©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Slow Calcium-dependent Inactivation of Depletion-activated Calcium Current
STORE-DEPENDENT AND -INDEPENDENT MECHANISMS (*)

Adam Zweifach (§) , Richard S. Lewis

From the (1)Department of Molecular and Cellular Physiology, Stanford University School of Medicine, Stanford, California 94305

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Feedback regulation of Ca release-activated Ca (CRAC) channels was studied in Jurkat leukemic T lymphocytes using whole cell recording and [Ca] measurement techniques. CRAC channels were activated by passively depleting intracellular Ca stores in the absence of extracellular Ca. Under conditions of moderate intracellular Ca buffering, elevating [Ca] to 22 mM initiated an inward current through CRAC channels that declined slowly with a half-time of 30 s. This slow inactivation was evoked by a rise in [Ca], as it was effectively suppressed by an elevated level of EGTA in the recording pipette that prevented increases in [Ca]. Blockade of Ca uptake into stores by thapsigargin with or without intracellular inositol 1,4,5-trisphosphate reduced the extent of slow inactivation by 50%, indicating that store refilling normally contributes significantly to this process. The store-independent (thapsigargin-insensitive) portion of slow inactivation was largely prevented by the protein phosphatase inhibitor, okadaic acid, and by a structurally related compound, 1-norokadaone, but not by calyculin A nor by cyclosporin A and FK506 at concentrations that fully inhibit calcineurin (protein phosphatase 2B) in T cells. These results argue against the involvement of protein phosphatases 1, 2A, 2B, or 3 in store-independent inactivation. We conclude that calcium acts through at least two slow negative feedback pathways to inhibit CRAC channels. Slow feedback inhibition of CRAC current is likely to play important roles in controlling the duration and dynamic behavior of receptor-generated Ca signals.


INTRODUCTION

In many nonexcitable cells, the depletion of intracellular Ca stores by inositol 1,4,5-trisphosphate (IP)()is the primary mechanism by which cell surface receptors activate Ca influx across the plasma membrane(1) . This phenomenon was first proposed by Putney and termed capacitative Ca entry(2) . Multiple types of ion channels underlying capacitative Ca entry have been identified in different cells on the basis of their Ca permeability and activation by agents that empty Ca stores (for review, see Ref. 3). The depletion-activated Ca channels present in mast cells and T cells, referred to as calcium release-activated Ca (CRAC) channels, are distinguished from other types of depletion-activated Ca channels by their high selectivity for Ca over monovalent and other divalent cations(4, 5, 6, 7, 8, 9) and their extremely small unitary conductance(7, 8) . CRAC channels have been shown to underlie the mitogen-stimulated Ca influx that is essential for T cell activation following T cell receptor engagement(7, 9) . Furthermore, periodic changes in CRAC channel activity have been shown to generate oscillations in the level of intracellular free Ca ([Ca])(4, 10) , which may serve to enhance signaling through the T cell receptor(11) .

The signal that couples store depletion to the activation of CRAC channels has not yet been identified, and in principle it could encompass diffusible and/or membrane-associated molecules. In a recent study, Randriamampita and Tsien (12) isolated a fraction from the cytoplasm of stimulated Jurkat leukemic human T cells that triggers Ca influx when applied to the exterior of astrocyte, neuroblastoma, and macrophage cell lines, apparently without releasing Ca from stores. This activity was attributed to calcium influx factor, a small (<500 M) nonproteinaceous phosphate-containing factor that they proposed as a diffusible messenger responsible for activating capacitative Ca entry. Additional activation mechanisms involving GTP-binding proteins(13, 14, 15) , tyrosine kinases(16) , cGMP(17, 18) , and direct physical coupling between the store's membrane and the plasma membrane (19) have been proposed (for reviews, see Refs. 1 and 3).

Little is known about how CRAC channels are turned off once activated, although several reports have demonstrated that increased [Ca] may play a role. Feedback inhibition by intracellular Ca appears to occur through multiple mechanisms. After entering the cell, Ca binds to sites probably residing on the CRAC channel itself(20) , eliciting rapid inactivation over tens of milliseconds(8, 20) . In addition, slow inactivation of I on a time scale of seconds has been observed following abrupt elevation of [Ca] or global rises in [Ca](4, 6, 7, 8) , but the underlying mechanisms have not been determined. A central tenet of the capacitative Ca entry hypothesis is that refilling of intracellular Ca stores should terminate Ca influx. This prediction has been confirmed in intact cells using fluorescent Ca indicators(10, 21, 22) . However, the ability of store refilling to close CRAC channels, which is an important test of the role of CRAC channels in capacitative Ca entry, has not yet been examined directly in patch-clamp experiments. Furthermore, the presence of additional negative feedback pathways through which Ca may turn off CRAC channels has not been explored.

This paper describes two mechanisms by which elevated [Ca] slowly feeds back to inactivate CRAC channels. One mechanism is dependent on store refilling, while the other operates even when refilling is completely prevented with thapsigargin. Thus, with the inclusion of fast inactivation(8, 20) , it appears that intracellular Ca feeds back by at least three independent pathways to control capacitative Ca entry. Autoregulatory feedback on CRAC channels by Ca is likely to be essential in determining the character of receptor-stimulated Ca signals in nonexcitable cells. A portion of this work has been reported previously in abstract form (23).


EXPERIMENTAL PROCEDURES

Cells and Reagents

Jurkat E6-1 human leukemic T cells were maintained in complete medium containing RPMI 1640 and 10% heat-inactivated fetal bovine serum, 2 mM glutamine, and 25 mM HEPES, in a 6% CO humidified atmosphere at 37 °C. Log phase cells (0.2-1.2 10/ml) were used in all experiments. Thapsigargin, okadaic acid, 1-norokadaone, and calyculin A (LC Pharmaceuticals, Woburn, MA) were prepared as stocks of 1 mM or 100 µM in MeSO. Cyclosporin A (2 mg/ml in 2% ethanol) and FK506 (50 µg/ml in ethanol) were the generous gift of Drs. N. Clipstone and G. Crabtree at Stanford University.

Patch-Clamp Recording

Patch-clamp experiments were conducted in the standard whole cell recording configuration(24) . Extracellular Ringer's solution contained the following: 155 mM NaCl, 4.5 mM KCl, 1 mM MgCl, 2 or 22 mM CaCl, 10 mMD-glucose, and 5 mM Na-HEPES (pH 7.4). Ca-free Ringer's contained 3 mM MgCl. Internal solutions contained the following: 140 mM cesium aspartate, 10 mM Cs-HEPES (pH 7.2), and either 0.66 mM CaCl, 11.68 mM EGTA, and 3.01 mM MgCl (12 mM EGTA solution) or 0.066 mM CaCl, 1.2 mM EGTA, and 2.01 mM MgCl (1.2 mM EGTA solution). Free [Ca] in both of these solutions as measured with indo-1 was 5 nM; free [Mg] was calculated to be 2 mM. Recording electrodes were pulled from 100-µl pipettes (VWR), coated with Sylgard near their tips, and fire-polished to a resistance of 2-6 megaohms when filled with cesium aspartate pipette solution. The patch-clamp output (Axopatch 200, Axon Instruments, Foster City, CA) was filtered at 1.5 kHz with an 8-pole Bessel filter (Frequency Devices, Haverhill, MA) and digitized at a rate of 5 kHz. Stimulation and recording were performed with an Apple Macintosh computer driving an ITC-16 interface (Instrutech, Elmont, NY) and using PulseControl software extensions (Jack Herrington and Richard Bookman, University of Miami) to Igor Pro (WaveMetrics, Inc., Lake Oswego, OR). Command potentials were corrected for the -12 mV junction potential that exists between the aspartate-based pipette solutions and Ringer's solution. Cells were allowed to settle onto but not firmly adhere to glass coverslip chambers shortly prior to each experiment. Adherent and nonadherent cells behaved identically in experiments with 1.2 mM EGTA; however, currents obtained with 12 mM EGTA were not as sustained in adherent cells as in nonadherent ones. The reason for this difference was not investigated further. I was induced through the store depletion protocol described previously(20) . Following formation of the gigaseal, each cell was exposed to Ca-free Ringer's, and the whole cell recording configuration was established. This procedure was sufficient to activate I maximally, because additional pretreatment with 1 µM thapsigargin did not further increase the average initial size of Ca current (). After 3 min, [Ca]was elevated to 2-22 mM, and steady-state I was measured in all experiments as a 5-10-ms average at the end of 200-ms pulses from -12 mV (the holding potential) to -132 mV delivered once every 2 s. In the experiment shown in Fig. 1, peak I was measured from a 1-ms average beginning 3 ms after the start of each hyperpolarizing pulse to minimize contributions from uncompensated capacitative current (time constant, <1 ms) and fast inactivation. All data are corrected for leak and residual capacitative current measured in the absence of Ca. Leak conductances were 20-100 picosiemens. Series resistance compensation was not employed, since the series resistance (4-25 megaohms) produced voltage errors of <3 mV. Cell capacitance was determined either from the settings of the whole cell capacitance-compensation circuitry, or by integrating currents elicited by 10-mV depolarizing steps. External solutions were changed by positioning the cell 1 mm inside one barrel of a perfusion tube array through which the desired solutions flowed (<0.1 ml/min). Experiments were conducted at 22-25 °C. Where normalized data are presented, current amplitudes were divided by the current's maximal value, and the time at which the maximum occurred was defined as time zero. Data are presented as mean ± S.E. Statistical significance of results was assessed using the Mann-Whitney U test, and differences are considered significant if p < 0.01. [Ca] Measurements with Indo-1-For experiments combining patch-clamp recording with [Ca] measurements, pipette solutions were supplemented with 100 µM indo-1 pentapotassium salt (Molecular Probes, Eugene, OR). Recording conditions have been described in detail previously(20) . Briefly, cells were illuminated using a 75W xenon arc lamp and a 360 ± 5-nm interference filter (Omega Optical, Brattleboro, VT) mounted on a Nikon Diaphot inverted microscope equipped with a Nikon Fluor 40 objective (numerical aperature, 1.3) and a transistor-transistor logic-controlled shutter to control the duration of illumination. The emission signal was collected from an area adjusted to be slightly larger than the cell. Emitted light was split with a 440-nm dichroic mirror and passed through 405 ± 15 and 480 ± 12.5 nm interference filters (Chroma Technology Corp., Brattleboro, VT) to two photomultiplier tubes (HC124-02, Hamamatsu Corp., Bridgewater, NJ). [Ca] was estimated from the relation [Ca] = K*(R - R)/(R - R). Background fluorescence was measured in the cell-attached mode and was subtracted from subsequent fluorescence signals before calculation of the 405/480 ratio, R. K*, R, and R were determined from in vivo calibrations as described previously(20) .


Figure 1: Fast and slow inactivation of I. A, after depletion of intracellular Ca stores, exposure of a Jurkat T cell to 22 mM Ca induces a slowly decaying inward current. Data points were measured every 2 s from the peak current () or steady-state current () elicited during brief hyperpolarizing voltage pulses shown in B (see ``Experimental Procedures''). Collection times for the traces in B and C are indicated by i-iii and a-c, respectively. B, selected current responses to 200-ms voltage pulses from -12 to -132 mV. Rapid inactivation of I is induced by the sudden increase in Ca influx upon hyperpolarization. Dashed line indicates zero current level. C, current/voltage relation for the inward current. Responses to voltage ramps (200-ms duration) from -120 to +50 mV show that current is inward in this voltage range as expected for I. Data in B and C are not corrected for leak or capacitative currents. Internal solution, cesium aspartate + 1.2 mM EGTA.




RESULTS

Fast and Slow Inactivation of I by Intracellular Ca

I was induced in Jurkat leukemic T cells by the passive depletion of intracellular Ca stores. Depletion was achieved in these experiments by incubating each cell in Ca-free Ringer's solution for 3 min while dialyzing its interior with a pipette solution in which [Ca] was buffered to 5 nM. After such treatment, elevation of [Ca] from 0 to 22 mM rapidly elicited an inward current (Fig. 1) whose properties identified it as I(4, 5, 6, 7, 8, 9, 20) . These properties included a dependence on Ca (Fig. 1A), rapid inactivation during hyperpolarizing voltage pulses (Fig. 1B), an inwardly rectifying current-voltage relation with no clear reversal potential up to +50 mV (Fig. 1C), voltage-independent gating, and a lack of significant current noise. As illustrated in Fig. 1A, I decayed slowly in cells dialyzed with a relatively low amount of EGTA (1.2 mM). In this paper, we refer to the slow decay in current as slow inactivation; the use of this terminology is not intended to imply any specific type of mechanism. In all experiments, we measured I during brief hyperpolarizing voltage pulses to -132 mV delivered every 2 s (as in Fig. 1B). This protocol optimizes the size of the current and allows independent measurement of fast and slow inactivation, as described previously(20) . The decay in the peak currents (Fig. 1A, squares) reflects the slow inactivation process alone, whereas decay of the steady-state currents (Fig. 1A, circles) is determined by both fast and slow inactivation. Thus, the fact that both measures of I decay with the same time course suggests that the fast and slow inactivation processes are independent. In further support of this conclusion, none of the pharmacological treatments described below affected fast inactivation.

[Ca] and current measurements were combined in indo-1-loaded cells to examine the calcium dependence of slow inactivation. When depleted cells dialyzed with low [EGTA](1.2 mM) were exposed to 22 mM Ca, [Ca] remained relatively constant for 10 s before climbing to micromolar levels, as would be expected since Ca influx via I should eventually overcome the capacity of EGTA to buffer Ca (Fig. 2A). The increase in [Ca] was associated with a progressive inactivation of I over a period of 100 s. As the current declined, [Ca] reached a peak and decreased, presumably as the rate of Ca entry fell below that of Ca efflux by Ca-ATPases in the plasma membrane. Increasing the buffering power of the pipette solution with 12 mM EGTA restricted [Ca] to levels below 100 nM for the duration of the experiment and reduced the slow decay of the current (Fig. 2B). The ability of increased intracellular Ca buffering to prolong the current is summarized for 10-17 experiments in Fig. 2C. These results demonstrate that slow inactivation is Ca-dependent.


Figure 2: Slow inactivation of I is Ca-dependent. A, in the presence of 1.2 mM EGTA, the induction and subsequent slow decline of I is associated with a delayed rise and fall of [Ca]. B, 12 mM EGTA in the pipette largely suppresses the [Ca] increase, and I is more sustained. All currents shown in this and subsequent figures were measured at the end of hyperpolarizing voltage pulses to -132 mV as described in Fig. 1B. C, the effect of intracellular buffering on slow inhibition of I from experiments like those in A and B. Current amplitude is normalized to the maximum amplitude reached in each experiment after exposure to 22 mM Ca, and time is plotted from this point onward. Plotted values are the mean ± S.E. of 10-17 cells.



During whole cell recording, soluble components diffuse out of the cell and may cause rundown of channel activity. To test whether slow inactivation of I is due to a washout phenomenon, we examined whether it can be reversed by removal of extracellular Ca. Inhibition of I in the presence of 22 mM Ca was allowed to reach steady state; at this point, Ca was removed for a variable period and then reapplied to measure the extent to which I had recovered. In the cell depicted in Fig. 3, a 60-s exposure to Ca-free conditions allowed nearly complete recovery of the current's amplitude. The extent of recovery varied among the cells tested. In six cells in which slow inactivation reduced I to a level of 11 ± 3%, subsequent incubation in 0 Ca for 100-150 s allowed recovery to 54 ± 14% of the initial peak amplitude. The source of the variation is not known. However, evidence presented below suggests that most of the inactivation in these experiments is due to reuptake of Ca by stores; thus, a variable degree of reemptying following removal of Ca may contribute to the different amounts of recovery that were observed. Regardless, these results further support the Ca dependence of slow inactivation and demonstrate that it is not simply due to nonspecific rundown or to washout of a factor essential for maintenance of I.


Figure 3: Slow inactivation is reversible. After I declined to a steady-state level in 22 mM Ca, the cell was bathed in Ca-free Ringer's solution for 60 s and then reexposed to 22 mM Ca. In this cell, the current recovered almost completely, indicating that inactivation is not due to washout of a diffusible factor.



Refilling of CaStores Contributes to Slow Inactivation

In the experiments described above, Ca stores were depleted passively, without increasing the permeability of the endoplasmic reticulum membrane to Ca or inhibiting its Ca-ATPases. Under these conditions, stores would be expected to refill efficiently following a rise in [Ca], thereby causing the slow inactivation of I. To test this hypothesis, we measured the time course of I in the presence of a high dose of TG (1 µM) that fully blocks Ca-ATPases in the endoplasmic reticulum (9, 10, 25) and hence prevents Ca reuptake. Cells were incubated in Ca-free Ringer's + TG for 3 min prior to exposure to 22 mM Ca. Under these conditions with 1.2 mM EGTA, slow inactivation of I was greatly reduced despite a large increase in [Ca] (Fig. 4A, ). In 21 cells treated with TG, I decayed over 100 s to a steady-state level of 50% of its initial value (Fig. 4B). Consistent with the time course of the current, [Ca] declined only partially over the same time period (Fig. 4A, ). These results demonstrate that store refilling contributes to slow Ca-dependent inactivation of I. The failure of TG to fully hinder this process is not due to an inability to prevent store refilling. Experiments with TG were repeated with 20 µM IP in the internal solution to attempt to increase the overall extent of store depletion. The release of stored Ca by 20 µM IP was confirmed in separate experiments by a large [Ca] spike occurring within seconds of breaking into cells in Ca-free Ringer's; such transients were not observed in the absence of IP (data not shown). Furthermore, IP alone prevented slow inactivation to about the same extent as TG alone (50%; n = 3 cells). As summarized in Fig. 4B, IP and TG together had the same effect as TG alone, indicating that store refilling is not occurring and therefore cannot explain the failure of TG to prevent I inactivation. Rather, these results reveal a second process of Ca-dependent slow inactivation occurring independently of changes in Ca store content.


Figure 4: Store refilling contributes to slow inactivation. A, 1 µM TG partially blocks the slow decline in I and [Ca] in one cell exposed to 22 mM Ca with 1.2 mM EGTA. TG was present during the 3-min preincubation in Ca-free Ringer's. B, TG limits slow inactivation to an average final extent of 50%. Addition of 20 µM IP to the internal solution produces no further effect, indicating that incomplete emptying of stores is not responsible for the partial effect of TG on current inhibition. Data are the average normalized currents ± S.E. from 14-21 cells in experiments like those shown in A.



Effects of Phosphatase Inhibitors on Slow Inactivation

Okadaic acid, a potent inhibitor of phosphatases 1 and 2A (26, 27) has been reported to enhance capacitative Ca influx in Xenopus oocytes(28) . We therefore tested the ability of okadaic acid and other phosphatase inhibitors to suppress the TG-insensitive component of slow inactivation. Okadaic acid inhibits protein phosphatases 1 and 2A in vitro with IC values of 1-50 nM(27, 29) . A 3-min preincubation with 100 nM okadaic acid + TG significantly reduced the extent of slow inactivation over that seen with TG alone (Fig. 5). On average, the current inactivated only 25% over 100 s, comparable with the amount the current declines when the [Ca] rise is mostly suppressed with 12 mM EGTA (Fig. 2C).


Figure 5: Effects of phosphatase inhibitors on store-independent I inactivation. Bars indicate the average (± S.E.) extent of current decline after 100 s in 22 mM Ca with 1.2 mM EGTA and various inhibitors. The dashed line shows the average extent of inactivation observed with 1.2 mM EGTA + 1 µM TG (defining the maximum possible level of store-independent inactivation). Conditions include 100 nM okadaic acid (n = 21), 100 nM 1-norokadaone (n = 6), 0.1-1.0 µM calyculin A (n = 11), 0.17 µM CsA (n = 14), 1.7 µM CsA (n = 20), and 62 nM FK506 (n = 10). Of these, only okadaic acid, 1-norokadaone, and 1.7 µM CsA reduce the extent of inactivation to a statistically significant degree (p < 0.01).



Okadaic acid pretreatment did not affect the maximum amplitude of I attained in the presence of TG (3.0 ± 0.2 pA/pF; n = 14) relative to control (), suggesting that an okadaic acid-sensitive process does not contribute to the activation of I. In addition, 100 nM okadaic acid applied in the absence of TG lacked any statistically significant effect on the time course or extent of slow inactivation (decay half-time, 36 s; final level, 0.26 ± 0.10; n = 14) compared with control (see Fig. 2C). This result implies that store refilling assumes the major role in turning off CRAC channels under the conditions of these experiments (i.e. passive store depletion and normal endoplasmic reticulum Ca-ATPase activity).

While effects of okadaic acid are often accepted as evidence for actions of protein phosphatase 1 or protein phosphatase 2A, the results of further experiments argue against such a conclusion. First, 100 nM 1-norokadaone, a compound similar in structure to okadaic acid but with little or no inhibitory activity against protein phosphatase 1 and protein phosphatase 2A(26, 29) , produced effects that were indistinguishable from those of okadaic acid (Fig. 5). Moreover, calyculin A (0.1-1 µM), an unrelated compound with subnanomolar efficacy for inhibiting protein phosphatase 1 and protein phosphatase 2A(27, 29) , lacked a significant effect on slow inactivation (Fig. 5). These results are inconsistent with a mechanism involving protein phosphatase 1 or protein phosphatase 2A.

The possible involvement of the Ca/calmodulin-dependent protein phosphatase 2B (calcineurin) in the store-independent inactivation of I was addressed using the immunosuppressants cyclosporin A and FK506. These compounds are extremely potent inhibitors of protein phosphatase 2B when complexed with their respective binding proteins, the cyclophilins and FKBP(30) . IC values for the inhibition of calcineurin in Jurkat cell lysates are 5 nM for CsA and 0.5 nM for FK506(31) . As summarized in Fig. 5, CsA prevented TG-insensitive slow inactivation to the same extent as okadaic acid when applied at 1.7 µM but not at 170 nM. Treatment of cells with 62 nM FK506 had no significant effect. These results argue against a necessary role for protein phosphatase 2B/calcineurin in the Ca-dependent inactivation of I.


DISCUSSION

Three Independent Mechanisms for Feedback Regulation of CRAC Channels

The results of this study and a previous one (20) demonstrate that intracellular Ca regulates CRAC channels by at least three distinct mechanisms. The three modes of feedback are readily distinguished by their kinetics, by their site of action relative to CRAC channels, and by their pharmacological profiles. Fast inactivation (Fig. 1B) occurs within milliseconds of Ca entry and is controlled by binding to sites located within several nanometers of the pore, probably residing on the CRAC channel itself(20) . The two forms of slow inactivation described in this study are roughly 1000-fold slower and are driven by a global rather than a local rise in [Ca] (see below). None of the pharmacological agents employed in this study had any effect on fast inactivation, whereas thapsigargin selectively inhibited the store-dependent component of slow inactivation, and okadaic acid, 1-norokadaone, and a high concentration of cyclosporin A inhibited the store-independent component. The independence of fast and slow inactivation is also consistent with the invariance of fast inactivation during the induction and decline of I as stores empty and refill(20) . For these reasons, we conclude that fast and slow inactivation occur through distinct mechanisms that operate in parallel to regulate CRAC channels. These studies provide the first evidence that capacitative Ca entry is controlled through mechanisms independent of Ca store content. Thus, the results raise the interesting possibility that additional signaling pathways may modulate I.

The Calcium Dependence of Slow Inactivation

The ability of high levels of an intracellular Ca buffer (12 mM EGTA) to reduce the extent of slow inactivation agrees with previous observations that intracellular Ca buffering enhances the amplitude and duration of I in whole-cell recordings(4, 6) . Due to its slow Ca binding rate, 12 mM EGTA cannot reduce [Ca] in microdomains near CRAC channels(20) , but it can suppress increases in bulk [Ca] (Fig. 2B). Thus, the effect of high [EGTA] suggests that most of the Ca binding sites subserving slow inactivation are farther from the channel than those underlying fast inactivation(20) .

Slow Inactivation by Store Refilling

A critical prediction of the capacitative Ca entry hypothesis is that store refilling should terminate influx. This prediction has been supported by previous reports that store refilling is temporally correlated with decreased influx of Ca or Mn (21, 22). However, the effect of refilling on the CRAC channels themselves, as well as the possible existence of additional inactivation pathways, has not been explored. Our findings that TG inhibits 50% of the slow inactivation of I therefore constitute an important confirmation that I underlies capacitative Ca influx in Jurkat T cells and additionally reveal a store-independent inactivation mechanism operating in parallel.

Because the mechanism by which depletion activates CRAC channels is not yet clear, we cannot infer how store refilling inhibits them. Although induction of an inhibitory signal by replete stores cannot be ruled out, the simplest explanation consistent with the capacitative entry hypothesis is that refilling terminates the channel activation signal, whether it be a diffusible activator or a protein-protein coupling mechanism. The time course of slow inactivation may reflect the lifetime of the activation signal if stores refill rapidly and the open channel lifetime is relatively brief. TG increases the peak [Ca] without affecting the peak current amplitudes (), suggesting that stores in passively depleted cells do in fact rapidly sequester Ca. The average current with 1.2 mM EGTA decays with a half-time of 32 s (Fig. 2C). These kinetics are similar to the time course with which depletion-induced Mn influx in neutrophils decays upon refilling of stores (t = 20 s at 25 °C)(22) . Thus, our results are consistent with the action of a relatively long-lived CRAC channel activator, a feature that may have important implications for the generation of [Ca] oscillations (see below).

Slow Inactivation by a Store-independent Mechanism

A second mechanism of slow Ca-dependent inactivation of I was revealed by the current's decline in the presence of 1 µM TG, a condition that precludes store refilling. Store-independent inactivation is largely inhibited by okadaic acid, an observation that may explain the potentiating effect of okadaic acid on capacitative Ca entry described previously in Xenopus oocytes(28) . In those studies, okadaic acid reversed the decay of a Ca-activated Cl current in 5-hydroxytryptamine-stimulated oocytes, leading to the speculation that a phosphatase acts to limit the lifetime of a diffusible CRAC channel activator(28) . In view of our present results, the gradual decline in Ca-activated Cl current seen in the oocytes may have resulted from the Ca-dependent inactivation of I by the store-independent (okadaic acid-sensitive) mechanism. Further experiments in oocytes using 1-norokadaone and calyculin A could test this hypothesis.

The pharmacological profile of store-independent slow inactivation argues against the involvement of several well known phosphatases. Although okadaic acid largely blocked inactivation and is known to be a potent inhibitor of phosphatases 1, 2A, and 3(29, 32) , calyculin A, an even more potent antagonist of these phosphatases(29) , lacked any effect on the inactivation process. Furthermore, 1-norokadaone, which lacks significant activity against all three phosphatases(29) , blocked slow inactivation with the same efficacy as okadaic acid. Calcineurin (protein phosphatase 2B) also does not appear to be required for slow inactivation, because cyclosporin A and FK506 lacked an effect even at doses 30-100-fold greater than their IC values for inhibition of protein phosphatase 2B in Jurkat cells. Taken together, our results argue against a role for protein phosphatases 1, 2A, 2B, and 3 in mediating slow inactivation of CRAC channels. The inhibitory effects we observed may be nonspecific. The mechanism underlying store-independent slow inactivation is unknown, but could in principle result from a decrease in the CRAC channel activation signal, production of a channel inhibitor, or a change in the channels themselves.

Physiological Role of Slow IInactivation

Slow inactivation is likely to play an essential role in the generation of [Ca]oscillations in T cells. Oscillations in Jurkat and human T cells are triggered by several stimuli that release moderate amounts of Ca from intracellular stores, such as cross-linking of the T cell antigen receptor and low doses of ionomycin or thapsigargin and other smooth endoplasmic reticulum Ca-ATPase inhibitors(4, 10, 33, 34) . Periodic Ca influx via I appears to be responsible for [Ca]oscillations in these cells(4, 10) , supporting a simple oscillation model in which fluctuations in store content and phasic opening and closing of CRAC channels are coupled through changes in [Ca](10) . To enable this model system to oscillate, time lags must exist between store depletion and CRAC channel opening and between [Ca] elevation, store refilling, and CRAC channel closure. Previous work has shown that CRAC channels open with a time constant of 20-30 s following store depletion with IP(6, 8) , demonstrating a lag between depletion and channel activation. This study describes two mechanisms that operate with similarly slow kinetics (half-times, 30 s) to control channel closure following [Ca]elevation.

Store-dependent and -independent mechanisms of slow inactivation constitute two parallel pathways by which intracellular Ca feeds back to control the amplitude and duration of capacitative Ca influx. The relative contribution of these two mechanisms to Ca signaling under physiological conditions is not known. The relatively small effect of okadaic acid observed in the absence of TG (Fig. 5) suggests that the store-dependent mechanism is dominant; however, it should be noted that the passive method of depletion employed in these experiments would be expected to favor store refilling. The store-independent inactivation mechanism is likely to play a greater role under physiological conditions, in which elevated intracellular IP promotes Ca release and hinders store refilling. The existence of a mechanism that regulates CRAC channels independently of store content reveals an unsuspected degree of complexity in the regulation of capacitative Ca entry.

  
Table: I and [Ca] following store depletion

Stores were depleted under the conditions listed, with 1 µM TG or 20 µM IP present where noted. Unless otherwise stated, all experiments were conducted with 1.2 mM EGTA. I was measured during voltage steps to -132 mV as described under ``Experimental Procedures.'' I is the maximal current observed after elevation of [Ca] from 0 to 22 mM; this occurred within 10 s of the solution change. Normalized I is the maximum current divided by membrane capacitance and reflects the surface density of open channels. Peak [Ca] is the highest value observed after elevation of [Ca] from 0 to 22 mM after store depletion. Steady-state [Ca] is the value measured 120 s after the solution change. [Ca] did not reach a peak or a steady-state value in the presence of 12 mM EGTA; for these experiments, peak [Ca] was measured 20 s after elevation of [Ca] from 0 to 22 mM. Data are presented as mean ± S.E. (n).



FOOTNOTES

*
This work was supported by National Institutes of Health postdoctoral fellowship AI08568 (to A. Z.) and National Institutes of Health Grant GM47354 (to R. S. L.). 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: Beckman Center B-003, Stanford University School of Medicine, Stanford, CA 94305. Tel.: 415-723-9623; Fax: 415-725-8021.

The abbreviations used are: IP, inositol 1,4,5-trisphosphate; [Ca], free intracellular Ca concentration; I, Ca release-activated Ca current; TG, thapsigargin; CsA, cyclosporin A.


ACKNOWLEDGEMENTS

We thank Ricardo Dolmetsch and Chris Fanger and Drs. Markus Hoth, Neil Clipstone, and Paul Driedger for helpful discussions; Supriya Kelkar for invaluable assistance with cell culture; and Dr. Markus Hoth for critical comments on the manuscript.


REFERENCES
  1. Putney, J. W., Jr., and Bird, G. S. J.(1993) Endocrinol. Rev.14, 610-631 [Medline] [Order article via Infotrieve]
  2. Putney, J. W., Jr.(1990) Cell Calcium11, 611-624 [Medline] [Order article via Infotrieve]
  3. Fasolato, C., Innocenti, B., and Pozzan, T.(1994) Trends Pharmacol. Sci.15, 77-83 [CrossRef][Medline] [Order article via Infotrieve]
  4. Lewis, R. S., and Cahalan, M. D.(1989) Cell Regul.1, 99-112 [Medline] [Order article via Infotrieve]
  5. Hoth, M., and Penner, R.(1992) Nature355, 353-356 [CrossRef][Medline] [Order article via Infotrieve]
  6. McDonald, T. V., Premack, B. A., and Gardner, P.(1993) J. Biol. Chem.268, 3889-3896 [Abstract/Free Full Text]
  7. Zweifach, A., and Lewis, R. S.(1993) Proc. Natl. Acad. Sci. U. S. A.90, 6295-6299 [Abstract]
  8. Hoth, M., and Penner, R.(1993) J. Physiol.(Lond.) 465, 359-386 [Abstract]
  9. Premack, B. A., McDonald, T. V., and Gardner, P.(1994) J. Immunol.152, 5226-5240 [Abstract/Free Full Text]
  10. Dolmetsch, R., and Lewis, R. S.(1994) J. Gen. Physiol.103, 365-388 [Abstract]
  11. Negulescu, P. A., Shastri, N., and Cahalan, M. D.(1994) Proc. Natl. Acad. Sci. U. S. A.91, 2873-2877 [Abstract]
  12. Randriamampita, C., and Tsien, R. Y.(1993) Nature364, 809-814 [CrossRef][Medline] [Order article via Infotrieve]
  13. Fasolato, C., Hoth, M., and Penner, R.(1993) J. Biol. Chem.268, 20737-20740 [Abstract/Free Full Text]
  14. Bird, G. S. J., and Putney, J. W., Jr.(1993) J. Biol. Chem.268, 21486-21488 [Abstract/Free Full Text]
  15. Jaconi, M. E. E., Lew, D. P., Monod, A., and Krause, K.-H.(1993) J. Biol. Chem.268, 26075-26078 [Abstract/Free Full Text]
  16. Sargeant, P., Farndale, R. W., and Sage, S. O.(1993) J. Biol. Chem.268, 18151-18156 [Abstract/Free Full Text]
  17. Bahnson, T. D., Pandol, S. J., and Dionne, V. E.(1993) J. Biol. Chem.268, 10808-10812 [Abstract/Free Full Text]
  18. Xu, X., Star, R. A., Tortorici, G., and Muallem, S.(1994) J. Biol. Chem.269, 12645-12653 [Abstract/Free Full Text]
  19. Berridge, M. J.(1993) Nature361, 315-325 [CrossRef][Medline] [Order article via Infotrieve]
  20. Zweifach, A., and Lewis, R. S.(1995) J. Gen. Physiol.105, 209-226 [Abstract]
  21. Jacob, R.(1990) J. Physiol.(Lond.) 421, 55-77 [Abstract]
  22. Montero, M., Alvarez, J., and Garca-Sancho, J.(1992) Biochem. J.288, 519-525 [Medline] [Order article via Infotrieve]
  23. Zweifach, A., and Lewis, R. S.(1994) Biophys. J.66, A153 [Abstract]
  24. Hamill, O. P., Marty, A., Neher, E., Sakmann, B., and Sigworth, F. J. (1981) Pflügers Arch.391, 85-100
  25. Lytton, J., Westlin, M., and Hanley, M. R.(1991) J. Biol. Chem.266, 17067-17071 [Abstract/Free Full Text]
  26. Nishiwaki, S., Fujiki, H., Suganuma, M., Furuya-Suguri, H., Matsushima, R., Iida, Y., Ojika, M., Yamada, K., Uemura, D., Yasumoto, T., Schmitz, F. J., and Sugimura, T.(1990) Carcinogenesis11, 1837-1841 [Abstract]
  27. Cohen, P., Holmes, C. F. B., and Tsukitani, Y.(1990) Trends Biochem. Sci.15, 98-102 [CrossRef][Medline] [Order article via Infotrieve]
  28. Parekh, A. B., Terlau, H., and Stühmer, W.(1993) Nature364, 814-818 [CrossRef][Medline] [Order article via Infotrieve]
  29. Honkanen, R. E., Codispoti, B. A., Tse, K., and Boynton, A. L.(1994) Toxicon32, 339-350 [Medline] [Order article via Infotrieve]
  30. Liu, J., Farmer, J. D., Jr., Lane, W. S., Friedman, J., Weissman, I., and Schreiber, S. L.(1991) Cell66, 807-815 [Medline] [Order article via Infotrieve]
  31. Fruman, D. A., Klee, C. B., Bierer, B. E., and Burakoff, S. J.(1992) Proc. Natl. Acad. Sci. U. S. A.89, 3686-3690 [Abstract]
  32. Cohen, P. T. W., Brewis, N. D., Hughes, V., and Mann, D. J.(1990) FEBS Lett.268, 355-359 [CrossRef][Medline] [Order article via Infotrieve]
  33. Donnadieu, E., Bismuth, G., and Trautmann, A.(1992) J. Biol. Chem.267, 25864-25872 [Abstract/Free Full Text]
  34. Hess, S. D., Oortgiesen, M., and Cahalan, M. D.(1993) J. Immunol.150, 2620-2633 [Abstract/Free Full Text]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.