INTRODUCTION

In many nonexcitable cells, depletion of intracellular Ca²⁺ stores by inositol 1,4,5-trisphosphate activates Ca²⁺ influx across the plasma membrane, a phenomenon termed ``capacitative Ca<sup>2+</sup> entry'' (Putney, 1986). This Ca<sup>2+</sup> entry pathway was first characterized electrophysiologically in mast cells and named ``calcium release-activated calcium current'' (I_CRAC)¹ (Hoth and Penner, 1992). Store depletion-activated currents with different properties from I_CRAC have been described in some cell types (Lückhoff and Clapham, 1994; Vaca and Kunze 1994; Vaca et al. 1994); however, I_CRAC appears to be the predominant Ca²⁺ influx pathway in nonexcitable cells (Hoth and Penner 1993; Zweifach and Lewis, 1993; Somasundaram and Mahaut-Smith 1994; for review, see Fasalato et al. 1994). I_CRAC is distinguishable from other store-dependent influx currents by being highly selective for Ca²⁺ and having a single channel conductance below the resolution of patch clamp recordings (Hoth and Penner, 1993; Zweifach and Lewis, 1993). The mechanism whereby depletion of Ca²⁺ stores is coupled to the activation and control of I_CRAC is still unclear. Several signaling mechanisms have been proposed, including direct coupling of the store and plasma membrane via protein-protein interaction (Irvine, 1992) or release of a small, nonproteinaceous phosphate-containing second messenger from the Ca²⁺ stores that diffuses to the plasma membrane (Randriamampita and Tsien, 1993; Kim et al., 1995). A number of other biochemical modulators have been implicated in the control of I_CRAC, including cytochrome P-450, phosphatases, tyrosine kinases, cGMP (for review, see Sargeant and Sage, 1994), and protein kinase C (Parekh and Penner, 1995).

It has also been postulated that store depletion may cause insertion of I_CRAC channels into the plasma membrane from specialized vesicles (Fasalato et al., 1994) following evidence of a role of small GTP-binding proteins in I_CRAC activation (Fasalato et al., 1993; Bird and Putney, 1993) and the well recognized role of these molecules in vesicular transport (Pryer et al., 1992). We recently provided evidence to further support this notion by showing that two established inhibitors of vesicle-mediated protein transport, the antimalarial amine primaquine and GTPS, block the appearance of I_CRAC in response to store depletion in rat megakaryocytes (Somasundaram et al., 1995). Primaquine is far less effective if added after I_CRAC has developed. Thus the channels responsible for I_CRAC or a molecule activating the channel may be held in a membrane compartment and transported to the plasma membrane following depletion of stores (Somasundaram et al., 1995). It is also well established that vesicular transport is temperature-sensitive and is blocked below temperatures ranging from 16 to 18 °C (Matlin and Simons 1983; Tartakoff, 1986; Saraste et al., 1986). Therefore, in this study, we have investigated the effect of temperature on the activation and maintenance of store-mediated Ca²⁺ influx in a human leukemic cell line, KU-812 (Nakazawa et al., 1989), using a combination of whole cell patch clamp recordings and fluorescent photoquench of fura-2. The KU-812 cell line was selected because it displayed a high I_CRAC channel density and an ability to load and retain the Ca²⁺-sensitive dye fura-2 and because of the availability of large numbers of cells for population studies of Mn²⁺ quench.

MATERIALS AND METHODS

KU-812 cells were obtained from the European Collection of Animal Cell Cultures (Centre for Applied Microbiology and Research, Wiltshire, United Kingdom) and cultured in a humidified atmosphere at 37 °C and 5% CO₂ in RPMI 1640 medium (Life Technologies, Inc.) supplemented with 10% fetal calf serum, 2 mM glutamine, 10 units/ml penicillin, and 10 mg/ml streptomycin. Cells were harvested by centrifugation, washed, and resuspended in a standard external solution (see below). Thapsigargin (TG), valinomycin, and ionomycin were obtained from Calbiochem, and fura-2/AM and pluronic acid F-127 were from Molecular Probes, Inc., (Eugene, OR). All these agents were prepared as stocks in dimethyl sulfoxide. Cs₄-BAPTA was from Molecular Probes (Eugene OR), and all other reagents were from Sigma.

Patch clamp experiments were performed in the conventional whole cell configuration (Hamill et al., 1981) by means of an Axopatch 200A patch clamp amplifier (Axon Instruments, Inc., Foster City, CA). Pipettes were pulled from borosilicate glass tubing (Clark Electromedical Instruments) and had filled resistances of 2-3 megaohms. Series resistances were in the range of 10-30 megaohms, and 40-70% series resistance compensation was used. The cells were stored at 23-24 °C in a standard external solution containing 140 mM NaCl, 5 mM KCl, 1 mM MgCl₂, 2 mM CaCl₂, 10 mM glucose, and 10 mM HEPES, pH 7.4 (adjusted with Tris) before transfer to the recording chamber. All recordings were conducted in an external solution in which Na⁺ and K⁺ ions were replaced by equal concentrations of positively charged n-methyl-D-glucamine and Cs⁺, respectively. The internal saline contained 60 mM cesium gluconate, 20 mM Cs₄-BAPTA, 5 mM NaCl, 2 mM MgCl₂, 2 mM CaCl₂, and 10 mM HEPES, pH 7.3, (adjusted with Tris). This Ca²⁺/BAPTA mixture maintained [Ca²⁺]_i near resting cytosolic levels and reduced the chance of spontaneous activation of I_CRAC due to leaking of Ca²⁺ from internal stores that often occurred with BAPTA alone. TG (3 µM) was applied from a puffer pipette placed 150 µm from the cell. A diffusible factor required for activation of I_CRAC is washed out during whole cell recordings from rat megakaryocytes (Somasundaram et al., 1995) and rat basophilic leukemic cells (Fasalato et al., 1993). The rate of dialysis of the cytoplasm will depend on the whole cell access resistance. Therefore, to allow comparison of I_CRAC in different cells, the time after breakthrough to the whole cell mode was normalized for series resistance, and thapsigargin was applied at a standard normalized time range of 9 s megaohm¹. Cells were held at 20 mV, and current-voltage relationships were obtained at regular intervals using 300-ms voltage ramps from 140 to +60 mV. The membrane currents during voltage ramps were low pass filtered at 2 kHz and sampled at 100 µs using a Digidata 1200 interface and pClamp software (Axon Instruments). A continuous record of whole cell current was also obtained using a Cairn Research computer interface and associated software (Cairn Research Ltd., Kent, UK) following acquisition at a rate of 60 Hz (low pass filtered at approximately 30 Hz). Liquid junction potentials were measured by reference to a 3 M KCl agar bridge, and membrane potentials were adjusted accordingly. Currents were normalized for cell capacitance, and I_CRAC was measured as the amount of inward current that developed in response to TG at a given voltage.

In the electrophysiological experiments the temperature of the recording chamber was adjusted by a combination of a heating block and a heat exchanger controlled by a Peltier element. A gravity-fed perfusion system was used to exchange the saline in the experimental chamber. With this system the temperature of the recording chamber could be varied from 14 to 30 °C. Temperature was measured with a thermistor positioned 1 mm from the cell. At each temperature selected, the cell was allowed to equilibrate for at least 2 min before whole cell configuration was established. In experiments in which the effect of temperature on the maintenance of I_CRAC was measured, the whole cell configuration was established at 24 °C, and the temperature was lowered at a rate of approximately 0.1 °C/s. In the fluorescence experiments the temperature was controlled by a water jacket connected to a heating-cooling system, which enabled the temperature to be varied between 10 and 37 °C. In the calcium electrode experiments the temperature was changed between 15 and 30 °C using a Peltier element.

Ca²⁺ was determined with a Ca²⁺-sensitive minielectrode suspended in 60 µl of permeabilized cell suspension solution stirred by a magnetic agitator. The Ca²⁺ minielectrodes were prepared using a protocol similar to that of Clapper and Lee (1985). The electrode membrane was made with a Ca²⁺ electrode mixture (Fluka Chemical Corp.) containing 4.15 mg of ETH 1001 Ca²⁺ ionophore, 37 mg of S-nitrophenyloctyl ether, 0.42 mg of sodium tetraphenyl borate, and 24 mg of polyvinyl chloride dissolved in 240 µl of tetrahydrofuran. The electrode was filled with a 10 mM CaCl₂ solution and connected to a Genway 3040 pH and ion analyzer. The minireference (3 M KCl) electrode used (ULTRAWICK^TM) was purchased from World Precision Instruments, Inc. The electrode was calibrated at 15, 20, and 30 °C using various Ca²⁺-EGTA buffers and displayed a linear response within the range of 0.1-100 µM free Ca²⁺ (25 mV/pCa²⁺ unit). Cells were washed in a solution containing 120 mM KCl, 25 mM HEPES, 3 mM MgCl₂, and 1 mM EGTA, washed again in the same solution without EGTA, and finally resuspended in this non-EGTA-containing solution at a final concentration of 2 × 10⁸ cells/ml. The cells were then stored on ice until use. For each experiment, 25 µl of the cell suspension was warmed to 37 °C, and digitonin was added to give a final concentration of 40 µM and incubated for 2 min. The cell suspension was made up to 60 µl with non-EGTA-containing solution, phosphocreatine (10 mM), creatine kinase (20 units/ml), and 3 mM ATP and then transferred to form a droplet into which the calcium electrode tip was placed. The temperature was adjusted as required before experimentation.

Fura-2 fluorescence measurements were carried out on cell populations in a final volume of 1.5 ml (10⁶ cells/ml) using a Cairn spectrophotometer system. Excitation wavelengths of 340, 360, and 380 nm were provided by a filter wheel rotating at 35 Hz in the light path. Emitted light was filtered by a 485 nm long pass filter and samples averaged to give a data point every 500 ms. The background-corrected 340:380 ratio was multiplied by the dissociation constants for fura-2 at different temperatures obtained from Shuttleworth and Thompson (1991) to give an indication of [Ca²⁺]_i. The isosbestic excitation wavelength (360 nm) was used to monitor the fura-2 photoquench by Mn²⁺; an established measure of influx through the Ca²⁺ store-activated influx pathway. Cells were loaded with fura-2 by incubation with 1 µM fura-2/AM and 0.025% Pluronic F127 for 30 min at 24 °C in standard external solution containing 140 mM NaCl, 5 mM KCl, 1 mM MgCl₂, 10 mM glucose, 2 mg/ml apyrase, 10 mM HEPES, pH 7.4 (adjusted with Tris), and 1 mM CaCl₂. The apyrase was used to minimize purinergic receptor activation, which may be caused by ATP or ADP released from damaged cells. The cells were spun down and resuspended in the standard external solution with 2 mM CaCl₂ and split into 100-ml aliquots each containing 10⁶ cells and stored at room temperature until use. Prior to experimentation the cells were spun and resuspended in 100 ml of nominally Ca²⁺-free standard external medium with the K⁺ ionophore valinomycin (1 mM). Valinomycin was then present throughout the experiment allowing the cell membrane potential to be ``clamped'' close to the reversal potential of K⁺ (approximately 80 mV). No differences in Mn²⁺ influx were observed (at 16 and 37 °C) when valinomycin was added immediately or 6 min before measurement of Mn²⁺ photoquench (n = 3). To study the effect of temperature on activation of store-dependent Mn²⁺ influx, the 100-µl cell aliquot was added to 1.4 ml of medium at the required temperature. After a 30-s delay, 100 nM TG was added, followed 6 min later by 100 µM Mn²⁺. To measure basal Mn²⁺ influx, 100 µM Mn²⁺ was added in place of TG, after a 30-s incubation at the required temperature. To study the effects of temperature on the maintenance of the store-mediated Mn²⁺ influx pathway, the stores were emptied first by adding 100 nM TG to the 100-µl cell suspension in nominally Ca²⁺-free standard external medium and incubated for 6 min at 37 °C. The cell suspension was then added to the cuvette containing 1.4 ml of nominally Ca²⁺-free standard external medium at the required temperature. After a 30-s delay, 100 µM Mn²⁺ was added, and the quench was measured. In controls, the 100-µl cell suspensions were incubated at 37 °C without TG for the same length of time but with 1 mM added Ca²⁺ to prevent any passive store depletion during incubation. To quantify Mn²⁺ entry, the slopes of the fura-2 photoquench over the first 15 s after Mn²⁺ addition were determined. The very initial, sharp drop in fluorescence observed in some experiments, due to external free dye, was not included in the calculations.

The degree of TG-induced store depletion was assessed at both 34 and 18 °C by comparing the ionomycin-induced (5 µM) Ca²⁺ rise before and after a 6-min incubation with TG. To eliminate the contribution of Ca²⁺ influx to these measurements, 0.1 mM EGTA was added to the external medium immediately before addition of ionomycin.

Cytosolic ATP concentrations were measured using an ATP assay kit (Calbiochem) based on the firefly luciferase-catalyzed oxidation of D-luciferin in the presence of an ATP-magnesium salt and oxygen. A cell suspension (3 × 10⁵ cells/ml) in standard external medium was incubated at the desired temperature for 8 min, spun, and resuspended in HEPES buffer, pH 7.75, containing a permeabilizing reagent (Calbiochem). To assess ATP released from the cytoplasm, luciferase was added, and the peak amount of photons emitted was measured using a photomultiplier tube (Thorn EMI Electron Tubes Ltd., Ruislip, Middlesex, UK) coupled to a spectrophotometer (Cairn). Luminescence following addition of luciferase was measured and subtracted from luminescence in the absence of cells. The cytosolic ATP concentration was calculated using a measured mean cell diameter of 10 µm and assuming the cytosolic volume to be 20% of the total cell volume.

Temperature coefficient (Q₁₀) values were determined for current densities and Mn²⁺ influx rates within a given temperature range using Q₁₀ = (P₁/ P₂)^{10/(T1 T2)}, where P₁ and P₂ are current densities at temperature T1 and T2, respectively. Activation energy (E_a) was determined from an equation describing the slope of the Arrhenius plot: E_a = R (lnP₁  lnP₂)/(1/T1  1/T2), where R is the gas constant.

RESULTS

To study I_CRAC in KU-812 cells, conditions were selected that have been shown to enhance this small current and largely eliminate contributions from other ionic currents (Somasundaram et al., 1995; see ``Materials and Methods''). Briefly, the external medium contained 2 mM Ca²⁺ and no K⁺ or Na⁺, and any outward K⁺ currents were blocked by replacing the internal K⁺ with Cs⁺. The [Ca²⁺]₁ was strongly buffered with 20 mM BAPTA. Under whole cell voltage clamp, depletion of internal Ca²⁺ stores by 3 µM TG, an endoplasmic Ca-ATPase inhibitor, evoked an inward current at a holding potential of 50 mV (Fig. 1A). The currents generated by voltage ramps from 140 to + 50 mV before TG application and at various times during the development of the inward current are shown in Fig. 1B. This inwardly rectifying current showed little or no reversal within the voltage range studied, developed without detectable single channel events, was selective for Ca²⁺ (data not shown), and was blocked by 1 mM Zn²⁺ (data not shown), all of which are characteristic of I_CRAC reported in other cells (Fasalato et al., 1994; Somasundaram et al., 1995). To study the effect of temperature on the activation of I_CRAC, cells were held at various temperatures ranging from 15 to 27 °C, and the current evoked by TG was recorded. The currents activated at a holding potential of 20 mV at four different temperatures are shown in Fig. 1C. The maximum TG-induced current density was reduced with decreasing temperature such that no measurable current was activated at or below 17 °C. The speed of development of the current was also reduced, and the times to half-maximum current were 82 ± 12, 72 ± 8, 55 ± 9, and 50 ± 7 s at 19, 20, 23, and 26 °C, respectively. At maximal activation, the current-voltage relationship generated by voltage ramps from 140 mV to +50 mV at the different temperatures did not show any obvious shift in the reversal potential along the x axis (Fig. 1D). It should be noted that it was not technically possible to perform whole cell patch clamp recordings from cells held at temperatures greater than 27 °C due to rapid deterioration of glass membrane seals.

The TG-evoked I_CRAC densities, measured at 120 mV, in individual cells at different temperatures are shown in Fig. 2A. Despite the considerable variation in current density between cells, there is a clear positive correlation between current density and temperature. This relationship is further illustrated in Fig. 2B, in which the mean current density values at 120, 70, and 20 mV are plotted against temperature. Over the temperature range of 27 to 21 °C the current density at all three voltages displayed little change, whereas further lowering of temperature caused a significant decrease in current, which became blocked completely at 17 °C. To describe quantitatively the temperature dependence of the current density, we have presented the same data in the form of Arrhenius plots (Fig. 2C). The current density decreases linearly as temperature is lowered from 27 to 22 °C, and from the slope, apparent E_a values of 4.3 kcal/mol (Q₁₀ = 1.11), 5.1 kcal/mol (Q₁₀ = 1.26), and 5.8 kcal/mol (Q₁₀ = 1.18) were calculated at 120, 70, and 20 mV, respectively. On lowering the temperature from 21 to 17 °C, there was an abrupt increase in the E_a by a factor of approximately 33 to give E_a values of 161 kcal/mol (Q₁₀ = 196), 178 kcal/mol (Q₁₀ = 67), and 167 kcal/mol (Q₁₀ = 180) at the respective voltages, reflecting a very high energy barrier at the lower temperatures. For both the high and low temperature ranges, the activation energies were similar at different membrane potentials, suggesting that these steps were not voltage-dependent.

Although depletion of Ca²⁺ stores at 17 °C failed to induce an inward current, an increase in temperature several min after TG application at 17 °C resulted in I_CRAC activation, as shown by the experiment in Fig. 3. The current-voltage relationships at 17 °C 2 min after TG application (Fig. 3B, trace b) showed no evidence of a store-dependent current, whereas this current slowly developed following a temperature increase to 20 °C (Fig. 3B, trace c). A further abrupt increase in temperature to 28 °C produced a parallel increase in I_CRAC (Fig. 3B, trace d), and furthermore, this temperature-dependent effect was reversible, as shown by the currents obtained at 20 °C before and after increasing the temperature to 28 °C (Fig. 3B, traces c and e).

We next studied the effect of temperature on I_CRAC following full activation at 24 °C, to determine whether the temperature dependence arose primarily from an effect on the activation mechanism of the current or via a direct effect on the influx pathway itself. In the experiments of Fig. 4, I_CRAC was maximally activated by TG-induced store depletion at 24.5 °C before the temperature was lowered to 14.5 °C at a rate of 0.1 °C/s and then returned to 24.5 °C. A continuous record of I_CRAC at a holding potential of 60 mV is shown in Fig. 4A, and current-voltage relationships at different temperatures are shown in Fig. 4B. Fig. 4C shows the average current density at 120 and 70 mV calculated from 10 cells at different temperatures following protocols similar to that of Fig. 4A and shows the complete reversibility of this temperature block.

When the temperature was dropped from 24.5 to 22 °C, the inhibition of current density was of a level (Fig. 5A) very similar to that observed when measuring temperature effects on I_CRAC activation (Fig. 2B). However, reduction in temperature from 22 to 14.5 °C resulted in incomplete inhibition of current. Even at 14.5 °C, the I_CRAC current density remained at 44% of maximum, in contrast to the complete inhibition at this temperature observed for I_CRAC activation (Fig. 2B). Indeed, even after prolonged incubations of up to 3 min at 14 °C, cells still maintained substantial amounts of I_CRAC (data not shown). This was in contrast to the complete block of I_CRAC activation observed at this temperature (Fig. 2A). To quantitatively describe the temperature dependence of I_CRAC maintenance, the mean current densities were plotted against temperature in the form of an Arrhenius plot (Fig. 5B) from which E_a values were calculated for temperatures from 24.5 to 22 °C of 5 kcal/mol (Q₁₀ = 1.4) at 120 mV and 2.2 kcal/mol (Q₁₀ = 1.1) at 70 mV. From 22 to 14.5 °C, the E_a values increased to 23 kcal/mol (Q₁₀ = 3.5) at 120 mV and 35 kcal/mol (Q₁₀ = 5.6) at 70 mV.

Although patch clamp recordings of I_CRAC give a direct measure of Ca²⁺ influx, the current is measured under nonphysiological ionic conditions. In addition, as a result of the whole cell patch configuration, dialysis of the cytosol may lead to removal of intracellular components controlling Ca²⁺ influx. These potential problems were minimized by investigating the effect of temperature on store-mediated Ca²⁺ influx in intact cells using Mn²⁺ as a surrogate permeable ion and measuring the rate of the Mn²⁺-induced photoquench of cytosolic fura-2 at different temperatures. The experimental protocol is illustrated in Fig. 6A and shows a greatly increased Mn²⁺ quench in cells depleted at 37 °C than in cells depleted at 16 °C. TG-induced store release led to a substantial increase in Mn²⁺ influx compared with nondepleted cells at 32 °C (Fig. 6B, traces c and d). At 10 °C, however, the photoquench in store-depleted cells was indistinguishable from that in nondepleted cells (Fig. 6B, traces a and b). The differences in the Mn²⁺ photoquench between store-depleted and nondepleted cells at 10, 16, 22, and 32 °C are shown in Fig. 6C. From this basal-subtracted photoquench, the rate of store-mediated Mn²⁺ entry at different temperatures was calculated (Fig. 6D). When the temperature was decreased from 37 to 28 °C, there was a small reduction in the rate of the Mn²⁺ quench, but on decreasing the temperature further from 28 to 10 °C, there was a drastic reduction in the rate of Mn²⁺ influx, with a complete block at 18 °C. To examine the temperature effects on Mn²⁺ influx quantitatively, the rate of the store-mediated Mn²⁺ quench was plotted as an Arrhenius plot (Fig. 6E). The store-mediated Mn²⁺ influx shows a linear decrease in the rate of influx from 37 to 25 °C with a E_a of 2 kcal/mol (Q₁₀ = 1.2). At about 25 °C there is an abrupt increase in the sensitivity of Mn²⁺ influx to temperature. Below this temperature, the E_a increases to 86 kcal/mol (Q₁₀ = 51), reflecting a much higher energy barrier and similar to the increase observed when Ca²⁺ influx was measured as I_CRAC.

To establish whether this temperature sensitivity was a result of its effect on the activation mechanism of influx or on the influx pathway itself, we depleted the stores at 37 °C prior to measuring the rate of Mn²⁺ influx at various temperatures. The difference in the rate of the Mn²⁺ photoquench between store-depleted and nondepleted cells measured in this way is shown in Fig. 7A. The maintenance of Mn²⁺ influx was affected by temperature in a manner similar to activation. The Arrhenius plot (Fig. 7B) of the rate of the store-mediated Mn²⁺ quench component shows a change of E_a from 1.3 kcal/mol (Q₁₀ = 1) between 37 and 23 °C to 119 kcal/mol (Q₁₀ = 61) between 23 and 10 °C. Thus, in contrast to the patch clamp recordings, in which there was found to be a temperature effect on both activation and maintenance, using the Mn²⁺ photoquench we could only detect a temperature-dependent block on the maintenance of the store-mediated influx pathway.

The observed temperature-dependent block of store-mediated Ca²⁺ entry may have been a consequence of temperature sensitivity in the ability of TG to release Ca²⁺ from internal stores. This appears unlikely, however, since the temperature effects on Mn²⁺ influx were equivalent whether TG was added at 37 °C or at lower temperatures (compare Fig. 6, D and E, with Fig. 7, A and B). To further assess whether Ca²⁺ stores were being emptied at low temperatures, we used both fura-2 and a Ca²⁺ minielectrode to measure TG-induced Ca²⁺ release at different temperatures. As illustrated in Fig. 6A, TG was added to cells in nominally Ca²⁺-free external medium at different temperatures, and the peak [Ca²⁺]_i during the subsequent 6 min was assessed using the 340:380-nm fura-2 ratio, corrected (see ``Materials and Methods'') for temperature effects on the K<sub>d</sub> of the dye (Fig. 8C). These results suggest that there was no significant difference in the level of Ca<sup>2+</sup> released at different temperatures by TG over 6 min. Further confirmation of this is illustrated in Fig. 8D. The initial pool size was assessed by measuring the peak [Ca<sup>2+</sup>]<sub>i</sub> following the addition of 5 µM ionomycin at both 18 and 34 °C. The degree of pool depletion (as a percentage of initial pool content) was measured by the ionomycin-induced [Ca<sup>2+</sup>]<sub>i</sub> rise following a 6-min incubation with TG and was not shown to be significantly different at these two temperatures. Since fura-2 measurements only indicate the net balance between Ca<sup>2+</sup> influx and efflux, we resorted to Ca<sup>2+</sup>-sensitive minielectrode measurements on digitonin-permeabilized cells to directly assess TG-induced Ca<sup>2+</sup> store release at different temperatures. The protocol by which this was carried out is shown in Fig. 8A (also see ``Materials and Methods''). The amount of Ca²⁺ released by TG at different temperatures was measured as a percentage of the Ca²⁺ released by ionomycin and is shown in Fig. 8B. There was no significant reduction in the percentage of Ca²⁺ released by TG at lower temperatures. However, this method may also have limitations, since ionomycin may show some temperature sensitivity. Taken together, these two independent methods of testing the temperature sensitivity of TG-induced store release suggest that there is no significant difference in the degree of store release within the temperature range studied, and, hence, the temperature effects on store-mediated Ca²⁺ and Mn²⁺ influx are unlikely to be due to incomplete store release.

Lowering the temperature of cells will slow down many metabolic processes, including ATP synthesis. Recent work suggests that ATP depletion by metabolic inhibitors can block store-dependent Ca²⁺ influx (Gamberucci et al., 1994; Marriott and Mason, 1995). To test whether the temperature-dependent inhibition of Ca²⁺ influx was due to a depletion of cytosolic ATP, we studied the effect of temperature on the cytosolic ATP concentration. Cytosolic ATP concentrations at different temperatures are shown in Fig. 9. Contrary to expectations, ATP levels increased at lower temperatures, possibly due to a reduction in ATP hydrolysis. It seems unlikely, therefore, that the temperature-dependent block of Ca²⁺ influx is due to depletion of cytosolic ATP.

DISCUSSION

The present study demonstrates that, in KU-812 cells, store-regulated Ca²⁺ and Mn²⁺ entry is strongly dependent on temperature. It is clear from patch clamp recordings that both the activation of I_CRAC and its maintenance are temperature-sensitive. The specific temperature effect on activation is apparent from the fact that development of I_CRAC is blocked at 17 °C, although once activated, lowering the temperature even to 14 °C does not totally inhibit the current. The temperature effects on the maintenance of I_CRAC are demonstrated by the nonlinear decrease when the temperature is lowered, with an abrupt transition around 22 °C. Although Mn²⁺ quench measurements also showed equivalent temperature-dependent decreases in influx with an abrupt transition temperature, we could only detect temperature effects on maintenance of influx, since these were equivalent regardless of whether the temperature was changed before or after store release.

This discrepancy between the two methods may be due to a number of factors. In contrast to Mn²⁺ influx measurements, patch clamp recordings will lead to dialysis of the cytosol with two possible consequences. The use of internal solutions low in Na⁺, K⁺, and Cl and high in Cs⁺ may alter temperature-dependent lipid rearrangements, which could explain the incomplete inhibition of I_CRAC seen on cooling following activation at 24 °C. In addition, any soluble factors that may inhibit I_CRAC will be dialyzed out. One such factor, ATP, which will be at much higher levels in intact cells, has recently been shown to inactivate I_CRAC possibly via protein kinase C activation (Parekh and Penner, 1995). This may explain the complete temperature-dependent inhibition of Mn²⁺ entry following store depletion at 37 °C.

The inhibitory effects of temperature on Ca²⁺ influx are unlikely to be due to partial emptying of stores by TG at lower temperatures. Both fura-2 and the Ca²⁺-sensitive minielectrode measurements demonstrate that, following a 6-min incubation with TG, the Ca²⁺ released from stores did not vary significantly over the temperature range studied (10-37 °C). Furthermore, the temperature-dependent inhibition of Mn²⁺ influx was equivalent whether TG addition was carried out at low temperatures or at 37 °C, suggesting equivalent store depletion after 6 min at different temperatures.

Another concern was whether the observed temperature-dependent block of store-dependent Ca²⁺ influx was due to depletion of cytosolic ATP, since Gamberucci et al. (1994) have shown that a 5% reduction in cytosolic ATP concentration can inhibit store-mediated Ca²⁺ influx by 50%. However, our results show that cytosolic ATP concentrations at temperatures at which I_CRAC and Mn²⁺ were blocked (4 mM at 17-18 °C) were in fact greater than at higher temperatures (3 mM at 37 °C), at which the influx was fully activated, implying a temperature-dependent reduction in ATP hydrolysis.

The main finding of the present study is the nonlinear reduction of I_CRAC and Mn²⁺ influx observed when the temperature was lowered. Most ion channels are known to be sensitive to temperature but exhibit a linear change in activity with change in temperature and have low activation energies (Bamberg and Laüger, 1974; Zeidel et al., 1992). Among Ca²⁺ channels, the voltage-gated channels from ventricular myocytes (Cavalie et al., 1985) and sensory neurons (Nobile et al., 1990) also show a linear increase in current with increasing temperature due to an increase in open channel probability. However, some ion channels, including the actylcholine channel (Fischbach and Lass, 1978) and Ca²⁺ channels in neuroblastoma cells (Narahashi et al., 1987), show a nonlinear decrease in conductance when the temperature is lowered, with a transition around 20 °C, and it has been suggested that such transition effects are indicative of membrane-dependent regulation of channel gating (Romey et al., 1980). The nonlinearity in the Arrhenius plots of various membrane functions have been correlated with phase transition or lateral phase separation of membrane phospholipids (Raison 1972; Linden et al., 1973; Warren et al., 1975; Chapman, 1975). In the present study, the observed transition of I_CRAC and Mn²⁺ influx at 21-24 °C indicates that this influx pathway may also be in close association with a lipid membrane environment. A similar transition of Mn²⁺ influx at 21 °C has also been observed in rat parotid acinar cells (Lockwich et al., 1994). The store-mediated influx pathway shows a much greater increase in E_a compared with the acetylcholine channel or the neuroblastoma channels. It should be also noted that such transitional changes also apply to other membrane transport systems, such as transporters and pumps (Rega, 1986), and in view of the fact that the store-dependent Ca²⁺ influx pathway has been shown to be of very low conductance with no single channel noise, the possibility of a transporter being involved in capacitative Ca²⁺ influx should not be ruled out. The possibility also exists that abrupt breaks in Arrhenius plots may reflect intrinsic changes in the protein conformation independent of changes in the membrane phospholipid (Dean and Tanford, 1978; Sondergaard, 1979; Hoffman et al., 1979). This may be occurring in the store-dependent Ca²⁺ influx pathway, in which a possible conformational coupling between stores and plasma membrane has been postulated (Irvine, 1992; Berridge, 1995).

The other interesting finding is that I_CRAC activation is completely inhibited at 17 °C but that, once activated, the current can be maintained even at lower temperatures. These observations suggest that, unlike the effect of temperature on Mn²⁺ entry, there is an additional effect of temperature specifically on I_CRAC activation. Although this could be a consequence of intracellular dialysis, it is consistent with the possible involvement of vesicular transport in the activation pathway of I_CRAC. It is established that low temperatures, between 15 and 22 °C, block the in vivo transport of membrane constituents at different points along the endoplasmic reticulum-golgi apparatus-plasma membrane pathway (Matlin and Simons 1983; Tartakoff, 1986; Saraste et al., 1986; Moreau and Cassagne, 1994). This transition temperature is dependent on the various chain lengths of the phospholipid, and typically in animal cells, the golgi apparatus-plasma membrane pathway mediates the transfer of C₂₀-C₂₄-containing phospholipids, which is blocked at 16 °C (Moreau and Cassagne, 1994). Furthermore, this may also explain the fact that the maintenance of I_CRAC, although reduced, is not totally inhibited by lowering the temperature. Once the activation of the influx pathway via a vesicular transport process has been achieved (recruitment of channels or regulators of channels to the plasma membrane), then blocking this process should not inhibit Ca²⁺ influx, since the channels, or channel regulators, are already on the plasma membrane.

In conclusion, we have provided the first direct evidence demonstrating that both activation and maintenance of the store-mediated Ca²⁺ influx pathway are exquisitely temperature-sensitive, suggesting a very intimate association with the lipid membrane environment.