Slow Feedback Inhibition of Calcium Release-activated Calcium Current by Calcium Entry
1998; Elsevier BV; Volume: 273; Issue: 24 Linguagem: Inglês
10.1074/jbc.273.24.14925
ISSN1083-351X
Autores Tópico(s)Nicotinic Acetylcholine Receptors Study
ResumoIn many nonexcitable cells, depletion of the inositol 1,4,5-trisphosphate-sensitive store activates Ca2+ influx, a process termed store-operated Ca2+ entry. In rat basophilic leukemia cells, emptying of the stores activates a highly selective Ca2+release-activated Ca2+ current (CRAC),I CRAC. We have recently found thatI CRAC activates in an essentially all-or-none manner when the current is evoked by receptor stimulation, dialysis with inositol 1,4,5-trisphosphate via the patch pipette, or through the Ca2+ATPase inhibitor thapsigargin (Parekh, A. B., Fleig, A., and Penner, R. (1997) Cell 89, 973–980). Regulatory mechanisms must therefore operate to control the overall amount of Ca2+ that enters through CRAC channels. Such mechanisms include membrane potential and protein kinase C. In the present study, we have investigated additional inhibitory pathways that serve to determine just how much Ca2+ can enter throughI CRAC. We have directly measured the current using the whole cell patch clamp technique. We report the presence of a slow Ca2+-dependent inactivation mechanism that curtails Ca2+ entry through CRAC channels. This inactivation mechanism is switched on by Ca2+ entering through CRAC channels, and therefore constitutes a slow negative feedback process. Although it requires a rise in intracellular Ca2+ for activation, it maintains CRAC channels inactive even under conditions that lower intracellular Ca2+ levels. The inactivation mechanism does not involve store refilling, protein phosphorylation, G proteins, nor Ca2+-dependent enzymes. It accounts for up to 70% of the total inactivation ofI CRAC, and therefore appears to be a dominant inhibitory mechanism. It is likely to be an important factor that shapes the profile of the Ca2+ signal in these nonexcitable cells. In many nonexcitable cells, depletion of the inositol 1,4,5-trisphosphate-sensitive store activates Ca2+ influx, a process termed store-operated Ca2+ entry. In rat basophilic leukemia cells, emptying of the stores activates a highly selective Ca2+release-activated Ca2+ current (CRAC),I CRAC. We have recently found thatI CRAC activates in an essentially all-or-none manner when the current is evoked by receptor stimulation, dialysis with inositol 1,4,5-trisphosphate via the patch pipette, or through the Ca2+ATPase inhibitor thapsigargin (Parekh, A. B., Fleig, A., and Penner, R. (1997) Cell 89, 973–980). Regulatory mechanisms must therefore operate to control the overall amount of Ca2+ that enters through CRAC channels. Such mechanisms include membrane potential and protein kinase C. In the present study, we have investigated additional inhibitory pathways that serve to determine just how much Ca2+ can enter throughI CRAC. We have directly measured the current using the whole cell patch clamp technique. We report the presence of a slow Ca2+-dependent inactivation mechanism that curtails Ca2+ entry through CRAC channels. This inactivation mechanism is switched on by Ca2+ entering through CRAC channels, and therefore constitutes a slow negative feedback process. Although it requires a rise in intracellular Ca2+ for activation, it maintains CRAC channels inactive even under conditions that lower intracellular Ca2+ levels. The inactivation mechanism does not involve store refilling, protein phosphorylation, G proteins, nor Ca2+-dependent enzymes. It accounts for up to 70% of the total inactivation ofI CRAC, and therefore appears to be a dominant inhibitory mechanism. It is likely to be an important factor that shapes the profile of the Ca2+ signal in these nonexcitable cells. In many nonexcitable cells, depletion of the inositol 1,4,5-trisphosphate (InsP3) 1The abbreviations used are: InsP3, inositol 1,4,5-trisphosphate; CRAC, calcium release-activated calcium current; RBL, rat basophilic leukemia; BAPTA, 1,2-bis(2-aminophenoxy)ethane-N ,N ,N ′,N ′-tetraacetic acid; ATPγS, adenosine 5′-O -(thiotriphosphate); GDPβS, guanosine 5′-O -2-(thiodiphosphate); GTPγS, guanosine 5′-3-O - (thiotriphosphate). -sensitive intracellular Ca2+ stores activates a Ca2+influx pathway in the plasma membrane (1Berridge M.J. Nature. 1993; 361: 315-325Crossref PubMed Scopus (6188) Google Scholar). This mechanism was originally proposed by Putney (2Putney J.W. Cell Calcium. 1986; 7: 1-12Crossref PubMed Scopus (2115) Google Scholar) and called capacitative Ca2+ influx. Patch-clamp experiments have identified a variety of Ca2+-permeable channels in the plasma membrane that seem to underlie capacitative Ca2+ influx (reviewed in Ref. 3Parekh A.B. Penner R. Physiol. Rev. 1997; 77: 901-930Crossref PubMed Scopus (1294) Google Scholar). These channels differ in their biophysical properties and are generically referred to as store-operated Ca2+ channels (3Parekh A.B. Penner R. Physiol. Rev. 1997; 77: 901-930Crossref PubMed Scopus (1294) Google Scholar,4Clapham D.E. Cell. 1995; 80: 259-268Abstract Full Text PDF PubMed Scopus (2272) Google Scholar). Of the store-operated Ca2+ currents, the best characterized is I CRAC, which was originally discovered in mast cells (5Hoth M. Penner R. Nature. 1992; 355: 353-356Crossref PubMed Scopus (1495) Google Scholar). I CRAC has subsequently been shown to exist in several different nonexcitable cells including basophils, T cells and megakaryocytes (3Parekh A.B. Penner R. Physiol. Rev. 1997; 77: 901-930Crossref PubMed Scopus (1294) Google Scholar). CRAC channels are remarkably selective for Ca2+ ions and have a low single-channel conductance (3Parekh A.B. Penner R. Physiol. Rev. 1997; 77: 901-930Crossref PubMed Scopus (1294) Google Scholar, 5Hoth M. Penner R. Nature. 1992; 355: 353-356Crossref PubMed Scopus (1495) Google Scholar). Just how depletion of the stores activates CRAC channels is still unclear. Several potential mechanisms have been proposed but the signal has not been unequivocably identified (reviewed in Refs. 3Parekh A.B. Penner R. Physiol. Rev. 1997; 77: 901-930Crossref PubMed Scopus (1294) Google Scholar and 4Clapham D.E. Cell. 1995; 80: 259-268Abstract Full Text PDF PubMed Scopus (2272) Google Scholar). One interesting aspect of I CRAC is that the current activates in an essentially all-or-none manner, irrespective of whether activation is evoked by dialysis with inositol 1,4,5-trisphosphate, receptor stimulation, or thapsigargin (6Parekh A.B. Fleig A. Penner R. Cell. 1997; 89: 973-980Abstract Full Text Full Text PDF PubMed Scopus (205) Google Scholar). One consequence of this is that, if I CRAC activates in an all-or-none manner, mechanisms must exist that control the amount of Ca2+ entering the cell through CRAC channels. This is required in order to achieve graded activation of Ca2+-dependent processes like secretion that correlate with the level of cell stimulation by receptors. We have recently reported that Ca2+ entry through CRAC channels in rat basophilic leukemia (RBL) cells can be graded, despite all-or-none activation, because of several regulatory mechanisms that serve to control CRAC channel activity (6Parekh A.B. Fleig A. Penner R. Cell. 1997; 89: 973-980Abstract Full Text Full Text PDF PubMed Scopus (205) Google Scholar, 7Parekh A.B. Penner R. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 7907-7911Crossref PubMed Scopus (155) Google Scholar). One way is by changing the membrane potential. Hyperpolarization increases the electrical gradient for Ca2+ entry, thus favoring further Ca2+influx, whereas depolarization decreases the driving force and hence reduces Ca2+ entry (6Parekh A.B. Fleig A. Penner R. Cell. 1997; 89: 973-980Abstract Full Text Full Text PDF PubMed Scopus (205) Google Scholar). I CRAC is also regulated by protein kinase C. Stimulation of this enzyme inactivates I CRAC. Since protein kinase C will be activated by diacylglycerol, which is produced following stimulation of receptors that engage the phosphoinositide pathway, it constitutes an important negative feedback mechanism on Ca2+ influx (7Parekh A.B. Penner R. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 7907-7911Crossref PubMed Scopus (155) Google Scholar). In mast cells and jurkat T lymphocytes, I CRAC is subjected to a fast inactivation process operating on a milliseconds time scale. This arises from Ca2+ ions entering the cell through CRAC channels and then binding to sites probably located on the channels themselves (5Hoth M. Penner R. Nature. 1992; 355: 353-356Crossref PubMed Scopus (1495) Google Scholar, 8Zweifach A. Lewis R.S. J. Gen. Physiol. 1995; 105: 209-226Crossref PubMed Scopus (336) Google Scholar). Here we report an additional mechanism that serves to regulate CRAC channels. We find that Ca2+ influx through CRAC channels exerts a slow feedback inhibition that curtails further Ca2+ entry and which is dependent on a rise in intracellular Ca2+ levels. Slow inactivation accounts for up to 70% of the inhibition of I CRAC. Once activated, this slow inactivation mechanism can maintain CRAC channels in an inactivated state for several minutes, even after intracellular Ca2+ levels have been reduced. Hence slow inactivation appears to be a dominant inhibitory mechanism that determines the time course of Ca2+ influx following store depletion in RBL cells. It also provides an additional mechanism whereby the amount of Ca2+ entering through CRAC channels can be regulated despite all-or-none Ca2+ entry. Rat basophilic leukemia cells (RBL-2H3) cells were kindly supplied by Michael Pilot, Max Planck Institute for Biophysical Chemistry, Goettingen, Germany, and were cultured essentially as described previously (6Parekh A.B. Fleig A. Penner R. Cell. 1997; 89: 973-980Abstract Full Text Full Text PDF PubMed Scopus (205) Google Scholar, 7Parekh A.B. Penner R. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 7907-7911Crossref PubMed Scopus (155) Google Scholar). Patch-clamp experiments were conducted in the tight seal whole cell configuration at room temperature (18–25 °C) as described previously (6Parekh A.B. Fleig A. Penner R. Cell. 1997; 89: 973-980Abstract Full Text Full Text PDF PubMed Scopus (205) Google Scholar, 7Parekh A.B. Penner R. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 7907-7911Crossref PubMed Scopus (155) Google Scholar). Patch pipettes were pulled from borosilicate glass (Hilgenberg), sylgard-coated, and fire-polished. Pipettes had d.c. resistances of 2.5–4 megohms when filled with standard internal solution that contained (in mm): cesium glutamate 145, NaCl 8, MgCl2 1, MgATP 1, InsP3 0.03, EGTA 1.4, HEPES 10, pH 7.2, with CsOH. A correction of +10 mV was applied for the subsequent liquid junction potential. In some experiments, the EGTA concentration was raised to 14 mm (indicated in text) or substituted with BAPTA. Drugs were added to this internal solution as described in the text. Extracellular solution contained (in mm): NaCl 145, KCl 2.8, CaCl2 10, MgCl2 2, CsCl 10, glucose 10, HEPES 10, pH 7.2 (NaOH). CsCl was present to block the activity of the inwardly rectifying potassium channel. High resolution current recordings were acquired by a computer-based patch-clamp amplifier system (EPC-9, HEKA Electronics, Germany). Capacitative currents were canceled before each voltage ramp using the automatic compensation of the EPC-9. Series resistance was between 5 and 15 megohms. Currents were filtered using an 8-pole Bessel filter at 2.5 kHz and digitized at 100 μs. I CRAC was measured using either voltage ramps (−100 to +100 mV in 50 ms) or voltage steps (pulses to −80 mV for 200 ms) applied every 2 s using PULSE software (HEKA Electronics) on a 9500 PowerMac. Cells were held at 0 mV between pulses. All currents were leak subtracted by averaging the first two to four ramps/steps after breaking in and then subtracting this from all subsequent traces. Several parameters (capacitance, series resistance, holding current) were displayed simultaneously on a second monitor at a slower rate (2 Hz) using the X-Chart display (HEKA Electronics). Data are presented as mean ± S.E., and statistical evaluation was carried out using Student's unpaired t test. Previous work has demonstrated that I CRAC inactivates partially when RBL cells are dialyzed with a patch pipette solution containing a high concentration of the slow Ca2+ chelator EGTA (10 mm), and this is due to a kinase-mediated phosphorylation (7Parekh A.B. Penner R. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 7907-7911Crossref PubMed Scopus (155) Google Scholar). In the present study, we have examined the effects of more moderate calcium buffering on the properties ofI CRAC by including 1.4 mm EGTA in the recording pipette. Fig.1 A (i ) shows an experiment in which the internal solution contained 30 μmInsP3 (a supramaximal concentration) (6Parekh A.B. Fleig A. Penner R. Cell. 1997; 89: 973-980Abstract Full Text Full Text PDF PubMed Scopus (205) Google Scholar) and 1.4 mm EGTA. Cells were voltage clamped at 0 mV andI CRAC was monitored using voltage ramps applied every two seconds (shown in Fig. 1 A (ii )). The current was measured at −80 mV. Fig. 1 A (i ) depicts the time course of the current during the experiment. Following the onset of whole cell recording, I CRACactivated as InsP3 diffused into the cell from the recording pipette and depleted the stores. The time constant for activation (τ) was 15.2 ± 1.6 s, similar to our previous measurements (19.4 ± 1.2 s) (9Parekh A.B. Penner R. J. Physiol. 1995; 489: 377-382Crossref PubMed Scopus (38) Google Scholar). The current peaked after 50–80 s and then inactivated substantially with time. For the cell shown in Fig. 1 A (i ), the current had fallen by 70% (relative to the peak amplitude) after 300 s. Fig.1 A (ii ) shows I–V relationships for the two time points indicated in Fig. 1 A (i ) (a andb ). The current measured in the ramps exhibited the classic features of I CRAC: voltage-independent activation, inward rectification, and a reversal potential close to +60 mV. We also monitored activity of I CRAC by applying voltage steps to −80 mV for 200 ms every 2 s from a holding potential of 0 mV. In Fig. 1 B (i ), the time course of I CRAC is plotted using this protocol, and Fig. 1 B (ii ) shows the inward current measured during these hyperpolarizing pulses. During the step,I CRAC inactivated by up to 40% with a time constant in the range of 50 ms (Fig. 1 B (ii )). This fast inactivation has been described in detail in T cells and arises from local feedback inactivation by Ca2+ ions in the vicinity of the channel pore (8Zweifach A. Lewis R.S. J. Gen. Physiol. 1995; 105: 209-226Crossref PubMed Scopus (336) Google Scholar). We measured the steady-state current during the step (indicated by the arrow in 1B (ii )), and this amplitude is plotted against time in Fig. 1 B (i ). As with the ramp protocol employed in Fig. 1 A , I CRACinactivated substantially during the experiment. For the cell shown in Fig. 1 B , the current amplitude at 300 s had fallen by 68% relative to the peak value. Fig. 1 C plots the time course ofI CRAC measured using either the ramp or step methods (six cells each). These experiments were carried out such that ramps or steps were applied to alternate cells on the same coverslips (henceforth referred to as paired experiments). There was no significant difference between ramp or step protocols with respect to τactivation (15.2 ± 1.6 versus 17.7 ± 2.0 s, respectively) peak amplitude (−2.98 ± 0.24versus −2.58 ± 0.19 pA/pF) or in the extent of steady-state inactivation (fallen by 53.4 ± 8.9%versus 56.1 ± 9.3%, respectively, Fig.1 C ). This inactivation in the presence of moderate Ca2+buffering will henceforth be referred to as slow inactivation to distinguish it from the fast inactivation seen during the hyperpolarizing step. In some cells, slow inactivation was very prominent and contributed to an almost complete inactivation ofI CRAC. Typical examples of this type of response are shown in Figs. 3 B and 5 A . In some other cells, slow inactivation contributed to around a 50% decrease in the amplitude of I CRAC (e.g. Fig.5 C ). Overall, there was little difference between preparations, with the average decrease being 64.8 ± 7.9% (n = 35, measured using the step protocol after 300 s).Figure 5Slow inactivation is not affected by a variety of intracellular signaling pathways. A , dialysis with the broad kinase inhibitor H-7 (200 μm) did not affect slow inactivation relative to a control cell from the same coverslip. See also Table II. B , preincubation for 30 min with the broad phosphatase inhibitor okadaic acid (1 μm) was ineffective on slow inactivation. Okadaic acid was also included in the recording pipette. Okadaic acid enhanced Ca2+ influx inXenopus oocytes after application of lysophosphatidic acid (n = 2), as described in Petersen and Berridge (19Petersen C.C.H. Berridge M.J. Pfluegers Arch. 1996; 432: 286-292Crossref PubMed Scopus (59) Google Scholar). Hence okadaic acid was active. C , dialysis with the nonhydrolyzable GDP analogue GDPβS (300 μm) did not interfere with slow inactivation relative to control cells. GDPβS prevented activation of I CRAC by the adenosine receptor agonist NECA in two cells, indicating it was active (7Parekh A.B. Penner R. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 7907-7911Crossref PubMed Scopus (155) Google Scholar).D , pretreatment with the Ca2-activated protease inhibitor calpeptin (2 μm), as well as its inclusion in the recording pipette, failed to alter slow inactivation.View Large Image Figure ViewerDownload Hi-res image Download (PPT) In experiments where drugs were used to interfere with slow inactivation, control responses were always obtained from the same preparation and the data were compared with these controls. To confirm that slow inactivation is dependent on a rise in intracellular Ca2+ levels, we dialyzed cells with a pipette solution in which the EGTA concentration had been increased 10-fold (to 14 mm). Pooled data from 10 cells in 14 mm EGTA and eight cells in 1.4 mm EGTA are summarized in Fig. 1 D (paired recordings). Slow inactivation was reduced by almost 2-fold in the presence of the higher EGTA concentration and appears therefore to require a rise in intracellular free Ca2+ because it can be suppressed by increasing the concentration of a slow mobile Ca2+ buffer in the cytosol. The inset of Fig. 1 D shows current traces taken from the voltage steps from two cells dialysed with 1.4 and 14 mmEGTA. Fast inactivation was the same in both cells, yet slow inactivation was less pronounced in the presence of higher EGTA. Fast and slow inactivation therefore reflect distinct processes. To assess the contribution of Ca2+ influx through CRAC channels to the slow Ca2+-dependent inactivation, we carried out a series of experiments in which the electrochemical gradient for Ca2+ influx was altered. First, we modified the electrical gradient by applying voltage pulses to potentials that either reduced Ca2+ entry (−40 mV) or enhanced it (−120 mV), relative to the responses obtained on pulsing to −80 mV. Typical results are shown in Fig. 2 A , and the data are summarized in Table I. Stepping the voltage to −40 mV reduced the rate and extent of inactivation almost 2-fold (Fig. 2 A (i ) and Table I). The current at −40 mV in Fig. 2 A (i ) has been scaled so that it has the same amplitude as that at −80 mV in order to clearly show the slower inactivation. Voltage pulses to −120 mV, however, did not increase the level of steady-state inactivation compared with that seen on stepping to −80 mV (Fig. 2 A (ii ), where the current at −80 mV has been scaled, see also Table I), which might suggest that sufficient Ca2+ enters at −80 mV to maximally activate the inhibitory process.Table IEffect of changing the electrochemical gradient for calcium entry on slow inactivationTreatmentNormalizedI maxSteady-state currentτactivationNo. of cellspA/pF% peaks1.4 BAPTA, −80 mV−1.90 ± 0.1748.9 ± 8.617.6 ± 1.261.4 EGTA, −40 mV−1.20 ± 0.0765.3 ± 3.818.6 ± 4.441.4 EGTA, −80 mV−1.60 ± 0.0535.7 ± 6.716.8 ± 1.841.4 EGTA, −120 mV−2.81 ± 0.3042.0 ± 5.813.9 ± 1.741.4 EGTA, 4 Ca2+ −80 mV−1.20 ± 0.1762.0 ± 6.518.6 ± 4.13In all experiments, 30 μm InsP3 was included in the recording pipette. 1.4 mm BAPTA/EGTA refers to 1.4 mm of total chelator. The voltage represents the potential to which the cells were stepped. All experiments were carried out in 10 mm external Ca2+, except the last row in which the bath solution contained 4 mm instead. The difference in steady-state current between 1.4 EGTA, −40 mV and 1.4 EGTA, −80 mV was significant (p < 0.02), whereas the difference between 1.4 EGTA, −80 mV and 1.4 EGTA, −120 mV was not (p > 0.4). The difference between 1.4 EGTA, −80 mV and 1.4 BAPTA, −80 mV was also not significant (p > 0.2). There were no significant differences between the τactivation values. Open table in a new tab In all experiments, 30 μm InsP3 was included in the recording pipette. 1.4 mm BAPTA/EGTA refers to 1.4 mm of total chelator. The voltage represents the potential to which the cells were stepped. All experiments were carried out in 10 mm external Ca2+, except the last row in which the bath solution contained 4 mm instead. The difference in steady-state current between 1.4 EGTA, −40 mV and 1.4 EGTA, −80 mV was significant (p < 0.02), whereas the difference between 1.4 EGTA, −80 mV and 1.4 EGTA, −120 mV was not (p > 0.4). The difference between 1.4 EGTA, −80 mV and 1.4 BAPTA, −80 mV was also not significant (p > 0.2). There were no significant differences between the τactivation values. In a second set of experiments, we reduced the concentration gradient for Ca2+ influx, while leaving the electrical driving force unchanged. This was accomplished by lowering the external Ca2+ concentration in the bath to 4 mm (from 10 mm). A typical example is shown in Fig. 2 B , in which the cell was hyperpolarized to −80 mV at 2-s intervals. Steady-state inactivation was less pronounced (22% for this cell, see also Table I). The fact that slow inactivation was reduced following a decrease only in the concentration gradient for Ca2+ entry while the electrical gradient was unchanged suggests that it is predominantly a Ca2+-dependent rather than a voltage-dependent phenomenon. The preceding results demonstrate that I CRAC is subject to a negative feedback mechanism in which Ca2+ influx through CRAC channels induces a slow Ca2+-dependent inactivation. Fast negative feedback inactivation of CRAC channels operates on a milliseconds time-scale and can be substantially reduced by inclusion of the fast Ca2+ chelator BAPTA in the recording pipette, whereas it is not altered by increasing the concentration of the slower Ca2+ chelator EGTA (8Zweifach A. Lewis R.S. J. Gen. Physiol. 1995; 105: 209-226Crossref PubMed Scopus (336) Google Scholar). Replacing EGTA with the fast chelator BAPTA (both at 1.4 mm) did not significantly affect the rate or extent of inactivation (six cells, Fig. 2 C and Table I). However, rapid inactivation of I CRAC was slightly slowed by this concentration of BAPTA (1.2–1.4-fold, data not shown), reinforcing the notion that the inactivation we observe is distinct from the fast inactivation process. Although Ca2+ entry is important for initiating slow inactivation, we set out to determine whether slow inactivation still required Ca2+ entry even after the inactivation process had started to develop. Fig.3 describes two types of experiment which were designed to address this. In Fig. 3 A , hyperpolarizing steps to −80 mV were repetitively applied every 2 s from a holding potential of 0 mV. Once inactivation had clearly developed (at 200 s), the cell was held continuously at +20 mV for 30 s. At this positive potential, very little Ca2+ enters through CRAC channels (6Parekh A.B. Fleig A. Penner R. Cell. 1997; 89: 973-980Abstract Full Text Full Text PDF PubMed Scopus (205) Google Scholar). After 30 s, the cell was held again at 0 mV and hyperpolarizing steps were resumed. The amplitude ofI CRAC increased only slightly relative to the level it had reached prior to clamping the cell at +20 mV (four of nine cells). In the other five cells, no recovery was observed at all. Fig. 3 B shows an experiment in whichI CRAC had inactivated fully by 300 s. Hyperpolarizing pulses were stopped and the cell was held at 0 mV for 120 s. Hyperpolarizing pulses were then resumed. No recovery ofI CRAC occurred (three of four cells). In one cell, the amplitude of I CRAC recovered by 9%. Taken together, these results suggest that, once initiated, Ca2+-dependent slow inactivation ofI CRAC is sustained despite using maneuvers that reduce Ca2+ influx. The following set of experiments were aimed at elucidating the molecular mechanism that gave rise to the slow inactivation ofI CRAC. In the presence of moderate concentrations of EGTA, it is conceivable that Ca2+ entry through CRAC channels might enable the intracellular Ca2+ stores to refill, a process that would turn off I CRAC. If such a mechanism were responsible for the inactivation of I CRAC, then one would predict that maneuvers directed toward reducing Ca2+ uptake into the stores should prevent slow inactivation from occurring. Thapsigargin is a specific inhibitor of the Ca2+ATPase on the endoplasmic reticulum and prevents store refilling (10Thastrup O. Cullen P.J. Drobak B.K. Hanley M.R. Dawson A.P. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 2466-2470Crossref PubMed Scopus (3010) Google Scholar). We therefore carried out paired recordings in which six control cells were dialyzed with internal solution containing InsP3 and 1.4 mm EGTA, whereas six other cells from the same preparations were dialyzed with this solution supplemented with 2 μm thapsigargin. Fig.4 A shows the averaged results. Slow inactivation was only slightly reduced by thapsigargin, but this was not statistically significant. Thapsigargin is lipophilic, so it is possible that the drug diffuses out of the cell into the bath solution. Because we included a high concentration of thapsigargin in the pipette (2 μm), one would be surprised if the steady-state cytoplasmic concentration was less than a few hundred nanomolar, a concentration that is sufficient to reduce Ca2+ATPase activity in a variety of cell types (10Thastrup O. Cullen P.J. Drobak B.K. Hanley M.R. Dawson A.P. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 2466-2470Crossref PubMed Scopus (3010) Google Scholar). In two cells, we applied thapsigargin (1 μm) from the outside just after the onset of I CRAC (thapsigargin was also included in the recording pipette). The current still inactivated (by 53 and 61%). Nevertheless, we sought additional ways to probe the effects of compromised store refilling on slow inactivation. One method would be to employ the Ca2+ ionophore ionomycin. Ionomycin increases the permeability of the store membrane to Ca2+, and this would enable any Ca2+ that had been pumped into the stores to diffuse back into the cytosol thereby preventing stores from refilling. We included 2 μm ionomycin in the recording pipette together with InsP3 and 1.4 mm EGTA. Fig. 4 B shows the effects of ionomycin.I CRAC activated slightly faster in ionomycin-treated cells (τ of 16.1 ± 1.7 sversus 22.7 ± 3.0 s in control paired cells), which might indicate that ionomycin is diffusing into the cells rather quickly and accelerating store depletion in combination with InsP3. Slow inactivation of I CRACwas still apparent in the presence of ionomycin, and the current declined to a value similar to that seen in the absence of ionomycin in paired recordings (six cells for ionomycin, five for controls). We tested the possible involvement of a kinase-mediated phosphorylation reaction in the Ca2+-dependent slow inactivation in RBL cells in two independent ways. First, we examined the effects of removing ATP from the pipette solution and then replacing it with ATPγS, and second, we tested the effects of broad kinase and phosphatase inhibitors on the inactivation process. Slow inactivation was still present when ATP was omitted from the pipette solution (three cells, not shown), suggesting that global ATP levels are probably not important for the inactivation process. We then replaced ATP with ATPγS in the pipette solution. ATPγS is a nonhydrolyzable analogue of ATP that is readily used by a variety of protein kinases resulting in "irreversible phosphorylation" of the target (11Eckstein F. Annu. Rev. Biochem. 1985; 54: 367-402Crossref PubMed Google Scholar). Slow inactivation was slightly increased by ATPγS (TableII). Importantly, it was not reduced, which one might have expected if recovery from inactivation required a phosphorylation (12Lewis R.S. Dolmetsch R. Zweifach A. Clapham D.E. Ehrlich B.E. Organellar Ion Channels and Transporters. The Rockefeller University Press, New York1996: 241-254Google Scholar).Table IIEffects of various treatments on slow inactivationTreatmentNormalized I maxSteady-state currentτactivationNo. of cellspA/pF% peaksATPγS−1.80 ± 0.4029.8 ± 4.213.5 ± 2.44H-7−1.77 ± 0.1528.4 ± 9.014.7 ± 1.47Bisindolylmaleimide−1.41 ± 0.1354.1 ± 6.017.8 ± 1.96Okadaic acid−1.13 ± 0.3038.3 ± 12.519.2 ± 1.24GDPβS−1.86 ± 2.0139.7 ± 10.222.5 ± 2.54Calpeptin−1.34 ± 0.1934.0 ± 9.316.0 ± 1.05Steady-state current in control recordings (i.e. absence of drug) (% peak), ATPγS 38.3 ± 9.8 (4 cells); H-7, 32.0 ± 11.2 (4 cells); bisindolylmaleimide, 32.5 ± 4.7 (4 cells); okadaic acid and calpeptin (same preparations), 36.7 ± 7.1 (6 cells); GDPβS, 49.0 ± 10 (3 cells). Only steady-state current in the presence of bisindolylmaleimide was significantly different to control (p < 0.05). Open table in a new tab Steady-state current in control recordings (i.e. absence of drug) (% peak), ATPγS 38.3 ± 9.8 (4 cells); H-7, 32.0 ± 11.2 (4 cells); bisindolylmaleimi
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