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Department of Integrative Medical Biology, Section for Histology and Cell Biology, Umeå University, Umeå S-901 87, Sweden

(Requests for offprints should be addressed to N Gustavsson; Email: natalia.gustavsson{at}histocel.umu.se)

	Abstract

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We recently reported that the timing and magnitude of the nutrient-induced Ca²⁺ response are specific and reproducible for each isolated ß-cell. We have now used tolbutamide and arginine to test if the cell specificity exists also for the response to non-nutrient stimulation of ß-cells and if so, whether it is disturbed in ß-cells from hyperglycemic ob/ob and db/db mice. Zn²⁺ outflow measurements were used to study the correlation between Ca²⁺ response and insulin secretion in individual ß-cells. Tolbutamide and arginine induced cell-specific Ca²⁺ responses in lean mouse ß-cells both with regard to lag times for [Ca²⁺]_i rise and peak [Ca²⁺]_i heights. ß-Cells within intact islets also showed cell-specific timing of their Ca²⁺ responses to tolbutamide. However, in tolbutamide- and arginine-stimulated single ß-cells from ob/ob and db/db mice only the magnitude of Ca²⁺ response was cell-specific, not the timing. The lag time of tolbutamide-induced insulin secretion was cell-specific in lean mouse ß-cells but not in ob/ob mouse cells. Therefore, cell specificity seems to be a robust mechanism, and probably important for an adequate ß-cell function. The loss of temporal cell specificity for the response to tolbutamide in single ß-cells from hyperglycemic mice may be a sign of K_ATP- or voltage-dependent calcium channel dysfunction.

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Despite a large number of studies on the pathogenesis of type-2 diabetes, the precise mechanism of ß-cell dysfunction remains unclear. It is known that insulin and Ca²⁺response is delayed and reduced in patients and animal models with type-2 diabetes and the coordination of function of ß-cells is disturbed (Cerasi et al. 1972, Porte 1991). ß-Cells form clusters in pancreatic islets, where they are functionally linked by gap junctions (Salomon & Meda 1986) and other types of cell communication (Hellman et al. 2004), which results in a spread of signal within an islet. Isolated ß-cells and small cell clusters show a high degree of functional heterogeneity (Pralong et al. 1990, Herchuelz et al. 1991, Pipeleers et al. 1994). We recently reported that nutrient-induced Ca²⁺and NADH response patterns are also cell-specific, i.e., reproducible with regard to timing and magnitude of response (Larsson-Nyrén et al. 2002, Pakhtusova et al. 2003, Gustavsson et al. 2005). However, the cell specificity of mitochondrial metabolism seems to be blunted in diabetes and obesity (Gustavsson et al. 2005). We have now used tolbutamide and arginine to test if cell-specific responses can be observed also with non-nutrient stimuli, which affect voltage-dependent plasma membrane ion channels and if so, whether they are disturbed in ob/ob and db/db mouse ß-cells. Tolbutamide blocks ß-cell K_ATP channels by a direct interaction with sulfonylurea (SUR1) receptors (Trube et al. 1986, Ashcroft & Ashcroft 1992) and arginine directly causes ß-cell depolarization (Charles et al. 1982, Herchuelz et al. 1984). In diabetic subjects, the ß-cell response to sulfonylureas is less impaired than the response to glucose (Del Guerra et al. 2005). db/db mice develop severe diabetes (Berglund et al. 1978) and their ß-cells show a defective Ca²⁺ response to glucose stimulation (Roe et al. 1994). ob/ob mice are hyperglycemic, but their ß-cells respond adequately in vitro to stimulators and inhibitors of insulin release (Hellman et al. 1974, Idahl et al. 1976). However, a disorganization of the response within islets has been reported in the ob/ob mouse (Ravier et al. 2002). We used recordings of Zn²⁺ outflow as a measure of insulin secretion (Qian et al. 2003) to study the correlation between Ca²⁺ and Zn²⁺ response patterns in individual ß-cells during the first and second stimulation.

	Materials and Methods

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Animals

Adult female lean and Umeå ob/ob mice were from a local colony and female db/db mice (BKS.Cg-m+/+Lepr^db) were from Taconic Europe, Ry, Denmark. db/db mice were brought to our animal care facility at the age of 6 weeks and were housed until 3–4 months old, when the diabetic condition becomes manifest, but not prolonged. Principles of laboratory animal care (National Institute of Health publication no. 83-25, revised 1985) were followed and the animal care and experimental procedures were carried out in accordance with the standards established by the European Communities Council Directive (86/609/EEC9). The experiments were also approved by the Northern Swedish Committee for Ethics in Animal Experiments.

In overnight-fasted db/db mice, the blood sugar level was 22±2 mM (mean±S.D., n=33). This is in line with earlier observations (Berglund et al. 1978, Gustavsson et al. 2005). The urinary glucose level was 17±2 mM (n=33). The blood sugar in overnight-fasted ob/ob mice was 7±0.4 mM (n=16), with negative urinary glucose. The blood sugar level of lean mice was 5±0.4 mM (n=15).

ß-Cell preparation

Single ß-cells were prepared by shaking collagenase-isolated islets in Ca²⁺-deficient medium supplemented with EGTA and DNAase (Roche, Mannheim, Germany) (Lernmark 1974). The cells were distributed on polylysine-coated cover glasses (Sigma Chemicals, Stockholm, Sweden) placed in Petri dishes and maintained in tissue culture for 24–48 h at 11.1 mM glucose in medium (RPMI 1640; Roche) supplemented with 10% heat-inactivated fetal calf serum (Gibco, Stockholm, Sweden), 60 mg/ml garamycin (Schering Co., Kenilworth, NJ, USA), 60 mg/ml benzylpenicillin (Schering Co.), and 2 mM L-glutamine (Labkemi AB, Stockholm, Sweden). Subsequent experimental handling was performed with a Krebs–Ringer medium (KRH, Gibco) having the following composition in mM: 130 NaCl, 4.7 KCl, 1.2 KH₂PO₄, 1.2 MgSO₄, and 2.56 CaCl₂ and supplemented with 1 mg/ml BSA (Sigma Chemicals) and 3 mM D-glucose (Larsson-Nyrén & Sehlin 1996). The medium was buffered with 20 mM HEPES (Roche) and NaOH to pH of 7.4 and equilibrated with ambient air.

Measurements of cytoplasmic Ca²⁺ with Fura-2

Isolated cells were loaded with 1 µM Fura-2 (Molecular Probes Inc., Eugene, Oregon, USA) for 40 min in KRH medium at 37 °C. After rinsing in KRH to remove extracellular Fura-2, the cover glasses were mounted as the bottom of an open chamber (Larsson-Nyrén & Sehlin 1996). Experiments were performed using an image analysis system Openlab (Improvision, Coventry, West Midlands, UK) on an inverted microscope (Zeiss Axiovert, Gottingen, Germany) equipped for epifluorescence measurements. The cells were continuously perifused at a flow rate of 0.6 ml/min. The diameter of the perifusion chamber was 5 mm and volume of fluid in the chamber during perifusion was ~80 µl, which means that the whole volume was replaced within 8 s. The inflow tube was mounted close to the bottom at a distance of 1 mm from the center of the chamber.

Fura-2 was successively excited at the wavelengths 340 and 380 nm during 23 ms each using a 75 W Xenon lamp and a Polychrome IV monochromator (TILL Photonics, Murtinsried, Germany). The interval between cycles of 340 and 380 nm excitation was 1.75 s. Images were acquired by a camera (Orca ER, Hamamatsu, Japan). Temperature control (37 °C) was obtained by heating the chamber holder and the objective separately. The Ca²⁺ response was recorded from single ß-cells without direct contact with other cells. ß-Cells were initially preperifused at 3 mM glucose for 15 min. They were then stimulated twice with 100, 20 µM tolbutamide (Hoechst, Frankfurt, Germany), or 10 mM arginine during 10 min with a resting period of 30 min at 3 mM glucose between stimulations. All test media contained 3 mM glucose except in studies with 20 µM tolbutamide, which was applied in KRH medium containing 5 mM glucose to increase the number of responding cells (Jonkers et al. 2001). The duration of stimulation and the resting period were chosen to minimize time-dependent effects after the first stimulation (Nesher et al. 1989), but with sufficient time to recognize individual profiles also from cells that respond late (Larsson-Nyrén et al. 2002, Pakhtusova et al. 2003). Before the second stimulation, the exposed cell was checked by visual inspection. [Ca²⁺]_i was calculated from the ratio of 340 and 380 nm signals after background subtraction using the equation described by Grynkiewicz et al.(1985), with a ^Kd of 224 nM.

Lag-time for [Ca²⁺]_i rise during the first and second stimulation (LTr1 and LTr2) was defined as the time from the addition of stimulus to the first [Ca²⁺]_i value above baseline average, calculated during the 3 min preceding the stimulation. Peak height during the first and second stimulation (PH1 and PH2) was calculated as the difference between the baseline and the highest [Ca²⁺]_i value during the first peak. Superimposed spikes on top of the peaks were not included in calculations of peak height.

Insulin secretion measurements

ß-Cells were prepared as described above. Zn²⁺ images with FluoZin-3 (Molecular Probes Inc.) wereacquiredwith 488 nm excitation and 510 nm emission using the Openlab image analysis system on the Zeiss Axiovert microscope (see Measurements of cytoplasmic Ca²⁺ in ß-cells with Fura–2). Buffer with a reduced background Zn²⁺ level was made as described by Qian et al.(2003). Briefly, KRH was prepared with all ingredients except calcium and magnesium salts. The buffer was then treated with 5 g/dl Chelex-100 (Bio-Rad) for 2 h. The pH was adjusted to 7.4 after Chelex treatment and puratronic grade CaCl₂ and MgSO₄ (Alfa Aesar, Karlsruhe, Germany) were added to the final concentrations. Glass containers were avoided to minimize metal contamination. The Zn²⁺ chelator 200 nM tetrakis-(2-pyridylmethyl) ethylenediamine (TPEN; Molecular Probes Inc.) was added. Glass slides for cell preparation were soaked in 2 mM EDTA for 2–4 days to minimize Zn²⁺ leaching during the experiments (Kay et al. 2004). Cover glasses with Fura-2 loaded cells were washed in 3 ml KRH containing 2 µM FluoZin-3 and then placed at the bottom of the open chamber and transferred to the microscope. KRH with FluoZin-3 (150 µl) was added to the chamber. To stimulate cells, 10 µl tolbutamide stock solution containing 2 µM FluoZin-3 were added to give a final concentration of 100 µM tolbutamide. Preliminary experiments showed that the final homogeneous concentration of the stimulator in the chamber was reached within 5 s. For controls, KRH buffer without tolbutamide was applied to the cells. FluoZin-3 fluorescence images were collected every 2 s and the average fluorescence intensities were measured after background subtraction in a region of interest (ROI) of ~20 µm around the cell periphery. Cells were identified by differential interference contrast (DIC) images before the measurement and by the low Zn–Fluozin background from the cell membrane. FluoZin-3 does not penetrate into the cell, presumably because of the three negative charges on the molecule at physiological pH (Gee et al. 2002, Kay et al. 2004). Cells were stimulated during 10 min, after which they were washed by perifusion with KRH containing 3 mM glucose during 30 min. Then, the cells were stimulated again, and [Ca²⁺]_i was measured as described above. The microscope system does not allow a change of excitation exposure during simultaneous measurements. Therefore, attempts were not made to measure Zn²⁺ efflux and Ca²⁺ response simultaneously, because Zn²⁺ measurements required higher excitation intensity than calcium measurements with Fura-2. The wavelengths used for calcium measurements (380 and 340 nm) can be harmful for cells during intense exposure.

Confocal microscopy and measurements of cytoplasmic Ca²⁺ with Fluo-3

Experiments were performed using the Leica SP2 spectral laser scanning confocal microscopy system (Leica Microsystems, Mannheim, Germany). Measurements were performed with the technique similar to that described by Nadal et al.(1999). Intact islets were loaded with the calcium dye Fluo-3 for 90 min at room temperature and then placed on coverslips on the bottom of an open perifusion chamber always in the same part of the chamber close to the inflow tube. The chamber was put on the stage of the microscope and maintained at 37 °C. Islets were continuously superfused at a flow rate of 0.6 ml/min. Cells were first visualized using transmission laser scanning microscopy. Fluo-3 was excited with the 488 nm line of an argon laser. Fluo-3 responses were recorded from peripheral cells in one randomly chosen cross-section in islets using a Leica x40 oil immersion lens with numerical aperture 1.3. The pinhole was optimized for the x40 objective. The resulting fluorescence was recorded in a channel set up to detect emitted light in the range 510–600 nm. Islets were stimulated twice with 20 µM tolbutamide as described above. Images were collected at 2-s intervals, and fluorescence signals from individual cells were measured as a function of time by using the Leica Confocal Software (Heidelberg, Germany) package. Each image frame was examined to adjust the regions of interest for slight movements in cellular location. Images were analyzed and results were plotted using Leica Confocal Software. Fluo-3 is a nonratiometric dye and the fluorescence intensity varies with even minor experimental change. The calcium-dependent change in fluorescence is therefore expressed as arbitrary units.

Chemicals

Collagenase type I, HEPES, and poly-L-lysine were purchased from Boehringer (Mannheim, Germany). BSA, fraction V, EGTA, arginine and DNAase were all from Sigma Chemical Co. Culture medium RPMI 1640 was obtained from Gibco, and fetal calf serum was from Labkemi AB (Stockholm, Sweden). Benzylpenicillin and garamycin were from Schering Co. (Kenilworth, NJ, USA). Tolbutamide was from Hoechst AG (Frankfurt, Germany). Fura 2-AM, Fluo-3, FluoZin-3, and TPEN were from Molecular Probes Inc. (Eugene, OR, USA). Puratronic CaCl₂ and MgSO₄ were from Alfa Aesar (Karlsruhe, Germany). All other reagents were of analytical grade.

Statistical analysis

The data are presented as means±S.E.M. Comparisons of responses from the same cell during first and second stimulation were made using Student’s t-test for paired data and calculated correlation coefficients (Pearson correlation). The linear representative of the correlation was calculated using the least square method. For comparison between sets of experiments, two-tailed Student’s t-test for independent data was used. The significance limit was set at P<0.05.

	Results

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Cell specificity of the ß-cell Ca²⁺ response to tolbutamide

Responses were most often registered in cells situated close to the center of the chamber. Control experiments have shown no differences in lag time for [Ca²⁺]_i rise in cells situated in the central area and on the periphery of the chamber when cells were stimulated with high glucose (133±24 s (n=7) vs 158±21 s (n=12) respectively).

The average lag time for tolbutamide-induced [Ca²⁺]_i rise is shorter than that for the response to glucose and unlike glucose, tolbutamide causes no initial lowering in [Ca²⁺]_i (Grapengiesser et al. 1988). We tested 100 µM, which gives near maximal stimulation and 20 µM tolbutamide to find out if the lag time is concentration dependent. The lower concentration may provide more heterogeneous responses with regard to timing and reduce the risk for persisting effects that may influence the second response. Tolbutamide at 20 µM induced no response in most cells when tested together with 3 mM glucose (data not shown). Therefore, the glucose concentration was increased to 5 mM to increase the number of responding cells (Frederickson & Bush 2001). The lag time for calcium rise was longer with 20 µM than with 100 µM tolbutamide during the first stimulation in lean and ob/ob mouse ß-cells (Table 1, P<0.001 for lean mouse, P<0.005 for ob/ob mouse, P=0.11 for db/db mouse ß-cells). Timing of the response was cell-specific in ß-cells from lean but not from ob/ob or db/db mice both with 100 and 20 µM (Table 1). The lag time was shorter in db/db mouse ß-cells than in lean mouse ß-cells (Table 1). ß-Cells from ob/ob mice also reacted more promptly to stimulation with 100 µM tolbutamide than lean mouse ß-cells.

View larger version (15K): [in this window] [in a new window]   Figure 1 Ca2+responses from a representative isolated ß-cell during two consecutive stimulations with 100 µM tolbutamide. (A, B and C) Recordings from the first (solid line) and second stimulations (dotted line) of (A) a lean, (B) ob/ob, and (C) db/db mouse ß-cell respectively. The resting period between stimulations was 30 min. (a–c) The correlation between lag times for [Ca2+]i rise during the first stimulation, LTr1 (x-axis) and second stimulation LTr2, (y-axis) for all studied cells in the three animal models respectively. Correlation coefficients, mean values, and numbers of experiments are shown in Table 1. Basal medium was switched to test medium at time 0 min.

View larger version (15K): [in this window] [in a new window]   Figure 2 Ca2+ responses from a representative isolated lean, ob/ob, and db/db mouse ß-cell during two consecutive stimulations with 20 µM tolbutamide. For experimental details, see Materials and Methods and legend of Fig. 1. Test medium contained 5 mM glucose. Correlation coefficients, mean values, and numbers of experiments are shown in Table 1.

Arginine induced cell-specific Ca²⁺ responses in lean mouse ß-cells with regard to both timing and magnitude (Table 1). There was no correlation between lag times for calcium rise during the first and second stimulation in ß-cells from db/db or ob/ob mice but peak heights were cell-specific in db/db mouse ß-cells (Table 1). As with tolbutamide, the response in db/db mouse ß-cells occurred with shorter lag times than in lean mouse ß-cells and reached a lower magnitude.

Cell specificity of the insulin secretion in isolated ß-cells

We have also tested to what extent temporal cell specificity of the Ca²⁺ response is related to insulin release. Zn²⁺ is co-released with insulin and Zn²⁺ efflux was measured as a marker of insulin secretion (Frederickson & Bush 2001, Qian et al. 2003). The secretory response to 100 µM tolbutamide was recorded during the first stimulation and the Ca²⁺ response was recorded during the second stimulation (Fig. 3A). We found that the time of onset of the Zn²⁺ outflow correlated with the lag time for calcium rise in lean mouse ß-cells (Table 1, Fig. 3B) but not in ob/ob mouse ß-cells (Table 1, Fig. 3C). We have not compared the magnitudes of Zn²⁺ and Ca²⁺ response because Zn²⁺ dissolves quickly in the surrounding medium and the rise in Zn²⁺ outflow is difficult to estimate.

View larger version (13K): [in this window] [in a new window]   Figure 3 Ca2+ response and Zn2+ outflow from isolated ß-cells during two consecutive stimulations with 100 µM tolbutamide. For experimental details, see Materials and Methods. (A) Recordings of the Zn2+ outflow from a lean mouse ß-cell during the first stimulation (solid line) and the Ca2+ response during the second stimulation (dotted line). The resting period between stimulations was 30 min. (B and C) Correlation between lag time for Zn2+ outflow LTr1 (x-axis) and lag time for [Ca2+]i rise LTr2, (y-axis) in (B) lean and (C) ob/ob mouse ß-cells. Test medium contained 3 mM glucose. Correlation coefficients, mean values, and numbers of experiments are shown in Table 1. a.u., arbitrary unit.

We studied responses from individual ß-cells on the periphery of Fluo-3-labeled intact islets (Fig. 4A) during two consecutive stimulations. Tolbutamide at 20 µM was chosen as the stimulus to get a larger variation in lag time for Ca²⁺rise. We tested 30 cells in six lean mouse islets and 87 cells in ten ob/ob mouse islets. Parameters were calculated as described for isolated ß-cells. ß-Cells within lean mouse islets showed cell-specific responses with a high degree of correlation between lag times during the first and second stimulation (Fig. 4C). Figure 4B shows representative traces from two cells. Within each islet, there was often only a small difference in lag time among individual cells and between the first and second stimulation (Table 1). PH1 and PH2 were correlated, but PH2 was lower than PH1 (Table 1). A lower PH2 could be due to dye degradation and/or leakage. However, we previously recorded a lower second response to glucose also using Fura-2 in intact islets (Pakhtusova et al. 2003).

View larger version (31K): [in this window] [in a new window]   Figure 4 Ca2+ responses from individual cells within lean and ob/ob mouse islets during two consecutive stimulations with 20 µM tolbutamide. (A) An image of a lean mouse islet loaded with Fluo-3 and stimulated with 20 µM tolbutamide+5 mM glucose; bar=20 µm. For clarity, the image was processed with a median filter (Leica Confocal Software) to minimize noise. (B) Recordings from two cells indicated by arrows in A. (C and D) Correlation between LTr1 (x-axis) and LTr2, (y-axis) in (C) lean and (D) ob/ob mouse ß-cells. For experimental details, see Materials and Methods and Results sections. Mean values, correlation coefficients, and numbers of experiments are presented in the Table 1.

	Discussion

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We found earlier that normal mouse ß-cells show cell-specific Ca²⁺ responses when stimulated with glycolytic as well as with mitochondrial substrates (Gustavsson et al. 2005). Here we have shown that normal ß-cells have unique and reproducible, i.e., cell-specific, Ca²⁺ responses also to the nonmetabolic stimulators tolbutamide and arginine. This indicates cell specificity not only of the stimulus–secretion coupling as a whole, but also of different parts of it. While cell specificity seems to be a robust phenomenon in normal ß-cells, ob/ob and db/db mouse ß-cells show no cell-specific timing of Ca²⁺ response when stimulated with tolbutamide or arginine. The mechanism of action of arginine is a direct depolarization of the ß-cell membrane similar to that of cations (Charles et al. 1982, Herchuelz et al. 1984) and tolbutamide causes closure of K⁺_ATP channels. Although we did not study ion channel function directly, the results suggest that the response of plasma membrane ion channels is also cell-specific in normal ß-cells. Cell-specific function of Ca²⁺ channels may be linked to the number of channels and/or differences in channel properties, such as the gating. Previous reports have shown different results regarding Ca²⁺ channel function in ß-cells from animals with conditions similar to type-2 diabetes. A reduction of L-type Ca²⁺ currents and a decreased expression of the 1-subunit of L-type Ca²⁺ channels was observed in the Zucker diabetic fatty rat (Roe et al. 1996). Other studies report an increased activity of L-type Ca²⁺ channels in the Goto Kakisaki rat (Kato et al. 1996). A lower expression of Ca²⁺ channels could possibly explain the reduced Ca²⁺ response but not the loss of temporal cell specificity, which we found in db/db and ob/ob mouse ß-cells. This indicates that other alterations of Ca²⁺ channels are present in ß-cells hyperglycemic mice.

The failure of db/db and ob/ob mouse ß-cells to show cell-specific timing of response to a K_ATP-channel blocker (tolbutamide) may be due to increased activity of K_ATP channels and impaired electrical activity (Ashcroft & Rorsman 2004). The cell-specific magnitude of the response to tolbutamide in ß-cells from hyperglycemic mouse indicates that timing and magnitude of the Ca²⁺ response are controlled by different mechanisms.

Type-2 diabetic ß-cells show a delayed Ca²⁺ response to glucose but not to arginine or sulfonylureas (Zaitsev et al. 1997, Del Guerra et al. 2005). In fact ß-cells from db/db mice responded to tolbutamide and arginine with a shorter lag time than lean mouse ß-cells and a similar tendency was found also in ob/ob mouse ß-cells. The magnitude of the response was lower in db/db mouse ß-cells with both nutrient (Roe et al. 1994, Gustavsson et al. 2005) and non-nutrient stimulators (this report). It may be that chronic hyperglycemia slows down the metabolic rate but increases the sensitivity of K⁺_ATP or Ca²⁺ channels (Kato et al. 1996).

The lag time for Ca²⁺ rise was longer in response to 20 µM tolbutamide when compared with 100 µM, but the magnitudes of response and the patterns of [Ca²⁺]_i changes were similar. There have been no studies aimed at comparing the timing of ß-cell depolarization when different concentrations of sulfonylureas are used. However, in the paper by Henquin (1998), recordings of electrical responses to 15, 25, and 100 µM tolbutamide were shown. From the data, it appears that the depolarization is slower with 15 µM, but does not differ between 25 and 100 µM tolbutamide (Henquin 1998). The longer lag time for calcium rise to 20 µM tolbutamide is therefore likely to be caused by a slower depolarization. The difference is probably not related to the difference in glucose concentration in the stimulatory solution, because an increase in glucose concentration would induce a quicker response.

The peak height during the second stimulation was smaller in ob/ob mouse ß-cells than in lean mouse cells (Table 1). Cells may be desensitized by the previous exposure to sulfonylureas (McClenaghan et al. 2000), but it is difficult to explain the fact that only mice with a milder hyperglycemic syndrome but not severely diabetic mice develop the desensitization.

The timing of Zn²⁺ outflow recorded during the first stimulation with tolbutamide in lean mouse ß-cells correlated with the timing of Ca²⁺response recorded during the second stimulation. We showed that ß-cells repeat their Ca²⁺ response pattern during consecutive stimulations with tolbutamide. It has also been found that cytoplasmic calcium changes and Zn²⁺release are linked with regard to both timing and magnitude (Qian et al. 2004). With the assumption that the timing of insulin release is correlated to the lag time for Ca²⁺ response also during consecutive stimulations we can, therefore, conclude that the timing of insulin release to tolbutamide is cell-specific in normal mouse ß-cells. No such correlation was found in ob/ob mouse ß-cells. Most likely, this is coupled to the lack of cell specificity for Ca²⁺ responses to tolbutamide.

It was recently claimed that cells within an islet show identical temporal responses (Kuznetsov et al. 2005). We could identify fast and slow responding cells within both lean and ob/ob mouse islets. The variation was rather small in lean mouse islets, but in ob/ob mouse islets the range of lag times was almost as large as in isolated ß-cells. This could be due to a disturbed stimulus–secretion coupling or to a damage of cell contacts and may explain the disorganization of [Ca²⁺]_i oscillations found in ob/ob mouse islets (Ravier et al. 2002). ß-Cells both within intact lean and ob/ob mouse islets showed cell-specific timing of their Ca²⁺ responses to tolbutamide. Thus, cell-specific timing of response to tolbutamide can be observed in functionally coupled, but not in isolated ß-cells from ob/ob mice. This supports the hypothesis that some ß-cells have a pacemaker function. ß-Cells may vary in their functional characteristics, because ß-cells within pancreatic islets originate not from a single stem cell but from a number of different low-differentiated precursors (Lechner & Habener 2003). We have previously reported that intact islets as a whole show a specific response (Pakhtusova et al. 2003). Taken together, this indicates that both the cell specificity of each ß-cell and the functional connection between cells is important for islet-specific behavior and adequate insulin release.

Normal ß-cells show cell-specific responses with all tested stimuli. ß-Cells from hyperglycemic mice have shown cell-specific timing and magnitude of responses to glucose and glyceraldehyde, but they have lost their temporal cell specificity when stimulated with a mitochondrial substrate, which suggested an altered mitochondrial function (Gustavsson et al. 2005). The loss of cell-specific timing of response also at the step of Ca²⁺ channel opening suggests that the mechanism(s) for setting the individual response characteristics is impaired in diabetic mouse ß-cells. Therefore, cell specificity for all steps of the stimulus–secretion coupling may be important for normal function.

ß-Cells constitute more than 90% of the cells islet ob/ob mice and about 70% in normal mouse islets. The proportion of ß-cells in islets from severely diabetic mice is reduced (Baetens et al. 1978). Therefore, it is possible that many of the cultured cells were non-ß-cells in the latter two mouse models. However, cells included in this study showed [Ca²⁺]_i responses to tolbutamide, tolbutamide in combination with glucose, and arginine that were all typical for ß-cells (Berts et al. 1996, Quesada et al. 1999).

In conclusion, normal ß-cells show cell-specific Ca²⁺ responses when stimulated with nonmetabolic agents. The lag time is not cell-specific in tolbutamide- and arginine-stimulated isolated ß-cells from hyperglycemic mice. Cell specificity seems to be a robust mechanism, which is probably necessary for an adequate ß-cell function.

	Acknowledgements

 
This work was supported by the Swedish Research Council (12X-4756), the Swedish Diabetes Association, the Sahlberg Foundation, the Lars Hiertas Foundation and the Medical Faculty, Umeå University. The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.

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Received in final form 10 April 2006
Accepted 25 April 2006
Made available online as an Accepted Preprint 10 May 2006

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