(Received for publication, January 2, 1997, and in revised form, June 20, 1997)
From the Department of Life Science, Pohang University of Science and Technology, Pohang 790-784, Republic of Korea
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.
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
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.
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.
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 FluorescenceCells 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-TrisphosphateIP3 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]cAMPIntracellular 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 ActivityPKA 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.
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.
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.
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.
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.
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.
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.
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.
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.
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.
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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
ATPS, and obtained the same results as in the stimulations with ATP
(data not shown). For example, ATP
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.
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.