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• W2017276599 abstract "The involvement of phosphatidylcholine-specific phospholipase C (PC-PLC) and D (PC-PLD) in the regulation of the thapsigargin-induced Ca2+ increase was investigated. Pretreatment of human lymphocytes with the PC-PLC inhibitors D609 or U73122 enhanced the thapsigargin-induced Ca2+ influx. By contrast, no effect was observed in the presence of phospholipase D inhibitor butanol. Addition of exogenous PC-PLC but not PC-PLD to lymphocytes prestimulated with thapsigargin led to a decrease of intracellular Ca2+. In addition, thapsigargin was shown to release diacylglycerol (DAG) from cellular phosphatidylcholine pools. The thapsigargin-induced DAG formation was inhibited by U73122 and D609 but not by butanol. Moreover, no formation of the PC-PLD activity marker phosphatidylbutanol was detected. Thapsigargin-induced DAG formation was dependent on the Ca2+ entry, as it was abolished in the absence of extracellular Ca2+ or in the presence of Ni2+. Further investigations demonstrated that the inhibition of the cellular DAG target, protein kinase C (PKC), enhanced thapsigargin-induced Ca2+ increase, whereas direct PKC activation had an inhibitory effect. Taken together, our results reveal the involvement of PC-PLC in the regulation of the thapsigargin-induced Ca2+ increase and point to the existence of a physiologic feedback mechanism activated by Ca2+ influx and acting via consecutive activation of PC-PLC and PKC to limit the rise of intracellular Ca2+. The involvement of phosphatidylcholine-specific phospholipase C (PC-PLC) and D (PC-PLD) in the regulation of the thapsigargin-induced Ca2+ increase was investigated. Pretreatment of human lymphocytes with the PC-PLC inhibitors D609 or U73122 enhanced the thapsigargin-induced Ca2+ influx. By contrast, no effect was observed in the presence of phospholipase D inhibitor butanol. Addition of exogenous PC-PLC but not PC-PLD to lymphocytes prestimulated with thapsigargin led to a decrease of intracellular Ca2+. In addition, thapsigargin was shown to release diacylglycerol (DAG) from cellular phosphatidylcholine pools. The thapsigargin-induced DAG formation was inhibited by U73122 and D609 but not by butanol. Moreover, no formation of the PC-PLD activity marker phosphatidylbutanol was detected. Thapsigargin-induced DAG formation was dependent on the Ca2+ entry, as it was abolished in the absence of extracellular Ca2+ or in the presence of Ni2+. Further investigations demonstrated that the inhibition of the cellular DAG target, protein kinase C (PKC), enhanced thapsigargin-induced Ca2+ increase, whereas direct PKC activation had an inhibitory effect. Taken together, our results reveal the involvement of PC-PLC in the regulation of the thapsigargin-induced Ca2+ increase and point to the existence of a physiologic feedback mechanism activated by Ca2+ influx and acting via consecutive activation of PC-PLC and PKC to limit the rise of intracellular Ca2+. In human lymphocytes, as well as in other electrically non-excitable cells, receptor stimulation results in a transient increase in the cytosolic free calcium concentration ([Ca2+] i). [Ca2+] i increase primarily occurs due to the calcium release from inositol 1,4,5-trisphosphate-sensitive intracellular pools and is generally accompanied by an entry of extracellular calcium mediated by the activation of receptor-operated or second messenger-activated Ca2+ channels (1Berridge M.J. Nature. 1993; 361: 315-325Crossref PubMed Scopus (6136) Google Scholar). In the hypothesis referred to as capacitative model, calcium influx is controlled by the filling state of intracellular calcium pools (2Putney J.W. Cell Calcium. 1990; 11: 611-624Crossref PubMed Scopus (1255) Google Scholar, 3Mason M.J. Mahaut -Smith M.P. Grinstein S. J. Biol. Chem. 1991; 266: 10872-10879Abstract Full Text PDF PubMed Google Scholar, 4Hoth M. Penner R. Nature. 1992; 355: 353-356Crossref PubMed Scopus (1471) Google Scholar). The mechanisms by which capacitative calcium influx is elicited are not clear. Depletion of cellular calcium pools may trigger release of a diffusible cytoplasmic messenger which in turn opens transmembrane Ca2+ channels (5Randriamampita C. Tsien R.Y. Nature. 1993; 364: 809-814Crossref PubMed Scopus (783) Google Scholar, 6Xu X. Star R.A. Tortorici G. Muallem S. J. Biol. Chem. 1994; 269: 12645-12653Abstract Full Text PDF PubMed Google Scholar). One approach to study the role of intracellular Ca2+ pools for the Ca2+ entry utilizes inhibitors of the Ca2+-ATPases in the store membrane. Thapsigargin has been shown to selectively inhibit Ca2+-ATPases in the endoplasmic reticulum without affecting Ca2+-ATPases in the plasma membrane (7Thastrup O. Cullen P.J. Drobak B.K. Hanley M.R. Dawson A.P. Proc. Natl. Acad. Sci. U. S. A. 1989; 87: 2466-2470Crossref Scopus (2978) Google Scholar, 8Sagara Y. Fernandez-Belda F. de Meis L. Inesi G. J. Biol. Chem. 1992; 267: 12606-12613Abstract Full Text PDF PubMed Google Scholar). In contrast to receptor agonists that initiate rapid formation of inositol 1,4,5-trisphosphate, thapsigargin depletes Ca2+ pools solely by preventing Ca2+ reuptake (9Takemura H. Hughes A.R. Thastrup O. Putney J.W. J. Biol. Chem. 1989; 264: 12266-12271Abstract Full Text PDF PubMed Google Scholar). In a variety of cells including human lymphocytes the thapsigargin-mediated depletion of calcium stores leads to a sustained elevation of [Ca2+] i supported by the entry of extracellular Ca2+ (10Kass G.E.N. Duddy S.K. Moore G.A. Orrenius S. J. Biol. Chem. 1989; 264: 15192-15198Abstract Full Text PDF PubMed Google Scholar, 11Kwan C.Y. Takemura H. Obie J. Thastrup O. Putney J.W. Am. J. Physiol. 1990; 258: C1006-C1015Crossref PubMed Google Scholar, 12Llopis J. Chow S.C. Kass G.E.N. Gahm A. Orrhenius S. Biochem. J. 1991; 277: 553-556Crossref PubMed Scopus (64) Google Scholar, 13Mason M.J. Garcia-Rodriguez C. Grinstein S. J. Biol. Chem. 1991; 266: 20856-20862Abstract Full Text PDF PubMed Google Scholar, 14Demaurex N. Lew D.P. Krause K.H. J. Biol. Chem. 1992; 267: 2318-2324Abstract Full Text PDF PubMed Google Scholar, 15Conroy L.A. Merrit J.E. Hallam T.J. Biochem. J. 1994; 303: 671-677Crossref PubMed Scopus (4) Google Scholar). Although cellular phospholipases are crucial for the regulation of numerous physiological processes, their role in maintaining intracellular calcium homeostasis is little understood. The early phase of Ca2+ increase due to Ca2+ release from inositol 1,4,5-trisphosphate-sensitive stores is thought to involve activation of the receptor-coupled, phosphoinositide-specific phospholipase C. The role of this enzyme in producing the sustained phase of Ca2+ increase which is caused by the transmembrane Ca2+ influx is, however, less clear (16Bianchini L. Todderud G. Grinstein S. J. Biol. Chem. 1993; 268: 3357-3363Abstract Full Text PDF PubMed Google Scholar, 17Yule D.I. Williams J.A. J. Biol. Chem. 1992; 267: 13830-13835Abstract Full Text PDF PubMed Google Scholar, 18Heemskerk J.W.N. Feijge M.A.H. Sage A.O. Walter U. Eur. J. Biochem. 1994; 223: 543-551Crossref PubMed Scopus (43) Google Scholar). There is also evidence pointing to the possible role of phospholipase A2in modulating Ca2+ entry following depletion of intracellular Ca2+ stores (19Tornquist K. Ekokoski E. Forss L. J. Cell. Physiol. 1994; 160: 40-46Crossref PubMed Scopus (12) Google Scholar, 20Wolf M.J. Wang J. Turk J. Gross R.W. J. Biol. Chem. 1997; 272: 1522-1526Abstract Full Text Full Text PDF PubMed Scopus (111) Google Scholar). To our knowledge, no data exist on the role of phosphatidylcholine-specific phospholipases in the regulation of intracellular Ca2+ homeostasis. Therefore, the aim of the present study was to investigate the involvement of phosphatidylcholine-specific phospholipase C (PC-PLC) 1The abbreviations used are: PC-PLC, phosphatidylcholine-specific phospholipase C; PC-PLD, phosphatidylcholine-specific phospholipase D; DAG, diacylglycerol; PtdOH, phosphatidic acid; PtdInsP2, phosphatidylinositol bisphosphate; PtdInsP, phosphatidylinositol monophosphate; PtdIns, phosphatidylinositol; PtdBut, phosphatidylbutanol; PtdChol, phosphatidylcholine; PMA, phorbol myristate acetate. 1The abbreviations used are: PC-PLC, phosphatidylcholine-specific phospholipase C; PC-PLD, phosphatidylcholine-specific phospholipase D; DAG, diacylglycerol; PtdOH, phosphatidic acid; PtdInsP2, phosphatidylinositol bisphosphate; PtdInsP, phosphatidylinositol monophosphate; PtdIns, phosphatidylinositol; PtdBut, phosphatidylbutanol; PtdChol, phosphatidylcholine; PMA, phorbol myristate acetate. and phosphatidylcholine-specific phospholipase D (PC-PLD) in the regulation of the calcium entry following depletion of Ca2+ stores with thapsigargin. Our results demonstrate that PC-PLC, but not PC-PLD, plays an important role in the down-regulation of Ca2+ influx in human lymphocytes. Tricyclodecan-9-yl xantogenate (D609), Fura-2-AM, propranolol, and phospholipid standards were from Sigma, Deisenhofen, Germany. {3-[1-[3-(Amidinothio)propyl-1H-indoyl-3-yl]-3-(1-methyl-1H-indoyl-3-yl)maleimide methane sulfonate} (Ro-31–8220), 1-(6((17β-3-methoxyestra-1,3,5(10)-trien-17-yl)amino)hexyl)-1H-pyrrole-2,5-dione (U73122), phorbol myristate acetate (PMA), and thapsigargin were purchased from Calbiochem, Bad Soden, Germany. 4α-PMA was obtained from Biomol, Hamburg, Germany. [1-14C]Arachidonic acid and l-lyso-3-phosphatidylcholine [1-14C]palmitoyl were from Amersham, Braunschweig, Germany, and [32P]orthophosphoric acid was from NEN Life Science Products, Dreieich, Germany. Phospholipase C (from Bacillus cereus) and phospholipase D (from Streptomyces chromofuscus) were purchased from Boehringer, Mannheim, Germany. Silica Gel 60 plates and solvents for thin layer chromatography were obtained from Merck, Darmstadt, Germany. Ficoll (Lymphoprep) was obtained from Nycomed, Uppsalla, Sweden. Autoradiography was performed with Kodak X-Omat film (Eastman Kodak). Ultima GOLD scintillation mixture was provided by Packard, Frankfurt, Germany. Lymphocytes were obtained form heparinized blood of 12 different donors according to previously described methods (21Tepel M. Schlotmann R. Barenbrock M. Kisters K. Klaus T. Spieker C. Walter M. Meyer C. Bretzel R.G. Zidek W. Circ. Res. 1995; 77: 1024-1029Crossref PubMed Scopus (27) Google Scholar). Briefly, blood was centrifuged at 240 ×g for 15 min, and the upper two-thirds of the supernatant were aspirated. The remaining blood was mixed 1:1 with Hanks' balanced salt solution containing 136 mm NaCl, 5.4 mmKCl, 0.44 mm KH2PO4, 0.34 mm Na2HPO4, 1.0 mmCaCl2, 5.6 mmd-glucose, and 10 mm HEPES, pH 7.4. Lymphocytes were prepared after centrifugation of blood on a Ficoll gradient (Ficoll 5.6% (w/v), density 1.077 g/ml). The lymphocyte interphase was carefully aspirated, washed three times (400 × g for 5 min), and resuspended in Hanks' balanced salt solution. The lymphocyte viability was greater than 95% as determined by the trypan blue exclusion test. Intracellular calcium measurements were performed according to the established method (22Tepel M. Kühnapfel S. Theilmeier G. Teupe C. Schlotmann R. Zidek W. J. Biol. Chem. 1994; 269: 26239-26242Abstract Full Text PDF PubMed Google Scholar). Lymphocytes were loaded with 2.5 μm Fura-2 AM for 45 min at 37 °C. The lymphocyte suspension was then washed twice (240 × g, 15 min) to remove unincorporated dye and adjusted to a final concentration of 1 × 106/ml. Fluorescence was recorded at 510 nm (bandwidth 10 nm) with excitation wavelengths of 340 and 380 nm (bandwidth 10 nm) using a fluorescence spectrophotometer (model 2000, Hitachi Ltd, Tokyo, Japan). The intracellular calcium concentration was calculated as described previously (22Tepel M. Kühnapfel S. Theilmeier G. Teupe C. Schlotmann R. Zidek W. J. Biol. Chem. 1994; 269: 26239-26242Abstract Full Text PDF PubMed Google Scholar). Briefly, the maximum fluorescence was obtained after addition of 1.0 mmdigitonin. The minimum fluorescence was obtained after addition of 5 mm EGTA. The ratio out of the measured fluorescence values was calculated. [Ca2+]] i was obtained according to Equation 1 by Grynkiewicz et al. (23Grynkiewicz G. Poenie M. Tsien R.Y. J. Biol. Chem. 1985; 260: 3440-3450Abstract Full Text PDF PubMed Scopus (80) Google Scholar). [Ca21]i=K∗(R−Rmin)/(Rmax−R)Equation 1 where R min stands for the ratio in calcium-free solution, R max for the ratio at calcium saturation, and K* for K d ×F min2/F max2, the latter representing the fluorescence maximum and minimum at 380 excitation.K d of Fura-2 was set to be 224 nmol/liter. Phospholipid analysis was done as described previously (24Nofer J.-R. Walter M. Kehrel B. Seedorf U. Assmann G. Biochem. Biophys. Res. Commun. 1995; 207: 148-154Crossref PubMed Scopus (33) Google Scholar, 25Walter M. Reinecke H. Nofer J.-R. Seedorf U. Assmann G. Arterioscler. Thromb. Vasc. Biol. 1995; 15: 1024-1029Crossref Scopus (53) Google Scholar). For metabolic lipid radiolabeling the lymphocyte suspension was incubated at 37 °C for 90 min with the following compounds: 0.2 μCi/ml [1-14C]arachidonic acid, 0.2 μCi/ml [1-14C]lyso-3-phosphatidylcholine, or 100 μCi/ml [32P]orthophosphoric acid. In the latter case, phosphate was replaced by carbonate in the incubation buffer. The lymphocytes were then washed, adjusted to a final concentration of 1 × 106 cells/ml, and stimulated with the desired agonist. At different time points 0.5-ml aliquots were withdrawn and added to 1.5 ml of ice-cold chloroform/methanol (2:1, v/v) or chloroform/methanol/hydrochloric acid (2:1:0.01, v/v) in the case of32P labeling. The phases were split by adding 0.5 ml of chloroform and 0.5 ml of water. The samples were centrifuged at 3000 rpm for 5 min, and lipid phases were collected. Water-soluble phases were extracted once more with 1.5 ml of chloroform. Lipid phases were then combined, dried under nitrogen, dissolved in 0.3 ml of hexane, and stored at −70 °C until analyzed. In most experiments, a double one-dimensional TLC as described by Gruchalla et al. (26Gruchalla R.S. Dingh T.T. Kennerley D.A. J. Immunol. 1990; 6: 2334-2342Google Scholar) was used to separate phospholipids and neutral lipids of interest. In this approach, a series of samples was spotted 12 cm from the bottom of the plate. To resolve labeled neutral lipids (DAG, fatty acids, and triglycerides) from phospholipids that remained at the origin, the plates were twice developed in toluene/ether/ethanol/triethylamine (100:80:4:2, by volume). After the first run to 20 cm, the plates were thoroughly dried and developed a second time with the same solution to 16 cm. Plates were then cut 0.8 cm above the origin (i.e. 12.8 cm above the lower edge of the plate), rotated 180°, and developed to the top with chloroform/methanol/ammonium hydroxide (65:35:5, by volume). After they were dried thoroughly, autoradiography was performed with Kodak X-Omat film for 10–14 days. Radioactive bands were cut from the silica plates, placed in scintillation vials containing 10 ml of Ultima-Gold scintillation fluid, and quantitated by liquid scintillation counting in a 1214 scintillation counter (LKB, Bromma, Sweden). The identities of labeled bands were determined based on R F values obtained for authentic neutral lipids and phospholipids visualized by iodine staining. For determination of phosphatidylinositols [32P]phospholipids were developed with chloroform/methanol/acetone/acetic acid/water (60:20:23:18:12, by volume) using potassium oxalate-impregnated Silica 60 plates. Bands corresponding to PtdIns, PtdInsP, PtdInsP2, or PtdChol were cut out from the silica plates, placed in scintillation vials, and quantitated as described above. The identities of labeled bands were determined based on R F values obtained for authentic neutral lipids and phospholipids. PC-PLD hydrolyzes phospholipids to yield the free polar head groups choline and PtdOH. In the presence of primary alcohols, however, PC-PLD catalyzes a phosphatidyl transfer reaction producing phosphatidyl alcohols. Since the transphosphatidylation is catalyzed solely by PC-PLD, the production of phosphatidyl alcohols is an unequivocal marker for involvement of this enzyme. For the examination of PC-PLD activity lymphocytes were labeled with 0.2 μCi/ml [1-14C]lyso-3-phosphatidylcholine. Butan-1-ol was added 5 min prior to agonist addition. Radiolabeled phospholipids were extracted as described above and analyzed using Silica 60 plates developed with ethyl acetate/2,2,4-trimethylpentane/acetic acid (9:5:2 by volume) as described by van der Meulen and Haslam (27van der Meulen F. Haslam R.J. Biochem. J. 1990; 271: 693-700Crossref PubMed Scopus (60) Google Scholar). In some experiments the amount of PtdBut formed was examined using the double one-dimensional system as described above. The optimal concentration of butanol was determined by performing a butanol concentration curve using 1 μm PMA as an agonist. As shown in Fig.1 the most effective PtdBut production was seen at butanol concentrations between 0.3 and 0.6% (v/v) and was accompanied by a decrease in PtdOH formation. Each experiment was performed in duplicate and repeated 3–5 times, as indicated in appropriate legends. Unless otherwise indicated, data represent the mean from duplicate determination in a representative experiment. To avoid possible bias due to the variability of lymphocyte populations obtained from different donors, control and drug responses were determined in the same cell preparations. For calcium measurements data are presented as means ± S.E. The groups were compared with the non-parametric Wilcoxon-Mann-Withney test using the computer software Instat 2.02 (GraphPAD). Two-tailed p values less than 0.05 were considered to be significant. [Ca2+] i was measured in intact human lymphocytes using the calcium-sensitive fluorescent dye Fura-2. The resting [Ca2+] i level in these cells averaged 94 ± 8 nm (n = 34). The resting [Ca2+] i level did not significantly change at least over 400 s (n = 23). As illustrated in Fig.2, addition of 5 μmthapsigargin resulted in a time-dependent increase in intracellular calcium. Within 200 s [Ca2+] i increased by 228 ± 27 nm (n = 34) over the resting level. Next, the effects of the PC-PLC-inhibitor D609, the unspecific PLC inhibitor U73122, the indirect PLD inhibitor butanol, and phosphatidic acid phosphohydrolase inhibitor propranolol on the thapsigargin-triggered [Ca2+] i elevation were tested. In the presence of 20 μm D609 or 10 μm U73122 the thapsigargin-induced [Ca2+] i increases were significantly enhanced and averaged 377 ± 54 nm (n = 19;p < 0.02) and 517 ± 99 nm(n = 16; p < 0.01) over the resting level, respectively (Fig. 2). By contrast, preincubation of cells with 0.3% butanol or 50 μm propranolol did not affect thapsigargin-induced [Ca2+] i elevation (Fig. 2). Under these experimental conditions [Ca2+] i rose by 188 ± 24 nm (n = 10; not significant versus thapsigargin alone) and 171 ± 15 nm (n = 6; not significant versus thapsigargin alone). The potentiating effects of D609 and U73122 on the thapsigargin-induced [Ca2+] i elevation were concentration-dependent (Fig. 2). For both compounds the significant enhancement of [Ca2+] i increases was seen at a concentration of about 10 μm. To test whether potentiating effects of U73122 and D609 on thapsigargin-stimulated intracellular calcium elevation occur as a consequence of increased Ca2+ entry, the unidirectional uptake of Mn2+, a Ca2+ surrogate, was determined. Mn2+, which quenches Fura-2 fluorescence, enters cells through physiological pathways, yet is not readily extruded at an appreciable rate. Thus, the rate of fluorescence decrease provides a relative measure of the divalent ion entry. As illustrated in Fig. 3, addition of 5 μm thapsigargin was found to accelerate the rate of Fura-2 quenching, indicating activation of the direct cation permeability pathway at the plasma membrane. When lymphocytes were first treated for 5 min with 20 μm D609 or 10 μm U73122, and subsequently stimulated with thapsigargin, the rate of Fura-2 quenching was considerably more pronounced (Fig. 3). These findings suggest that U73122 and D609 exert their potentiating effects on the thapsigargin-induced [Ca2+] i elevation by increasing trans-plasma membrane calcium influx. Since diacylglycerol is the principal product released upon activation of PLC, we next investigated whether it is formed following stimulation of lymphocytes with 5 μm thapsigargin. The lymphocytes were labeled for 2 h with 0.2 μCi/ml [1-14C]arachidonic acid. Upon this treatment phosphatidylinositol (PtdIns) and phosphatidylcholine (PtdChol) incorporated 42 and 34% of the total phospholipid radioactivity, respectively. As demonstrated in Fig.4, the [14C]DAG level increased by 98 ± 15% (n = 5) within 120 s following addition of 10 μm thapsigargin. Thereafter, [14C]DAG remained elevated over the basal level for at least 300 s. Preincubation of lymphocytes with 20 μmD609 for 5 min reduced the thapsigargin-induced [14C]DAG accumulation to 48 ± 7% (n = 3) at 120 s after stimulation (Fig. 4). Similarly, the reduction of the thapsigargin-induced [14C]DAG formation to 37 ± 6% (n = 3) at 120 s after stimulation was observed in lymphocytes pretreated with 10 μm U73122 (Fig. 4). In contrast to D609 and U73122, no effect on the thapsigargin-induced [14C]DAG formation was noted in lymphocytes pretreated with 0.3% butanol (Fig. 4). Due to the uniform radioactivity incorporation in all major phospholipids, the experiments with [1-14C]arachidonic acid labeling do not allow us to conclude whether [14C]DAG is primarily derived from PtdIns or PtdChol. To elucidate the phospholipid substrate for thapsigargin-stimulated phospholipase activation, we labeled lymphocytes with 0.1 mCi/ml [32P]orthophosphoric acid. As shown in Table I, the radioactivity associated with PtdInsP2, PtdInsP, and PtdIns was not significantly altered following stimulation of lymphocytes with 5 μm thapsigargin. By contrast, 5 μmthapsigargin induced decrease of the radioactivity associated with PtdChol (Fig. 5). This effect was abolished in the presence of 20 μm D609. These results suggest that phosphatidylcholine rather than phosphatidylinositol is the main source for the thapsigargin-induced DAG production.Table IEffect of thapsigargin on phosphatidylinositols in human lymphocytes[32P]PtdlnsP2[32P]PtdlnsPmindlp/ml/106 cells02084 ± 1712507 ± 20512290 ± 2052831 ± 31522203 ± 2542804 ± 30151935 ± 982292 ± 237 Open table in a new tab To investigate further which phospholipid pool serves as a source for DAG during lymphocyte stimulation with thapsigargin, cells were labeled for 2 h with 0.2 μCi/ml [1-14C]lyso-phosphatidylcholine. Under this condition, the majority of the radioactivity incorporated into phospholipids was found in PtdChol with less than 2% incorporated into PtdIns. Addition of 5 μm thapsigargin to the lymphocyte suspension resulted in a substantial increase in [14C]DAG. The maximum response (74 ± 4.1% over the basal level (n = 6)) was attained 60 s after stimulation. The [14C]DAG level remained elevated above the basal value from 60 s onwards (Fig.6 A). In the presence of 0.3% butanol or 50 μm propranolol, [14C]DAG increased by 54 ± 7.6% (n = 3) and 71 ± 11.3%, respectively, within 120 s after stimulation. The amount of [14C]DAG then declined within the next 180 s but remained elevated above the basal level. Preincubation of lymphocytes with 20 μm D609 or 10 μm U73122 virtually abolished the thapsigargin-induced accumulation of [14C]DAG. As shown in Fig. 6 B, the inhibition of the thapsigargin-stimulated [14C]DAG formation by D609 and U73122 was concentration-dependent. For both compounds the inhibitory effect was noted at concentrations above 10 μm. In contrast to [14C]DAG, no increase in [14C]PtdOH was detected following stimulation of [14C]lyso-phosphatidylcholine-labeled lymphocytes with thapsigargin (not shown). In the presence of primary alcohols PC-PLD catalyzes a phosphatidyl transfer reaction yielding poorly metabolized phosphatidyl alcohols. Since trans-phosphatidylation is catalyzed solely by PLD, synthesis of phosphatidyl alcohols is considered to be an unequivocal marker of PLD activation. To confirm the lack of PC-PLD involvement in the thapsigargin-stimulated DAG production, lymphocytes were stimulated with various concentrations of thapsigargin in the presence of 0.3% (v/v) butanol. Phosphatidylbutanol (PtdBut) formation was measured 5 min after stimulation. No significant [14C]PtdBut formation was observed in cells stimulated with thapsigargin at concentrations up to 10 μm (Fig.7). By contrast, marked [14C]PtdBut formation within 5 min after stimulation was observed in the same experiment when PMA was used instead of thapsigargin (Fig. 7). The latter compound stimulates PC-PLD via PKC activation. To study further the role of PC-PLC and PC-PLD in the regulation of the store-operated Ca2+ influx, we examined the effects of exogenous PC-PLC and PC-PLD on the thapsigargin-induced Ca2+ elevation. Fig.8 A demonstrates that the addition of 5 units/ml PC-PLC to lymphocytes resulted in a rapid formation of [14C]DAG, whereas the addition of PC-PLD led to the accumulation of [14C]PtdOH. As shown in Fig.8 B, depletion of intracellular Ca2+ stores with 5 μm thapsigargin for 10 min resulted in an increase of [Ca2+] i by 587 ± 77 nm(n = 24). Subsequent addition of 5 units/ml PC-PLC led to a significant decrease of [Ca2+] i by 309 ± 48 nm (n = 14; p < 0.001 versus control) (Fig. 8, B and C). This effect was completely abolished in the presence of 20 μm D609 (Fig. 8, B and C) or when heat-treated PC-PLC was used instead of the native one (not shown). In contrast to PC-PLC, no significant effect on [Ca2+] i was noted when 10 units/ml PC-PLD were added to lymphocytes prestimulated with thapsigargin (Fig. 8,B and C). Treatment of cells with thapsigargin causes depletion of intracellular calcium stores and triggers influx of extracellular calcium. In the absence of extracellular calcium the thapsigargin-induced [Ca2+] i increase is due to calcium release from intracellular stores. A typical tracing of thapsigargin-triggered [Ca2+] i elevation in lymphocytes resuspended in Ca2+-free medium is shown in Fig.9 A. Under this experimental condition [Ca2+] i rose by 20 ± 4 nm (n = 18; p < 0.01). Similarly, in the presence of 5 mm Ni2+, which blocks divalent cation entry pathways, thapsigargin-induced [Ca2+] i increase was significantly reduced and averaged 83 ± 20 nm (n = 14,p < 0.001). We next investigated whether the thapsigargin-induced DAG formation depends on the store-operated calcium influx. [14C]Arachidonic acid-labeled lymphocytes were suspended in Ca2+-free medium or in Ca2+-containing medium in the presence of 5 mm Ni2+ and stimulated with 5 μm thapsigargin. Fig. 9 Bdemonstrates that under these experimental conditions the thapsigargin-triggered DAG formation was markedly reduced indicating that PC-PLC activation depends on the extracellular calcium entry. Since DAG liberated by PC-PLC is the main physiological activator of PKC, we next investigated the effect of PKC modulation on the thapsigargin-induced Ca2+ increase. This was accomplished using the direct PKC activator, PMA, or the selective PKC inhibitor Ro31-8220. As shown in Fig.10 A, 10 μm PMA markedly inhibited Ca2+ increase induced by 5 μm thapsigargin. Under this experimental condition [Ca2+] i increased by 65 ± 18 nm(n = 6, p < 0.01). By contrast, in the presence of 10 μm Ro31-8220 the thapsigargin-induced [Ca2+] i increase was significantly enhanced and averaged 348 ± 41 nm (n = 12,p < 0.01). The effect of PKC activation on the Ca2+ elevation was further examined in cells prestimulated for 10 min with 5 μm thapsigargin. Under this experimental condition, direct stimulation of PKC with 10 μm PMA led to a gradual decrease of [Ca2+] i by 202 ± 22 nm(n = 6, p < 0.01 versuscontrol) (Fig. 10 B). By contrast, inactive PMA analogue 4α-PMA (10 μm) failed to affect [Ca2+] i in cells prestimulated with thapsigargin (Δ[Ca2+] i 26 ± 29 nm,n = 4, not significant). Furthermore, the PC-PLC-induced decrease of [Ca2+] i was significantly inhibited by the PKC inhibitor Ro31-8220 (Fig.10 B). Under this experimental condition [Ca2+] i decreased by 152 ± 28 nm (n = 9, p < 0.02versus decrease by PC-PLC alone). In the present study we examined the role of phosphatidylcholine-specific phospholipases in the regulation of the store-operated calcium influx. For this purpose we utilized two structurally unrelated inhibitors, D609 and U73122. Whereas D609 is a PC-PLC inhibitor (28Schutze S. Potthof K. Machleidt T. Berkowitz D. Wiegman K. Kronke M. Cell. 1992; 71: 765-776Abstract Full Text PDF PubMed Scopus (964) Google Scholar), the specificity of U73122 is less well defined, and additional effects distinct from inhibition of phospholipases have been observed in the presence of this agent (29Vickers J.D. Pharmacol. Exp. Ther. 1993; 266: 1156-1163PubMed Google Scholar, 30Willems P.H.G.M. Van de Put F.H.M.M. Engbersen R. Bosch R.R. Van Hoof H.J.M. de Pont J.J.H.H.M. Pfluegers. Arch. Eur. J. Physiol. 1994; 427: 233-243Crossref PubMed Scopus (46) Google Scholar). The possible role of PC-PLD for [Ca2+] regulation was studied with butanol. In the presence of butanol PC-PLD forms metabolically inactive phosphatidylbutanol instead of its physiologically relevant product PtdOH. In several experimental systems primary alcohols were shown to blunt PC-PLD-mediated physiological responses (31Haslam R.J. Coorssen J.R. Adv. Exp. Med. Biol. 1993; 344: 149-164Crossref PubMed Scopus (34) Google Scholar, 32Sciorra V.A. Daniels L.W. J. Biol. Chem. 1996; 271: 14226-14232Abstract Full Text Full Text PDF PubMed Scopus (21) Google Scholar, 33Cross M.J. Roberts S. Ridley A.J. Hodgkin M.N. Stewart A. Claesson-Welsh L. Wakelam M.J.O. Curr. Biol. 1996; 5: 588-597Abstract Full Text Full Text PDF Googl" @default.
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• W2017276599 cites W1220599151 @default.
• W2017276599 cites W137310981 @default.
• W2017276599 cites W1451010299 @default.
• W2017276599 cites W145796029 @default.
• W2017276599 cites W1479942921 @default.
• W2017276599 cites W1488868281 @default.
• W2017276599 cites W1490988370 @default.
• W2017276599 cites W1497010214 @default.
• W2017276599 cites W1509232294 @default.
• W2017276599 cites W1511600524 @default.
• W2017276599 cites W1512762418 @default.
• W2017276599 cites W1519576356 @default.
• W2017276599 cites W1531469735 @default.
• W2017276599 cites W155164777 @default.
• W2017276599 cites W1556795890 @default.
• W2017276599 cites W1557233801 @default.
• W2017276599 cites W1590365700 @default.
• W2017276599 cites W1597049622 @default.
• W2017276599 cites W1597612174 @default.
• W2017276599 cites W1598760385 @default.
• W2017276599 cites W1600993549 @default.
• W2017276599 cites W1602363216 @default.
• W2017276599 cites W1615731496 @default.
• W2017276599 cites W1665071864 @default.
• W2017276599 cites W1863780064 @default.
• W2017276599 cites W1905601533 @default.
• W2017276599 cites W1966734816 @default.
• W2017276599 cites W1967791417 @default.
• W2017276599 cites W1970842110 @default.
• W2017276599 cites W1980166477 @default.
• W2017276599 cites W1981793725 @default.
• W2017276599 cites W1983672484 @default.
• W2017276599 cites W1990050800 @default.
• W2017276599 cites W1998606603 @default.
• W2017276599 cites W2002848696 @default.
• W2017276599 cites W2004008970 @default.
• W2017276599 cites W2004668833 @default.
• W2017276599 cites W2006541068 @default.
• W2017276599 cites W2015480034 @default.
• W2017276599 cites W2037672487 @default.
• W2017276599 cites W2041576624 @default.
• W2017276599 cites W2045521563 @default.
• W2017276599 cites W2058301131 @default.
• W2017276599 cites W2063535351 @default.
• W2017276599 cites W2063916296 @default.
• W2017276599 cites W2073619652 @default.
• W2017276599 cites W2106988040 @default.
• W2017276599 cites W2123820258 @default.
• W2017276599 cites W2132811374 @default.
• W2017276599 cites W2133910858 @default.
• W2017276599 cites W2141260882 @default.
• W2017276599 cites W2146506131 @default.
• W2017276599 cites W2163428550 @default.
• W2017276599 cites W2203858401 @default.
• W2017276599 cites W2244677359 @default.
• W2017276599 cites W2252441433 @default.
• W2017276599 cites W2306576403 @default.
• W2017276599 cites W2332590873 @default.
• W2017276599 cites W2338214652 @default.
• W2017276599 cites W2340246652 @default.
• W2017276599 cites W2397741376 @default.
• W2017276599 cites W2403629777 @default.
• W2017276599 cites W2409213589 @default.
• W2017276599 cites W2414393060 @default.
• W2017276599 cites W2799503982 @default.
• W2017276599 cites W567122488 @default.
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