©1996 by The American Society for Biochemistry and Molecular Biology, Inc.
Regulation of the Inositol 1,4,5-Trisphosphate-activated Ca Channel by Activation of G Proteins (*)

(Received for publication, December 26, 1995; and in revised form, February 19, 1996)

Xin Xu Weizhong Zeng Shmuel Muallem (§)

From the Department of Physiology, University of Texas Southwestern Medical School, Dallas, Texas 75235

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Streptolysin O-permeable pancreatic acini were used to study the regulation of the inositol 1,4,5-trisphosphate (IP(3))-activated Ca channel (IPACC) by agonists and antagonists. Measurements of the apparent affinity for IP(3) (KIP(3)) showed that the IPACC is dynamically controlled during cell stimulation and inhibition, i.e. agonists decreased and antagonists increased KIP(3). KIP(3) was also independently regulated by thimerosal, Ca content of the stores, the incubation temperature, activation of protein kinases, and inhibition of protein phosphatases, but none of these mechanisms contributed to the regulation by agonists and antagonists. Incubating the cells with low concentration of GTPS or AlF(3) reproduced the effect of the agonist on KIP(3). Moreover, low [GTPS] allowed activation of the IPACC by agonists at basal levels of IP(3) and markedly impaired channel inactivation by antagonists. Channel sensitization by GTPS also restored the ability of thimerosal to mobilize Ca from internal stores with no change in cellular IP(3) levels. The combination of low [GTPS] and thimerosal locked the channel in an open, antagonist-insensitive state. All modulatory effects of GTPS are independent of phospholipase C activation and IP(3) production. We propose that the dynamic regulation of the IPACC by a G protein-dependent mechanism can play a major role in triggering and maintaining Ca oscillations at low agonist concentrations when minimal or no changes in IP(3) level take place.


INTRODUCTION

Ca mobilizing agonists stimulate the production of inositol 1,4,5-triphosphate (IP(3)), (^1)which releases Ca from intracellular stores located in the endoplasmic reticulum (ER)(1) . Stimulation with high agonist concentration leads to a persistent activation of the IP(3)-dependent Ca channel, which results in a single transient change in free cytosolic Ca concentration ([Ca])(2) . On the other hand, at low agonist concentrations usually oscillations in [Ca]are observed(1, 2) . Only partial correlation exists between the type of the [Ca] signal generated and IP(3) production. In most cases, high agonist concentration increases IP(3) to a supermaximal concentration, while at low agonist concentration it is difficult to demonstrate stimulated production of IP(3)(1, 2) . In addition, several agonists such as parathyroid hormone acting on osteoblasts (3, 4) or bradykinin acting on 3T6 fibroblasts (5) cause substantial Ca release from intracellular stores with no apparent increase in IP(3). However, basal levels of IP(3), although varying widely among different cell types, are almost always higher than that needed for maximal Ca release(1, 6, 7, 8, 9) .

A plausible explanation for Ca release in the absence of IP(3) production and in the face of high basal levels of IP(3) is that low [agonists] trigger Ca release by regulating the affinity of the IP(3)-activated Ca channel (IPACC) for IP(3). Studies in several cellular systems and the isolated and reconstituted IPACC showed the presence of multiple mechanism of channel regulation. Besides activation by IP(3), the channel can be regulated by Ca in a biphasic manner(10, 11, 12, 13) . ATP can regulate the channel directly (10, 11, 14) or by an indirect mechanism(13, 15) , which may involve phosphorylation/dephosphorylation reactions(13) . Recent elegant work by Cameron et al.(16) showed the intimate association of the cerebellar IPACC with calcineurin. The channel's affinity to IP(3) was regulated by the combined action of protein kinase C and calcineurin(16) . The same system was used to show that protein kinase A reduces the apparent affinity of the IPACC to IP(3)(14, 16) . In hepatocytes, protein kinase A increased the affinity for IP(3) in inducing Ca release(17, 18) , which was further augmented by inhibition of protein phosphatases with okadaic acid(18) . The affinity of the IPACC for IP(3) was also shown to be regulated by thiol-reactive agents such as thimerosal (TMS)(19, 20, 21, 22, 23, 24) and by temperature(25) . The relationship between all the different modes of regulating IP(3) affinity and Ca release are not known.

The above regulatory modes offer many mechanisms for the agonists to effect Ca release and initiate Ca oscillations without the need to increase IP(3) levels. Indeed, in rat liver, prestimulation with agonists was shown to modify the behavior and/or affinity for IP(3)(26) . More recently we showed that agonists can activate and antagonists inactivate the IPACC independent of IP(3) metabolism(27, 28) . How such a regulation is achieved, what mechanism the agonists use to modulate Ca release and the antagonists to terminate the release is not known.

In the present studies we used agonist/antagonist responsive, streptolysin O (SLO)-permeabilized pancreatic acini to demonstrate that in the same cells the IPACC can be regulated by multiple and independent mechanisms. More importantly, a G protein-dependent mechanism is used by agonists to activate the channel at basal levels of IP(3). Such activation dramatically impairs the ability of the antagonist in inactivating the IPACC. Once activated by the G protein-dependent mechanism, the channel can be locked in the open state by TMS. These findings point to the possible mechanism by which agonists can initiate Ca release and Ca oscillations with minimal changes in IP(3) concentrations.


EXPERIMENTAL PROCEDURES

Preparation of Pancreatic Acini

Acini were prepared from a rat pancreas using standard collagenase digestion(27) . In brief, the pancreas was removed, minced, and incubated for 5-6 min at 37 °C in a solution composed of (in mM) 140 NaCl, 5 KCl, 1 MgCl(2), 1 CaCl(2), 10 Hepes (pH 7.4 with NaOH), 10 glucose, 10 pyruvate, 0.1% bovine serum albumin, 0.02% soybean trypsin inhibitor (solution A) and supplemented with 165 µg/ml collagenase P. After digestion, the acini were collected and washed by centrifugation and kept in solution A on ice until use.

Measurement of Ca Uptake and Release in Permeabilized Cells

This procedure was identical with that described before(27, 28) . About 100 mg acini were washed twice with a solution containing 145 mM KCl and 10 mM Hepes (pH 7.4 with NaOH) and once with the same solution that was treated with Chelex 100. The acini were transferred to a fluorimeter cuvette containing a warm (37 °C) incubation medium composed of the Chelex-treated solution, 3 mM ATP, 5 mM MgCl(2), 10 mM creatine phosphate, 5 units/ml creatine kinase, 10 µM antimycin A, 10 µM oligomycin, 0.02% soybean trypsin inhibitor, 1 µM Fluo 3, and 0.4 unit/ml SLO. Fluo 3 fluorescence was recorded at an excitation wavelength of 488 nm and emission wavelength of 530 nm. Calibration of Fluo 3 signals was as described before (27, 28) using a K(d) of 370 nM at 37 °C. In several experiments, IP(3)-mediated Ca release was measured at 0 °C. For these experiments, the K(d) for Fluo 3 was measured to be 215 nM, using EGTA to buffer and set the different [Ca]. The K(d) values for EGTA at 0 °C were taken from Martell and Smith(29) .

Measurement of IP(3)

Acini were washed and incubated in permeabilization medium as for measurement of Ca uptake and release. After a 1.5-min incubation at 37 °C, if required, TMS or U73122 was added, and, after a 2.5-min incubation, the cells were stimulated with agonists. At the indicated times, 100-µl samples were transferred to 100 µl of solution containing 15% perchloric acid, mixed and kept on ice for about 20 min to allow complete protein precipitation. After a 2-min centrifugation at 10,000 times g, the supernatants were transferred to clean tubes. Standards of IP(3) were prepared in permeabilization medium and processed in a similar manner. Perchloric acid was precipitated, and IP(3) was extracted by the addition of 0.2 ml of tri-n-octylamine and 0.2 ml of freon. The samples were mixed vigorously, centrifuged for 30 s at 10,000 times g to separate the phases, and 15-25 µl of the upper layer was used for mass measurement of IP(3) by a standard radioligand binding assay(13) .


RESULTS

Regulation of K for IP(3)

The maintained intact signaling system in the SLO-permeabilized cells allowed examination of the acute effect of agonists and antagonists on the IP(3)-activated Ca channel (IPACC). Fig. 1shows the properties and demonstrates the advantages of this experimental system. Pancreatic acinar cells are fully permeabilized to small molecules within 10-15 s of exposure to the concentration of SLO used and reduced [Ca] of the incubation medium to the 50-80 nM range within 2 min of incubation at 37 °C. Addition of increasing concentrations of IP(3) resulted in discrete events of [Ca] increase, typical of the quantal behavior of Ca release (30) . Plotting the increments in medium [Ca] as a function of IP(3) concentration showed a saturation kinetic with an apparent affinity for IP(3) (KIP(3)) of 0.43 ± 0.02 µM (Fig. 1a, Table 1) and a Hill coefficient of 1.42 ± 0.026 (n = 33). Stimulating the cells with low concentrations of the muscarinic agonist, carbachol, caused small Ca release. Titration of Ca release in these cells showed that carbachol stimulation decreases the KIP(3) by about 2.53-fold to 0.17 µM (Fig. 1b, Table 1), without changing the Hill coefficient. On the other hand, stimulation by maximal [carbachol] and inactivation by the antagonist atropine was followed by a 1.42-fold increase in the KIP(3) to about 0.61 ± 0.02 µM (Fig. 1c, Table 1), again without an apparent change in the Hill coefficient. Thus, agonist stimulation increases whereas antagonist inhibition decreases the apparent affinity of the IPACC to its ligand IP(3). These changes in KIP(3) are independent of the total amount of Ca release since the release was measured at the same Ca buffering conditions.


Figure 1: Effect of agonist stimulation and antagonist inhibition on KIP(3). Pancreatic acini were allowed to reduce [Ca] of the permeabilization medium to about 60 nM (a) before stimulation with 10 µM (b) or 2 mM (c) carbachol. In experiment c, where indicated, the cells were inhibited with 0.1 mM atropine. After stabilization of medium [Ca] in a-c, the KIP(3) was measured by incremental additions of [IP(3)] between 0.05 and 2.55 µM. The numbers shown in trace a also apply for experiments b and c. The bottom panel shows the relationship between [IP(3)] and the increase in medium [Ca] above that measured before the additions of IP(3). The results of this and all other similar experiments are summarized in Table 1.





To study the possible mechanism by which the agonist and antagonist regulate the KIP(3), we characterized the effect of different compounds reported to affect IP(3)-mediated Ca release and the relationship between them. Examples of individual experiments are shown in Fig. 2and Fig. 3, and the combined results are summarized in Table 1. Treating the cells with relatively low concentrations of GTPS, which by itself caused no or minimal Ca release, decreased KIP(3) in a concentration- and time-dependent manner (Fig. 2b, Table 1). Incubation with 10 µM GTPS for 30 s increased the affinity for IP(3) by about 2.4-fold. Thimerosal (TMS) was shown to induce [Ca](i) oscillations in pancreatic acini (21) and increase the affinity for IP(3) in several cell types(19, 20, 21, 22, 23, 24) . Fig. 2c shows that unlike the case in intact cells, TMS (up to 500 µM) alone was unable to cause Ca release in permeable cells. However, as little as 100 µM TMS decreased KIP(3) by about 4-fold (Table 1). Fig. 2c and Table 1show the results with 100 µM TMS since this concentration was sufficient to maximally modify the effects of the agonist and GTPS on IPACC (see below).


Figure 2: Effect of GTPS, TMS, Ca load, and temperature on KIP(3). After completion of Ca uptake and stabilization of medium [Ca] at about 55 nM (a), the cells were treated with 5 µM GTPS (b) or 100 µM TMS (c). In experiment d, where indicated, two additions of 5 µM Ca to the incubation medium were made. In experiment e, the acini were first incubated for 2.5 min at 37 °C, then transferred to a thermostated cuvette and incubated for 3 min at 0 °C before the first addition of IP(3). The trace shows the last part of the incubation at 0 °C under control conditions. After each treatment, the KIP(3) was measured by incremental additions of IP(3). In experiments a-d, additions of IP(3) were as those shown in trace a. The numbers in trace e indicate the concentrations of IP(3) used for this titration. The results of all similar experiments are summarized in Table 1.




Figure 3: Additivity in the regulation of KIP(3). Experimental protocols were similar to those described in the legend to Fig. 2. In experiments a and b, the stores were overloaded by two additions of 5 µM CaCl(2). For clarity, the loading traces are shown only for experiment a. After stabilization of [Ca], the cells were treated with 5 µM GTPS (a) or 100 µM TMS (b) before additions of IP(3). The numbers in trace a indicate the concentrations of IP(3) used for experiments a and b. In experiments c and d, the acini were incubated for 2 or 1.5 min at 37 °C before addition of 5 µM GTPS (c, GTPS-treated) or 100 µM TMS (d, TMS-treated). After an additional 0.5- or 1-min incubation at 37 °C, respectively, the cells were cooled and KIP(3) was measured by the addition of the IP(3) concentrations indicated by the numbers in trace c. The results of all similar experiments are summarized in Table 1.



The effect of Ca content in the IP(3)-mobilizable Ca pool on IPACC has been studied extensively in several cell types(31, 32, 33, 34, 35, 36, 37, 38) . Fig. 2d shows that overloading the IP(3)-sensitive pool with Ca caused small, but significant increase in the affinity to IP(3) (see also Table 1). Similar KIP(3) was measured after one (not shown), two (Fig. 2d), and four pulses of 5 µM Ca (not shown). Finally, the temperature of the incubation medium had a profound effect on KIP(3) but without changing the quantal nature of Ca release. Thus, Fig. 2e shows that after 2.5 min of cell permeabilization and Ca loading at 37 °C and a subsequent 3-min incubation at 0 °C, increasing concentrations of IP(3) induce discrete and finite Ca release events. The release was much slower than that at 37 °C, which allowed a clear resolution of each Ca release event. In separate experiments, Ca release at submaximal [IP(3)] was followed up to 15 min with no sign of deviation from quantal behavior (not shown). Table 1shows that at 0 °C KIP(3) was reduced by about 8-fold.

An important question concerning the various modes of regulation of KIP(3) was the relationship between them to determine whether they regulate the IPACC by the same or by independent mechanisms. Fig. 3and Table 1show the additive and independent effect of the different modes. Thus, overloading the Ca pool did not prevent or reduce the effect of GTPS (Fig. 3a), TMS (Fig. 3b), or low concentration of carbachol (Table 1) on the KIP(3). Similarly, the effect of low temperature on KIP(3) was additive with the effect of GTPS (Fig. 3c), TMS (Fig. 3d), and carbachol (Table 1). Interestingly, addition of all agents after cooling to 0 °C had no effect on KIP(3). The cells had to be treated with carbachol, GTPS, or TMS at 37 °C before cooling to observe their effect on KIP(3), indicating activation of biochemical pathways by all agents, including TMS, to regulate KIP(3).

Several previous studies reported regulation of KIP(3) by activation of protein kinases(13, 14, 16, 17, 18) . To evaluate the role of protein kinases in the SLO-permeabilized system, we tested the effect of several protein kinase activators (12-O-tetradecanoylphorbol 13-acetate, cAMP, cGMP), protein kinase inhibitors (genistein, H7), and phosphatase inhibitors (KT62, okadaic acid, cyclosporine) on KIP(3). None of these agents affected KIP(3) in unstimulated SLO-permeabilized cells. Further, these agents did not prevent or augment the effect of TMS, Ca load, or low temperature (not shown). Surprisingly, all kinase activators and phosphatase inhibitors tested reduced or prevented the effect of GTPS and carbachol, whereas the kinase inhibitors augmented the effect of the agonists. Table 1lists the effect of the most effective compounds, okadaic acid and H7. Treatment with 0.12 µM okadaic acid largely prevented the effect of 30 s of treatment with 10 µM GTPS. H7 at 0.5 mM, a concentration sufficient to inhibit most protein kinases(38) , had no effect in unstimulated cells but it increased the effect of submaximal [GTPS]. The lack of selectivity in the effect of kinase and phosphatase activators/inhibitors raises questions as to the physiological significance of these findings. However, their value for the present studies is in showing the similarity between their effects on GTPS and carbachol, which were different from those of TMS, Ca load, and low temperature.

Regulation of KIP(3) Is Independent of PLC Activation

At this stage of the study, the possibility arose that the effects of carbachol and GTPS on KIP(3) were simply due to partial stimulation of PLC and a persistent, small global or local increase in [IP(3)]. Three types of experiments were performed to exclude this possibility. The first is shown in Fig. 4, a-c, in which U73122 was used to inhibit PLC. U73122 did not reduce the effect of either carbachol (Fig. 4b), GTPS (Fig. 4c), or TMS (not shown) on KIP(3), although it inhibited the small Ca release evoked by low concentrations of carbachol. Table 2shows that stimulation with 10 µM carbachol had a small effect on IP(3), whereas U73122 slightly reduced the basal level of IP(3). In the presence of U73122, IP(3) level was not changed by carbachol or GTPS. The second evidence is provided by the finding that prestimulation with low [GTPS] had a marked effect on KIP(3) with minimal or no effect on IP(3) levels, Ca release, or the size of the IP(3)-mobilizable Ca pool.


Figure 4: Effect of carbachol and GTPS on KIP(3) is independent of PLC. Acini incubated at 37 °C were exposed to 10 µM U73122 (a-c) or 25 µg/ml heparin (d and e) before stimulation with 10 µM carbachol (b) or 5 µM GTPS (c and e). a and d are the respective controls. Next, KIP(3) was measured by incremental additions of the indicated [IP(3)]. Note the high [IP(3)] used in experiments d and e.





The third and most convincing evidence against stimulation of PLC by carbachol or GTPS as the cause of the change in KIP(3) is shown in Fig. 4, d and e. In these experiments, 25 µg/ml IP(3) competitive inhibitor heparin (39, 40) were added to the incubation medium to increase the KIP(3) under control conditions from 0.43 µM (Table 1) to 1.67 µM (Fig. 4d). This should dilute the effect of any IP(3) generated by GTPS or carbachol stimulation by about 4-fold (1.67/0.43) to virtually eliminate the effect of the agents on KIP(3). Fig. 4e shows that this is not the case. Pretreatment with GTPS in the presence of 25 µg/ml heparin had the same effect on KIP(3) as in control cells. Similar results were obtained with heparin concentrations between 5 and 50 µg/ml and with cells stimulated with 10 µM carbachol (not shown). Hence, together the three protocols indicate that carbachol and GTPS modified KIP(3) independent of PLC stimulations.

G Protein-dependent Regulation of IPACC

The finding that GTPS can modulate KIP(3) independent of PLC activation suggested that activation of a G protein(s) by GTPS was sufficient to modulate interaction of IPACC with its ligand. Further support for this notion was obtained by testing the effect of AlF(3), which activates mainly heterotrimeric but not small G proteins(41) . All the effects of GTPS presented above and in subsequent figures could be initiated by 1 mM NaF + 0.2 µM AlCl(3) (not shown). 10 mM NaF + 2 µM AlCl(3) induced strong Ca release comparable with that observed with 100 µM GTPS (see below) or 2 mM carbachol.

More dramatic evidence for an effect of G protein(s) activation on the activity of the IPACC was obtained when the effect of a low concentration of GTPS on the response to carbachol, atropine, and TMS was studied. Fig. 5a shows that exposure of control or stimulated cells to 100 µM TMS in the absence of GTPS augmented the effect of low concentrations of carbachol on Ca release (Fig. 5, b and c), without affecting IP(3) levels during the first 20 s of cell stimulation (Table 2). Performing complete concentration dependence of both IP(3) production and Ca release showed that pretreatment with 100 µM TMS had no effect on IP(3) production while increasing the apparent affinity for carbachol-mediated Ca release by about 6.3-fold. On the other hand, GTPS profoundly modified the effect of carbachol. Treating the cells with as little as 2 µM GTPS for 30 s was sufficient to cause maximal Ca release by 10 µM carbachol (Fig. 5d).


Figure 5: Effect of TMS and GTPS on carbachol-triggered Ca release. Control acini (a and b) and acini treated with 100 µM TMS (c) or 2 µM GTPS (d) were stimulated with 2 mM (a) or 10 µM (c and d) carbachol. The cells in b were exposed to TMS after stimulation with 10 µM carbachol.



That the effect of GTPS was independent of PLC activation became even more evident when the effect of 2 µM GTPS on the dose response to carbachol and atropine was measured. Fig. 6shows that under control conditions the concentration dependence curves for carbachol stimulation of IP(3) production and Ca release were identical. Half-maximal stimulation (EC) of Ca release was at 210 ± 13 µM (n = 8), and the EC for IP(3) production was 236 ± 27 µM (n = 4). In the presence of 2 µM GTPS, the EC for IP(3) production was reduced by about 2.9-fold to 82 ± 11 µM (n = 4), whereas the EC for Ca release was reduced by 90-fold to 2.35 ± 0.13 µM (n = 8).


Figure 6: Effect of GTPS on carbachol-induced IP(3) production and Ca release. For measurement of Ca release (circle, bullet), the protocols shown in Fig. 5, a and d, were used, except that the cells were stimulated with different [carbachol] and in the presence (bullet) or absence (circle) of 2 µM GTPS. Resting [Ca] was subtracted from the peak increase in [Ca] at each carbachol concentration. For measurement of IP(3) levels (up triangle, ), cells were incubated for 2.5 min at 37 °C in permeabilization medium and then stimulated for 20 s with the indicated concentration of carbachol and in the presence () or absence (up triangle) of 2 µM GTPS. After 20 s of stimulation, the reactions were stopped with perchloric acid, and the levels of IP(3) were evaluated as described under ``Experimental Procedures.''



The effect of low [GTPS] on signal termination by atropine is shown in Fig. 7. In the absence of GTPS, atropine inhibited the IPACC of cells stimulated with 2 mM carbachol (measured from the rate of [Ca] reduction in carbachol-stimulated cells, see Fig. 1and (27) ) with an IC of 0.33 ± 0.02 µM (n = 5). Under the same conditions, atropine accelerated IP(3) hydrolysis (measured from the rate of IP(3) reduction relative to continuously stimulated cells as in Fig. 11below and (27) ) with an IC of 0.21 ± 0.03 µM (n = 3). Including 2 µM GTPS in the incubation medium shifted the IC for IP(3) production 4-fold, to 0.83 ± 0.09 µM (n = 3), while the IC for inhibition of Ca release was increased about 2600-fold to 860 ± 65 µM (n = 5). The fact that atropine inhibited PLC activation in the presence of GTPS clearly shows that the effect of GTPS was independent of PLC activation, since binding of nonhydrolyzable GTP analogues to the alpha subunit of G proteins, including G(41, 42) and G(43) , irreversibly stimulates the alpha subunits and prevents inhibition by antagonists.


Figure 7: Effect of GTPS on atropine-dependent channel inactivation and IP(3) hydrolysis. Inactivation of Ca uptake was measured from the initial rate of reduction in [Ca] measured after addition of atropine (see Fig. 1and Fig. 8for examples) to acini stimulated with 2 mM carbachol (bullet) or 2 mM carbachol and 2 µM GTPS (circle). The rationale and procedure for calculation of the rate of reduction in [Ca] are detailed in (27) . For measurement of rates of IP(3) hydrolysis, acini incubated in permeabilization medium as described in the legend to Fig. 6were stimulated with 2 mM carbachol () or 2 mM carbachol and 2 µM GTPS (up triangle). After 20 s of stimulation, the cells were inhibited with the indicated concentration of atropine, and the levels of IP(3) were followed for the subsequent 3 min (see also Fig. 11).




Figure 11: TMS accelerates atropine-induced IP(3) hydrolysis. Acini incubated in permeabilization medium for 2.5 min at 37 °C () were stimulated with 100 µM carbachol and 2 µM GTPS (bullet, circle) and in the presence of 100 µM TMS (circle). After 20 s of stimulation, a portion of cells stimulated in the absence () or the presence of TMS (box) were transferred to vials containing atropine to give a final concentration of 1 mM. At the indicated times, samples were removed to assay the levels of cellular IP(3). This experiment represents 2 others with similar results. The same behavior was found in two experiments where 10 µM carbachol was used. In the absence of GTPS, TMS augmented IP(3) hydrolysis even more than that shown in the figure (three independent experiments).




Figure 8: The antagonist inhibits the effect of GTPS but not of TMS on KIP(3). In all experiments (a-c), the acini were stimulated with 2 mM carbachol and then inhibited with 0.1 mM atropine. After carbachol and atropine treatment, the acini were treated with 10 µM GTPS (b) or 100 µM TMS (c) before addition of IP(3) at the concentrations indicated by the numbers next to each trace. The results of this and all similar experiments are summarized in Table 1.



Stabilization of IPACC in an Active State

In previous studies, we reported that antagonists inactivate the IPACC independent of IP(3) metabolism(27, 28) . In Fig. 1and Table 1, we showed that termination of carbachol stimulation with atropine significantly increased KIP(3). To better understand the mechanism of channel inactivation, we first tested the effect of atropine on the various mechanisms shown to affect KIP(3). Fig. 8illustrates the effect of carbachol stimulation and atropine inhibition of the ability of GTPS and TMS to modify KIP(3). Similar experiments were performed to test the effect of Ca load and low temperature. Carbachol stimulation followed by atropine inhibition did not prevent the reduction in KIP(3) caused by TMS (Fig. 8c), Ca load, or low temperature (not shown), while completely preventing the effect of GTPS (Fig. 8b). Incubating the cells with atropine alone without carbachol stimulation had no effect on the ability of GTPS to reduce KIP(3) (not shown). Table 1shows that after carbachol and atropine treatment the KIP(3) was 0.61 µM. It remained the same (0.62 µM) after a 1.5-min incubation with 10 µM GTPS. This value should be compared to the KIP(3) of 0.18 µM measured in control cells incubated with 10 µM GTPS for 30 s (Table 1). These experiments indicate that, in permeabilized cells, once the IPACC was inactivated by atropine, the inactivation could not be relieved by activation of G proteins.

Because the effect of carbachol was modified by both GTPS and TMS, to understand how they may modify channel activity we tested the effect of TMS on the modulation of IPACC by GTPS. Interestingly, GTPS markedly sensitized the effect of TMS to cause maximal discharge of the Ca stores. This is illustrated in Fig. 9. Fig. 9a shows that addition of 100 µM TMS to cells treated with 5 µM GTPS induced rapid and maximal Ca release. Similar results were obtained when GTPS was added to TMS-treated cells, but the time course of Ca release was significantly slower. Accordingly, as all other effects of GTPS, the effect shown in Fig. 9was time- (not shown) and concentration-dependent (Fig. 9c). In the absence of TMS, high concentrations of GTPS could release Ca, which was probably due to stimulation of PLC to generate IP(3). TMS actually partially inhibited the production of IP(3) generated by all concentrations of GTPS while sensitizing activation of Ca release by GTPS. It is important to note that despite the absence of an increase in IP(3) the effect of TMS and GTPS on Ca release was still inhibited by heparin (Fig. 9c). This would suggest that TMS + GTPS sensitized the IPACC to a level that maximal and rapid Ca release was observed at the level of IP(3) present in unstimulated cells.


Figure 9: GTPS together with TMS induces maximal Ca release. Experiment a in the upper panel shows that 100 µM TMS causes rapid and maximal Ca release from acini treated with 5 µM GTPS for 30 s. This Ca release can be inhibited largely by 50 µg/ml heparin (b). Experiment c in the lower panel shows the dependence of Ca release (circle, bullet) and IP(3) production (up triangle, ) on GTPS concentration in the absence (circle, up triangle) or presence (bullet, ) of 100 µM TMS. For Ca release, the protocol in experiment a was used except that the cells were incubated with or without 100 µM TMS for 1 min before addition of GTPS. IP(3) production was measured as described in the legend to Fig. 6except that the cells were treated with or without 100 µM TMS for 1 min before being stimulated with the indicated concentration of GTPS for 1 min.



In the next stage, we tested the effect of channel sensitization by GTPS and TMS on the inactivation induced by atropine. Fig. 10a shows that 1 mM atropine completely inactivated channels activated by 10 µM carbachol and 2 µM GTPS. However, treating the cells with 100 µM TMS prior to stimulation with carbachol and GTPS completely prevented channel inactivation by atropine. The small reduction in medium [Ca] due to atropine probably reflects the activity of channels that were not accessed by GTPS. The channels were permanently stabilized in an active state since medium [Ca] remained elevated for at least 15 min with no sign of decline. To show that TMS and GTPS did not inhibit the SERCA pumps and that the maintained high level of medium [Ca] was due to stabilizing the IPACC in an active state, the channel was inhibited by heparin. Addition of heparin resulted in channel inhibition and, consequently, rapid Ca uptake into the IP-sensitive pool at a rate comparable to that measured in Fig. 10a after addition of atropine.


Figure 10: TMS stabilizes the IPACC in an open state. Acini incubated in permeabilization medium were treated with (b, d) or without (a, c) 100 µM TMS before stimulation with 10 µM carbachol and 2 µM GTPS (a, b) or 100 µM GTPS (c, d). Where indicated, all cells were inhibited with 1 mM atropine. In experiments (b, d) after atropine inhibition, the cells were exposed to 100 µg/ml heparin. Results similar to those in experiment d were obtained with 2 and 5 µM GTPS. The experiment shown and the one using 5 µM GTPS were repeated at least seven times with different cell preparations.



As indicated above, TMS alone (not shown) or GTPS alone (Fig. 10a and Fig. 7), although reducing the affinity for atropine, never prevented channel inactivation by atropine. That activation of G proteins by GTPS and channel sensitization by TMS was required to prevent channel inactivation is further emphasized in the experiments shown in Fig. 10, c and d. As we showed before(28) , stimulation of Ca release with 100 µM GTPS did not prevent channel inactivation by atropine (Fig. 10c), even though IP(3) levels under these conditions were very high(28) . On the other hand, treating the cells with 2 µM (not shown) or 100 µM GTPS (Fig. 10d) and 100 µM TMS in the absence of agonist stimulation was sufficient to prevent channel inactivation by atropine.

An important control for the experiments in Fig. 10is to show that TMS did not prevent the hydrolysis of IP(3) initiated by atropine. The results of such experiments are shown in Fig. 11. Even in the presence of 100 µM carbachol and 5 µM GTPS, TMS accelerated, rather than inhibited, the hydrolysis of IP(3). After 2 min of exposure to atropine, IP(3) was reduced to basal levels, while the channel was fully activated (Fig. 10b).


DISCUSSION

A long standing question in understanding agonist-evoked [Ca](i) oscillations is how low concentrations of agonists induce oscillations without an apparent or only small change in [IP(3)](1) . Modulation of the IP(3)-activated Ca channel (IPACC) can be potentially important in view of the high basal IP(3) levels in most cells(1, 2, 6, 7, 8, 9) . Although several regulatory mechanisms of IPACC have been reported (10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26) , the relationship between them and their role in agonist-dependent regulation of Ca release is not known. The present study suggests the existence of multiple and independent mechanisms for regulation of IPACC, prominent among which is regulation by G protein activation. The latter is used by agonists and antagonists to modulate the KIP(3) in a reciprocal manner and thus facilitate Ca release during agonist stimulation and impair the release during antagonist inhibition. Such a mechanism can contribute to the cyclical activation and inactivation of the IPACC during Ca oscillations(44) . Below we discuss the evidence in support of these findings.

Comparing the effect of many agents and conditions in the same cell type and experimental system showed that KIP(3) can be modulated by several independent mechanisms (Table 1). Thus, additive effects were found between TMS, Ca load, low temperature, and the agonists. The important implication of these findings is that although the IPACC can be regulated by various mechanisms, none of them appear to contribute to the regulation by agonists. Of course, regulation by Ca load can have important physiological significance in that when the stores are loaded, they are primed for release by small additional change in KIP(3). It is likely that in empty stores the IPACC has the lowest affinity for IP(3), which will facilitate channel inactivation at the termination of cell stimulation. Nonetheless, the regulation of KIP(3) by Ca content in the ER is relatively modest and occurs by a mechanism different from that used by agonists.

Interestingly, despite the fact that variations in KIP(3) between compartmentalized Ca pools account in large part for quantal Ca release(28) , none of the modulators of KIP(3) changed the quantal properties of Ca release. This includes low temperature ( Fig. 2and Fig. 3). We particularly studied the effect of low temperature in detail since a previous report suggested that quantal Ca release becomes continuous at low temperature, and this process is reversible(25) . However, we failed to convert the quantal to a continuous Ca release by short or long incubation at 25 or 0 °C or by any other modulator of KIP(3). In the earlier studies by Kindman and Meyer(25) , one concentration of IP(3) was used to demonstrate submaximal Ca release at 37 °C and maximal release at 0 °C, without considering the effect of the temperature on KIP(3). Such an effect as demonstrated in the present study (Table 1) can account well for the differences between the two studies. Our results of maintained quantal behavior under all conditions suggest that all modulators of KIP(3), including agonists, affect all IPACC equally rather than equalize KIP(3) of channels of different compartments.

The present studies show that KIP(3) is dynamically controlled. Low concentrations of agonists, which minimally activate PLC, markedly reduced KIP(3) of the IPACC. Moreover, termination of cell stimulation with antagonists increased KIP(3) to a level above that measured in control cells. The antagonist was effective only if the cells were first stimulated with carbachol. The antagonist had no effect in control cells, and, when added before GTPS, it did not prevent the GTPS-dependent reduction in KIP(3). The combined effects of the agonist and antagonist indicate that the KIP(3) of the IPACC is dynamically controlled during cell stimulation/inhibition cycles. One advantage of such a dynamic control is that small changes in IP(3) levels can lead to maximal opening or closing of the channel. This will be translated into a high cooperativity for interaction of IP(3) with the IPACC and in channel activation/inactivation.

Probably the most interesting finding of the present studies is that agonists and antagonists appear to modulate KIP(3) by a mechanism dependent on activation of G proteins. The first indication of this was the similarity between the effect of the agonist and preincubation with GTPS on KIP(3). Both affected KIP(3) in cells treated with TMS, high Ca load or incubated at low temperature. Both effects were inhibited similarly by activators of protein kinases, inhibitors of protein phosphatases, and augmented by inhibitors of protein kinases. Inhibition of agonist-activated cells by atropine to increase KIP(3) inhibited the effect of GTPS, but not of any other agent or treatment. Together, these observations strongly suggest that agonist stimulation and GTPS reduced the KIP(3) by the same mechanism. It is possible that the agonists and GTPS activated heterotrimeric, rather than small G proteins, since all effects of GTPS could be reproduced with low concentrations of AlF(3). The effect of GTPS described in the present study is different from the previously described modification of the size of the IP(3)-sensitive Ca pool by GTP(45, 46, 47) . GTPS inhibited the GTP-induced expansion of the Ca pool, whereas GTPS increased the KIP(3) and millimolar concentrations of GTP were required to mimic the effect of low concentrations of GTPS and AlF(3).

That G proteins are involved in the effect of the agonist/antagonist and their full impact becomes more evident when the effect of their activation on agonist/antagonist-dependent changes in Ca release and IP(3) levels are considered. Even in the absence of preincubation, low concentrations of GTPS (0.2-2 µM) markedly increased the potency of the agonist and decreased the potency of the antagonist in affecting Ca release. Thus, when irreversible activation of G proteins was allowed by the presence of GTPS, the agonists exceedingly sensitized the IPACC to trigger Ca release at basal [IP(3)]. Measurement of IP(3) levels showed that GTPS had a small effect on IP(3) production during agonist stimulation and did not prevent initiation of IP(3) hydrolysis by the antagonist. The latter excludes the possibility that the effects of GTPS, or for that matter the agonist and the antagonist, on Ca release was dependent on PLC activity (41, 42, 43) . In fact, in cells stimulated with carbachol and GTPS, 10-100 µM atropine completely inactivated PLC without having any inactivation effect on the IPACC (Fig. 7). These experiments, therefore, indicate that agonists use G protein activation to reduce KIP(3) and stabilize the channel in an open state. Once the channel is activated, complete inactivation of G proteins by the antagonist beyond (or different from?) that required to modulate PLC activity, is needed for channel inactivation. Indeed, preliminary experiments showed that muscarinic, bombesin, and cholecystokinin antagonists were between 100- and 1000-fold more effective in preventing cell stimulation when added before the respective agonist than in reversing agonist effects when added to stimulated cells. (^2)

Further insight into the regulation of IPACC by G proteins was obtained when the effect of TMS on GTPS-, agonist-, and antagonist-dependent Ca release was studied. TMS and low GTPS caused maximal Ca release at basal IP(3) concentration. It is interesting that in intact cells 100 µM TMS caused significant Ca release (21 and data not shown), whereas in permeabilized cells up to 500 µM TMS did not release Ca but only decreased KIP(3). Incubating the cells with as little as 0.2 µM GTPS was sufficient to restore the ability of TMS to cause Ca release with no change in IP(3) levels. Together, these observations indicate that: (a) TMS and GTPS modulate the IPACC by different mechanisms, (b) the IPACC is regulated by G proteins in permeabilized and probably in intact cells, and (c) TMS appears to modulate the interaction of the G protein-dependent mechanism with the IPACC to stabilize the channel in an open state. Indeed, treating the cells with TMS and GTPS, with or without agonist, completely prevents channel inactivation by the antagonist. This effect was absolutely dependent on the combined action of TMS and GTPS to the extent that incubating the cells with very high GTPS (100 µM) did not prevent the effect of atropine, while stimulation with 2 µM GTPS and 100 µM TMS locked the channel in an antagonist-insensitive state. It is important to note that TMS increased, rather than decreased, the rate of IP(3) hydrolysis under all conditions, including incubation with GTPS.

The implication of the studies with TMS, GTPS, agonist, and antagonist is that all point to the involvement of G proteins in the regulation of KIP(3) by agonist stimulation and antagonist inhibition. They also point to the broad extent of such regulation. Such a regulatory mechanism can be very attractive in explaining how agonists can stimulate Ca oscillations in the absence of stimulated IP(3) production. Our results suggest that minimal activation of G proteins (low GTPS, AlF(3), or agonist concentrations), which is not sufficient to appreciably activate PLC, is sufficient to markedly decrease the KIP(3) and trigger large Ca release at basal [IP(3)]. This regulation is a dynamic process as revealed by the antagonist-induced increase in KIP(3) beyond that found in resting cells. It is well established in many cell systems that Ca oscillations occur only at low agonist concentration, when usually no change in IP(3) is observed. It is easy to see how the dynamic regulation of KIP(3) described here can contribute or even dominate the mechanism by which agonists signal Ca oscillations. Understanding how activation of G proteins modulate KIP(3) is essential to evaluate its role in Ca oscillation in particular and Ca signaling in general. This should be the challenge for future studies.


FOOTNOTES

*
This work was supported by National Institutes of Health Grants DK 38938 and DK 46591. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Dept. of Physiology, University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd., Dallas, TX 75235. Tel.: 214-648-2593; Fax: 214-648-8685; smuall{at}mednet.swmed.edu.

(^1)
The abbreviations used are: IP(3), inositol, 1,4,5-trisphosphate; ER, endoplasmic reticulum; IPACC, IP(3)-activated Ca channel; TMS, thimerosal; SLO, streptolysin O; KIP(3), apparent affinity for IP(3); H7, (1-(5-isoquinolinesulfonyl)-2 methylpiperazine dihydrochloride; PLC, phospholipase C; GTPS, guanosine 5`-O-(thiotriphosphate).

(^2)
X. Xu and S. Muallem, unpublished observations.


ACKNOWLEDGEMENTS

We thank Mary Vaughn for expert administrative support.


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