INTRODUCTION

Extracellular ATP evokes many physiological effects such as platelet aggregation, neurotransmission, inflammation, and muscle contraction in numerous cell types (1). These various effects of ATP are mediated by plasma membrane P₂ purinergic receptors (2). Six subtypes of P₂ purinergic receptors, P_2X, P_2Y, P_2D, P_2T, P_2Z, and P_2U, were identified in pharmacological and functional studies and supported by cloning data (3). It has been reported that in HL-60 cells extracellular ATP increases the intracellular free Ca²⁺ concentration ([Ca²⁺]_i)¹ via plasma membrane P_2U and P_2X1 type receptors (4, 5). We have also shown that extracellular ATP elevates cAMP through a novel type of receptor (6). The P_2U receptor is functionally coupled to phospholipase C (PLC) through pertussis toxin-sensitive and pertussis toxin-insensitive G proteins. PLC hydrolyzes phosphatidylinositol 4,5-bisphosphate to generate inositol 1,4,5-trisphosphate (IP₃) and diacylglycerol. The IP₃ produced increases the [Ca²⁺]_i by mobilizing Ca²⁺ from the intracellular Ca²⁺ stores. This Ca²⁺ mobilization activates the plasma membrane Ca²⁺ influx pathway through Ca²⁺ release-activated Ca²⁺ channels (CRAC) and is termed capacitative Ca²⁺ entry (7, 8). The degree of Ca²⁺ entry is determined by the filling status of the intracellular Ca²⁺ store. The P_2X1 receptor triggers entry of cations; however, it has been shown that the activity is very weak in undifferentiated HL-60 cells. Thus, ATP increases intracellular Ca²⁺ in HL-60 cells by mobilizing it from the intracellular stores and by influx from the extracellular space. We observed a different rate of decrease in the Ca²⁺ response, while the peak level remained the same, when HL-60 cells were stimulated with supramaximal concentrations of ATP. There are several mechanisms responsible for Ca²⁺ removal from the cytosol after the elevation of the [Ca²⁺]_i. These mechanisms include sequestering of Ca²⁺ into intracellular stores, binding to various Ca²⁺-binding proteins, and actions by the Ca²⁺ pump and Na⁺/Ca²⁺ exchanger (9). Among these, the Ca²⁺ pump, which transports ions across the plasma membrane and into intracellular stores, plays a critical role in reducing the elevated [Ca²⁺]_i. The plasma membrane Na⁺/Ca²⁺ exchanger also plays an important role in the control of the intracellular free Ca²⁺ concentration, exchanging three Na⁺ for one Ca²⁺. It appears to have a lower affinity for Ca²⁺ than the plasma membrane Ca²⁺ pump and a high capacity for removing increased Ca²⁺. Thus it operates efficiently when [Ca²⁺]_i is increased beyond 10⁸ M. A number of Ca²⁺-binding proteins are also involved in buffering the cytosolic Ca²⁺ concentration. We studied the mechanism by which the different patterns of decrease in Ca²⁺ occur upon stimulation with supramaximal concentrations of ATP in HL-60 cells. Our results suggest that this difference is not due to the cytosolic Ca²⁺ removal system, but that it is instead mainly due to changes in capacitative Ca²⁺ entry by actions of PKA and PKC, which are differentially activated by ATP itself.

EXPERIMENTAL PROCEDURES

ATP, UTP, thapsigargin, Triton X-100, Trizma (Tris base), trichloroacetic acid, EGTA, EDTA, sulfinpyrazone, MOPS, sodium fluoride, sodium orthovanadate, sodium pyrophosphate, -glycerophosphate, leupeptin, pepstatin A, aprotinin, phenylmethylsulfonyl fluoride, and IP₃ were purchased from Sigma. Fura-2 pentaacetoxymethyl ester was from Molecular Probes (Eugene, OR), and [³H]IP₃ and [-³²P]ATP were from NEN Life Science Products. PMA, chelerythrine, Rp-cAMPS, GF 109203X, KN62, and benzamil were obtained from Research Biochemicals Inc. (Natick, MA), and 1.4-dithiothreitol was from Boehringer Mannheim (Mannheim, Germany). H89 was purchased from Seikagaku Co. (Tokyo, Japan), and P81 phosphocellulose paper was purchased from Whatman. Nonidet P-40 was purchased from U. S. Biochemical Corp.

Human promyelocytic leukemia HL-60 cells were maintained in RPMI 1640 medium (Life Technologies, Inc.) supplemented with 20% (v/v) heat-inactivated bovine calf serum (Hyclone, Logan, UT) plus 1% (v/v) penicillin/streptomycin (Life Technologies, Inc.) under a humidified atmosphere of 5% CO₂ at 37 °C.

[Ca²⁺]_i level was determined using the fluorescent Ca²⁺ indicator fura-2 as reported previously (10). HL-60 cells were incubated with 3 µM fura-2/AM in complete medium at 37 °C with stirring for 60 min. The final concentration of dimethyl sulfoxide (Me₂SO) in the incubation medium was 0.3%. After the loading, cells were washed twice with Locke's solution (154 mM NaCl, 2.2 mM CaCl₂, 5.6 mM KCl, 5.0 mM HEPES, 10 mM glucose, 1.2 mM MgCl₂, pH 7.4) to remove extracellular dye. Sulfinpyrazone was added to the washing solution to a final concentration of 250 µM to prevent dye leakage (11). The fluorescence ratio was recorded at excitation wavelengths of 340 and 380 nm and at an emission wavelength of 500 nm. [Ca²⁺]_i was calculated according to Grynkiewicz et al. (12). In Ca²⁺-free experiments, cells were bathed in Ca²⁺-free Locke's solution (156.2 mM NaCl, 5.6 mM KCl, 5.0 mM HEPES, 10 mM glucose, 1.2 mM MgCl₂, pH 7.4) instead of Ca²⁺-containing Locke's solution.

Cells loaded with fura-2/AM as described above were stimulated with ATP in the presence of 2 mM Mn²⁺, and fluorescence quenching was measured at an excitation wavelength of 360 nm, which is an isosbestic wavelength, and at an emission wavelength of 500 nm (13).

IP₃ mobilization was determined by competition assay with [³H]IP₃ in binding to IP₃-binding protein as described previously (14). To determine IP₃ production, 2 × 10⁶ cells per sample were harvested and stimulated with ATP. The reaction was terminated by the addition of ice-cold 15% (w/v) trichloroacetic acid containing 10 mM EGTA. After centrifugation at 2,000 × g for 5 min, the supernatant was obtained. The trichloroacetic acid was removed by three extractions with diethyl ether. The final extract was neutralized with 200 mM Trizma base and its pH adjusted to about 7.4. 20 µl of the cell extract was added to 20 µl of assay buffer (0.1 M Tris buffer containing 4 mM EDTA) and 20 µl of [³H]IP₃ (0.1 µCi/ml). Finally, 20 µl of binding protein solution was added. The IP₃-binding protein was prepared from bovine adrenal cortex according to the method of Challiss et al. (15). The mixture was incubated for 15 min on ice and then centrifuged at 2,000 × g for 10 min. 100 µl of water and 1 ml of scintillation mixture were added to the pellet to measure the radioactivity. The IP₃ concentration of the sample was determined by comparison with a standard curve and expressed as picomoles/mg of protein. The total cellular protein concentration was measured by the Bradford method after sonication of 2 × 10⁶ cells.

Intracellular cAMP was determined by measuring the formation of [³H]cAMP from [³H]adenine nucleotide pools as we have described previously (16). The cells were grown in complete medium and loaded with [³H]adenine (2 µCi/ml) for 24 h. After the loading, cells were washed twice with Locke's solution and then stimulated with agonist. The reaction was stopped by adding ice-cold 5% (v/v) trichloroacetic acid containing 1 µM cold cAMP. [³H]cAMP and [³H]ATP were separated by sequential chromatography on Dowex AG50W-X4 (200-400 mesh) cation exchanger and neutral alumina column. The increase in intracellular cAMP was calculated as [³H]cAMP/([³H]ATP + [³H]cAMP) × 10³.

PKA activity was determined by measuring the incorporation of ³²P from [-³²P]ATP into the PKA-specific peptide, Leu-Arg-Arg-Ala-Ser-Leu-Gly (Kemptide), using a procedure described previously (17, 18), with some modifications. Briefly, HL-60 cells (1 × 10⁷ cells/tube) were harvested and treated with inhibitor mixture containing 1 µM GF 109203X and 1 µM KN62, inhibitors of PKC and Ca²⁺/calmodulin-dependent protein kinase, respectively, for 5 min. They were then stimulated with 30 or 300 µM ATP for 3 min. After the stimulation, the cells were washed twice with Locke's solution within another 2 min, then resuspended in 100 µl of buffer I containing 20 mM Tris-HCl, pH 7.5, 0.25 M sucrose, 10 mM EGTA, 2 mM EDTA, 0.5 mM phenylmethylsulfonyl fluoride, 10 µg/ml leupeptin, 10 µg/ml pepstatin A, 10 µg/ml aprotinin, 200 µM sodium pyrophosphate, 200 µM sodium fluoride, 1 mM dithiothreitol. The cells were sonicated and centrifuged at 10,000 × g for 10 min at 4 °C. The supernatant was saved as the PKA fraction and used for in vitro PKA activity measurements. All of the following procedures were performed on ice unless stated otherwise. The reaction was initiated by the addition of 10 µl of cell extract to the 30 µl of a test mixture consisting of 10 µl of Mg²⁺/ATP mixture containing 75 mM MgCl₂, 500 µM ATP, 50 µCi of [-³²P]ATP (3,000 Ci/mmol), 10 µl of 500 µM Kemptide, and 10 µl of inhibitor mixture containing 0.02 µM GF 109203X, 0.9 µM KN62. 10 µM cAMP were added to the reaction mixture with Kemptide for a positive control, and 10 µl of buffer II instead of Kemptide was added to determine the endogenous PKA substrate. All assay components were prepared by using buffer II that contained 20 mM MOPS, pH 7.2, 25 mM -glycerol phosphate, 5 mM EGTA, 1 mM sodium orthovanadate, and 1 mM dithiothreitol. The reaction mixture was gently vortexed and placed in a 30 °C water bath for 10 min. Then 25 µl of the reaction mixture was transferred to 1 × 3-cm P81 phosphocellulose strips, which were immediately immersed into 0.75% phosphoric acid. The strips were washed three times with 0.75% phosphoric acid and then dehydrated in 95% ethanol, air-dried, and placed into liquid scintillation vials. The radioactivity was quantified in a Beckman LS 8000 liquid scintillation counter.

PKC activity was measured by determining the incorporation of ³²P from [-³²P]ATP into histone IIIS as described previously (17, 19), with some modifications. HL-60 cells were harvested and treated with inhibitor mixture containing 10 µM H89 and 1 µM KN62 and then stimulated with 30 or 300 µM ATP and 100 nM PMA. After the stimulation, the cells were washed three times with Locke's solution and then resuspended in 200 µl of buffer I, which is described in the PKA assay. The cells were sonicated and centrifuged at 100,000 × g for 1 h at 4 °C, and the pellet was saved as the membrane fraction and then was solubilized with the above buffer I containing 1% Nonidet P-40. The reaction was initiated by the addition of 10 µl of solubilized membrane fraction to the 40 µl of reaction mixture containing 10 µl of 500 µM histone IIIS, 10 µl of inhibitor mixture containing 2 µM PKI, PKA inhibitor peptide and 0.9 µM KN62, 10 µl of 500 nM PMA, and 10 µl of the Mg²⁺/ATP mixture containing 75 mM MgCl₂, 500 µM ATP, and 100 µCi of [-³²P]ATP. All assay components were prepared using buffer II described in the assay of PKA. The reaction mixture was incubated at 30 °C for 10 min, and 25 µl of the reaction mixture was transferred to the P81 phosphocellulose strips. The strips were immersed into the 0.75% phosphoric acid and washed three times for 10 min. After washing, they were rinsed in 95% ethanol, air-dried, and quantified by measuring the radioactivity in a liquid scintillation counter.

Data are summarized as the means ± S.E. EC₅₀ was calculated with the AllFit program (20). We considered differences significant at p < 0.05.

RESULTS

In HL-60 cells, ATP increased the [Ca²⁺]_i in a concentration-dependent manner with maximal and half-maximal effective concentrations (EC₅₀) seen at approximately 10 µM and 85 nM, respectively (Fig. 1A). Fig. 1B illustrates the typical changes in [Ca²⁺]_i observed in fura-2-loaded HL-60 cells stimulated with maximal concentrations of ATP. Initially, the [Ca²⁺]_i increased rapidly to a peak level and then completely returned to the basal Ca²⁺ level, even if the stimulant remained present. Notably, the changes in cytosolic Ca²⁺ exhibited a different desensitization pattern in response to supramaximal concentrations of ATP as compared with the lower concentrations. Although the peak levels were similar, the rate of return to the basal [Ca²⁺]_i level was faster in cells treated with the higher concentration of ATP. This phenomenon is clearly seen in Fig. 1C. We measured the time from peak response of the Ca²⁺ signal to the 70% desensitized [Ca²⁺]_i level as indicated in the inset of Fig. 1C. The data show that the times for return to 30% of the peak level became less in stimulations with increasing concentration of ATP. In other words, the higher the ATP concentration, the faster the return rate. We further analyzed whether changes in the desensitization rate are elicited by Ca²⁺ release from intracellular stores or Ca²⁺ influx from the extracellular space, since ATP increases [Ca²⁺]_i via both pathways.

To measure Ca²⁺ release from the intracellular stores, cells were stimulated with ATP in the absence of extracellular Ca²⁺. Subsequently, 3 mM Ca²⁺ was introduced to the medium to assess the activity of the Ca²⁺ influx. As shown in Fig. 2A, when cells were stimulated with 30, 100, and 300 µM ATP in Ca²⁺-free medium, there were no significant differences in Ca²⁺ release, but large differences occurred in Ca²⁺ influx between the different ATP concentrations. Ca²⁺ influx stimulated by 300 µM ATP was 62.5% less than when stimulated with 30 µM ATP. The data indicate that the differences in the falling state of the Ca²⁺ responses caused by the supramaximal concentration of ATP resulted from changes in Ca²⁺ influx. Since the amount of Ca²⁺ release was small, it is possible that undetectable differences could exist between those stimulations. Thus, we measured the IP₃ production of cells stimulated with ATP. Fig. 2B shows the time course for IP₃ production when cells were stimulated with 30 and 300 µM ATP. At both concentrations, maximal IP₃ generation was obtained after 15 s. At that time, 300 µM ATP generated approximately 2.3 times the amount of IP₃ than 30 µM ATP. Furthermore, the intracellular IP₃ level was more sustained in the stimulation with 300 µM as compared with the stimulation with 30 µM. Thus, from the result of IP₃ production, it seems likely that 300 µM ATP stimulation can cause more Ca²⁺ release than 30 µM ATP or, at least, can trigger the release of a similar amount of Ca²⁺ from the internal stores, while IP₃ produced by 30 µM ATP is enough to maximally mobilize Ca²⁺. Therefore, we conclude that the 300 µM ATP-induced Ca²⁺ release is not less than the 30 µM ATP-induced one and that the difference detected in the Ca²⁺ decay rate is due to changes in the Ca²⁺ influx from the extracellular space.

To test whether differences in the falling state of the Ca²⁺ response are due to modulation of the Na⁺/Ca²⁺ exchanger and Ca²⁺/ATPase activity, we measured Mn²⁺ and Ba²⁺ influx after the addition of ATP. Mn²⁺ and Ba²⁺ are good Ca²⁺ surrogates, since they are not pumped out of the cell, so they can be considered as selective tracers for entry (21, 22). Mn²⁺ uptake was estimated by the quenching of the fura-2 fluorescence when excited at the 360-nm wavelength, which is an isosbestic wavelength and insensitive to variations in Ca²⁺ concentration. Ba²⁺ uptake was estimated by the increase in the fura-2 fluorescence ratio when excited at the 340- and 380-nm wavelength. Fig. 3A shows the fluorescence quenching by Mn²⁺ influx when cells were stimulated with 30, 100, and 300 µM ATP. As 2 mM Mn²⁺ was applied to the medium, it entered the cell slowly. Subsequent stimulation of the cells with 30 µM ATP accelerated the Mn²⁺ entry as compared with the untreated control (dotted trace). 100 µM ATP also accelerated the entry, however, with a slower rate than 30 µM ATP. In contrast, 300 µM ATP stimulation had little effect on fluorescence quenching in comparison with the untreated control. The data indicate that the lower concentration of ATP activates the divalent cation influx. This result was also supported by the Ba²⁺ uptake. To measure Ba²⁺ influx, cells were stimulated with ATP in the absence of external Ca²⁺. When the Ba²⁺ was added to the medium, it caused an increase in the fluorescence intensity reflecting Ba²⁺ uptake. The influx of Ba²⁺ elicited by ATP shows a concentration-dependent pattern with the higher concentrations of ATP triggering less Ba²⁺ uptake. This is similar to the result of Ca²⁺ influx as shown in Fig. 2A. These results suggest that ATP regulates the amount of Ca²⁺ influx, but does not modulate the activity of Na⁺/Ca²⁺ exchanger and Ca²⁺/ATPase. To investigate the involvement of CRAC, we tested the effect of various metal ions on ATP-induced Ca²⁺ signaling, because metal ions are known to block CRAC. Cells were treated with 30 µM La³⁺, Cd²⁺, Co²⁺, or Ni²⁺ for 1 min and then stimulated with ATP in Ca²⁺-containing medium. The difference between the 30 and 300 µM ATP-induced Ca²⁺ signals disappeared in the presence of 30 µM La³⁺, whereas the other metal ions had little or negligible effects (data not shown). The data, thus, suggested that changes in CRAC activity could be the main cause for the rapid desensitization of the Ca²⁺ response induced by higher concentrations of ATP.

In the above experiments, we found that different Ca²⁺ decay rates were caused by changes in Ca²⁺ influx. ATP induces capacitative Ca²⁺ entry through CRAC, which is stimulated by a Ca²⁺ influx factor liberated from the depleted intracellular Ca²⁺ stores by action of IP₃. To study the regulation of capacitative Ca²⁺ entry, we used thapsigargin, which depletes intracellular Ca²⁺ stores by inhibiting the microsomal Ca²⁺/ATPase and induces Ca²⁺ influx (23). The differences in Ca²⁺ influx could also be demonstrated when we measured the effect of ATP pretreatment on thapsigargin-induced capacitative Ca²⁺ entry. As shown in Fig. 4, cells were incubated with 100 nM thapsigargin in a Ca²⁺-free medium, which resulted in a transient [Ca²⁺]_i elevation. After reaching a peak, the [Ca²⁺]_i decreased slowly to the basal level, which reflects the emptying of the intracellular Ca²⁺ stores. The subsequent addition of 3 mM Ca²⁺ to the medium induced a marked and sustained Ca²⁺ rise (dotted trace). Treatment with 300 µM ATP for 1 min prior to the extracellular Ca²⁺ application significantly diminished the thapsigargin-induced capacitative Ca²⁺ entry by 25.7% as compared with the untreated control (dotted trace). 100 µM ATP had also a slightly inhibitory effect. However, 30 µM ATP substantially potentiated the thapsigargin-induced capacitative Ca²⁺ entry to 152.8% over the control cells. These results indicate that ATP has a biphasic effect on the thapsigargin-induced capacitative Ca²⁺ entry linked to its concentration. Therefore, we speculated that ATP itself might potentiate and inhibit capacitative Ca²⁺ entry that it evokes, forming both a positive and a negative feedback loop. At 30 µM, ATP potentiates Ca²⁺ influx, which slows down the desensitization of the [Ca²⁺]_i. Whereas, at 300 µM, ATP inhibits Ca²⁺ influx, which speeds up the desensitization of the [Ca²⁺]_i.

ATP activates PLC and produces IP₃ and diacylglycerol, which subsequently activates PKC. We have also shown that extracellular ATP triggers elevation of cAMP in HL-60 cells (6). To assess the involvement of PKC and PKA in modulation of capacitative Ca²⁺ entry, we used inhibitors specific for those kinases. GF 109203X and chelerythrine, selective PKC inhibitors, were used to characterize the inhibitory or stimulatory effect of ATP on the capacitative Ca²⁺ entry. Fig. 5A shows what effect pretreatment with protein kinase inhibitors has on the Ca²⁺ transient elicited by thapsigargin. 300 µM ATP has a substantial inhibitory effect on thapsigargin-induced capacitative Ca²⁺ entry as seen in Fig. 4. This inhibitory action was antagonized by pretreatment with 1 µM GF 109203X, and Ca²⁺ influx was even potentiated in the presence of GF 109203X. Similar effects were obtained when 1 µM chelerythrine was used in place of GF 109203X. The results suggest that PKC, when activated by 300 µM ATP, inhibits thapsigargin-induced capacitative Ca²⁺ entry.

The involvement of PKA in the 30 µM ATP-induced potentiation of capacitative Ca²⁺ entry was investigated by testing the effect of the PKA inhibitors H89 and Rp-cAMPS. Fig. 5B shows the effect that H89 and Rp-cAMPS has on the enhancement of the thapsigargin-induced capacitative Ca²⁺ entry by 30 µM ATP. In cells treated with H89, this potentiation was blocked. The potentiation also disappeared in 20 µM Rp-cAMP-treated cells. These results suggest the involvement of PKA in the 30 µM ATP-induced enhancement of the capacitative Ca²⁺ entry.

The effects of protein kinase inhibitors on ATP activity in the desensitization pattern of [Ca²⁺]_i were also investigated. Inhibition of PKA by pretreatment with 2 µM H89 significantly accelerated the decay rate of the 30 µM ATP-induced [Ca²⁺]_i level with little effect on the peak Ca²⁺ level (Fig. 6A). In contrast, pretreatment with 1 µM GF 109203X slowed the decay rate induced by 300 µM ATP, resulting in a Ca²⁺ response similar to the 30 µM ATP-evoked response (Fig. 6B). The inhibitory action of 300 µM ATP was slightly enhanced in the presence of PKA inhibitors, while the potentiating effect of 30 µM ATP became even more activated in the presence of PKC inhibitors (data not shown). Thus, the slower decay rate of the 30 µM ATP-induced Ca²⁺ signal may be the result of the potentiating action of PKA as it increases the capacitative Ca²⁺ entry induced by ATP, whereas the rapid decay rate in the 300 µM ATP-induced Ca²⁺ signal might be the result of an inhibitory action by PKC as it blocks the ATP-induced capacitative Ca²⁺ entry. Taken together, the different desensitization rates of the ATP-induced Ca²⁺ signals after peak level could be the result of an interplay between inhibition by PKC and activation by PKA of the capacitative Ca²⁺ entry.

Since it has been shown that P_2U receptors were present and coupled to PLC in HL-60 cells, we treated the cells with UTP and measured the return rate of the [Ca²⁺]_i level. Fig. 7A illustrates the times for return to 30% of the peak level in response to supramaximal concentrations of UTP: the higher the UTP concentration, the faster the return rate. However, although the phenomenon was similar to that of ATP, the rate of the return to the basal [Ca²⁺]_i level was not as remarkable compared with that induced by ATP in Fig. 1C. We further analyzed whether changes in the desensitization rate involve PKC and PKA in the modulation of the UTP-induced capacitative Ca²⁺ entry using kinase inhibitors. Fig. 7B shows that pretreatment with GF 109203X slowed the decay rate induced by 300 µM UTP. However, inhibition of PKA by pretreatment with H89 had no effect on the return rate of the 30 µM UTP-induced [Ca²⁺]_i level (data not shown). The difference in the desensitization pattern between ATP and UTP might result from different activations of effector enzymes, such as PLC and adenylyl cyclase. In HL-60 cells, UTP has no effect on cAMP production (6). Therefore, the results suggest that the effect of PKA increasing the capacitative Ca²⁺ entry is not involved in UTP-treated cells and that PKC alone acts in the desensitization of the UTP-induced Ca²⁺ response.

To assess agonist concentration-dependent differential activation of PKC and PKA, we measured the production of cAMP and IP₃. Fig. 8 shows the production of cAMP and IP₃ induced by various concentrations of ATP. The maximal increase of cAMP was obtained with 300 µM ATP. The EC₅₀ value was 19.2 µM. Particularly, for 30 and 300 µM ATP, respectively, the cAMP levels reached 137.2 ± 9.3 and 190.2 ± 7.7 over the basal cAMP level of 25.3 ± 4.2. The amounts of IP₃ caused by 30 and 300 µM ATP were 27.7 ± 5.7 and 67.0 ± 5.5 pmol/mg of protein, respectively, while the basal IP₃ level was 18.0 ± 3.1. There was only a slight increase in the IP₃ level in the response to 30 µM ATP, whereas the cAMP level was already significantly increased at that concentration. On the other hand, during stimulation with 300 µM ATP, cAMP was produced maximally, and IP₃ was also dramatically increased over the basal IP₃ level, suggesting that both PKA and PKC might be highly activated. We directly measured the activities of the protein kinases induced by different concentrations of ATP. Fig. 9A shows that stimulation with 30 µM ATP induced strong activation of PKA similar to 300 µM ATP. However, stimulation with 30 µM ATP produced a relatively weak activation of PKC (Fig. 9B), indicating that PKA was more significantly activated than PKC during the 30 µM ATP stimulation. However, PKC seems to have a dominant effect during the highly activated state of PKA and PKC as occurs with the 300 µM ATP treatment, because inhibition of the capacitative Ca²⁺ entry was only exhibited during the stimulation with 300 µM ATP.

To investigate the interrelation between PKA and PKC, thapsigargin-induced capacitative Ca²⁺ entry was measured in cells treated simultaneously for 1 min with 100 nM PMA and 30 µM ATP. As shown in Table I, PMA by itself has an inhibitory effect on the thapsigargin-induced capacitative Ca²⁺ entry. Moreover, the 30 µM ATP-induced potentiation of the capacitative Ca²⁺ entry also disappeared when cells were simultaneously treated with PMA. Thus it seems likely that strong activation of PKC has a dominant effect on the capacitative Ca²⁺ entry, even during a state of strongly activated PKA. Similarly, the dominant effect of PKC might cause the rapid decline of the 300 µM ATP-induced Ca²⁺ response.

Table I. The effect of PMA on 30 µM ATP-induced Ca2+ influx HL-60 cells were incubated with 100 nM thapsigargin for 10 min in Ca2+-free medium, then treated with 3 mM CaCl2, and the peak level of [Ca2+]i was measured (control). 30 µM ATP, 100 nM PMA, or 30 µM ATP + 100 nM PMA were added to the medium for 1 min prior to addition of 3 mM CaCl2, respectively, or simultaneously (ATP + PMA). The result is presented as the means ± S.E. Net increase of [Ca2+]i Percentage of control nM % Control 804  ± 28 100 ATP 1229  ± 53a 153 PMA 628  ± 22a 78 ATP + PMA 658  ± 34a 82 a p < 0.01, compared with the control.

DISCUSSION

The present study demonstrates that ATP stimulation with maximal concentrations of ATP causes different decay rates for the Ca²⁺ signal after obtaining similar peak levels if supramaximal concentrations of ATP are used. We suggest that this is a result of homologous feedback regulation of PKC and PKA activation. ATP increases intracellular Ca²⁺ by the release of Ca²⁺ from the intracellular stores and by influx from the extracellular space. Most of the Ca²⁺ increase was caused by capacitative Ca²⁺ entry activated by the store depletion. It has also been reported that ATP activates nonselective cation channels permeable for Ca²⁺ and Na⁺ in dibutyryl cAMP-differentiated HL-60 cells (24). Recently, Buell et al. (5) demonstrated the presence of P_2X1 in HL-60 cells. The current through the P_2X1 was barely detected in undifferentiated HL-60 cells. However, the current was markedly increased in differentiated cells. We cannot exclude the involvement of this nonselective cation channel in the homologous desensitization of the ATP-induced Ca²⁺ response; however, its contribution would be small, since we used undifferentiated cells.

Our data indicate that the differences in the Ca²⁺ signal were caused not by Ca²⁺ release but by influx. However, it is possible that the different rates of desensitization could also be the result of a differential activity of the cytosolic Ca²⁺ removing system. There are two major pathways by which to decrease the [Ca²⁺]_i. One is the pumping out of Ca²⁺ from the cytosol to the cell exterior by Na⁺/Ca²⁺ exchanger and/or by plasma membrane Ca²⁺/ATPase. The other is the pumping of Ca²⁺ into the intracellular stores by Ca²⁺/ATPase. The Na⁺/Ca²⁺ exchanger may not be directly involved in the phenomena of the present study, because stimulation of cells with ATP in Na⁺-free medium or in the presence of the Na⁺/Ca²⁺ exchanger blocker benzamil did not affect the desensitization pattern of the Ca²⁺ responses elicited by supramaximal concentrations of ATP (data not shown). It has been reported that PKC stimulates Ca²⁺ efflux by activation of plasma membrane Ca²⁺/ATPase in neutrophils (25). It seems unlikely that the activation of the Ca²⁺ efflux was involved in the fast return to basal level at higher concentrations of ATP, because unidirectional Ca²⁺ surrogates, Mn²⁺ and Ba²⁺, showed a similar pattern as Ca²⁺.

It has been reported that the capacitative Ca²⁺ entry is blocked by metal ions with an efficiency order of La³⁺ > Zn²⁺ > Cd²⁺ > Be²⁺ = Co²⁺ = Mn²⁺ > Ni²⁺ > Sr²⁺ > Ba²⁺ (7). We found the same desensitization pattern between stimulations with 30 and 300 µM ATP in the presence of La³⁺. This is consistent with the notion that the different desensitization rates of the ATP-induced Ca²⁺ signals resulted from differential feedback regulation of the capacitative Ca²⁺ entry evoked by ATP itself.

Using ATP as an agonist, we had to consider whether a particular cellular response was caused by the activation of a P₂ purinergic receptor per se. ATP is rapidly hydrolyzed to adenosine by extracellular ATPase and nucleotidase (26). But adenosine itself is a potent signaling substance. To assess this potential problem, we tested the hydrolysis-resistant ATP analog, adenosine ATPS, and obtained the same results as in the stimulations with ATP (data not shown). For example, ATPS also showed the differences in the Ca²⁺ decay rate and the biphasic effect on the thapsigargin-induced capacitative Ca²⁺ entry. These results indicate that the changes in the decay rate of elevated [Ca²⁺]_i were due to ATP and not to its metabolite.

In the experiments with UTP, the rate of return to the basal Ca²⁺ level increased as the UTP concentration was raised, but the decay rate was not as prominent as with ATP. Recently, we (6) showed that ATP, but not UTP, elevates the cAMP level in HL-60 cells, maybe through a novel subtype of P₂ receptor. Therefore, the difference in the desensitization pattern between ATP and UTP might be due to the effects of the nucleotides on the activity of the protein kinases. While ATP activates both PKC and PKA, UTP activates only PKC, which would result in a difference in Ca²⁺ signaling.

Until now, little is known about the signaling between the intracellular Ca²⁺ store and the plasma membrane CRAC. It has been proposed that the signal is mediated via Ca²⁺ entry factors, which include calcium influx factor (27, 28), heterotrimeric G protein (29), and small G protein (30), or is mediated via direct interaction between the IP₃ receptor and the plasma membrane Ca²⁺ channel (31). Recently, there were some reports describing the cloning and functional expression of a mammalian homologue to the Drosophila eye-specific trp gene (32-34). It was identified as a Ca²⁺-permeable cation channel that is activated by calcium store depletion. Although it was not clearly shown that TRP is the I_CRAC protein, there is a possibility that TRP should be classified as one of the CRAC family (35). The molecular regulatory mechanism of signaling between the Ca²⁺ store depletion and CRAC is controversial and complicated. There is some evidence that protein phosphorylation is involved in the regulation of capacitative Ca²⁺ entry. In Xenopus oocytes and lymphocytes, protein phosphatase inhibitor potentiates Ca²⁺ influx (36). Tyrosine kinase inhibitor blocks the bradykinin- and thapsigargin-induced Ca²⁺ influx in lymphocytes and in human foreskin fibroblast cells (37, 38). Protein kinase C-dependent phosphorylation plays a key role in the modulation of the capacitative Ca²⁺ entry, too. In the insulin-secreting cell line RINm5F, PKC activates capacitative Ca²⁺ entry (39). On the contrary, PKC stimulation has been shown to inhibit capacitative Ca²⁺ entry in thyroid cells (40). In human neutrophils, formyl-methionyl-leucyl-phenylalanine (41) and PMA (42) inhibited capacitative Ca²⁺ entry, which was mediated by PKC. Capacitative Ca²⁺ entry caused by Drosophila photoreceptor activation is inhibited by PKC as well (43). Here, we suggest that 300 µM ATP preferentially inhibits capacitative Ca²⁺ entry by PKC activation. However, little is known about the involvement of PKA in the capacitative Ca²⁺ entry. It has been reported that activation of PKA had a biphasic effect on Ca²⁺ entry-evoked currents in thapsigargin-treated Xenopus oocytes. Application of dibutyryl cAMP at 1-10 µM inhibited the current, whereas at 1-10 mM potentiated the current (44). We show here that 30 µM ATP preferentially activates capacitative Ca²⁺ entry by relatively strong PKA activation rather than PKC activation. This activation is not the result of the further emptying the intracellular stores, because Ca²⁺ stores may be fully depleted after thapsigargin treatment for 10 min, and no Ca²⁺ increase was detectable upon subsequent ATP treatment. We also tested the effect of prostaglandin E₂ on the thapsigargin-induced capacitative Ca²⁺ entry. Prostaglandin E₂ activates adenylyl cyclase and increases intracellular cAMP concentration, but there were no detectable changes in the Ca²⁺ signal and IP₃ generation in HL-60 cells.² Prostaglandin E₂ also potentiates the thapsigargin-induced capacitative Ca²⁺ entry as shown in the stimulation with 30 µM ATP. This result also suggests that PKA activates the capacitative Ca²⁺ entry in HL-60 cells.

In many cell types, functional effects elicited by extracellular ATP are related to a Ca²⁺ increase. Therefore, the Ca²⁺ increase must be tightly regulated to maintain cellular homeostasis and to exert physiological effects. The fine regulation of the ATP-induced Ca²⁺ signal could be achieved in a feedback mode with PKC and PKA, which are differentially activated according to the extent of stimulation caused by different ATP concentrations.