help button home button Endocrine Society Endocrinology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Endocrinology, doi:10.1210/en.2003-1099
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
145/2/667    most recent
Author Manuscript (PDF)
Right arrow Purchase Article
Right arrow View Shopping Cart
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Merglen, A.
Right arrow Articles by Maechler, P.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Merglen, A.
Right arrow Articles by Maechler, P.
Endocrinology Vol. 145, No. 2 667-678
Copyright © 2004 by The Endocrine Society

Glucose Sensitivity and Metabolism-Secretion Coupling Studied during Two-Year Continuous Culture in INS-1E Insulinoma Cells

Arnaud Merglen, Sten Theander, Blanca Rubi, Gaelle Chaffard, Claes B. Wollheim and Pierre Maechler

Division of Clinical Biochemistry, Department of Internal Medicine, University Medical Center, 1211 Geneva 4, Switzerland

Address all correspondence and requests for reprints to: Pierre Maechler, Ph.D., DBC-9100, University Medical Center, 1 rue Michel-Servet, CH-1211 Geneva 4, Switzerland. E-mail: pierre.maechler{at}medecine.unige.ch.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Rat insulinoma-derived INS-1 cells constitute a widely used ß-cell surrogate. However, due to their nonclonal nature, INS-1 cells are heterogeneous and are not stable over extended culture periods. We have isolated clonal INS-1E cells from parental INS-1 based on both their insulin content and their secretory responses to glucose. Here we describe the stable differentiated INS-1E ß-cell phenotype over 116 passages (no. 27–142) representing a 2.2-yr continuous follow-up. INS-1E cells can be safely cultured and used within passages 40–100 with average insulin contents of 2.30 ± 0.11 µg/million cells. Glucose-induced insulin secretion was dose-related and similar to rat islet responses. Secretion saturated with a 6.2-fold increase at 15 mM glucose, showing a 50% effective concentration of 10.4 mM. Secretory responses to amino acids and sulfonylurea were similar to those of islets. Moreover, INS-1E cells retained the amplifying pathway, as judged by glucose-evoked augmentation of insulin release in a depolarized state. Regarding metabolic parameters, INS-1E cells exhibited glucose dose-dependent elevations of NAD(P)H, cytosolic Ca2+, and mitochondrial Ca2+ levels. In contrast, mitochondrial membrane potential, ATP levels, and cell membrane potential were all fully activated by 7.5 mM glucose. Using the perforated patch clamp technique, 7.5 and 15 mM glucose elicited electrical activity to a similar degree. A KATP current was identified in whole cell voltage clamp using diazoxide and tolbutamide. As in native ß-cells, tolbutamide induced electrical activity, indicating that the KATPconductance is important in setting the resting potential. Therefore, INS-1E cells represent a stable and valuable ß-cell model.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
RELIABLE ß-CELL MODELS are of paramount importance for diabetes research. It is generally accepted that the use of primary cells is preferable. However, this requires large quantities of isolated pancreatic islets, which is work-intensive and has the inherent inconvenience of representing a mixed population of ß-, {alpha}-, {delta}-, and F cells. Consequently, rodent ß-cell lines have proven their usefulness, and their continuous development is still essential until clonal human ß-cells become available. The intrinsic challenge when establishing a ß-cell line is the maintenance of tissue-specific differentiation combined with adequate cell proliferation. As a result, only a limited number of ß-cell lines are available to date, all of rodent origin.

Among the mouse derived ß-cell lines (1, 2), MIN6 cells (1, 3) represent a valuable model, which was further improved by isolation of the clonal subline MIN6m9 (4). Of rat origin, RINm5F cells do not respond to glucose in the physiological concentration range (5, 6), and BRIN-BD11 cells are poorly differentiated, exhibiting low insulin content and weak secretory responses to glucose (7).

In the early nineties, our laboratory established the insulin-secreting cell line INS-1 isolated from a radiation-induced rat insulinoma (8). Due to the nonclonal nature of these cells, limited stability over passages probably explains some of the discrepancies observed among the numerous laboratories using parental INS-1 cells worldwide. To circumvent this problem, Hohmeier and colleagues (9) stably transfected INS-1 cells with the human proinsulin gene, followed by selection of clones based on robust glucose-stimulated insulin secretion. The resulting INS-1-832/13 cells are highly glucose responsive. However, these cells overexpress human insulin driven by the ubiquitous cytomegalovirus promoter in addition to the endogenous rat insulin, rendering it impossible to judge the differentiated state based on insulin content. An alternative approach would be the cloning of well differentiated INS-1 cells without genetic manipulation. Hence, we isolated clonal INS-1E cells from the parental cells based on both their insulin content and their secretory response to glucose (10). This new INS-1E ß-cell line has been provided to many investigators, although thorough characterization of these cells has not been documented to date.

The aim of the present study was to provide an extensive and long-term characterization of INS-1E cells for a series of parameters recognized to play a key role in metabolism-secretion coupling. In the consensus model of glucose-stimulated insulin secretion, glucose equilibrates across the plasma membrane and is phosphorylated by glucokinase, initiating glycolysis (11). Subsequently, mitochondrial metabolism generates ATP, which promotes the closure of ATP-sensitive K+ channels (KATP channel) and, as a consequence, depolarization of the plasma membrane (12). This leads to Ca2+ influx through voltage-gated Ca2+ channels and a rise in cytosolic Ca2+ ([Ca2+]c), triggering insulin exocytosis (12, 13). Additional signals are necessary to reproduce the sustained secretion elicited by glucose. They participate in the amplifying pathway (14), formerly referred to as the KATP channel-independent stimulation of insulin secretion.

Here we describe the stable differentiated INS-1E ß-cell phenotype over more than 100 passages corresponding to a 2-yr continuous follow-up. INS-1E cells exhibited secretory responses in the physiological glucose concentration range as well as expressed the amplifying pathway. We measured several metabolic parameters and electrophysiological correlates. Finally, spheroid clusters, sometimes referred to as pseudo-islets (15), composed of reaggregated INS-1E cells were prepared and tested for their ability to respond to secretagogues.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell culture
Maintenance.
The clonal ß-cell line INS-1E, derived from parental INS-1 cells (8), was selected for its insulin content and adequate proliferation (10). INS-1E cells were cultured in a humidified atmosphere containing 5% CO2 in complete medium composed of RPMI 1640 supplemented with 5% heat-inactivated fetal calf serum, 1 mM sodium pyruvate, 50 µM 2-mercaptoethanol, 2 mM glutamine, 10 mM HEPES, 100 U/ml penicillin, and 100 µg/ml streptomycin. The maintenance culture was passaged once a week by gentle trypsinization, and cells were seeded at a density of 4 x 104 cells/cm2, i.e. 3 x 106 cells, in 75-cm2 Falcon bottles (BD Biosciences Labware, Franklin Lakes, NJ) with 20 ml complete medium. The potential presence of mycoplasma was regularly checked using a photometric enzyme immunoassay for the detection of PCR-amplified mycoplasma DNA (Roche, Penzberg, Germany). For most experiments, unless otherwise mentioned, INS-1E were seeded at 2 x 105 cells/1 ml in Falcon 24-well plates and used 4–5 d thereafter, with one medium change on d 3 or 4.

INS-1E cell clusters.
INS-1E cells were seeded in nonadherent bacterial 10-cm petri dishes and cultured in complete medium for 5–6 d before use.

Insulin secretion
Attached cells.
The secretory responses to glucose and other secretagogues were tested in INS-1E cells between passages 54–95. Before the experiments, cells were maintained for 2 h in glucose-free culture medium. The cells were then washed twice and preincubated for 30 min at 37 C in glucose-free Krebs-Ringer bicarbonate HEPES buffer (KRBH) of the following composition: 135 mM NaCl, 3.6 mM KCl, 5 mM NaHCO3, 0.5 mM NaH2PO4, 0.5 mM MgCl2, 1.5 mM CaCl2, and 10 mM HEPES, pH 7.4. BSA (0.1%) was added as an insulin carrier. Next, cells were washed once with glucose-free KRBH and then incubated for 30 min in KRBH and stimuli as indicated. Incubation was stopped by putting the plates on ice, the supernatants were collected for insulin secretion, and cellular insulin contents were determined from acid-ethanol extracts. Insulin secretion was measured by RIA using rat insulin as standard (16).

Perifusion of INS-1E cell clusters.
Spheroid clusters composed of INS-1E cells were preincubated for 2 h in glucose-free culture medium. After low speed centrifugation (500 rpm), cell clusters were resuspended in glucose-free KRBH and counted, and about 500 spheroids were distributed per chamber in a 250-µl volume thermostated at 37 C (Brandel, Gaithersburg, MD). The flow rate was set at 0.5 ml/min, and fractions were collected every minute after a 20-min washing period at basal 2.5 mM glucose. At the end of the perifusion, cell clusters were collected, and insulin contents were determined from acid-ethanol extracts.

Fluorescence and luminescence measurements
Several metabolic parameters were measured using either fluorescence or bioluminescence assays. In such experiments, unless otherwise mentioned, INS-1E cells cultured for 4–5 d in 24-well plates were preincubated for 2 h in glucose-free culture medium, washed, and further preincubated in glucose-free KRBH for 30 min. The 24-well plates were then transferred to a thermostated (37 C) plate reader set, according to the probe, in either fluorescence or luminescence mode (Fluostar Optima, BMG Labtechnologies, Offenburg, Germany). Individual wells of the 24-well plate were recorded sequentially over a 40-min period, and test compounds were added at the indicated times through automated built-in microinjectors.

NAD(P)H.
Autofluorescence of NAD(P)H was monitored with excitation and emission filters set at 320 and 460 nm, respectively. After stabilization of the signal for 10 min in 2.5 mM glucose KRBH, increasing glucose concentrations were administered as indicated. NAD(P)H autofluorescence was normalized over a 10-min stimulation period by setting the fluorescence at 100% for cells maintained in basal 2.5 mM glucose. Controls of maximal fluorescence changes were performed after the addition of 5 µM rotenone plus 2 µM antimycin, added as inhibitors of the electron transport chain.

Mitochondrial membrane potential.
Mitochondrial membrane potential ({Delta}{psi}m) was measured using the fluorescent probe, rhodamine 123. Cells cultured in 24-well plates were maintained for 2 h in 2.5 mM glucose medium at 37 C before loading with 10 µg/ml rhodamine 123 (Molecular Probes, Eugene, OR) for 20 min at 37 C in KRBH (17). The {Delta}{psi}m was monitored with excitation and emission filters set at 485 and 520 nm, respectively. Glucose (additions on top of basal 2.5 mM) and then the protonophore carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) were added to each well.

Cytosolic ATP levels.
Cytosolic ATP levels were monitored in cells expressing the ATP-sensitive bioluminescent probe luciferase 1 d after transduction with the specific AdRIP-Luc viral construct (18, 19). After the preincubation steps described above, the 24-well plates were transferred to the plate reader in the luminometer mode. The luciferase substrate, 100 µM beetle luciferin (Promega Corp., Madison, WI), was added to the KRBH. After a 10-min period in basal 2.5 mM glucose, cells were stimulated with the indicated glucose concentrations, and 20 min later the mitochondrial poison NaN3 (2 mM) was added.

Cell membrane potential.
Cell membrane potential was monitored using 100 nM of the fluorescent probe bis-oxonol [bis-(1,3-diethylthiobarbituric acid) trimethine oxonol; Molecular Probes]. Filters used for excitation and emission had wavelength optima at 544 and 590 nm, respectively.

Calcium levels.
Calcium levels were monitored in cells transduced the day before the experiment with the calcium-sensitive photoprotein aequorin. Measurements of mitochondrial and cytosolic calcium (20, 21) were performed using the corresponding adenovirus constructs, AdCA-mtAeq and AdCA-cyAeq, respectively. The 2-h preincubation period also served in this case to load cells with the aequorin prosthetic group, i.e. 5 µM coelenterazine [2-(p-hydroxybenzyl)-6-(p-hydroxyphenyl)-8-benzylimidazo[1,2-{alpha}] pyrazin-3-(7H)-one; Calbiochem, San Diego, CA]. Calibration was calculated based on the total counts obtained at the end of the trace following permeabilization of the cells with 50 µM digitonin and 10 mM CaCl2 exposure (22, 23).

Electrophysiology
Cells were seeded on glass coverslips and maintained in culture for 4–5 d. For patch-clamp recordings a glass coverslip was transferred to a temperature-controlled perfusion chamber. The bath was perfused, using a gravity-driven perfusion system, with extracellular solution at a rate of 1.3 ml/min.

For the study of electrical activity and voltage-dependent current, we used the perforated patch whole cell recording mode, whereas KATP current was analyzed in the standard whole cell configuration. Patch-clamp recordings were performed with an EPC 9 amplifier (HEKA, Darmstadt, Germany) and Pulse software (version 8.53, HEKA). Patch pipettes were pulled from borosilicate glass capillaries (GC150F-10, Clark Instruments, Reading, UK) on a model P-97 puller from Sutter Instruments (Novato, CA). Pipette resistance was between 4–6 MOhm. Zero-voltage currents were cancelled electronically before seal formation. For perforated patch recordings, 0.24 mg/ml of the pore-forming agent amphotericin B (24) was added to the pipette solution (Sigma-Aldrich Corp., Buchs, Switzerland).

The action of amphotericin was seen as a continuous decrease in serial resistance. Recordings began when the serial resistance had attained values below 25 MOhm. In experiments using the perforated patch configuration, the bath temperature was maintained at 33 C to facilitate action potential firing. Whole cell recordings were made at room temperature.

Solutions for electrophysiology
The standard extracellular medium consisted of 140 mM NaCl, 3.6 mM KCl, 2 mM NaHCO3, 0.5 mM NaH2PO4, 0.5 mM MgSO4, 5 mM HEPES (pH 7,4 with NaOH), and 1.5 mM CaCl2. In some experiments, 20 mM tetraethyl ammonium (TEA) was used. In this case extracellular NaCl was lowered to 120 mM to keep the osmolarity constant. The pipette solution for perforated patch recordings was 76 mM K2SO4, 10 mM NaCl, 10 mM KCl, 1 mM MgCl2, and 5 mM HEPES (pH 7.35 with KOH). For the recording of whole cell KATP current, the pipette solution consisted of 125 mM KCl, 30 mM KOH, 10 mM EGTA, 1 mM MgCl2, 5 mM HEPES, 0.3 mM MgATP, and 0.3 mM K-ADP (pH 7.15 with KOH). Glucose was present at a concentration of 2.5 mM in all experiments unless stated otherwise. Diazoxide, tolbutamide, and 4-aminopuridine were dissolved in dimethylsulfoxide and added to the extracellular bath solution at the indicated concentrations. Tetrodotoxin (Latoxan, Valence, France) was dissolved in a 100-mM sodium citrate buffer.

Statistical analysis
Values are given as the mean ± SEM, and n values refer to the number of independent experiments. In the case of representative experiments, values are the mean ± SD. The significance of differences was assessed by t test for unpaired data.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Stability of INS-1E cells over passages
Cultured in plastic dishes, INS-1E cells attached firmly and spread well (Fig. 1AGo). We analyzed INS-1E cells in terms of insulin contents and secretory responses over 116 passages (no. 27–142), corresponding to 2.2-yr continuous culture, assuming 1 passage/wk. Passages below 27 correspond to the cloning period of parental INS-1 cells as well as the subsequent amplification phase of the isolated clones with very small number of cells available at each passage. Stocks of frozen cells in liquid nitrogen were subsequently prepared from passages 24–27, and INS-1E cells were extensively analyzed starting from this period. Insulin contents averaged 2.15 ± 0.11 µg insulin/106 cells (Fig. 1BGo). More than 1 value for the same passage corresponds to the collection of data from noncontemporary cultures. Measurements of insulin contents over 116 passages exhibited a bell-shaped distribution, with the highest content in the upper part of the curve reaching 3.46 ± 0.63 µg insulin/106 cells at passage 64.



View larger version (54K):
[in this window]
[in a new window]
  		FIG. 1. Long-term characterization of INS-1E cells. A, Phase contrast light microscopy of INS-1E cells (passage 99) in culture 4 d after seeding (magnification, x200). B, Insulin content of INS-1E cells over 116 passages). The dotted line shows the average of insulin contents (2.15 ± 0.11 µg insulin/106 cells) calculated from all passages. Values are the mean ± SD for individual experiments. More than one value for the same passage corresponds to the collection of data from noncontemporary cultures.

 
Insulin secretion
We tested the secretory responses of INS-1E cells (passages 54–95) to various concentrations of glucose within the physiological range over a 30-min period (Fig. 2AGo). Compared with basal insulin release at 2.5 mM glucose, exposure to glucose concentrations up to 20 mM resulted in a dose-dependent elevation of insulin secretion with a plateau phase at about 15–20 mM glucose. The averaged response to 15 mM glucose was 6.2-fold (P < 0.001 vs. 2.5 mM glucose), and the calculated 50% effective concentration was 10.4 mM glucose (r2 = 0.989). Secretory responses to 15 mM glucose averaged 4.2 ± 1.2-fold over passages 40–50 (n = 5 independent experiments) and 4.3 ± 1.1-fold between passages 90–100 (n = 7 independent experiments). As a comparison, 15 mM glucose elicited a 6.6-fold secretory response at passage 64, which also exhibited the highest insulin content. Insulin secretion was also stimulated by nonnutrient secretagogues eliciting cell membrane depolarization (Fig. 2BGo). Accordingly, 250 µM tolbutamide and 30 mM KCl resulted in 4.8-fold (P < 0.001) and 4.7-fold (P < 0.001) secretory responses, respectively. This is lower than glucose-induced secretion. We next investigated whether INS-1E cells retained the amplifying pathway of secretion, classically observed upon glucose stimulation in islets treated with depolarizing concentrations of KCl plus the KATP channel opener diazoxide (25). At basal 2.5 mM glucose, 30 mM KCl combined with 250 µM diazoxide increased insulin release 5.3-fold (P < 0.05). Under these experimental conditions, raising glucose to 20 mM further markedly augmented the secretory response by 86% (P < 0.01), demonstrating the presence of the amplifying pathway. We also tested two key amino acids on insulin secretion. Glutamine alone is not a secretagogue unless glutamate dehydrogenase is activated by L-leucine or its nonmetabolizable analog 2-aminobicyclo-[2,2,1]heptane-2-carboxylic acid (BCH), as previously demonstrated in rat islets (26, 27). In INS-1E cells, such a pattern of secretory responses was observed, as shown in Fig. 2BGo. Finally, 1 µM epinephrine caused a 56% (P < 0.05) inhibition of glucose-stimulated insulin release (28).



View larger version (21K):
[in this window]
[in a new window]
  		FIG. 2. Insulin secretion in response to glucose, amino acids, and nonnutrient secretagogues in INS-1E cells. A, The glucose dose response was tested over a 30-min stimulation period. B, Cells were treated with 250 µM tolbutamide (TBM), 30 mM KCl (KCl), KCl plus 250 µM diazoxide (KCl+DZ), KCl+DZ in the presence of 20 mM glucose (G20+KCl+DZ), 1 µM epinephrine at 15 mM glucose (G15+Epi), 5 mM glutamine (Gln), 20 mM L-leucine (Leu), 10 mM BCH, and their combinations as indicated. Values are the mean ± SEM of individual experiments performed at passages 54, 55, 60, 61, 63, 64, 82, 94, and 95. *, P < 0.001 vs. G2.5; §, P < 0.001 vs. G20; ¶, P < 0.001 vs. G15; #, P < 0.01 vs. Leu; &, P < 0.001 vs. BCH.

 
NAD(P)H
Glucose generates reduced forms of NAD(P)H through both glycolysis and mitochondrial metabolism. Electrons from reduced NADH in the cytosol are transferred into the mitochondria through the glycerol phosphate and the malate-aspartate shuttles. The autofluorescence of NAD(P)H in INS-1E cells was monitored to assess a crucial step in glucose metabolism (Fig. 3AGo). Raising glucose from 2.5 to 7.5 and 15.5 mM caused dose-dependent increases in NAD(P)H by 19% (P < 0.001) and 41% (P < 0.001) respectively.



View larger version (21K):
[in this window]
[in a new window]
  		FIG. 3. Effects of glucose on NAD(P)H autofluorescence and mitochondrial membrane potential in INS-1E cells (passage 91). A, Dose-dependent effect of glucose on NAD(P)H autofluorescence. The NAD(P)H autofluorescence of cells stimulated with different glucose (Glc) concentrations is compared with the 100% for basal 2.5 mM glucose over a 10-min period. B, Mitochondrial membrane potential was monitored as rhodamine 123 fluorescence. INS-1E cells were stimulated with glucose for 10 min before depolarization was induced by 1 µM FCCP. Values are the mean ± SD for representative experiments. *, P < 0.001 vs. 2.5 mM glucose.

 
Mitochondrial membrane potential ({Delta}{psi}m)
Mitochondrial metabolism leads to the activation of the electron transport chain with resulting hyperpolarization of the mitochondrial membrane, the prerequisite for ATP generation. The {Delta}{psi}m measured in cells loaded with rhodamine 123 showed that glucose elicited hyperpolarization, with a plateau phase reached after 1–2 min (Fig. 3BGo). Maximal effects were obtained at 7.5 mM glucose, and high glucose did not elicit stronger hyperpolarization. As expected, the protonophore FCCP (1 µM) rapidly depolarized {Delta}{psi}m.

ATP generation
Cytosolic ATP levels were monitored as luminescence in INS-1E cells expressing luciferase. Elevation of glucose from 2.5 mM to stimulatory concentrations increased ATP levels in a biphasic manner (Fig. 4AGo). The first peak was observed after about 2 min, followed by sustained elevation 4 min after stimulation. In accordance with {Delta}{psi}m measurements, maximal ATP levels were reached at 7.5 mM glucose. Sodium azide (2 mM) added subsequently disrupted ATP generation with a concomitant drop in ATP levels.



View larger version (28K):
[in this window]
[in a new window]
  		FIG. 4. Effects of glucose on ATP generation and [Ca2+]m levels in INS-1E cells. A, Cytosolic ATP levels were monitored in cells expressing luciferase. Cells (passage 83) were stimulated with the indicated glucose (Glc) concentrations for 10 min before abrogation of ATP generation by the addition of 2 mM NaN3. B, Mitochondrial Ca2+ concentrations were monitored in cells (passage 84) expressing aequorin targeted to the mitochondria. Values are the mean ± SD for representative experiments (n = 6).

 
Mitochondrial calcium levels ([Ca2+]m)
To further assess mitochondrial function, measurements of [Ca2+]m were performed in INS-1E cells expressing the photoprotein aequorin targeted to the mitochondria. As shown in Fig. 4BGo, basal [Ca2+]m was about 100 nM. Glucose stimulation resulted in biphasic elevations of [Ca2+]m, with the first peak lasting 2 min, and a second phase 3 min after glucose addition (Fig. 4BGo). The initial peaks of [Ca2+]m induced by glucose elevation to 7.5, 15.5, and 20.5 mM reached 807 ± 80, 1069 ± 242, and 1156 ± 141 nM, respectively. In contrast to the first phase, amplitudes of the second phases (500–600 nM) were similar at all tested stimulatory glucose concentrations.

Cell membrane potential
The cell membrane potential was measured as bis-oxonol fluorescence. Increasing glucose from 2.5 to 7.5, 15.5, or 20.5 mM resulted in sustained cell depolarization, further augmented by the subsequent addition of 30 mM KCl (Fig. 5AGo). Interestingly, 20.5 mM glucose induced the first significant transient hyperpolarization of the cell membrane, lasting about 1 min before the establishment of the expected depolarized state.



View larger version (34K):
[in this window]
[in a new window]
  		FIG. 5. Effects of glucose on cell membrane potential and cytosolic Ca2+ levels in INS-1E cells (passage 101). A, Cell membrane potential monitored as bis-oxonol fluorescence. Cells were stimulated with the indicated glucose (Glc) concentrations for 10 min before control depolarization induced by 30 mM KCl. B, Cytosolic Ca2+ concentrations were monitored in cells expressing aequorin. Values are the mean ± SD for representative experiments (n = 6).

 
[Ca2+]c
Basal [Ca2+]c levels, measured as calibrated aequorin luminescence, were in the 100-nM range (Fig. 5BGo). Stimulation with 7.5, 15.5, and 20.5 mM glucose induced dose-dependent, first phase [Ca2+]c responses of 234 ± 43, 396 ± 83, and 449 ± 66 nM, respectively. The second phase of [Ca2+]c, appearing 2–3 min after the glucose rise, was uniformly in the range of 250 nM.

Electrophysiology
Using the perforated patch configuration, we studied glucose-induced electrical activity in INS-1E cells. Of 18 tested cells, 16 responded to glucose by eliciting trains of action potentials. There was no obvious difference in the intensity of action potential firing or in plateau potential between cells stimulated with 7.5 or 15 mM glucose (Fig. 6AGo). A few (n = 3) cells were spontaneously active also at low glucose values. This activity, however, subsided after a few minutes. We attribute this spontaneous activity to the high (11 mM) glucose in the culture medium.



View larger version (23K):
[in this window]
[in a new window]
  		FIG. 6. INS-1E cells respond to glucose by eliciting action potentials. A, Recording of membrane voltage using the perforated patch-clamp configuration. At the right, a part (at the arrow) of the trace is shown at a higher time resolution. B, Effect of 100 µM tolbutamide on membrane potential. C, Whole cell recording of KATP current. Current responses from a cell to brief (500-ms) 10 mV hyper- and depolarizing voltage steps and superfused with tolbutamide (100 µM), diazoxide (200 µM), or both as indicated. D, Voltage-dependent currents, as revealed by 50-ms depolarizing voltage steps from -70 to -40, -20, 0, and +20 mV in a cell in control solution. E, Voltage response of a cell to a voltage step to 0 mV, from a holding voltage of -70, in control solution, in the presence of 0.5 mM 4-AP, and in the presence of both 0.5 mM 4-AP and 20 mM TEA. F, Response to a step to 0 mV from a holding voltage of -70 in the presence of 0.5 mM 4-AP and 20 mM TEA (upper trace) and in the additional presence of 0.5 µM tetrodotoxin (lower trace). D and E, Recordings from the same cell (passage 111; F, recording from a different cell (passage 87). The experiments shown in B–F are representative of at least three experiments (passages 87–111).

 
When comparing the electrical activity of INS-1E cells with that of native ß-cells, it is clear that the voltage span of the action potentials is larger in INS-1E cells than in native ß-cells, and the peak of the action potentials was frequently overshooting zero by 10–20 mV. This was always the case at the beginning of stimulation with glucose (or tolbutamide, see below), whereas the amplitude declined thereafter. The extent and time course of this amplitude attenuation, however, were quite variable. The sample trace in Fig. 6AGo is an example of rapid attenuation, whereas traces very similar to that shown in Fig. 6CGo, displaying less attenuation, were also observed using glucose as a stimulus. We did not observe any true periodicity in the electrical activity characteristic of primary ß-cells, studied in isolated cells (29) or whole islets (30). The electrical activity was mostly continuous. Only occasionally could brief periods of bursting be observed, i.e. episodes of electrical activity from a distinct plateau voltage interrupted by silent phases at a voltage lower than the plateau (Fig. 6BGo).

To investigate whether INS-1E cells, like primary ß-cells as well as other ß-cell clones, contain ATP-dependent K+ current and diazoxide- and tolbutamide-sensitive KATP current, we studied cells in the whole cell configuration. Cells were stimulated with a short 10-mV depolarizing and hyperpolarizing pulse from a holding potential of -70 mV. As shown in Fig. 6DGo, diazoxide caused a large increase in membrane conductance, which was antagonized by tolbutamide. These experiments indicate that INS-1E cells possess a tolbutamide- and diazoxide-sensitive current. This current, furthermore, seems to control the resting potential, because tolbutamide, applied at low glucose concentrations, elicited vigorous electrical activity.

We also investigated cells with respect to voltage-dependent currents using the perforated patch-clamp configuration. A representative example of current responses to depolarizing voltage steps of 50-msec duration is shown in Fig. 6CGo. These revealed the existence of inward as well as outward voltage-dependent currents. A rapidly activating and inactivating inward current was seen at the onset of the depolarizing pulse. Within a few milliseconds the inward current was outstripped by an outward current. The outward current appeared to be composed of both transient and sustained components. To further identify these components, we first applied 0.5 mM 4-aminopyridine (4-AP), a classical blocker of the so-called A current, a transient voltage-dependent K+ current. As shown in Fig. 6FGo, 4Go-AP substantially blocked the first transient phase of outward current, whereas it had only minor effects on the sustained current. This maneuver revealed a current with a slower activation pattern. This current was readily blocked by 20 mM TEA, confirming its identity as a delayed rectifier current. In the combined presence of 4-AP and TEA, only inward current components remained (see Fig. 6GGo for better visualization). The rapidly activating and inactivating component was fully blocked by 0.5 mM tetrodotoxin, thus establishing that this current is a voltage-dependent Na+ current. Finally, the current remaining after blockage of voltage-dependent K+ and Na+ currents was very likely a Ca2+ current.

Spheroid clusters made of INS-1E cells
INS-1E cells were cultured in nonadherent petri dishes, promoting the formation of cell aggregates hereafter referred to as INS-1E spheroids (see Fig. 7AGo). INS-1E spheroids were distributed into perifusion chambers, and effluent was collected every minute for insulin release measurements. At the end of the perifusion period, INS-1E spheroids were recovered from the chambers for the determination of insulin content. According to the number of INS-1E spheroids distributed per chamber and the INS-1E insulin content, the estimated average number of cells per spheroid was 4000.



View larger version (23K):
[in this window]
[in a new window]
  		FIG. 7. Secretory responses of spheroid clusters composed of INS-1E cells. A, Phase contrast light microscopy of INS-1E cells (passage 64) forming spheroids in culture (magnification, x200). B, Clusters of INS-1E cells (passage 66) were perifused at 0.5 ml/min with basal 2.5 mM glucose (Glc) before stimulation with 15 mM glucose, resulting in sustained insulin secretion. C, Addition of 30 mM KCl at 2.5 mM glucose resulted in transient insulin release. Values are the mean ± SD for representative experiments (n = 3).

 
Basal insulin release at 2.5 mM glucose was less than 0.2% the insulin content of INS-1E spheroids. Raising glucose from 2.5 to 15 mM resulted in a 6.6-fold stimulation of insulin secretion at the first phase peak, followed by the establishment of a second sustained phase (Fig. 7BGo). KCl-mediated cell membrane depolarization resulted in rapid, but transient, secretory responses lacking the second sustained phase (Fig. 7CGo). Dispersed INS-1E cells obtained by trypsinization and loaded into the same perifusion chambers responded very poorly to secretagogues (not shown).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The INS-1E clone isolated from parental INS-1 cells was analyzed continuously over an extended period of more than 2 yr, corresponding to 116 passages. Cells exhibited remarkable stability over time and passages. INS-1E at passages earlier than 40 and later than 100 contained less insulin, although the secretory responses to glucose were maintained. This unique collection of data indicates that INS-1E cells can be safely cultured and used within passages 40–100, corresponding to approximately a 1-yr period. The average insulin contents of cells between passages 40–100 was 2.30 ± 0.11 µg/million cells, corresponding to approximately 10% of the content of native rat ß-cells (31).

Glucose-stimulated insulin secretion in isolated rat islets is fairly variable. For instance, secretory responses to 16.7 mM glucose reported from the same experienced laboratory range from 4.2-fold (32) to 12.3-fold (33). Accordingly, an adequate rat ß-cell line should ideally respond within that range. On the average for passages 54–95, 15 mM glucose stimulated insulin release 6.2-fold in INS-1E cells, and the calculated 50% effective concentration for glucose effect was 10.4 mM, in agreement with the value for rat islets (11). At the extremes of the recommended passage range, i.e. 40 and 100, 15 mM glucose elicited secretory responses of 4.2- and 4.3-fold, respectively. It is of interest that both insulin content and glucose-evoked insulin release displayed bell-shaped characteristics as a function of passage number.

The cells exhibited typical features observed in primary ß-cells regarding stimulation of insulin release by various secretagogues. INS-1E cells responded to sulfonylurea to a lesser extent than to high glucose. Such a difference might be explained by the contribution of additive factors participating in the stimulation of insulin secretion upon glucose stimulation, an effect also referred to as the amplifying pathway (14). Combination of KCl with diazoxide at basal vs. high glucose demonstrated the presence of the amplifying pathway in INS-1E cells. Another characteristic of ß-cells is a weak secretory response to glutamine unless glutamate dehydrogenase is activated by L-leucine or its nonmetabolizable analog BCH (26, 27). INS-1E cells retained such characteristics, rendering these cells a valuable model for the study of such complex metabolic pathways (34).

Cell to cell communication is required for proper ß-cell function, and transgenic mice with altered ß-cell coupling exhibit reduced glucose-evoked insulin secretion (35). In vitro, a preparation of so-called pseudo-islets composed of MIN6 cells improved the functional responsiveness of this mouse-derived ß-cell line (15). In the present study, strong dynamic secretory responses were obtained in perifusion from INS-1E cells aggregated as spheroid clusters, or pseudo-islets, but not from dispersed floating cells. This indicates that INS-1E cells retained cell to cell communication properties.

The glucose-induced NAD(P)H increase in ß-cells is remarkable and permits autofluorescence-activated cell sorting of pancreatic islet cells, i.e. purification of ß-cells according to glucose-induced changes in cellular redox state (36). It has previously been reported that 20 mM glucose stimulation caused a 50–80% increase in NAD(P)H autofluorescence measured in single rat ß-cells (37) and a 200–300% elevation in adult rat islets (38, 39, 40). NAD(P)H increases of 8% at 12 mM glucose (41) and 27% at 22.2 mM glucose (42) were reported in mouse islets. Similarly, in INS-1E cells, glucose increased NAD(P)H dose-dependently, with a 41% elevation at 15 mM glucose.

ATP generation depends on activation of the electron transport chain revealed by hyperpolarization of the mitochondrial membrane. Interestingly, glucose stimulation ranging from 7.5–20.5 mM exhibited similar effects on both parameters. Indeed, we observed hyperpolarization of the mitochondrial membrane as well as ATP generation, with maximal effects at 7.5 mM glucose. The thus-formed ATP promotes the closure of KATP channels, leading to cell depolarization. In accordance with ATP levels, there was a lack of glucose dose-response on cell membrane potential. These data indicate that the electron transport is fully activated and saturated at glucose concentrations lower than those eliciting optimal secretory responses, in accordance with previous observations (43).

As opposed to ATP changes, increases in [Ca2+]c and [Ca2+]m were glucose dose dependent, at least during the first phase response. This indicates that the [Ca2+]c elevation mediated by opening of voltage-gated Ca2+ channels, as a consequence of KATPchannel closure, does not account for the full effect of glucose on cellular Ca2+ modulation. It is of interest to note that voltage-independent Ca2+ influx has been observed in pancreatic ß-cells (44). Moreover, Ca2+-induced Ca2+ release might contribute to the augmentation of cytosolic Ca2+ signals in ß-cells (45, 46), although this specific pathway remains to be investigated in INS-1E cells. Elevations of Ca2+ levels in the mitochondrial compartment depend on both a [Ca2+]c rise and hyperpolarization of the mitochondrial membrane, which drives Ca2+ into the mitochondria through the uniporter (17, 47, 48). As shown in this study, kinetic comparisons of [Ca2+]c, [Ca2+]m, and {Delta}{psi}m suggest that during the first 1–2 min of glucose stimulation, [Ca2+]m changes were driven by [Ca2+]c and, as a result, were glucose dose dependent. Meanwhile, {Delta}{psi}m was fully activated by glucose in a dose-independent manner, leading to similar second phases between glucose stimulations. It should be noted that the calibrated values of [Ca2+]c and [Ca2+]m measured here in a newly developed multiwell plate mode are in good agreement with previously reported values (20).

Cell membrane potential was depolarized upon glucose stimulation as a result of the closure of KATP channels. However, prominent glucose levels (20.5 mM) induced transient hyperpolarization for about 1 min. This paradoxical effect might be explained by high activity of glucokinase consuming ATP upon glucose entry into the cell. This would cause a transient drop in ATP levels in the vicinity of the plasma membrane before sustained generation of ATP from the mitochondria leading to the expected depolarization.

It is clear that INS-1E cells retain many of the electrophysiological properties of native ß-cells. Importantly, they are stimulated by glucose. Furthermore, a tolbutamide- and diazoxide-sensitive current is present. Tests in current clamp mode also indicated that this current may set the resting potential, because its blockage with tolbutamide promptly resulted in electrical activity. It seems therefore likely that, similar to native ß-cells, stimulus-response coupling in INS-1E cells is largely controlled by an ATP-sensitive K+ conductance, KATP. INS-1E cells do, however, also display notable differences in comparison with native ß-cells. Most striking is the large voltage span of the action potential, which in native ß-cells does not overshoot 0 mV. It is likely that this phenomenon involves activation of the Na+ current in INS-1E cells in the physiological voltage range. This is also the case in, for example, RINm5f cells (49), but appears not to be the case at least in mouse ß-cells in which a Na+ current is present but fully inactivated at the resting voltage, thereby being inconsequential for the electrical activity (30).

Although INS-1E cells do not consistently display the typical bursting pattern of electrical activity, burst-like firing was occasionally observed. Ulrich et al. (50) also found bursting in a small number (15%) of INS-1 cells. Thus, similar to single isolated primary ß-cells (29) and in contrast to hamster insulinoma tumor cells (29), INS-1 cells are capable of displaying bursting. We conclude that INS-1E cells represent a valuable tool for the study of electrical transduction mechanisms in insulin-secreting cells.

In the present study the newly developed clonal INS-1E insulinoma cells have been characterized over a long-term continuous culture period. This unique collection of data should help the many researchers who already obtained INS-1E cells for various investigations. Although these cells do not substitute for primary ß-cells, INS-1E cells represent a significant improvement in terms of ß-cell differentiation and stability.


   Acknowledgments
 
We are indebted to H. Ishihara for sharing some of the adenovirus constructs used in the present report.


   Footnotes
 
This work was supported by the Swiss National Science Foundation (Grants 31-67023.01 to P.M. and 32-66907.01 to C.B.W.) and grants (to P.M.) from the Dr. Max Cloetta Foundation, the European Foundation for the Study of Diabetes/Johnson & Johnson, and the Leenaards Foundation. This study was part of the Geneva Program for Metabolic Disorders.

A.M. is the recipient of a research program fellowship (Faculty of Medicine, Geneva) in the laboratory of P.M.

Abbreviations: 4-AP, 4-Aminopyridine; BCH, 2-aminobicyclo-[2,2,1]heptane-2-carboxylic acid; [Ca2+]c, cytosolic Ca2+; [Ca2+]m, mitochondrial calcium concentration; {Delta}{psi}m, mitochondrial membrane potential; FCCP, carbonyl cyanide p-trifluoromethoxyphenylhydrazone; KRBH, Krebs-Ringer bicarbonate HEPES buffer; TEA, tetraethyl ammonium.

Received August 22, 2003.

Accepted for publication October 21, 2003.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Miyazaki J, Araki K, Yamato E, Ikegami H, Asano T, Shibasaki Y, Oka Y, Yamamura K 1990 Establishment of a pancreatic ß cell line that retains glucose-inducible insulin secretion: special reference to expression of glucose transporter isoforms. Endocrinology 127:126–132[Abstract]
  2. Knaack D, Fiore DM, Surana M, Leiser M, Laurance M, Fusco-DeMane D, Hegre OD, Fleischer N, Efrat S 1994 Clonal insulinoma cell line that stably maintains correct glucose responsiveness. Diabetes 43:1413–1417[Abstract]
  3. Ishihara H, Asano T, Tsukuda K, Katagiri H, Inukai K, Anai M, Kikuchi M, Yazaki Y, Miyazaki JI, Oka Y 1993 Pancreatic ß cell line MIN6 exhibits characteristics of glucose metabolism and glucose-stimulated insulin secretion similar to those of normal islets. Diabetologia 36:1139–1145[CrossRef][Medline]
  4. Minami K, Yano H, Miki T, Nagashima K, Wang CZ, Tanaka H, Miyazaki JI, Seino S 2000 Insulin secretion and differential gene expression in glucose-responsive and -unresponsive MIN6 sublines. Am J Physiol 279:E773–E781
  5. Praz GA, Halban PA, Wollheim CB, Blondel B, Strauss AJ, Renold AE 1983 Regulation of immunoreactive-insulin release from a rat cell line (RINm5F). Biochem J 210:345–352[Medline]
  6. McClenaghan NH, Elsner M, Tiedge M, Lenzen S 1998 Molecular characterization of the glucose-sensing mechanism in the clonal insulin-secreting BRIN-BD11 cell line. Biochem Biophys Res Commun 242:262–266[CrossRef][Medline]
  7. McClenaghan NH, Flatt PR 1999 Engineering cultured insulin-secreting pancreatic B-cell lines. J Mol Med 77:235–243[CrossRef][Medline]
  8. Asfari M, Janjic D, Meda P, Li G, Halban PA, Wollheim CB 1992 Establishment of 2-mercaptoethanol-dependent differentiated insulin-secreting cell lines. Endocrinology 130:167–178[Abstract]
  9. Hohmeier HE, Mulder H, Chen G, Henkel-Rieger R, Prentki M, Newgard CB 2000 Isolation of INS-1-derived cell lines with robust ATP-sensitive K+ channel-dependent and -independent glucose-stimulated insulin secretion. Diabetes 49:424–430[Abstract]
  10. Janjic D, Maechler P, Sekine N, Bartley C, Annen AS, Wolheim CB 1999 Free radical modulation of insulin release in INS-1 cells exposed to alloxan. Biochem Pharmacol 57:639–648[CrossRef][Medline]
  11. Matschinsky FM 1996 Banting lecture 1995. A lesson in metabolic regulation inspired by the glucokinase glucose sensor paradigm. Diabetes 45:223–241[Abstract]
  12. Rorsman P 1997 The pancreatic ß-cell as a fuel sensor: an electrophysiologist’s viewpoint. Diabetologia 40:487–495[CrossRef][Medline]
  13. Lang J 1999 Molecular mechanisms and regulation of insulin exocytosis as a paradigm of endocrine secretion. Eur J Biochem 259:3–17[Medline]
  14. Henquin JC 2000 Triggering and amplifying pathways of regulation of insulin secretion by glucose. Diabetes 49:1751–1760[Abstract]
  15. Hauge-Evans AC, Squires PE, Persaud SJ, Jones PM 1999 Pancreatic ß-cell-to-ß-cell interactions are required for integrated responses to nutrient stimuli: enhanced Ca2+ and insulin secretory responses of MIN6 pseudoislets. Diabetes 48:1402–1408[Abstract]
  16. Maechler P, Wollheim CB 1999 Mitochondrial glutamate acts as a messenger in glucose-induced insulin exocytosis. Nature 402:685–689[CrossRef][Medline]
  17. Maechler P, Kennedy ED, Pozzan T, Wollheim CB 1997 Mitochondrial activation directly triggers the exocytosis of insulin in permeabilized pancreatic ß-cells. EMBO J 16:3833–3841[CrossRef][Medline]
  18. Maechler P, Wang H, Wollheim CB 1998 Continuous monitoring of ATP levels in living insulin secreting cells expressing cytosolic firefly luciferase. FEBS Lett 422:328–332[CrossRef][Medline]
  19. Rubi B, Ishihara H, Hegardt FG, Wollheim CB, Maechler P 2001 GAD65-mediated glutamate decarboxylation reduces glucose-stimulated insulin secretion in pancreatic ß cells. J Biol Chem 276:36391–36396[Abstract/Free Full Text]
  20. Kennedy ED, Rizzuto R, Theler JM, Pralong WF, Bastianutto C, Pozzan T, Wollheim CB 1996 Glucose-stimulated insulin secretion correlates with changes in mitochondrial and cytosolic Ca2+ in aequorin-expressing INS-1 cells. J Clin Invest 98:2524–2538[Medline]
  21. Ishihara H, Maechler P, Gjinovci A, Herrera PL, Wollheim CB 2003 Islet ß-cell secretion determines glucagon release from neighbouring {alpha}-cells. Nat Cell Biol 5:330–335[CrossRef][Medline]
  22. Rizzuto R, Simpson AW, Brini M, Pozzan T 1992 Rapid changes of mitochondrial Ca2+ revealed by specifically targeted recombinant aequorin. Nature 358:325–327[CrossRef][Medline]
  23. Brini M, Marsault R, Bastianutto C, Alvarez J, Pozzan T, Rizzuto R 1995 Transfected aequorin in the measurement of cytosolic Ca2+ concentration ([Ca2+]c). A critical evaluation. J Biol Chem 270:9896–9903[Abstract/Free Full Text]
  24. Rae J, Cooper K, Gates P, Watsky M 1991 Low access resistance perforated patch recordings using amphotericin B. J Neurosci Methods 37:15–26[CrossRef][Medline]
  25. Gembal M, Gilon P, Henquin JC 1992 Evidence that glucose can control insulin release independently from its action on ATP-sensitive K+ channels in mouse B cells. J Clin Invest 89:1288–1295
  26. Malaisse WJ, Sener A, Malaisse-Lagae F, Welsh M, Matthews DE, Bier DM, Hellerstrom C 1982 The stimulus-secretion coupling of amino acid-induced insulin release. Metabolic response of pancreatic islets of L-glutamine and L-leucine. J Biol Chem 257:8731–8737[Abstract/Free Full Text]
  27. Malaisse-Lagae F, Sener A, Garcia-Morales P, Valverde I, Malaisse WJ 1982 The stimulus-secretion coupling of amino acid-induced insulin release. Influence of a nonmetabolized analog of leucine on the metabolism of glutamine in pancreatic islets. J Biol Chem 257:3754–3758[Abstract/Free Full Text]
  28. Wollheim CB, Lang J, Regazzi R 1996 The exocytotic process of insulin secretion and its regulation by Ca2+ and G-proteins. Diabetes Rev 4:276–297
  29. Kinard TA, de Vries G, Sherman A, Satin LS 1999 Modulation of the bursting properties of single mouse pancreatic ß-cells by artificial conductances. Biophys J 76:1423–1435[Abstract/Free Full Text]
  30. Gopel S, Kanno T, Barg S, Galvanovskis J, Rorsman P 1999 Voltage-gated and resting membrane currents recorded from B-cells in intact mouse pancreatic islets. J Physiol 521:717–728[Abstract/Free Full Text]
  31. Pipeleers DG, in’t Veld PA, Van de Winkel M, Maes E, Schuit FC, Gepts W 1985 A new in vitro model for the study of pancreatic A and B cells. Endocrinology 117:806–816[Abstract]
  32. Sener A, Conget I, Rasschaert J, Leclercq-Meyer V, Villanueva-Penacarrillo ML, Valverde I, Malaisse WJ 1994 Insulinotropic action of glutamic acid dimethyl ester. Am J Physiol 267:E573–E584
  33. Sener A, Malaisse WJ 2002 The stimulus-secretion coupling of amino acid-induced insulin release. Insulinotropic action of L-alanine. Biochim Biophys Acta 1573:100–104[Medline]
  34. Maechler P 2002 Mitochondria as the conductor of metabolic signals for insulin exocytosis in pancreatic ß-cells. Cell Mol Life Sci 59:1803–1818[CrossRef][Medline]
  35. Charollais A, Gjinovci A, Huarte J, Bauquis J, Nadal A, Martin F, Andreu E, Sanchez-Andres JV, Calabrese A, Bosco D, Soria B, Wollheim CB, Herrera PL, Meda P 2000 Junctional communication of pancreatic ß cells contributes to the control of insulin secretion and glucose tolerance. J Clin Invest 106:235–243[Medline]
  36. Van De Winkel M, Pipeleers D 1983 Autofluorescence-activated cell sorting of pancreatic islet cells: purification of insulin-containing B-cells according to glucose-induced changes in cellular redox state. Biochem Biophys Res Commun 114:835–842[CrossRef][Medline]
  37. Pralong WF, Bartley C, Wollheim CB 1990 Single islet ß-cell stimulation by nutrients: relationship between pyridine nucleotides, cytosolic Ca2+ and secretion. EMBO J 9:53–60[Medline]
  38. Bennett BD, Jetton TL, Ying G, Magnuson MA, Piston DW 1996 Quantitative subcellular imaging of glucose metabolism within intact pancreatic islets. J Biol Chem 271:3647–3651[Abstract/Free Full Text]
  39. Patterson GH, Knobel SM, Arkhammar P, Thastrup O, Piston DW 2000 Separation of the glucose-stimulated cytoplasmic and mitochondrial NAD(P)H responses in pancreatic islet ß cells. Proc Natl Acad Sci USA 97:5203–5207[Abstract/Free Full Text]
  40. Tan C, Tuch BE, Tu J, Brown SA 2002 Role of NADH shuttles in glucose-induced insulin secretion from fetal ß-cells. Diabetes 51:2989–2996[Abstract/Free Full Text]
  41. Zhou YP, Sreenan S, Pan CY, Currie KP, Bindokas VP, Horikawa Y, Lee JP, Ostrega D, Ahmed N, Baldwin AC, Cox NJ, Fox AP, Miller RJ, Bell GI, Polonsky KS 2003 A 48-hour exposure of pancreatic islets to calpain inhibitors impairs mitochondrial fuel metabolism and the exocytosis of insulin. Metabolism 52:528–534[CrossRef][Medline]
  42. Eto K, Suga S, Wakui M, Tsubamoto Y, Terauchi Y, Taka J, Aizawa S, Noda M, Kimura S, Kasai H, Kadowaki T 1999 NADH shuttle system regulates K(ATP) channel-dependent pathway and steps distal to cytosolic Ca2+ concentration elevation in glucose-induced insulin secretion. J Biol Chem 274:25386–25392[Abstract/Free Full Text]
  43. Antinozzi PA, Ishihara H, Newgard CB, Wollheim CB 2002 Mitochondrial metabolism sets the maximal limit of fuel-stimulated insulin secretion in a model pancreatic ß cell: a survey of four fuel secretagogues. J Biol Chem 277:11746–11755[Abstract/Free Full Text]
  44. Leech CA, Holz GGt, Habener JF 1994 Voltage-independent calcium channels mediate slow oscillations of cytosolic calcium that are glucose dependent in pancreatic ß-cells. Endocrinology 135:365–372[Abstract]
  45. Kang G, Holz GG 2003 Amplification of exocytosis by Ca2+-induced Ca2+ release in INS-1 pancreatic ß cells. J Physiol 546:175–189[Abstract/Free Full Text]
  46. Graves TK, Hinkle PM 2003 Ca2+-induced Ca2+ release in the pancreatic ß-cell: direct evidence of endoplasmic reticulum Ca2+ release. Endocrinology 144:3565–3574[Abstract/Free Full Text]
  47. Gunter TE, Gunter KK, Sheu SS, Gavin CE 1994 Mitochondrial calcium transport: physiological and pathological relevance. Am J Physiol 267:C313–C339
  48. Duchen MR 1999 Contributions of mitochondria to animal physiology: from homeostatic sensor to calcium signalling and cell death. J Physiol 516:1–17[Abstract/Free Full Text]
  49. Rorsman P, Arkhammar P, Berggren PO 1986 Voltage-activated Na+ currents and their suppression by phorbol ester in clonal insulin-producing RINm5F cells. Am J Physiol 251:C912–C919
  50. Ullrich S, Abel KB, Lehr S, Greger R 1996 Effects of glucose, forskolin and tolbutamide on membrane potential and insulin secretion in the insulin-secreting cell line INS-1. Pflugers Arch 432:63063–63066



This article has been cited by other articles:


Home page
 EndocrinologyHome page
N. Hou, S. Torii, N. Saito, M. Hosaka, and T. Takeuchi
Reactive Oxygen Species-Mediated Pancreatic {beta}-Cell Death Is Regulated by Interactions between Stress-Activated Protein Kinases, p38 and c-Jun N-Terminal Kinase, and Mitogen-Activated Protein Kinase Phosphatases
Endocrinology, April 1, 2008; 149(4): 1654 - 1665.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
F. Allagnat, F. Alonso, D. Martin, A. Abderrahmani, G. Waeber, and J.-A. Haefliger
ICER-1{gamma} Overexpression Drives Palmitate-mediated Connexin36 Down-regulation in Insulin-secreting Cells
J. Biol. Chem., February 29, 2008; 283(9): 5226 - 5234.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
A. V. Zhdanov, M. W. Ward, J. H. M. Prehn, and D. B. Papkovsky
Dynamics of Intracellular Oxygen in PC12 Cells upon Stimulation of Neurotransmission
J. Biol. Chem., February 29, 2008; 283(9): 5650 - 5661.
[Abstract] [Full Text] [PDF]

Home page
 EndocrinologyHome page
A. Dov, E. Abramovitch, N. Warwar, and R. Nesher
Diminished Phosphodiesterase-8B Potentiates Biphasic Insulin Response to Glucose
Endocrinology, February 1, 2008; 149(2): 741 - 748.
[Abstract] [Full Text] [PDF]

Home page
 FASEB J.Home page
B. R. Gauthier, D. L. Duhamel, M. Iezzi, S. Theander, F. Saltel, M. Fukuda, B. Wehrle-Haller, and C. B. Wollheim
Synaptotagmin VII splice variants {alpha}, , and {delta} are expressed in pancreatic -cells and regulate insulin exocytosis
FASEB J, January 1, 2008; 22(1): 194 - 206.
[Abstract] [Full Text] [PDF]

Home page
 Mol. Cell. ProteomicsHome page
W. D'Hertog, L. Overbergh, K. Lage, G. B. Ferreira, M. Maris, C. Gysemans, D. Flamez, A. K. Cardozo, G. Van den Bergh, L. Schoofs, et al.
Proteomics Analysis of Cytokine-induced Dysfunction and Death in Insulin-producing INS-1E Cells: New Insights into the Pathways Involved
Mol. Cell. Proteomics, December 1, 2007; 6(12): 2180 - 2199.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
N. Ahuja, B. Schwer, S. Carobbio, D. Waltregny, B. J. North, V. Castronovo, P. Maechler, and E. Verdin
Regulation of Insulin Secretion by SIRT4, a Mitochondrial ADP-ribosyltransferase
J. Biol. Chem., November 16, 2007; 282(46): 33583 - 33592.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
N. Baroukh, M. A. Ravier, M. K. Loder, E. V. Hill, A. Bounacer, R. Scharfmann, G. A. Rutter, and E. Van Obberghen
MicroRNA-124a Regulates Foxa2 Expression and Intracellular Signaling in Pancreatic beta-Cell Lines
J. Biol. Chem., July 6, 2007; 282(27): 19575 - 19588.
[Abstract] [Full Text] [PDF]

Home page
 DiabetesHome page
J. Diaz-Delfin, M. Morales, and C. Caelles
Hypoglycemic Action of Thiazolidinediones/Peroxisome Proliferator-Activated Receptor {gamma} by Inhibition of the c-Jun NH2-Terminal Kinase Pathway
Diabetes, July 1, 2007; 56(7): 1865 - 1871.
[Abstract] [Full Text] [PDF]