Ca2+ Depletion from Granules Inhibits Exocytosis
A STUDY WITH INSULIN-SECRETING CELLS*

Wim J. J. M. ScheenenDagger §, Claes B. Wollheim, Tullio PozzanDagger , and Cristina FasolatoDagger

From the Dagger  Department of Biomedical Sciences, Consiglio Nazionale delle Ricerche, Center of Biomembranes, University of Padova, Via G. Colombo 3, 35100 Padova Italy and  Division of Clinical Biochemistry, Department of Internal Medicine, University Medical Center, CH-1211, Geneva 4, Switzerland

    ABSTRACT
Top
Abstract
Introduction
Procedures
Results
Discussion
References

The secretory compartment is characterized by low luminal pH and high Ca2+ content. Previous studies in several cell types have shown that the size of the acidic Ca2+ pool, of which secretory granules represent a major portion, could be estimated by applying first a Ca2+ ionophore followed by agents that collapse acidic pH gradients. In the present study we have employed this protocol in the insulin-secreting cell line Ins-1 to determine whether the Ca2+ trapped in the secretory granules plays a role in exocytosis. The results demonstrate that a high proportion of ionophore-mobilizable Ca2+ in Ins-1 cells resides in the acidic compartment. The latter pool, however, does not significantly contribute to the [Ca2+]i changes elicited by thapsigargin and the inositol trisphosphate-producing agonist carbachol. By monitoring membrane capacitance at the single cell level or by measuring insulin release in cell populations, we show that Ca2+ mobilization from nonacidic Ca2+ pools causes a profound and long lasting increase in depolarization-induced secretion, whereas breakdown of granule pH had no significant effect. In contrast, releasing Ca2+ from the acidic pool markedly reduces secretion. It is suggested that a high Ca2+ concentration in the secretory compartment is needed to sustain optimal exocytosis.

    INTRODUCTION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

A rise in intracellular Ca2+ concentration ([Ca2+]i) is necessary to induce regulated secretion in most cell types (1, 2). In neurons, [Ca2+]i increases up to several hundred µM are needed to trigger vesicle fusion, whereas in endocrine cells, granule exocytosis appears to require lower [Ca2+]i rises (3-7). The time course of exocytosis also appears different in the two cell types. Synaptic vesicle fusion is very fast (µs) and abrupt, whereas granule fusion is slower and more sustained (3, 8, 9).

Aside from these differences, important similarities exist between secretory vesicles and granules. Both secretory vesicles and granules contain large amounts of Ca2+ ions (10-13). The function traditionally attributed to the high Ca2+ content in the secretory compartment is the packaging and processing of intravesicular content (14, 15). More recently a granular localization of the type 3 InsP31 receptor has been suggested, based on high resolution immunocytochemistry of pancreatic beta -cells (16). In the exocrine pancreas, evidence has been provided suggesting that the intragranular Ca2+ content is released by opening of low affinity InsP3 receptors (17). These conclusions, however, have recently been challenged (18, 19), and the role of granular Ca2+ remains elusive. Another line of evidence suggesting that intragranular Ca2+ is implicated in secretion comes from the recent identification of an acidic Ca2+-binding protein, granule lattice Protein 1 (Grl1p), in dense core secretory granules of Tetrahymena thermophila that appears essential for regulated secretion (20).

Another common feature between secretory vesicles and granules is their low luminal pH. They share this characteristic with the lysosomal/endosomal compartment and the trans-Golgi network (21, 22). Indeed, the low pH of the lumen has proven a reliable means for determining the Ca2+ content of the so-called "acidic Ca2+ pool." Since ionophores such as ionomycin or A23187 are largely ineffective in transporting Ca2+ from an acidic environment (28), the pH gradient between lumen and cytosol must be collapsed before they can effectively release the Ca2+ content of this pool into the cytoplasm (23-26).

The aim of the present study was to establish whether the Ca2+ stored within the acidic pool is important in the late steps of exocytosis. For this purpose we employed as a model system the beta -cell line Ins-1, an insulin-secreting cell line established from a rat insulinoma that, among different lines, best retains the phenotype of beta -cells (27-29). Among other properties, Ins-1 cells display temperature-dependent and glucose-responsive secretion and, as shown here, temperature-dependent increases in membrane capacitance upon depolarizing pulses. Therefore, they can be used as an alternative to beta -cells for studying secretion at the single cell level. By using capacitance measurements in combination with agents that mobilize Ca2+ and/or collapse intracellular pH gradients, the role of different intracellular Ca2+ pools in secretion has been assessed. We here demonstrate that, although a low pH in the granules is not required for the late steps of secretion, the level of intragranular Ca2+ significantly affects the secretory profile.

    EXPERIMENTAL PROCEDURES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Cells-- Ins-1 cells were cultured as described previously (27, 29). Two days before the experiment, cells were trypsinized and plated on poly(L-lysine)-coated coverslips (diameter 24 mm; 105 cells/coverslip).

Ca2+ Measurements in Ins-1 Cells-- Cells were loaded for 30 min at 37  °C with 2 µM fura-2/AM as described previously (26). Coverslips were then bathed in 1 ml of Ringer's solution containing 140 mM NaCl, 2.8 mM KCl, 1 mM MgCl2, 2 mM CaCl2, 5.6 mM glucose, 10 mM HEPES, pH 7.4, at 33 °C. Cells were placed on the stage of an inverted microscope equipped with a 40× oil immersion objective (Zeiss, Germany) and connected to a digital video imaging system (Georgia Instruments, Roswell, GA). All experiments were performed at 31-33 °C. Excitation wavelengths were set at 340 and 380 nm. Fluorescence emission at 510 ± 15 nm was collected by a CCD camera, and 8 images were averaged/time point. Time series were acquired with a frame interval of 4 s, and images at both excitation wavelengths were stored on an optomagnetic disc recorder (Panasonic, Japan). All data were normalized to the first min base-line ratio.

The total content of cellular Ca2+ under different conditions was assayed by atomic absorption spectrophotometry. Cells (20 × 107 cells/ml) were suspended in a Ca2+-free medium containing 1 mM EGTA and challenged with ionomycin (10 µM) alone or in combination with monensin (10 µM). Cell aliquots (107) were then centrifuged for 5 min at 14,000 rpm in Eppendorf tubes containing 100 µl of sucrose 12.5% and 400 µl of silicon oil. The pellet was resuspended in 1 ml of a solution containing Triton (0.02%) and NaOH (0.2 N) before measurement.

Membrane Capacitance Measurements-- Unless otherwise specified, during electrophysiological recordings, cells were perfused at 31-33  °C with an external solution containing 118 mM NaCl, 20 mM tetraethylammonium chloride, 5.6 mM CsCl, 1.2 mM MgCl2, 2 mM CaCl2, 5.7 mM glucose, 10 mM HEPES at pH 7.4. Electrophysiological recordings and fura-2 fluorescence were performed in the whole-cell configuration of the patch-clamp technique with a computer-based patch-clamp amplifier (EPC-9, HEKA, Lambrecht, Germany) controlled by the Pulse software (HEKA). The internal solution consisted of 145 mM glutamic acid, 155 mM CsOH, 1 mM MgCl2, 8 mM NaCl, 2 mM MgATP, 0.1 mM cAMP, 0.1 mM fura-2 (or 0.1 mM BCECF), and 10 mM HEPES at pH 7.2. Membrane capacitance (Cm) was measured with the "sine+dc" mode of the "lock-in" extension of the Pulse software, based on the Lindau-Neher algorithm (30). An 800 Hz, 40 mV peak-to-peak sinusoid stimulus was applied to the DC holding potential of -80 mV. During a depolarizing pulse and 5 ms before and after the pulse, no sinewave was applied. No leak subtraction was performed on the evoked currents in the calculations used. After the whole-cell configuration was established, Cm was recorded and canceled by the automatic capacitance compensation of the EPC-9. The procedure was repeated every 180 s to prevent a possible saturation of the lock-in signal (31).

For fast capacitance changes after depolarizing pulses, it has been reported that activation of a Na+ current can lead to transient increases in Cm not linked to exocytosis (Delta Ct) (32). Delta Ct contributed maximally 10% of the initial Delta Cm and returned to zero within 100 ms after the depolarization. To reduce its contribution on Delta Cm measurements, the first 100 ms of the trace was discarded. The capacitance values of the following 200 ms were averaged and compared with the capacitance values before the depolarization to obtain the Delta Cm reported in Figs. 6 and 7.

The [Ca2+]i was monitored by a photometry equipment (T. I. L. L. Photonics, Germany) controlled by the fura-2 extension of the Pulse software (HEKA) as described previously (33). Calculation of [Ca2+]i was performed on the calibrated ratio values (360 nm/380 nm), where Rmin (0.86), Rmax (6.07), and K-factor (1.68 ×10-3) were obtained by an internal calibration procedure. The fura-2 fluorescence, the holding current, the lock-in, and other parameters were synchronously recorded also at low resolution (3 Hz) by the X-Chart extension of the Pulse software (HEKA). Time courses of Cm as shown in Figs. 3-5 were obtained from the Cm trace, recorded at low frequency by a point-by-point subtraction. Positive and negative values are indicative of, respectively, exocytotic and endocytotic events. The cytosolic pH was monitored by ratioing the BCECF fluorescence signal (510 nm), excited at 440 and 490 nm.

Ionomycin and monensin were prepared from stock solution in Me2SO (or ethanol) (0.4% final concentration) in the Ca2+-free external solution containing 5 mM EGTA to prevent cells from loading with extracellular Ca2+ in the presence of ionomycin. All drugs were applied by local pressure from a wide-tipped micropipette (5-10 µm) positioned close to the cell.

Insulin Secretion Studies-- Superfusion experiments on Ins-1 cell suspensions were performed as described previously (34). In short, cells were brought into suspension and placed in 1-ml superfusion chambers at a density of 106 cells/chamber. The cells were superfused at a rate of 1 ml/min at 37 °C, and test substances were introduced with the buffer. One-min fractions were collected and subjected to an insulin radioimmunoassay. For presentation, the KCl-induced stimulation of the second pulse was integrated and normalized to that of the first pulse.

Materials-- fura-2/AM, fura-2 free acid, BCECF free acid, and Pluronic F-127 were obtained from Molecular Probes (Eugene, OR, USA); culture media and sera were from Technogenetics (Milan, Italy); and other chemicals were from Sigma.

    RESULTS
Top
Abstract
Introduction
Procedures
Results
Discussion
References

[Ca2+]i Dynamics in Ins-1 Cells-- When [Ca2+]i was monitored by fura-2 in intact cells bathed in a medium containing CaCl2 (2 mM) and glucose (5.6 mM), approximately 50% of the cells displayed asynchronous oscillations of [Ca2+]i, whose frequency and amplitude largely depended on the cell batch (29). In Fig. 1, the [Ca2+]i kinetics from several individual cells were averaged, leading to a partial masking of the initial oscillations. The addition of EGTA immediately abolished these oscillations and caused a decrease in the base-line ratio, indicating that they depend on Ca2+ influx. The presence of acidic Ca2+ pools and their contribution to [Ca2+]i rises were evaluated with the protocol previously employed in other cell lines (23-26). In Fig. 1A, after EGTA addition, the fast-exchangeable Ca2+ pool was released by the Ca2+ pump inhibitor thapsigargin (Tg) (1 µM). The subsequent addition of the Ca2+ ionophore ionomycin (1 µM) led to a small, further increase in [Ca2+]i, indicating that in these cells the large majority of the ionomycin-sensitive pool is represented by the Tg-sensitive one. Release of the acidic Ca2+ pool was then achieved by the addition of the Na+/H+ exchanger monensin (2 µM). Qualitatively similar data have been obtained by addition of the weak base chloroquine (40 µM), used in place of monensin to dissipate the intraluminal pH gradients. The increase in [Ca2+]i after monensin (or chloroquine) application requires the pretreatment with ionomycin (26). In fact, addition of either drug alone was without appreciable effect on [Ca2+]i (data not shown). Integrating peak areas showed that the amount of Ca2+ residing in acidic compartments was, on average, 51.3 ± 3.6% (n = 4) of total releasable Ca2+. The amount of Ca2+ released from the different Ca2+ pools was also assayed by atomic absorption spectrophotometry. In controls (unstimulated conditions), the total content of cellular Ca2+ was estimated to be 5.7 nmol of Ca2+/mg of protein (n = 3). Ionomycin alone or ionomycin and monensin together released 2.4 ± 1.1 and 4.5 ± 0.4 nmol of Ca2+/mg of protein, respectively (n = 3). Thus, of the total releasable Ca2+, about 54% was released by ionomycin alone; the remaining 46% was then attributed to the Ca2+content selectively released from the acidic pool.


View larger version (11K):
[in this window]
[in a new window]
 
Fig. 1.   InsP3 production neither reduces nor releases Ca2+ from acidic Ca2+ pools. The [Ca2+]i was monitored in individual cells loaded with fura-2/AM. The traces shown are means of 15 to 20 cells. A, Tg treatment (1 µM, T) in Ringer's solution containing EGTA (5 mM, E) causes a transient increase in [Ca2+]i. The subsequent addition of ionomycin (1 µM, I) led to a small, transient increase in [Ca2+]i, whereas subsequent application of monensin (2 µM, M) induced a strong Ca2+ release. B, addition of 0.5 mM carbachol (C) in the presence of 5 mM EGTA induces a transient increase in [Ca2+]i. The subsequent addition of Tg (1 µM) shows an additional, smaller increase. This treatment did not affect the ionomycin (1 µM) or monensin response (2 µM). C, when Tg (1 µM) was applied before CCh (0.5 mM), the [Ca2+]i increase induced by CCh was completely abolished, whereas the peak induced by ionomycin (1 µM) and monensin (2 µM) remained unchanged.

We functionally tested for the existence of InsP3 receptors on granules. Fig. 1B shows that InsP3 production, induced by the muscarinic agonist carbachol (CCh, 0.5 mM), reduced the Tg-sensitive pool but had no effect on the size of the peak induced by monensin application. In fact, in the presence of CCh, the acidic pool represented 49 ± 1.7% (n = 7) of total mobilizable Ca2+. Finally, Fig. 1C shows that pretreatment with Tg abolished the peak in [Ca2+]i induced by CCh, indicating that in this cell type, InsP3- and Tg-sensitive pools fully overlap. Altogether the data demonstrate that acidic Ca2+ compartments in Ins-1 cells are depleted neither by InsP3 produced by receptor stimulation nor by inhibition of Tg-sensitive pumps.

Capacitance Changes After Activation of Voltage Operated Ca2+ Channels-- Fig. 2 shows fast changes in Cm, membrane conductance (Gm) as well as series conductance (Gs) before and after a 200-ms depolarizing pulse from -80 mV holding potential to 0 mV. With 200-ms pulses, individual cells displayed Delta Cm ranging from 20 to 400 fF with an average of 49.2 ± 4.4 fF (mean ±S.E., n = 76). Increasing the pulse duration to 400 ms led to an increase in Delta Cm of 60% when compared with a 200-ms depolarizing pulse in 4 of 9 cells (data not shown). However, these longer depolarizing pulses caused a rapid rundown of the evoked Ca2+ currents. We therefore decided to perform the experiments with 200-ms pulse duration.


View larger version (21K):
[in this window]
[in a new window]
 
Fig. 2.   Cm changes induced by depolarization in Ins-1 cells. Activation of a Ca2+ current by a 200-ms depolarizing step from -80 to 0 mV (bottom panels) induces fast changes in Cm (upper panel), recorded as described under "Experimental Procedures." Monitoring Gs and Gm (second and third panels) shows that, after the depolarizing pulse, Gm transiently increases and returns to basal values within 100 ms, whereas Gs does not change. The figure also shows the time used to calculate the Delta Cm (shaded area).

By following Cm at a lower frequency (3 Hz) it can be seen that upon such a depolarization Delta Cm remained constant for up to 4 min before a rundown in secretion was observed, as long as these pulses were at least 20 s apart (Fig. 3A). This result indicates that secretion in Ins-1 cells is not easily "exhaustible" by successive 200-ms depolarizing pulses. With this protocol, maximum secretion increased the initial membrane capacitance by about 10% and was equivalent to fusion of approximately 300 granules (n = 13) (assuming the mean diameter of the insulin-containing granules to be 250 nm and a membrane capacitance of 1 µF/cm2). However, since an interval of 20 s is too short to apply test substances between subsequent depolarizations, the interpulse duration was increased to 90 s. From the time course of Delta Cm, we also noticed that, upon depolarization, the majority of the cells (65%) displayed only an increase in Cm; in the remaining 35%, slow endocytotic processes were observed after the first and, occasionally, the second pulse (see Fig. 3B), whereas large, abrupt endocytotic events were never observed under our experimental conditions.


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 3.   Exocytosis in Ins-1 cells is not easily "exhaustible." A, changes in Cm (Delta Cm) after repetitive depolarizing pulses of 200-ms duration, delivered every 20 s. The time course of Delta Cm was obtained by subtracting from each point of the Cm trace (recorded at low frequency (3 Hz)) the preceding value. B, enlargement of an exocytotic event that is followed by endocytosis. The shaded areas are an indication of the exocytosis (light gray) and endocytosis (dark gray).

In addition to Cm and Ca2+ current, the [Ca2+]i was monitored as described under "Experimental Procedures." As can be seen in Fig. 4A, the amplitude of the [Ca2+]i peaks decreased upon subsequent depolarizations, whereas the integrated [Ca2+]i peaks and Ca2+ currents (Delta Ip) (Fig. 4C) as well as Delta Cm (Fig. 4B) remained unchanged. The decrease in [Ca2+]i peak amplitude is probably due to the relatively slow influx of fura-2 from the pipette, leading to changes in the Ca2+ buffering capacity of the intracellular medium during prolonged incubations (35).


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 4.   Measurements of [Ca2+]i, Delta Cm, and Ca2+ charge in control cells. A and B, [Ca2+]i and Cm changes were monitored as described under "Experimental Procedures." Delta Cm is plotted as described in Fig. 3. C, the integrated Ca2+ peak was obtained from the trace shown in panel A, whereas the Ca2+ charge (q) was obtained by integrating the Ca2+ current.

Role of Granular Ca2+ Content in Secretion-- We first tested if drug application by itself induced Delta Cm. From the low frequency recording of Cm it can be seen that during application of ionomycin, monensin, or the combination of the two, an increase in Cm occurred (Fig. 5A-C, lower panels). Since similar increases were observed when the solvents ethanol or Me2SO were tested and when experiments were performed at room temperature to inhibit regulated secretion (36), we conclude that to a large extent the observed changes are due to the solvent. Chloroquine, on the other hand, being dissolved in Ringer's solution, had no effect. We also tested whether drug application changed the intracellular pH; therefore in some experiments BCECF was included in the intracellular solution instead of fura-2. These experiments showed that application of none of the drugs, applied alone or in combination, significantly altered the cytosolic pH when cells were kept in the whole-cell configuration (data not shown).


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 5.   [Ca2+]i and Delta Cm after application of test substances during depolarizing pulses. Depolarizing pulses of 200-ms duration were delivered every 90 s. During the first and the second pulses, test substances were puffed onto the cell for 20 s. [Ca2+]i (upper panels) and Cm changes (lower panels) were measured as described in Fig. 4 in cells challenged, respectively, with 2 µM monensin/5 mM EGTA (A), 1 µM ionomycin/5 mM EGTA (B), 1 µM ionomycin/2 µM monensin/5 mM EGTA (C).

To determine the role of the granular Ca2+ content in granule fusion, the increases in Cm in response to 200-ms depolarizing pulses were thus monitored immediately before and after application of the different drugs. Changes in Cm at the second and subsequent depolarizing pulses were normalized to the change obtained in the first pulse. When Delta Cm was monitored in untreated, control cells after this normalization protocol, it remained stable during the second depolarization (Fig. 6, n = 40). Figs. 5B and 6 show that, after a brief application of ionomycin, the subsequent depolarizing pulse caused a consistently larger increase in Delta Cm (47 ± 12%, n = 10). The increase in Delta Cm after the ionomycin pulse was relatively long lasting since it was maintained for at least 3 min and did not depend on larger Ca2+ currents during subsequent depolarizations (compare Figs. 7, A and B).


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 6.   Comparison of Cm changes in control and treated cells. Cells were repeatedly depolarized and challenged with test substances as described in Fig. 5. Changes in Cm immediately after the depolarizing pulses were monitored as described in Fig. 2. Values obtained in the second pulse were then normalized to the first pulse. *, p < 0.05; 2-tailed Student's t test. The effect of chloroquine (40 µM) and monensin (2 µM) alone was not statistically significant.


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 7.   Long lasting effect of the ionomycin-enhanced stimulation of regulated secretion. Cells were treated as described in Fig. 5. Between the first and the second depolarizing pulses, test substances were applied: black-square, control; black-down-triangle , 1 µM ionomycin/5 mM EGTA; bullet , 2 µM monensin/5 mM EGTA; ×, 1 µM ionomycin/2 µM monensin/5 mM EGTA. Changes in peak currents (Delta Ip) (A) and Delta Cm (B), recorded as described in Fig. 2, were normalized to the changes obtained during the first depolarizing pulse. *, p < 0.05; 2-tailed Student's t test.

Application of chloroquine between the first and the second pulse had no effect (n = 6), whereas monensin led to a quite variable stimulation (Figs. 5A and 6). On average, however, the stimulation caused by monensin treatment between two successive depolarizing steps was not statistically significant (23 ± 21%, n = 9; see also Fig. 7).

A completely different pattern was observed when the Ca2+ ionophore and chloroquine (or monensin) were applied together in order to discharge the Ca2+ content of the acidic pools. The Delta Cm increase following the depolarizing step, elicited after the discharge of the acidic Ca2+ pools (Figs. 5C and 6), was reduced not only with respect to the potentiation caused by ionomycin (46 ± 13%, n = 17, monensin/ionomycin; 54 ± 9%, chloroquine/ionomycin, n = 6) but was also reduced with respect to untreated control cells (21 ± 9%, monensin/ionomycin; 31 ± 2%, chloroquine/ionomycin).

The reduction in Delta Cm following the depolarizing step was also observed with further test pulses, i.e. it was prolonged for up to 3 min (Fig. 7B). It is worth mentioning that there was no significant change in either Gm or Gs when the second depolarizing pulse was compared with the first pulse either in controls or treated cells (data not shown). Moreover, application of ionomycin, monensin, or the combination of ionomycin and monensin had no significant effect on changes in [Ca2+]i (Delta Ca) (Fig. 5) and Delta Ip of the subsequent depolarizing pulses (Fig. 7A).

The inhibition of secretion did not exceed more than 30% of the control values. A possible explanation for this incomplete inhibition is that a complete alkalinization by monensin or chloroquine (and therefore complete discharge of granule Ca2+) takes longer than the application time of 20 s used in our electrophysiological experiments. To test this possibility, studies were performed where the H+ exchanger was present for 5 min before the first depolarizing pulse was given, to ensure a complete breakdown of the pH gradient. After this prolonged incubation, secretion during the first depolarizing pulse was within the expected range of variability (43 ± 7.2 fF; n = 7). However, the prolonged treatment with monensin did not further increase the level of inhibition obtained when ionomycin was applied between the first and second pulses (24 ± 12%, n = 7; Fig. 6).

Insulin Secretion Studies-- To determine whether the reduction in Delta Cm caused by acidic Ca2+ pool depletion was attributable, at least in part, to fusion of insulin-containing granules, we followed the release of insulin in populations of cells treated with protocols that mimic those used in Fig. 5. Cell suspensions obtained from monolayers were challenged with two pulses of 30 mM KCl of 1-min duration, 5 min apart. As summarized in Fig. 8, secretion during the second depolarizing pulse was 25 ± 4% (n = 3) of that obtained during the first challenge. This reduction in secretion probably reflects a reduction in readily releasable insulin granules, although a rundown of the Ca2+ peak after depolarization may also contribute to this effect (34). One-min stimulation with 1 µM ionomycin between the first and second KCl pulses resulted in a less drastic reduction of insulin secretion (62 ± 24% that of initially released, n = 3). When a combination of ionomycin and chloroquine (or monensin) was employed, secretion during the second KCl pulse was 25 ± 11 and 24 ± 4% (n = 3), respectively; i.e. the potentiating effect of ionomycin was completely abolished.


View larger version (15K):
[in this window]
[in a new window]
 
Fig. 8.   Effect of the Ca2+-depleting protocols on insulin secretion induced by KCl in cell populations. Cell suspensions (106/ml) were depolarized with two pulses of 1 min duration of 30 mM KCl applied 5 min apart in Ringer's solution. Insulin secretion during the second KCl challenge was measured by radioimmunoassay as described (52) and was normalized to that obtained during the first pulse. 1 µM ionomycin alone or a combination of 1 µM ionomycin with 2 µM monensin or 40 µM chloroquine in Ca2+-free Ringer's solution containing EGTA (1 mM) was applied for 1 min between the first and second depolarizing pulses. * p < 0.05; 2-tailed Student's t test.

    DISCUSSION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

In the beta -cell line Ins-1, as in other secretory cells, a relatively high proportion of intracellular Ca2+ appears to be stored in acidic structures. In fact, these cells respond with a large [Ca2+]i increase to the protocol previously employed to reveal this compartment, i.e. the application of drugs that collapse internal acidic pH gradients (monensin or chloroquine) after addition of the Ca2+ ionophore ionomycin. The subcellular localization of acidic Ca2+ pools has not been determined with certainty, although it is likely that in Ins-1 cells, as in other cell types, it is heterogeneous. A rough estimation of the contribution of insulin granules to the Ca2+ content of the acidic pool can be obtained by considering the total releasable Ca2+ of Ins-1 cells (4.5 nmol/mg of protein, this work), the cell volume occupied by the granules (1.2%),2 and the releasable granule Ca2+ (about 125 nmol/mg of granule protein, Ref. 10). By using these parameters, intragranular Ca2+ mobilization could be as high as 1.5 nmol/mg of protein. We have shown here that in Ins-1 cells 46% (i.e. 2.4 nmol/mg of protein) of the total releasable Ca2+ is due to the acidic pool. Although based on a number of assumptions, these values indicate that insulin granules represent a major part (more than 60%) of the acidic compartment in this cell type.

The main goal of this investigation was to establish the role played by Ca2+ trapped in the secretory compartment in the process of secretion. To address this question, we first investigated whether or not the acidic compartment could contribute to (i) the [Ca2+]i changes induced in Ins-1 cells by the muscarinic agonist CCh and (ii) the [Ca2+]i changes induced by depolarization. The finding that Ca2+ mobilization induced by thapsigargin or InsP3 production through activation of muscarinic receptors does not affect the acidic pool is meaningful. In fact, given that insulin granules represent a large proportion of that pool, it confirms by a functional approach the conclusion of Ravazzola et al. (18) that InsP3 receptors are not expressed on the membrane of the secretory granules of beta -cells. Similarly, a role for the acidic pool (and thus for insulin granules) in Ca2+-induced Ca2+ release is unlikely since the increase in [Ca2+]i caused by depolarization was indistinguishable in controls and cells whose acidic pool had been depleted (Fig. 5).

We next tested the possibility that intragranular Ca2+ plays a role in the secretory process by monitoring membrane capacitance in single Ins-1 cells under different experimental conditions. In untreated cells, the magnitude of Delta Cm has a tendency to decrease during a series of successive pulses; however up to the fourth pulse (i.e. 300 s), Delta Cm is fairly constant. On the contrary, manipulation of intracellular Ca2+ in the time interval between the first and the second depolarizing pulses significantly changed the extent of secretion. In fact, depletion of Ca2+ from nonacidic stores led to a prolonged stimulation of secretion up to 50%. Such a priming action of ionomycin has been described previously (4), but the fact that it can last for several min at resting [Ca2+]i is a novel observation. Releasing Ca2+ from the ionomycin-sensitive compartments may favor granule recruitment from a distant cytoskeletal-anchored pool (38) or by promoting priming of granules at a late, post-docking step (39). Such a priming has been previously described by mechanisms that cause long lasting phases of moderately elevated [Ca2+]i (31).

In marked contrast with the potentiating effect of a brief increase in [Ca2+]i, releasing Ca2+ from the acidic compartments led to inhibition of secretion that reached 50% when compared with cells treated only with ionomycin. Since breakdown of the intracellular pH gradients by itself was without effect and the inhibition was observed with both monensin and chloroquine (two agents that act on pH gradients by different mechanisms), it can be concluded that the inhibitory effect is due to the release of Ca2+ from the acidic organelles, including insulin granules. Since our alkalinization protocol is by no means specific for the granules, the question can be raised as to whether the reduction in secretion is due to the decrease in Ca2+ within the granules themselves or in other acidic compartments (trans-Golgi network or lysosomes). The observation that inhibition is maximal within a few tens of seconds after acidic Ca2+ pool discharge argues for a distal site of action, i.e. the granules themselves.

It is probable that both insulin-containing granules and delta -aminobutyric acid-containing vesicles contribute to the increases in Cm monitored by us and by other groups (7, 40, 41). However, the fact that Ca2+ depletion from nonacidic and acidic compartments affects both Cm increases and insulin secretion in radioimmunoassay suggests that at least part of the effects seen in our study reflects fusion of insulin-containing granules.

A high intragranular Ca2+ concentration may be important for docking or priming of the granules for the fusion process itself or for all of the steps. It is noteworthy that a protein called Grl1p, abundant in secretory granules of T. thermophila (20) is sensitive to both Ca2+ and pH.

In conclusion, our experiments show that in Ins-1 cells, a relatively large amount of the stored Ca2+ resides in acidic compartments. A high [Ca2+] in this compartment but not a low pH is needed for optimal exocytosis.

    ACKNOWLEDGEMENTS

We are grateful to Clarissa Bartley for performing the superfusion experiments and the radioimmunoassays, M. Mancon for total Ca2+ measurements, and G. Ronconi and M. Santato for skillful assistance. We thank Drs. Aldebaran M. Hofer and Bruce G. Jenks for discussion and for critical reading of the manuscript.

    FOOTNOTES

* This work was supported by grants from Telethon number 845, European Union Programs Human Capital Mobility Network CHRXCT940500, Human Frontier Science Program RG520/95, the Armenise Foundation grant (Harvard) (to T. P.), and in part by Swiss National Science Foundation Grant 32-49755.96 (to C. B. W.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§ Supported by EU Grant CHRXCT940500. To whom correspondence should be addressed: Dept. of Cellular Animal Physiology, University of Nijmegen, Toernooiveld 1, 6525 ED Nijmegen, The Netherlands. Tel.: 00.31.24.3653335; Fax: 00.31.24.3652714; E-mail: scheenen{at}sci.kun.nl.

1 The abbreviations used are: InsP3, inositol trisphosphate; BCECF, 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein; Cm, membrane capacitance; Gm, membrane conductance; Gs, series conductance; Tg, thapsigargin; CCh, carbachol; F, farads.

2 This value has been estimated considering that Ins-1 cells contain 10% of the insulin content of beta -cells (27) and that in the latter cell type, the percentage volume occupied by granules is 11.5% based on morphometric analysis (37).

    REFERENCES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

  1. Smith, S. J., and Augustine, G. J. (1988) Trends Neurosci. 11, 458-464[CrossRef][Medline] [Order article via Infotrieve]
  2. Zucker, R. S. (1996) Neuron 17, 1049-1055[Medline] [Order article via Infotrieve]
  3. Heidelberger, R., Heinemann, C., Neher, E., and Matthews, G. (1994) Nature 371, 513-515[CrossRef][Medline] [Order article via Infotrieve]
  4. Augustine, G. J., and Neher, E. (1992) J. Physiol. 450, 247-271[Abstract]
  5. Thomas, P., Wong, J. G., Lee, A. K., and Almers, W. (1993) Neuron 11, 93-104[Medline] [Order article via Infotrieve]
  6. Bokvist, K., Eliasson, L., Ämmälä, C., Renström, E., and Rorsman, P. (1995) EMBO J. 14, 50-57[Abstract]
  7. Wollheim, C. B., Lang, J., and Regazzi, R. (1996) Diabetes Rev. 4, 276-296
  8. Chow, R. H., Klingauf, J., and Neher, E. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 12765-12769[Abstract/Free Full Text]
  9. Heinemann, C., Chow, R. H., Neher, E., and Zucker, R. S. (1994) Biophys. J. 17, 2546-2557
  10. Hutton, J. C., Penn, E. J., and Peshavaria, M. (1983) Biochem. J. 210, 297-305[Medline] [Order article via Infotrieve]
  11. Nicaise, G., Maggio, K., Thirion, S., Horoyan, M., and Keicher, E. (1992) Biol. Cell 75, 89-99[Medline] [Order article via Infotrieve]
  12. Thirion, S., Stuenkel, E. L., and Nicaise, G. (1995) Neuroscience 64, 125-137[CrossRef][Medline] [Order article via Infotrieve]
  13. Grohovaz, F., Bossi, M., Pezzati, R., Meldolesi, J., and Torri Tarelli, F. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 4799-4803[Abstract/Free Full Text]
  14. Austin, C. D., and Shields, D. (1996) J. Biol. Chem. 271, 1194-1199[Abstract/Free Full Text]
  15. Lang, I. M., and Schleef, R. R. (1996) J. Biol. Chem. 271, 2754-2761[Abstract/Free Full Text]
  16. Blondel, O., Moody, M. M., Depaoli, A. M., Sharp, A. H., Ross, C. A., Swift, H., and Bell, G. I. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 7777-7781[Abstract]
  17. Petersen, O. H. (1996) Trends Neurosci. 19, 411-413[CrossRef][Medline] [Order article via Infotrieve]
  18. Ravazzola, M., Halban, P. A., and Orci, L. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 2745-2748[Abstract/Free Full Text]
  19. Yule, D. I., Ernst, S. A., Ohnishi, H., and Wojcikiewicz, R. J. H. (1997) J. Biol. Chem. 272, 9093-9098[Abstract/Free Full Text]
  20. Chilcoat, N. D., Melia, S. M., Haddad, A., and Turkewitz, A. P. (1996) J. Cell Biol. 135, 1775-1787[Abstract]
  21. Carnell, L., and Moore, H. P. (1994) J. Cell Biol. 127, 693-705[Abstract]
  22. Calakos, N., and Scheller, R. H. (1996) Physiol. Rev. 76, 1-30[Abstract/Free Full Text]
  23. Fasolato, C., Hoth, M., and Penner, R. (1993) J. Biol. Chem. 268, 20737-20740[Abstract/Free Full Text]
  24. Bastianutto, C., Clementi, E., Codazzi, F., Podini, P., De Giorgi, F., Rizzuto, R., Meldolesi, J., and Pozzan, T. (1995) J. Cell Biol. 130, 847-855[Abstract]
  25. Martinez, J. R., Willis, S., Puente, S., Wells, J., Helmke, R., and Zhang, G. H. (1996) Biochem. J. 320, 627-634[Medline] [Order article via Infotrieve]
  26. Pizzo, P., Fasolato, C., and Pozzan, T. (1997) J. Cell Biol. 136, 355-366[Abstract/Free Full Text]
  27. Asfari, M., Janjic, D., Meda, P., Li, G., Halban, P. A., and Wollheim, C. B. (1992) Endocrinology 130, 167-178[Abstract]
  28. Sekine, N., Cirulli, V., Regazzi, R., Brown, L. J., Gine, E., Tamarit-Rodriguez, J., Girotti, M., Marie, S., Macdonald, M. J., Wollheim, C. B., and Rutter, G. A. (1994) J. Biol. Chem. 269, 4895-4902[Abstract/Free Full Text]
  29. Sekine, N., Fasolato, C., Pralong, W. F., Theler, J. M., and Wollheim, C. B. (1997) Diabetes 46, 1424-1433[Abstract]
  30. Lindau, M., and Neher, E. (1988) Pfluegers Arch. Eur. J. Physiol 411, 137-146[Medline] [Order article via Infotrieve]
  31. Von Rüden, L., and Neher, E. (1993) Science 262, 1061-1065[Medline] [Order article via Infotrieve]
  32. Horrigan, F. T., and Bookman, R. J. (1993) Biophys. J. 64, 101 (abstr.)
  33. Innocenti, B., Pozzan, T., and Fasolato, C. (1996) J. Biol. Chem. 271, 8582-8587[Abstract/Free Full Text]
  34. Kennedy, E. D., Rizzuto, R., Theler, J. M., Pralong, W. F., Bastianutto, C., Pozzan, T., and Wollheim, C. B. (1996) J. Clin. Invest. 98, 2524-2538[Abstract/Free Full Text]
  35. Helmchen, F., Borst, J. G. G., and Sakmann, B. (1997) Biophys. J. 72, 1458-1471[Abstract]
  36. Renström, E., Eliasson, L., Bokvist, K., and Rorsman, P. (1996) J. Physiol. 494, 41-52[Abstract]
  37. Dean, P. M. (1973) Diabetologia 9, 115-119[Medline] [Order article via Infotrieve]
  38. Li, G., Rungger-Brandle, E., Just, I., Jonas, J-C., Aktories, K., and Wollheim, C. B. (1994) Mol. Biol. Cell 5, 1199-1213[Abstract]
  39. Martin, T. F. J. (1997) Trends Cell Biol. 7, 271-276[CrossRef]
  40. Takahashi, N., Kadowaki, T., Yazaki, Y., Miyashita, Y., and Kasai, H. (1997) J. Cell Biol. 138, 55-64[Abstract/Free Full Text]
  41. Thomas-Reetz, A. C., and De Camilli, P. (1994) FASEB J. 8, 209-216[Abstract/Free Full Text]


Copyright © 1998 by The American Society for Biochemistry and Molecular Biology, Inc.