights, IL) and quantitated densitometrically by
use of Visage 2000. The results were expressed as the optical density
of the PLC band in the fraction divided by the total optical density,
where the total optical density is the sum of the PLC band in the
cytosol plus the PLC band in the membrane.
Reagents.
Hanks' solution was used for the islet isolation. The KRB incubation
and perifusion medium consisted of 115 mM NaCl, 5 mM KCl, 2.2 mM
CaCl2, 1 mM
MgCl2, 24 mM
NaHCO3, and 0.17 g/dl bovine serum
albumin (BSA). Other compounds were added where indicated, and the
solution was gassed with a mixture of 95%
O2-5%
CO2. The 125I-labeled insulin used for the
insulin assay and the
[5-3H]glucose and
3H2O
used for the metabolic analysis were purchased from New England Nuclear
(Boston, MA).
myo[2-3H]inositol
was purchased from Amersham. BSA (RIA grade), glucose, and the salts
used to make the Hanks' solution and perifusion medium were purchased
from Sigma (St. Louis, MO). Rat insulin standard (lot no. 615-ZS-157)
was the generous gift of Dr. Gerald Gold, Eli Lilly (Indianapolis, IN).
Collagenase (type P) was obtained from Boehringer Mannheim Biochemicals
(Indianapolis, IN). Forskolin was obtained from Calbiochem (LaJolla,
CA). Rabbit polyclonal PLC-1 and anti-mouse and anti-rabbit IgG-HRP
secondary antibodies were obtained from Santa Cruz Biotechnology (Santa
Cruz, CA.) Mouse monoclonal PLC-
1 antibody was purchased from
Upstate Biotechnology.
Statistics.
Statistical significance was determined using the Student's
t-test for unpaired data or analysis
of variance in conjunction with the Newman-Keuls test for unpaired
data. A P value 0.05 was taken as
significant. Values represent means ± SE of at least three observations.
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RESULTS |
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Studies with rat islets.
The insulin secretory responses of rat islets previously incubated for
180 min with 5 mM glucose and subsequently stimulated with 20 mM
glucose are given in Fig. 1. Before the
onset of stimulation with 20 mM glucose, release rates from these
islets averaged 39 ± 6 (n = 12)
pg · islet1 · min
1
in the presence of 3 mM glucose. Peak first-phase release averaged 190 ± 27 pg · islet
1 · min
1,
and 25-30 min after the onset of stimulation with 20 mM glucose, these rates increased to 563 ± 57 pg · islet
1 · min
1.
When measured at the end of the perifusion, insulin content of these
islets averaged 115 ± 9 ng/islet.
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Studies with mouse islets.
A different picture emerged if mouse islets were studied under the same
conditions as were employed with rat islets. After a 180-min incubation
with 5 mM glucose, mouse islets were perifused with 3 mM glucose for 20 min to establish stable secretion rates and then stimulated for 30 min
with 20 mM glucose (Fig. 3,
left). An initial spike of secretion
that peaked at 103 ± 11 pg · islet1 · min
1
(n = 10) was observed. With
continued exposure to the hexose, release rates remained stable and
flat, unlike the response of rat islets but in agreement with previous
studies using mouse islets (3, 29, 42, 53). Insulin release rates
25-30 min after the onset of 20 mM glucose stimulation averaged
only 77 ± 15 pg · islet
1 · min
1.
Insulin content measured after the perifusion averaged 93 ± 8 ng/islet.
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Metabolic studies.
We considered the possibility that the decline in secretory
responsiveness of rat islets exposed to 20 mM glucose may be a consequence of impaired hexose metabolism. To address this issue, the
usage of 20 mM glucose was determined in rat islets after a 180-min
incubation in either 5 mM or 20 mM glucose. Islets previously incubated
in 5 mM glucose utilized 20 mM glucose at rates of 168.9 ± 13 (n = 5)
pmol · islet1 · h
1.
Comparable glucose usage rates, 174.7 ± 6 (n = 5)
pmol · islet
1 · h
1,
were observed with islets incubated for 180 min with 20 mM glucose before the metabolic studies.
IP measurements. We have suggested previously (43, 50) that impaired activation of islet PLC in response to glucose stimulation was a characteristic of desensitized islets regardless of the compound used to induce impaired secretion. On the basis of this consideration, additional studies were conducted with control and glucose-desensitized islets to examine IP accumulation, a sensitive index of PLC activation (12, 30). The results are presented in Table 1. Control rat islets incubated for 180 min with 5 mM glucose displayed low levels of IPs in the presence of 3 mM glucose. When these islets were stimulated with 20 mM glucose, a greater than fivefold enhancement of IP accumulation was observed. Slightly higher basal IP levels were measured in rat islets previously incubated with 20 mM glucose for 180 min. Most striking, however, was the reduced response of these islets to subsequent restimulation with 20 mM glucose (compare Table 1, lines 2 and 4).
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PLC isozyme analysis.
The three major classes of PLC isozymes have been identified in rat
islets (53). PLC-1 isozyme activation is dependent on GTP-binding
proteins, whereas PLC-
1 isozyme activation is most sensitive to
calcium (2). PLC-
1 activation is dependent on growth factors (12,
34). We considered the possibility that the decline in IP accumulation
in glucose-desensitized islets may be a consequence of decrease in PLC
isozyme content in these islets. Because glucose-induced PLC activation
requires both calcium and a metabolic signal (24), perhaps GTP (39), we
measured rat islet contents of the
1 and
1 PLC isozymes and their
cellular distribution by use of quantitative densitometric measurements of Western blot bands. In control islets incubated for 180 min in 5 mM
glucose, the partitioning of PLC-
1 was evident with 57 ± 2.4% of
the enzyme in the cytosolic fraction and 43 ± 2.4% associated with
the membrane fraction (Fig. 5
top,
n = 3). PLC-
1 was more unevenly
distributed in control islets; only 22 ± 2.1% was associated with
the membrane compartment (Fig. 5,
bottom,
n = 3). After a 180-min
incubation in 20 mM glucose, there was no effect on the distribution
(membrane vs. cytosolic) of these two PLC isozymes (Fig. 5). When the
total optical densities of the membrane and cytosolic bands were added
together, there were no significant differences between control and
glucose-desensitized islets. Compared with control islets (taken as
100%), the total optical density of the PLC-
1 and PLC-
1 bands in
glucose-desensitized islets averaged 84 ± 18 and 89 ± 15%,
respectively.
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DISCUSSION |
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There was one major goal of the present series of experiments. We
wanted to establish whether the anomalous behavior of mouse islets,
compared with either rat or human islets, extended to the capacity of
the hexose to desensitize -cell insulin secretory responses. To
address this issue, it was necessary for us to repeat previous seminal
observations with regard to the capacity of sustained high glucose
exposure to impair insulin secretion from islets, an alteration that
appears to play an essential role in the pathogenesis of type II
diabetes (14).
Perhaps the first data clearly documenting the adverse effects of
sustained glucose exposure on -cell responses from rat islets were
reported by Curry (13), who used the perfused rat pancreas preparation.
Similar to the findings reported herein, insulin release rates peaked
after ~40 min of stimulation and then fell slowly as the perfusion
progressed. After 180 min of stimulation with 16.7 mM glucose, insulin
release rates had fallen ~50% compared with peak secretion rates
observed earlier in the perfusion. In the present series of
experiments, insulin secretion was more severely affected, a difference
most likely attributable to the stronger (20 mM) glucose stimulus used
in our studies.
A series of cogent reports by Bolaffi, Grodsky, and co-workers
(8-10, 21) followed shortly thereafter. The concept that emerged
from these and other studies (6, 23, 40, 47, 55) is that
desensitization of the -cell is not related to a depletion of
insulin content and that its induction is not confined to glucose. At
least in rat islets, it can be induced by a wide variety of
structurally diverse compounds. For example, in addition to glucose,
prior chronic exposure to
-ketoisocaproate, monomethylsuccinate, carbachol, N-acetylglucosamine,
interleukin-1, or forskolin desensitizes rat islets to subsequent
restimulation (8, 40, 47, 49, 52, 57). Regardless of the nature of the
compound used to induce it, however, a uniform observation is that
desensitization is characterized by the failure of PLC to be activated
appropriately on restimulation (43, 50).
The time-dependent actions of stimulatory glucose on rat and human
-cell responses have been well documented and, in rat islets, their
biochemical mechanisms have been explored. However, a series of studies
with mouse islets, initially reported almost 20 years ago (3, 4) and
confirmed in more recent experiments (29, 53), proved most informative
and provided possible clues to the mechanisms involved. It was observed
that in response to glucose stimulation, mouse islets, in sharp
contrast to rat or human islets, exhibit neither a rising second-phase
response to glucose stimulation nor the capability of being primed or
sensitized by short-term exposure to the hexose. Berglund (3,
4) suggested a possible relationship between these two
aberrant (compared with rat and human islets) responses from mouse
islets. No biochemical explanation was forthcoming in these studies.
The subsequent findings, that the activation of PLC by glucose is
minimal in mouse islets compared with rat islets, may provide a
reasonable biochemical explanation for this species divergence (53).
If there is linkage between the biochemical mechanisms responsible for the rising second-phase insulin secretory response of rat islets (and human islets as well) to glucose stimulation and the capacity of the hexose to induce time-dependent potentiation and time-dependent suppression of insulin release, then it might be predicted that mouse islets, which fail to demonstrate the former two phenomena, might be less susceptible to the toxic or adverse effect of sustained glucose exposure. This prediction was verified in these studies. Unlike rat and human islets, which exhibit all of these three temporal response patterns to glucose stimulation, sustained 180-min exposure of mouse islets to 20 mM glucose had no adverse effect on the response to subsequent restimulation with the hexose. Prolonged exposure of mouse islets to 20 mM glucose had no deleterious effect on total insulin content, suggesting that during the 180-min incubation period little insulin was being continuously secreted by islets from this species.
Additional studies were conducted to explore the biochemical mechanisms
responsible for the reduction in insulin secretion from
glucose-desensitized rat islets. The impaired secretory response to a
subsequent 20 mM glucose stimulus was not due to depletion of
releasable insulin pools. The addition of 500 nM forskolin to
glucose-desensitized islets was accompanied by an insulin secretory response virtually identical to that observed from control islets previously incubated for 180 min with 5 mM glucose and subsequently stimulated with 20 mM glucose alone. We did not observe any obvious derangement in glucose usage rates by glucose-desensitized rat islets,
a result which suggests that glucose transport into the -cell and
its phosphorylation by glucokinase are both normal. What we did observe
was that the subsequent activation of PLC by 20 mM glucose was severely
compromised in glucose-desensitized rat islets.
We measured the cellular distribution of PLC-1 or PLC-
1 in rat
islets. We focused on these two PLC isozymes because in other tissues
their activation is dependent on either GTP or calcium or both (12). In
agreement with other observations made concerning other cells (12), the
cellular distribution was not uniform in control islets. Most of the
immunoreactive PLC-
1 and -
1 isozymes were located in the soluble
cytosolic compartment. Under conditions in which the activation of PLC
is suppressed, we could not uncover any adverse effect of sustained 20 mM glucose exposure on either the content or cellular distribution of
PLC-
1 or PLC-
1. If either one of these PLC isozymes is activated
by glucose, then these findings suggest that the decline in PLC
activation, monitored by IP accumulation, is not a consequence of
reduced islet PLC content or maldistribution. A more subtle alteration
has to be invoked. Because the precise identity of the
glucose-activated PLC isozyme remains to be established, it is possible
that a yet-to-be-described isozyme is involved.
Unlike rat islets, mouse islets respond poorly to the activation of PLC by glucose (43, 53) (See also Table 1). The failure of high glucose to generate significant phosphoinositide-derived second messenger molecules may account not only for the failure of glucose to evoke a rising second-phase insulin secretory response from mouse islets but also for its failure to induce time-dependent potentiation and time-dependent suppression of insulin secretion. These and other findings with both rat and mouse islets suggest that all three of these temporal response patterns are linked phenomena, a result, perhaps, of the underexpression in mouse islets of the nutrient-activated PLC isozyme. The observation that carbachol, which activates an isozyme of PLC different from the one activated by nutrient stimulants (25), evokes a markedly enhanced sustained insulin secretory response, time-dependent potentiation, and time-dependent suppression of insulin release is consistent with this concept (43, 50, 51, 53).
In an attempt to provide both a unifying concept to understand these observations and a framework for future investigation, we propose that glucose-induced desensitization in rat islets is a consequence of altered activation of islet PLC. This family of isozymes, three of which have been identified in islets (53), is inhibited by both protein kinase (PK) C- and PKA-mediated phosphorylation events (34). Agonists that induce desensitization may do so by increasing the phosphorylation state of PLC. We suggest further that mouse islets are immune to desensitization for several reasons. First and foremost, the nutrient-regulated isozyme of PLC is underexpressed in mouse islets. When acutely stimulated with 20 mM glucose, failure to generate PI-derived signals to the same quantitative extent as rat islets contributes to the failure of glucose to cause a rising second-phase response. The flat but sustained modest second-phase response of mouse islets to 20 mM glucose is largely independent of PLC activation. When exposed to 20 mM glucose for prolonged periods, as in the present studies, the failure of glucose to increase PLC-mediated PI hydrolysis is accompanied by the reduced generation of signals with the potential to induce desensitization. Although this concept remains to be established, it is consistent with what is known about the regulation of this important enzyme in other tissues (34-36) and with the rapid reversal of glucose toxicity in rat islets (26, 27). This issue is also amenable to future investigation. Finally, if this is a reasonably accurate portrayal of biochemical events in a glucose-desensitized islet, then the manipulation of the enzymes such as phosphatases, which regulate dephosphorylation events, might be appropriate pharmacological targets.
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ACKNOWLEDGEMENTS |
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These studies were supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant 41230.
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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. §1734 solely to indicate this fact.
Address for reprint requests: W. S. Zawalich, Yale Univ. School of Nursing, 100 Church St. South, New Haven, CT 06536-0740.
Received 6 July 1998; accepted in final form 6 August 1998.
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