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The Role of P2Y1 Purinergic Receptors and Cytosolic Ca2+ in Hypotonically Activated Osmolyte Efflux from a Rat Hepatoma Cell Line

2002; Elsevier BV; Volume: 277; Issue: 43 Linguagem: Inglês

10.1074/jbc.m204712200

1083-351X

Pauline R. Junankar, Ari Karjalainen, Kiaran Kirk,

Neonatal Health and Biochemistry

Exposure of HTC rat hepatoma cells to a 33% decrease in extracellular osmolality caused the cytosolic Ca2+ concentration ([Ca2+]i) to increase transiently by ∼90 nm. This rise in [Ca2+]i was inhibited strongly by apyrase, grade VII (which has a low ATP/ADPase ratio) but not by apyrase grade VI (which has a high ATP/ADPase ratio) or hexokinase, indicating that extracellular ADP and/or ATP play a role in the [Ca2+]i increase. The hypotonically induced rise in [Ca2+]i was prevented by the prior discharge of the intracellular Ca2+ store of the cells by thapsigargin. Removal of extracellular Ca2+ or inhibition of Ca2+ influx by 1–10 μm Gd3+depleted the thapsigargin-sensitive Ca2+ stores and thereby diminished the rise in [Ca2+]i. The hypotonically induced rise in [Ca2+]i was prevented by adenosine 2′-phosphate-5′-phosphate (A2P5P) and pyridoxyl-5′-phosphate-6-azophenyl-2′,4′-disulfonate, inhibitors of purinergic P2Y1 receptors for which ADP is a major agonist. Both inhibitors also blocked the rise in [Ca2+]ielicited by addition of ADP to cells in isotonic medium, whereas A2P5P had no effect on the rise in [Ca2+]i elicited by the addition of the P2Y2 and P2Y4 receptor agonist, UTP. HTC cells were shown to express mRNA encoding for rat P2Y1, P2Y2, and P2Y6 receptors. Inhibition of the hypotonically induced rise in [Ca2+]i blocked hypotonically induced K+ (86Rb+) efflux, modulated the hypotonically induced efflux of taurine, but had no significant effect on Cl− (125I−) efflux. The interaction of extracellular ATP and/or ADP with P2Y1purinergic receptors therefore plays a role in the response of HTC cells to osmotic swelling but does not account for activation of all the efflux pathways involved in the volume-regulatory response. Exposure of HTC rat hepatoma cells to a 33% decrease in extracellular osmolality caused the cytosolic Ca2+ concentration ([Ca2+]i) to increase transiently by ∼90 nm. This rise in [Ca2+]i was inhibited strongly by apyrase, grade VII (which has a low ATP/ADPase ratio) but not by apyrase grade VI (which has a high ATP/ADPase ratio) or hexokinase, indicating that extracellular ADP and/or ATP play a role in the [Ca2+]i increase. The hypotonically induced rise in [Ca2+]i was prevented by the prior discharge of the intracellular Ca2+ store of the cells by thapsigargin. Removal of extracellular Ca2+ or inhibition of Ca2+ influx by 1–10 μm Gd3+depleted the thapsigargin-sensitive Ca2+ stores and thereby diminished the rise in [Ca2+]i. The hypotonically induced rise in [Ca2+]i was prevented by adenosine 2′-phosphate-5′-phosphate (A2P5P) and pyridoxyl-5′-phosphate-6-azophenyl-2′,4′-disulfonate, inhibitors of purinergic P2Y1 receptors for which ADP is a major agonist. Both inhibitors also blocked the rise in [Ca2+]ielicited by addition of ADP to cells in isotonic medium, whereas A2P5P had no effect on the rise in [Ca2+]i elicited by the addition of the P2Y2 and P2Y4 receptor agonist, UTP. HTC cells were shown to express mRNA encoding for rat P2Y1, P2Y2, and P2Y6 receptors. Inhibition of the hypotonically induced rise in [Ca2+]i blocked hypotonically induced K+ (86Rb+) efflux, modulated the hypotonically induced efflux of taurine, but had no significant effect on Cl− (125I−) efflux. The interaction of extracellular ATP and/or ADP with P2Y1purinergic receptors therefore plays a role in the response of HTC cells to osmotic swelling but does not account for activation of all the efflux pathways involved in the volume-regulatory response. Cells swollen by a decrease in the osmolality of the external medium return to their original volume by a process known as regulatory volume decrease (RVD). 1The abbreviations used are: RVD, regulatory volume decrease; AM, acetoxymethyl ester; A2P5P, adenosine 2′-phosphate-5′-phosphate; A3P5P, adenosine 3′-phosphate-5′-phosphate; Δ[Ca2+]i, maximum increase in cytosolic Ca2+ concentration; Me2SO, dimethyl sulfoxide; FBS, fetal bovine serum; HBS, HEPES-buffered saline; MAPK, mitogen-activated protein kinase; 2-MeSADP, 2-methylthioadenosine-5′-diphosphate; PPADS, pyridoxyl-5′-phosphate-6-azophenyl-2′,4′-disulfonate; RT, reverse transcriptase; ER, endoplasmic reticulum. The signaling pathways activated during RVD have not yet been elucidated fully but result in the efflux of the inorganic ions, K+and Cl−, and organic molecules, with a concomitant efflux of water and a decrease in cell volume. Transient increases in cytoplasmic Ca2+ concentration ([Ca2+]i) have been observed in response to osmotic swelling in many, but not all, cell types (1Foskett J.K. Strange K. Cellular and Molecular Physiology of Cell Volume Regulation. CRC Press, Inc., Boca Raton, FL1994: 259-277Google Scholar, 2Lang F. Busch G.L. Volkl H. Cell. Physiol. Biochem. 1998; 8: 1-45Google Scholar, 3Pasantes-Morales H. Morales Mulia S. Nephron. 2000; 86: 414-427Google Scholar) and may contribute to RVD by direct activation or modulation of K+, Cl−, or organic osmolyte channels and/or by activating signaling pathways involved in RVD. Recent reports (4–16) have demonstrated that cells respond to mechanical stimuli, including osmotic swelling, by releasing ATP. ATP is known to bind to two main classes of purinergic receptors present on the cell plasma membrane; P2Y receptors are G-protein-linked receptors, and P2X receptors are intrinsic ion channels (17Ralevic V. Burnstock G. Pharmacol. Rev. 1998; 50: 413-492Google Scholar). Activation of either class of receptor can result in a transient increase in [Ca2+]i. In a number of cell types apyrase (an ATP/ADP phosphatase) and P2 receptor antagonists such as suramin and PPADS have been shown to inhibit, at least partially, swelling and mechanically stimulated cytoplasmic Ca2+ signals (5Schlosser S.F. Burgstahler A.D. Nathanson M.H. Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 9948-9953Google Scholar, 13Oike M. Kimura C. Koyama T. Yoshikawa M. Ito Y. Am. J. Physiol. 2000; 279: H630-H638Google Scholar,15Dezaki K. Tsumura T. Maeno E. Okada Y. Jpn. J. Physiol. 2000; 50: 235-241Google Scholar, 16Shinozuka K. Tanaka N. Kawasaki K. Mizuno H. Kubota Y. Nakamura K. Hashimoto M. Kunitomo M. Clin. Exp. Pharmacol. Physiol. 2001; 28: 799-803Google Scholar), thus providing evidence for a link between extracellular ATP release and transient increases in [Ca2+]i. In a study of HTC rat hepatoma cells, Fitz and co-workers (4Wang Y. Roman R. Lidofsky S.D. Fitz J.G. Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 12020-12025Google Scholar, 6Roman R.M. Wang Y. Lidofsky S.D. Feranchak A.P. Lomri N. Scharschmidt B.F. Fitz J.G. J. Biol. Chem. 1997; 272: 21970-21976Google Scholar) proposed that extracellular ATP, released in response to a decrease in the extracellular osmolality, could act in an autocrine manner by binding to purinergic receptors and thereby stimulating signaling pathways responsible for activating the Cl− efflux required for RVD. A recent report (18Roe M.W. Moore A.L. Lidofsky S.D. J. Biol. Chem. 2001; 276: 30871-30877Google Scholar) demonstrated a rise in [Ca2+]i in HTC cells in response to osmotic swelling and showed that depletion of thapsigargin-sensitive intracellular stores of Ca2+ inhibited both swelling-activated K+ and Cl− currents. In contrast to the findings in other cell types (13Oike M. Kimura C. Koyama T. Yoshikawa M. Ito Y. Am. J. Physiol. 2000; 279: H630-H638Google Scholar, 15Dezaki K. Tsumura T. Maeno E. Okada Y. Jpn. J. Physiol. 2000; 50: 235-241Google Scholar, 16Shinozuka K. Tanaka N. Kawasaki K. Mizuno H. Kubota Y. Nakamura K. Hashimoto M. Kunitomo M. Clin. Exp. Pharmacol. Physiol. 2001; 28: 799-803Google Scholar), however, the rise in [Ca2+]i was not inhibited by either apyrase or inhibitors of purinergic receptors, from which it was concluded that the swelling-induced release of ATP was not responsible for the rise in [Ca2+]i (18Roe M.W. Moore A.L. Lidofsky S.D. J. Biol. Chem. 2001; 276: 30871-30877Google Scholar). In this study we have investigated the relationship between a decrease in extracellular osmolality, extracellular nucleotides, [Ca2+]i, and ion and organic osmolyte efflux in HTC cells. In experiments comparing the actions of three enzyme preparations that hydrolyze ATP, we found that the hypotonically induced rise in [Ca2+]i was inhibited significantly only under conditions in which extracellular ADP as well as ATP was removed efficiently. The hypotonically induced rise in [Ca2+]i in HTC cells was compared with the increase in [Ca2+]i elicited by the addition of ATP, UTP, or ADP to cells in isotonic medium, and the effect of submicromolar concentrations of ATP on the efflux of K+(86Rb+), Cl−(125I−), and the organic osmolyte taurine from cells in isotonic medium was compared with the effect of a decrease in extracellular osmolality. Our results confirm the earlier finding by Roe and colleagues (18Roe M.W. Moore A.L. Lidofsky S.D. J. Biol. Chem. 2001; 276: 30871-30877Google Scholar) that the hypotonically induced rise in [Ca2+]i seen in HTC cells is a consequence of the release of Ca2+ from intracellular stores. However, our results also show that, in contrast to the conclusion drawn by Roe and colleagues (18Roe M.W. Moore A.L. Lidofsky S.D. J. Biol. Chem. 2001; 276: 30871-30877Google Scholar), extracellular adenine nucleotides do play a role in this rise in [Ca2+]i and in activating/modulating the consequent volume-regulatory efflux of K+(86Rb+) and taurine, but not Cl−(125I−), from the cells. Fura-2 AM and pluronic F127 were obtained from Molecular Probes (Eugene, OR). Ionomycin was fromCalbiochem-Novabiochem or Sigma. All other biochemicals were from Sigma. Radionuclides 86RbCl, Na125I, and [3H]taurine were obtained from Amersham Biosciences. HTC cells, derived from Morris hepatoma 7288C cells (19Morris H.P. Adv. Cancer Res. 1965; 9: 227-302Google Scholar), were maintained as a monolayer culture in Dulbecco's modified Eagle's medium supplemented with fetal bovine serum (FBS, 10%, v/v), glutamine (2 mm), penicillin (0.012% w/v), and streptomycin (0.02% w/v) in a humidified 5% CO2atmosphere at 37 °C. Isotonic HEPES-buffered saline (HBS) was composed of the following (in mm): HEPES (10Roman R.M. Feranchak A.P. Davison A.K. Schwiebert E.M. Fitz J.G. Am. J. Physiol. 1999; 277: G1222-G1230Google Scholar); NaCl (140); KCl (5Schlosser S.F. Burgstahler A.D. Nathanson M.H. Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 9948-9953Google Scholar); CaCl2 (1.3); MgCl2 (0.5); glucose (5Schlosser S.F. Burgstahler A.D. Nathanson M.H. Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 9948-9953Google Scholar), adjusted to pH 7.4 with NaOH. The osmolality was measured by freezing point depression in an Advanced Osmometer, model 3D3 (Advanced Instruments, Norwood, MA) and adjusted to 300 mOsm kg−1 (range of 299–301 mOsm kg−1) with NaCl. The hypotonic (200 mOsm kg−1) solutions used in efflux experiments were prepared as above, except that the NaCl was reduced to 90 mm. For the measurement of [Ca2+]i cells were subjected to a decrease in the extracellular osmolality by dilution of the isotonic HBS with a low-osmolality solution (35 mOsm kg−1) of the same composition as HBS but from which the NaCl was omitted. The final (measured) osmolality of the hypotonic medium was, in all cases, within the range 198–204 mOsm kg−1. In experiments in which CaCl2 was omitted from the buffers, the osmolality was adjusted to the required value with NaCl. In experiments in which extracellular Ca2+ was chelated with EGTA at the time of dilution, the low-osmolality solution used to reduce the osmolality to 200 mOsm kg−1 contained 5 mm EGTA, and the concentration of the other components was reduced by 50% so that the osmolality of the diluting solution remained in the range 33–35 mOsm kg−1. The final concentration of EGTA in the extracellular solution was 1.84 mm and that of free Ca2+ (estimated using a program based on the equations described in Ref. 20Marks P.W. Maxfield F.R. Anal. Biochem. 1991; 193: 61-71Google Scholar) was 50 nm. Apyrase (grade VII and VI) and hexokinase (type F) were each dissolved in water at a concentration of 2000 units/ml and stored as aliquots at −20 °C. Solutions of 100× the final required concentration were prepared in HBS and in the appropriate NaCl-reduced HBS solutions on the day of the experiment. ATP, ADP, 2-MeSADP, PPADS, and A2P5P were prepared as stock solutions in water at 5–50 mm. Frozen aliquots were thawed on the day of use and kept on ice. ADP was preincubated at 37 °C for 1 h with hexokinase (20 units/ml) and glucose (20 mm) in order to remove contaminating ATP. Ionomycin and thapsigargin were prepared as 10 and 2 mm stocks, respectively, in dimethyl sulfoxide (Me2SO) and stored as aliquots at −20 °C. Fura-2 AM was prepared on the day of the experiment as a 10 mmstock in Me2SO. Pluronic F127 (20% w/v) was dissolved in Me2SO and stored at room temperature. HTC cells were grown on glass coverslips (11 × 18 mm, size 0) in 35-mm culture dishes for 24 h at 37 °C, as for cell culture. The cells were loaded with Fura-2 AM (10 μm) in HBS containing 10% (v/v) FBS, 0.02% (w/v) pluronic F127, and 0.02% (v/v) Me2SO for 1 h at room temperature (20–23 °C). The loaded cells were kept for up to 2 h at room temperature in HBS plus 10% FBS for de-esterification of the Fura-2 AM before carrying out fluorescence measurements. The coverslips were washed 4× with HBS before being clamped into a coverslip holder (PerkinElmer Life Sciences) and placed in a quartz cuvette, thermostatted at 25 °C. Solutions in the cuvette were stirred throughout the experiment using a magnetic stirrer. Ratiometric measurements of fluorescence were made by exciting Fura-2 at 340 and 380 nm and measuring the emission at 510 nm in a PerkinElmer Life Sciences LS-50B luminescence spectrometer fitted with a fast-filter accessory. The fluorescence signals at each wavelength were corrected for autofluorescence (i.e. the signal that remained after quenching the Fura-2 fluorescence with 10 mm MnCl2 at the end of each experiment). [Ca2+]i was estimated as described by Grynkiewiczet al. (21Grynkiewicz G. Poenie M. Tsien R.Y. J. Biol. Chem. 1985; 260: 3440-3450Google Scholar) using the following equation: [Ca2+]i =S f,2/S b,2× K d(R −R min)/(R max −R), and assuming a K d of 224 nm (21Grynkiewicz G. Poenie M. Tsien R.Y. J. Biol. Chem. 1985; 260: 3440-3450Google Scholar). The parametersS f,2,S b,2, R min, andR max were all estimated in cells treated with 10 μm ionomycin, using the internal calibration procedure described by Kao (22Kao J.P. Methods Cell Biol. 1994; 40: 155-181Google Scholar). S f,2 andS b,2 are the fluorescence emission (at 510 nm) obtained on excitation at 380 nm, after chelation of Ca2+ with EGTA and in the presence of excess Ca2+, respectively. R min andR max are the ratios of the fluorescence emission (at 510 nm) obtained on excitation at 340 nm to that obtained on excitation at 380 nm, following chelation of Ca2+ with EGTA, and in the presence of excess Ca2+, respectively. The maximum increase in [Ca2+]i(Δ[Ca2+]i), occurring in response to different stimuli was calculated by subtracting an average of 15–20 data points acquired immediately prior to the stimulus from an average of 15 data points acquired during the period when the rise in [Ca2+]i was maximal. Total RNA was prepared from HTC cells (∼5 × 106) using the NucleoSpin RNA II kit (Macherey-Nagel, Duren, Germany) according to the manufacturer's instructions. 2 μg of RNA was incubated with oligo(dT)15primer (0.5 μg, Invitrogen) for 10 min at 65 °C in a total volume of 12 μl and then chilled on ice. Dithiothreitol (10 mm, final), dNTPs (500 μm, final), 5× first-strand buffer (Invitrogen) and RNaseOUT ribonuclease inhibitor (20 units, Invitrogen) were added (total volume, 20 μl) and incubated for 2 min at 42 °C. In order to prepare first strand cDNA, Superscript II-RT (200 units, Invitrogen) was added, and the mixture was incubated for 2 h at 42 °C, before terminating the reaction by heating to 75 °C for 15 min. In order to detect any contamination of the RNA by genomic DNA, control incubations were performed in samples in which the reverse transcriptase was omitted. PCRs were carried out for four rat P2Y receptor subtypes, rP2Y1, rP2Y2, rP2Y4 and rP2Y6, as well as for the housekeeping gene actin (using the 5′-primer set, Gene Checker Kit, Invitrogen). The P2Y primers pairs, designed to amplify a 400-bp fragment from each receptor subtype, are shown in Table I. The PCR mixtures contained 2% (v/v) first strand cDNA solution, 100 pmol of each of the sense and antisense primers, 2 units of Taqpolymerase (Qiagen), 20% (v/v) Q solution (Qiagen), and 200 μm dNTPs. The amplification conditions were 2 min at 94 °C, then 30 cycles (30 s at 94 °C, 1 min at 50 °C, 1 min at 72 °C), and then 10 min at 72 °C. Amplification products were separated on a 1% agarose gel by electrophoresis.Table IPrimer sequences used to amplify 400-bp fragments of rat P2Y receptor subtypesSubtypesAccession no.SequencePrimerNucleotide sequencesrP2Y1U22830984–1003Sense5′-TTATGTGCAAGCTGCAGAGG-3′1364–1383Antisense5′-CGGAGAGGAGAGTTGTCCAG-3′rP2Y2U56839503–522Sense5′-TCCTCTTCCTCACCTGCATC-3′883–902Antisense5′-GCGAAGACGGCCAGTACTAA-3′rP2Y4Y14706597–616Sense5′-CATCAACCTGGTGGTGACTG-3′968–987Antisense5′-ACACATGATACGGCCTGTGA-3′rP2Y6NM 057124776–795Sense5′-AGCATCCTGTTCCTCACCTG-3′1156–1175Antisense5′-CTGCTACCACGACAGCCATA-3′ Open table in a new tab HTC cells (∼2 × 105 cells) were plated in 35-mm dishes and grown for ∼24 h under normal culture conditions before being loaded with radioisotopes for 1–2 h at 37 °C. In some cases86Rb+ and 125I− efflux measurements were carried out using cells dual-labeled with both isotopes (0.3 μCi/ml for 86Rb+ and 5 μCi/ml for 125I−). Taurine efflux measurements were carried out using separate dishes, prepared on the same day, and labeled with [3H]taurine (0.1 μCi/ml). At the end of the loading period extracellular radiolabel was removed by repeated addition then removal of 1 ml of HBS (300 mOsm kg−1; 8–10 times). The flux experiment was then commenced by the careful addition to the cells of 1 ml of HBS (300 mOsm kg−1). After 4–5 min the extracellular solution was transferred to a vial for scintillation counting, and the extracellular solution was replaced by HBS (300 mOsm kg−1) with or without grade VII apyrase or inhibitors. After a further 4–5 min, the procedure was repeated using hypotonic HBS (200 mOsm kg−1) that also contained the appropriate agent. The radioactivity remaining in the cells at the end of the 4–5-min period in hypotonic medium was estimated by lysing the cells with 0.5 m NaOH. Efflux measurements were carried out at room temperature (23 °C) except where specified otherwise. Unidirectional efflux rate constants (k) were calculated using the expression: k = ln(Xt/Xt−1)/Δt, whereX t is the fraction of isotope remaining in the cells at time point (t), X t−1 is the fraction of isotope remaining at the previous time point, andΔt is the interval between the two time points. Data are expressed as mean ± S.E. For the measurement of [Ca2+]i, "n" refers to the number of coverslips analyzed. For efflux experiments,n refers to the number of different days on which86Rb+ and 125I− efflux were measured (in triplicate), and [3H]taurine efflux was measured (in duplicate). A two-tailed Student's t test of unpaired samples was used to calculate p values for the comparison of [Ca2+]i measurements and of paired samples for efflux rate constants. Results were considered significant if p ≤ 0.05. Estimates of [Ca2+]i in resting HTC cells loaded with Fura-2 were in the range 50–140 nm. When the cells were subjected to a decrease in osmolality, from 300 to 200 mOsm kg−1, [Ca2+]i rose significantly from a mean value of 72 ± 3 nm to a peak of 165 ± 7 nm (n = 42;p < 0.001) at ∼1 min after the reduction in osmolality. The mean maximum increase in [Ca2+]i(Δ[Ca2+]i) was 93 ± 5 nm. Within 6 min the [Ca2+]i levels returned to approximately the pre-stimulus levels. A representative [Ca2+]i trace is shown in Fig.1 A. Smaller decreases in the osmolality gave rise to smaller rises in [Ca2+]i, with significant increases still being observed in response to reducing the osmolality from 300 to 250 mOsm kg−1 (not shown). Grade VII apyrase is an ATP/ADP phosphatase with a low ATP/ADPase ratio. Inclusion of 3 units/ml of this enzyme preparation in the extracellular solution 5 min prior to and during a decrease in the osmolality caused a marked reduction in the hypotonically induced Δ[Ca2+]i to 12 ± 2 nm(n = 10), 13% of the corresponding same-day controls (91 ± 12 nm, Fig. 1 B). In experiments in which the initial resting [Ca2+]i was found to be above 100 nm (in isotonic medium), the addition of apyrase (grade VII) also caused a significant decrease in the resting [Ca2+]i (results not shown), implicating extracellular nucleotides in the elevated resting [Ca2+]i. Grade VI apyrase is an ATP/ADP phosphatase with a high ATP/ADPase ratio. In contrast to the results obtained with grade VII apyrase, inclusion of grade VI apyrase (3 units/ml) did not decrease the hypotonically induced Δ[Ca2+]i. The Δ[Ca2+]i in the presence of grade VI apyrase was 100 ± 15 nm (n = 5), not significantly different from the value of 79 ± 6 nmobserved for the same-day controls (Fig. 1 B). Hexokinase, which reacts with ATP to phosphorylate glucose and form ADP, was also without significant effect on the hypotonically induced Δ[Ca2+]i observed in response to a reduction in the extracellular osmolality (Fig. 1 B). The Δ[Ca2+]i in the presence of hexokinase was 125 ± 20 nm, not significantly different from the 115 ± 20 nm increase seen for the same-day controls (n = 3). The transient increase in [Ca2+]i observed on reduction of the osmolality of the extracellular solution was not due to mechanical disturbance or to the decrease in ionic strength. Addition of an identical volume of isotonic buffer did not alter the measured resting [Ca2+]i nor did the addition of an isotonic mannitol buffer, which maintained osmolality but reduced ionic strength of the extracellular solution (results not shown). The role of extracellular Ca2+ in the hypotonically induced rise in [Ca2+]i was investigated by the following: 1) omitting Ca2+ from the extracellular solution; 2) chelating extracellular Ca2+simultaneously with the reduction in osmolality; and 3) pretreating cells with thapsigargin in order to deplete the endoplasmic reticulum (ER) Ca2+ stores (23Thastrup O. Cullen P.J. Drobak B.K. Hanley M.R. Dawson A.P. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 2466-2470Google Scholar) prior to the reduction in osmolality. When HTC cells were placed for 15–20 min, in solutions from which Ca2+ had been omitted, the resting [Ca2+]i decreased to 21 ± 4 nm(n = 10). Under these conditions the Δ[Ca2+]i observed in response to a decrease in the extracellular osmolality from 300 to 200 mOsm kg−1 was reduced to 12 ± 2 nm (n = 6), compared with 94 ± 16 nm (n = 8) for same-day controls in solutions containing 1.3 mmextracellular Ca2+. One possible explanation for this reduction in Δ[Ca2+]i is that extracellular Ca2+ is the major source of Ca2+ required for the Δ[Ca2+]i observed on osmotic swelling. However, when cells that had been subjected to a decrease in osmolality were treated subsequently with thapsigargin in order to release any Ca2+ remaining in the ER Ca2+ stores (23Thastrup O. Cullen P.J. Drobak B.K. Hanley M.R. Dawson A.P. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 2466-2470Google Scholar), the thapsigargin-induced Δ[Ca2+]i in cells bathed in the Ca2+-free solution (57 ± 6 nm,n = 6) was ∼10-fold lower than that in cells bathed in solution containing 1.3 mm extracellular Ca2+ (680 ± 90 nm, n = 8). Exposure of HTC cells to a Ca2+-free solution therefore caused a marked decrease in the size of the intracellular Ca2+ pool. Representative [Ca2+]itraces are shown in Fig. 2, Aand B. In Fig. 2 C the Δ[Ca2+]i observed in response to a decrease in the osmolality (in cells in both Ca2+-free and Ca2+-containing media) is expressed relative to the Δ[Ca2+]i observed in response to the subsequent addition of thapsigargin. For cells in both solutions the ratio is the same. These data are consistent with the hypothesis that the Ca2+ giving rise to the hypotonically induced increase in [Ca2+]i was derived from the ER Ca2+ store. The amount of Ca2+ in this store was reduced substantially when the cells were transferred from a Ca2+-containing to a Ca2+-free medium. Nevertheless, the proportion of the total amount of ER store Ca2+ store that was released in response to a hypotonic shock was the same in each case. In order to confirm that entry of extracellular Ca2+ is not required for the hypotonically induced Δ[Ca2+]i, the normal (1.3 mmextracellular Ca2+) isotonic solution was diluted with a low osmolality solution containing 5 mm EGTA, so that the extracellular concentration of free Ca2+ was reduced from 1.3 mm to ∼50 nm simultaneously with the reduction in osmolality from 300 to 200 mOsm kg−1. A typical trace is shown in Fig.3 A. Under these conditions the decrease in osmolality caused [Ca2+]i to undergo an initial rise of 104 ± 16 nm (n = 6), not significantly different from same-day controls in cells exposed to 1.3 mm free Ca2+ (Fig. 3 B). The initial rise was followed by a decrease in [Ca2+]i, to ∼20 nm. When an isotonic solution containing EGTA was added (such that the final free Ca2+concentration was ∼50 nm) there was no increase in [Ca2+]i; [Ca2+]i simply decreased to ∼20 nm(Fig. 3 C), close to the value measured in cells exposed to solutions containing no added Ca2+ (Fig. 2 A). For cells in the EGTA-containing solutions (containing ∼50 nm free Ca2+) the size of the thapsigargin-sensitive Ca2+ store was reduced ∼10-fold relative to that seen in cells in media containing 1.3 mmfree Ca2+ (compare Fig. 3, A and C, with Fig. 2 B), as was the case when Ca2+ was omitted from the solutions (Fig. 2 A). When thapsigargin (200 nm) was added to HTC cells bathed in isotonic medium containing 1.3 mm Ca2+, [Ca2+]i increased to a maximum of 1000 ± 40 nm (n = 5) above resting levels and then gradually declined to a value of ∼400 nm after 30 min. A representative Ca2+ trace is shown in Fig. 3 D. When the cells were subjected to a decrease in the extracellular osmolality after 30 min or more in the presence of thapsigargin, [Ca2+]i showed no sign of increasing, again consistent with the thapsigargin-sensitive ER Ca2+ store providing the source of Ca2+ for the hypotonically induced increase in [Ca2+]i. Instead the decrease in osmolality caused [Ca2+]i to decline rapidly to ∼200 nm, as would be expected from a rapid increase in cell volume and consequent dilution of the cytoplasm. The initial decline was followed by a slower increase, probably a consequence of the cells undergoing RVD (Fig. 3 D). Gd3+, a nonspecific inhibitor of stretch-activated non-selective cation channels, as well as of various Ca2+ channels (24Hamill O.P. McBride Jr., D.W. Pharmacol. Rev. 1996; 48: 231-252Google Scholar) has been found to inhibit RVD in a number of hypotonically swollen cells including HTC cells (10Roman R.M. Feranchak A.P. Davison A.K. Schwiebert E.M. Fitz J.G. Am. J. Physiol. 1999; 277: G1222-G1230Google Scholar,25Lippmann B.J. Yang R. Barnett D.W. Misler S. Brain Res. 1995; 686: 29-36Google Scholar, 26Bergeron L.J. Stever A.J. Light D.B. Am. J. Physiol. 1996; 270: R801-R810Google Scholar). It has also been proposed that Gd3+ inhibits swelling-activated ATP release with an IC50 of between 10 and 100 μm (8Taylor A.L. Kudlow B.A. Marrs K.L. Gruenert D.C. Guggino W.B. Schwiebert E.M. Am. J. Physiol. 1998; 275: C1391-C1406Google Scholar, 10Roman R.M. Feranchak A.P. Davison A.K. Schwiebert E.M. Fitz J.G. Am. J. Physiol. 1999; 277: G1222-G1230Google Scholar, 11Hazama A. Shimizu T. Ando-Akatsuka Y. Hayashi S. Tanaka S. Maeno E. Okada Y. J. Gen. Physiol. 1999; 114: 525-533Google Scholar, 27Hazama A. Fan H.T. Abdullaev I. Maeno E. Tanaka S. Ando-Akatsuka Y. Okada Y. J. Physiol. (Lond.). 2000; 523: 1-11Google Scholar), although the extent to which this is true has recently been questioned. Gd3+ has been found to interfere with the luciferase assay for ATP (28Boudreault F. Grygorczyk R. Am. J. Physiol. 2002; 282: C219-C226Google Scholar), and after accounting for this inhibitory action, Maroto and Hamill (29Maroto R. Hamill O.P. J. Biol. Chem. 2001; 276: 23867-23872Google Scholar) found that 10 μm Gd3+ reduced mechanosensitive release of ATP from Xenopus oocytes by only 7%. In this study Gd3+, at concentrations as low as 1 μm, reduced the resting [Ca2+]i in HTC cells and inhibited the hypotonically induced increase in [Ca2+]i. Representative traces are shown in Fig.4 A and averaged data in Fig.4 B. 1 μm Gd3+ reduced the hypotonically induced Δ[Ca2+]i to ∼20% of the control value, whereas 10 μm Gd3+completely prevented the hypotonically induced rise in [Ca2+]i. The observation that 1 μm Gd3+ reduced the resting [Ca2+]i in HTC cells is consistent with Gd3+-sensitive Ca2+ influx pathways playing a role in maintaining the normal resting [Ca2+]i of the cells under the conditions used throughout this study. In separate experiments (results not shown) Gd3+ was found to inhibit the ability of extracellular Mn2+ to quench Fura-2 fluorescence in HTC cells under isotonic conditions, consistent with Gd3+ blocking an entry pathway for divalent cations. Gd3+ was also found to reduce the amount of Ca2+ that could be released from thapsigargin-sensitive Ca2+ stores. The Δ[Ca2+]i observed on addition of thapsigargin, 4 min after Gd3+ treatment, was 128 ± 7 nm (n = 3) compared with 1100 ± 80 nm for the untreated control cells (results not shown). These data are consistent with the effect of Gd3+ on the hypotonically induced increase in [Ca2+]i being a consequence of the depletion of the (thapsigargin-sensitive) intracellular Ca2+ stores. 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