Feedback Regulation of ATP-induced Ca2+ Signaling in HL-60 Cells Is Mediated by Protein Kinase A- and C-mediated Changes in Capacitative Ca2+ Entry*

(Received for publication, January 2, 1997, and in revised form, June 20, 1997)

Hyosang Lee , Byung-Chang Suh and Kyong-Tai Kim Dagger

From the Department of Life Science, Pohang University of Science and Technology, Pohang 790-784, Republic of Korea

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES


ABSTRACT

Extracellular ATP increases intracellular Ca2+ ([Ca2+]i) in HL-60 cells. When cells are stimulated with supramaximal concentrations of ATP, although the initial [Ca2+]i increase is similar over a range of 30, 100, and 300 µM ATP, the rate of the return to basal [Ca2+]i level is faster in cells treated with higher concentrations of ATP. This probably results from differences in Ca2+ influx rather than Ca2+ release, since the influx of the unidirectional Ca2+ surrogates Ba2+ and Mn2+ also exhibit similar responses. Furthermore, while 300 µM ATP had an inhibitory effect on the thapsigargin-induced capacitative Ca2+ entry, 30 µM ATP potentiated the response. However, the inhibitory action of 300 µM ATP was blocked by protein kinase C (PKC) inhibitors, such as GF 109203X and chelerythrine, and the potentiating action of 30 µM ATP was blocked by protein kinase A (PKA) inhibitors H89 and Rp-cAMPS. The PKC inhibitors also slowed the decay rate of the Ca2+ response induced by 300 µM ATP, and the PKA inhibitors increased it when induced by 30 µM ATP. In the measurements of PKA and PKC activity, 30 µM ATP activates only PKA, while 300 µM ATP activates both kinases. Taken together, these data suggest that the changes in the ATP-induced Ca2+ response result from differential modulation of ATP-induced capacitative Ca2+ entry by PKC and PKA in HL-60 cells.


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, P2X, P2Y, P2D, P2T, P2Z, and P2U, 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 P2U and P2X1 type receptors (4, 5). We have also shown that extracellular ATP elevates cAMP through a novel type of receptor (6). The P2U 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 P2X1 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 10-8 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

Materials

ATP, UTP, thapsigargin, Triton X-100, Trizma (Tris base), trichloroacetic acid, EGTA, EDTA, sulfinpyrazone, MOPS, sodium fluoride, sodium orthovanadate, sodium pyrophosphate, beta -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 [gamma -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.

Cell Culture

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.

Measurement of Intracellular Ca2+ Level

[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.

Mn2+ Quenching of Fura-2 Fluorescence

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).

Measurement of Inositol 1,4,5-Trisphosphate

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.

Measurement of [3H]cAMP

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.

Assay of PKA Activity

PKA activity was determined by measuring the incorporation of 32P from [gamma -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 [gamma -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 beta -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.

Assay of PKC Activity

PKC activity was measured by determining the incorporation of 32P from [gamma -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 [gamma -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.

Analysis of Data

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

Effect of Extracellular ATP on Cytosolic [Ca2+]i in HL-60 Cells

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.


Fig. 1. ATP-induced intracellular Ca2+ rise in HL-60 cells. A, concentration-dependent curve of Ca2+ increase. Fura-2-loaded cells were treated with various concentrations of ATP in the presence of 2.2 mM extracellular Ca2+, and the peaks of the Ca2+ increases were measured. The experiments were performed three times and the values are the means ± S.E. B, typical changes in Ca2+ rise after treatment with maximal concentrations of ATP in the presence of 2.2 mM extracellular Ca2+. Arrows indicate the addition of various concentrations of ATP to the medium. The data are representative of 15 experiments with similar results. C, Ca2+ decrease time curve of ATP-induced stimulation. We measured the time period in which the Ca2+ level returned to 30% of the peak starting from the initial ATP treatment as indicated in the inset. The experiments were carried out three times and the values are the means ± S.E.
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Effect of Extracellular ATP on Ca2+ Release and IP3 Generation

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.


Fig. 2. ATP-induced Ca2+ release and IP3 generation in HL-60 cells. A, fura-2-loaded cells were treated with 30 µM, 100 µM, and 300 µM ATP as indicated for 90 s in Ca2+-free medium after which 3 mM CaCl2 was added to the medium. The data are representative of 10 experiments with similar results. B, time course of IP3 generation. Cells were treated with 30 µM (open circle) or 300 µM (closed circle) ATP for the indicated times, and the reactions were stopped by the addition of 15% (w/v) trichloroacetic acid containing 10 mM EGTA. IP3 production was measured by competition assay as described in the experimental procedures. The experiments were done three times and the values are the means ± S.E.
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Effect of Extracellular ATP on Mn2+ Quenching and Ba2+ Uptake

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.


Fig. 3. Effect of ATP on Mn2+ quenching and Ba2+ influx. A, fura-2-loaded cells were incubated with 2 mM Mn2+ for 2 min prior to 30, 100, and 300 µM ATP treatment as indicated. The influx of the Mn2+ was measured by the quenching of fura-2 fluorescence excited at 360 nm and emitted at 500 nm. Traces shown are representative of three similar experiments. B, fura-2-loaded cells were stimulated with ATP for 90 s in Ca2+-free medium, then 10 mM Ba2+ was added to the medium. The data are representative of four experiments with similar results.
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Effect of Extracellular ATP on Thapsigargin-induced Capacitative Ca2+ Entry

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.


Fig. 4. Effect of ATP on thapsigargin-induced capacitative Ca2+ entry. Fura-2-loaded cells were treated with 100 nM thapsigargin (Tg) for 10 min in Ca2+-free medium followed by the addition of 3 mM CaCl2 (dotted trace). In solid traces, the treatment with the designated concentrations of ATP was for 1 min prior to the CaCl2 addition. The data are representative of seven experiments with similar results.
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Effect of PKC and PKA Inhibitors on the ATP Activities in [Ca2+]i Rise and Thapsigargin-induced Capacitative Ca2+ Entry

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.


Fig. 5. Effect of PKC and PKA inhibitors on the thapsigargin-induced capacitative Ca2+ entry induced by ATP. A, effects of GF 109203X and chelerythrine on the thapsigargin-induced capacitative Ca2+ entry induced by 300 µM ATP. Fura-2-loaded HL-60 cells were incubated with vehicle (Cont: Me2SO) or the PKC inhibitors 1 µM GF 109203X (GFX) or 1 µM chelerythrine (Chel) for 1 min and then stimulated with 100 nM thapsigargin for 10 min in Ca2+-free medium. Subsequently, 3 mM CaCl2 was added to the medium with (hatched bar) or without (open bar) 300 µM ATP pretreatment for 1 min before the addition of 3 mM CaCl2, and changes in [Ca2+]i were measured. B, effect of H89 and Rp-cAMPS on the thapsigargin-induced capacitative Ca2+ entry induced by 30 µM ATP. Fura-2-loaded HL-60 cells were incubated with vehicle (Cont: Me2SO) or PKA inhibitors, 10 µM H89, or 20 µM Rp-cAMPS for 1 min and then stimulated with 100 nM thapsigargin for 10 min in Ca2+-free medium. Subsequently, 3 mM CaCl2 was added to the medium with (hatched bar) or without (open bar) 30 µM ATP pretreatment for 1 min, and changes in [Ca2+]i were measured. The values are the means ± S.E. A Student's t test was used for comparing the 300 and 30 µM ATP treatment with the untreated control. p < 0.01, compared with the untreated control.
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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.


Fig. 6. Effect of PKC and PKA inhibitors on the ATP-induced capacitative Ca2+ entry. A, effect of H89 on the 30 µM ATP-induced [Ca2+]i rise. Fura-2-loaded HL-60 cells were incubated with vehicle (Cont: Me2SO) or 2 µM H89 for 5 min and then stimulated with 30 µM ATP. The 300 µM ATP-induced control response is presented as a dashed line. B, effects of GF 109203X on the 300 µM ATP-induced [Ca2+]i rise. Fura-2-loaded HL-60 cells were incubated with vehicle (Cont: Me2SO) or 1 µM GF 109203X (GFX) for 5 min and then stimulated with 300 µM ATP. The 30 µM ATP-induced control response is presented as a dashed line. The data are representative of six experiments with similar results.
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Effect of Extracellular UTP on Cytosolic [Ca2+]i in HL-60 Cells

Since it has been shown that P2U 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.


Fig. 7. UTP-induced intracellular Ca2+ rise in HL-60 cells. A, time curve of Ca2+ decrease in UTP-induced stimulation. We measured the time period in which the Ca2+ level returned to 30% of the peak starting from the initial UTP treatment as indicated in Fig. 1C. The experiments were carried out three times and the values are the means ± S.E. B, effects of GF 109203X on the 300 µM UTP-induced [Ca2+]i rise. Fura-2-loaded HL-60 cells were incubated with vehicle (Cont: Me2SO) or 1 µM GF 109203X (GFX) for 5 min and then stimulated with 300 µM UTP. The 30 µM UTP-induced control response is presented as a dashed line. The data are representative of six experiments with similar results.
[View Larger Version of this Image (13K GIF file)]

Effects of ATP on cAMP Generation, IP3 Production, and Protein Kinase Activity

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.


Fig. 8. ATP concentration dependence curve of ATP-induced cAMP (open circles) and IP3 (closed circles) production. [3H]Adenine-loaded cells were stimulated with various concentrations of ATP for 3 min. cAMP production was measured as described under "Experimental Procedures." The experiments were done three times, and the values are the means ± S.E. For the measurement of IP3, HL-60 cells were stimulated with various concentration of ATP for 15 s. The reaction was stopped by the addition of 15% trichloroacetic acid containing 10 mM EGTA. IP3 generation was measured as described under "Experimental Procedures." Three separate experiments were done, and values are the means ± S.E.
[View Larger Version of this Image (23K GIF file)]


Fig. 9. Effects of ATP on protein kinase activity. A, PKA activity induced by different concentrations of ATP. PKA activity was determined after stimulation of cells with 30 or 300 µM ATP for 3 min in Locke's solution as described under "Experimental Procedures." 10 µM cAMP was added to the reaction mixture with Kemptide to determine positive control. The experiments were performed three times in triplicate, and the values are the mean ± S.E. *, p < 0.01, compared with the control. B, PKC activity in the membrane fraction induced by different concentrations of ATP. Cells were treated with 30 and 300 µM ATP or 100 nM PMA for 3 min in Locke's solution, and membrane fraction was prepared to measure the PKC activity as described under "Experimental Procedures." The experiments were carried out twice in triplicate and the values are the means ± S.E. **, p < 0.01, compared with the control.
[View Larger Version of this Image (38K GIF file)]

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 P2X1 in HL-60 cells. The current through the P2X1 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 ATPgamma S, and obtained the same results as in the stimulations with ATP (data not shown). For example, ATPgamma S 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 ICRAC 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.


FOOTNOTES

*   This work was supported by grants from Pohang University of Science and Technology/BSRI Special Fund, the Hallym Academy of Sciences, Hallym University, the Korea Science and Engineering Foundation (KOSEF 95-0401-02), and the Basic Science Research Institute Program (Project BSRI-96-4435) from the Ministry of Education.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Dagger    To whom correspondence should be addressed: Dept. Life Science, POSTECH, San 31, Hyoja dong, Pohang 790-784, Republic of Korea. Tel.: 562-279-2297; Fax: 562-279-2199; E-mail: ktk{at}vision.postech.ac.kr.
1   The abbreviations used are: [Ca2+]i, intracellular free Ca2+ concentration; CRAC, Ca2+ release-activated Ca2+ channel; fura-2/AM, fura-2 pentaacetoxymethyl ester; IP3, inositol 1,4,5-trisphosphate; PKA, protein kinase A; PKC, protein kinase C; PLC, phospholipase C; PMA, phorbol 12-myristate 13-acetate; MOPS, 3-[N-morpholino]propanesulfonic acid; ATPgamma S, 5'-O-(3-thiotriphosphate); Rp-cAMPS, 3',5'-cyclic monophosphothioate.
2   H. Lee, B.-C. Suh, and K.-T. Kim, unpublished data.

ACKNOWLEDGEMENTS

We are grateful to Dr. J. S. Chun, H. D. Chae, and M. J. Park for valuable discussion of the assays of protein kinase activity. We also thank G. Hoschek and H. M. Kim for editing this manuscript.


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