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Journal of Physiology (2002), 541.1, pp. 91-101
© Copyright 2002 The Physiological Society
DOI: 10.1113/jphysiol.2001.013199

Mg²⁺-sensitive non-capacitative basolateral Ca²⁺ entry secondary to cell swelling in the polarized renal A6 epithelium

Danny Jans *, Paul De Weer †, S. P. Srinivas ‡, Els Larivière *, Jeannine Simaels * and Willy Van Driessche *

	ABSTRACT

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Polarized renal A6 epithelia respond to hyposmotic shock with an increase in transepithelial capacitance (C_T) that is inhibited by extracellular Mg²⁺. Elevation of free cytosolic [Ca²⁺] ([Ca²⁺]_i) is known to increase C_T. Therefore, we examined [Ca²⁺]_i dynamics and their sensitivity to extracellular Mg²⁺ during hyposmotic conditions. Fura-2-loaded A6 monolayers, cultured on permeable supports were subjected to a sudden reduction in osmolality at both the basolateral and apical membranes from 260 to 140 mosmol (kg H₂O)^-1. Reduction of apical osmolality alone did not affect [Ca²⁺]_i. In the absence of extracellular Mg²⁺, the hyposmotic shock induced a biphasic rise in [Ca²⁺]_i. The first phase peaked within 40 s and [Ca²⁺]_i increased from 245 ± 12 to 606 ± 24 nM. This phase was unaffected by removal of extracellular Ca²⁺, but was abolished by activating P2Y receptors with basolateral ATP or by exposing the cells to the phospholipase C (PLC) inhibitor U73122 prior to the osmotic shock. Suramin also severely attenuated this first phase, suggesting that the first phase of the [Ca²⁺]_i rise followed swelling-induced ATP release. The PLC inhibitor, the ATP treatment or suramin did not affect a second rise of [Ca²⁺]_i to a maximum of 628 ± 31 nM. The second phase depended on Ca²⁺ in the basolateral perfusate and was largely suppressed by 2 mM basolateral Mg²⁺. Acute exposure of the basolateral membrane to Mg²⁺ during the upstroke of the second phase caused a rapid decline in [Ca²⁺]_i. Basolateral Mg²⁺ inhibited Ca²⁺ entry in a dose-dependent manner with an inhibition constant (K_i) of 0.60 mM. These results show that polarized A6 epithelia respond to hyposmotic shock by Ca²⁺ release from inositol trisphosphate-sensitive stores, followed by basolateral Ca²⁺ influx through a Mg²⁺-sensitive pathway. The second phase of the [Ca²⁺]_i response is independent of the initial intracellular Ca²⁺ release and therefore constitutes non-capacitative Ca²⁺ entry.

(Received 27 August 2001; accepted after revision 25 February 2002)
Corresponding author W. Van Driessche: Laboratory of Physiology, K. U. Leuven, Campus Gasthuisberg O/N, B-3000 Leuven, Belgium. Email: willy.vandriessche{at}med.kuleuven.ac.be

	INTRODUCTION

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Calcium is an important second messenger for many cellular processes. Several agonists increase [Ca²⁺]_i after activating receptor-linked phosphatidylinositol- phospholipase C (PI-PLC) (Rebecchi & Pentyala, 2000). PI-PLC generates inositol 1,4,5-trisphosphate (InsP₃), necessary for the release of Ca²⁺ from InsP₃-sensitive intracellular stores. In addition, Ca²⁺ entry is activated by diverse mechanisms. Gating of voltage-dependent Ca²⁺ channels is affected by membrane potential (Hofmann et al. 1999). Depletion of Ca²⁺ stores induces Ca²⁺ influx through capacitative Ca²⁺ entry pathways (Putney, 1999). Stretching the plasma membrane by hyposmotic-induced cell swelling or flow-induced shear stress opens putative mechanosensitive or stretch-activated Ca²⁺ channels leading to Ca²⁺ influx (Kawahara & Matsuzaki, 1992). These latter channels comprise the non-capacitative Ca²⁺ entry pathways since they are activated independent of the depletion of InsP₃-sensitive Ca²⁺ stores, involving either the activation of protein kinase C (PKC) (Rosado & Sage, 2000) or the generation of arachidonic acid (Broad et al. 1999).

In many cell types, cell swelling induces an elevation of [Ca²⁺]_i that is thought to activate mechanisms underlying regulatory volume decrease (Hazama & Okada, 1990; Tinel et al. 1994). Osmo-mechanical stimuli also elevate [Ca²⁺]_i in A6 epithelia (Brochiero & Ehrenfeld, 1997; Yu & Sokabe, 1997; Urbach et al. 1999). Hyposmotic stimulation of whole-cell voltage clamped cells produced [Ca²⁺]_i oscillations involving both Ca²⁺ entry and Ca²⁺ release from intracellular InsP₃-sensitive stores (Yu & Sokabe, 1997). The Ca²⁺ entry was shown to occur partly through stretch-activated channels (Urbach et al. 1999) and nifedipine-sensitive pathways (Brochiero & Ehrenfeld, 1997). Consistent with this observation, another study has demonstrated stretch-activated Ca²⁺ entry in A6 cells by direct application of shear stress (Kawahara & Matsuzaki, 1992). In our laboratory, we previously observed a transient increase of about 57 % in transepithelial capacitance (C_T) during hypotonicity, presumably due to an increase in apical cell membrane surface area (Jans et al. 2000). Such a process requires fusion of membrane vesicles, which is a [Ca²⁺]_i-dependent mechanism (Fernandez-Chacon & Alvarez de Toledo, 1995). Interestingly, the observed increase in C_T was significantly attenuated when extracellular Mg²⁺ was present in the basolateral hyposmotic Ringer solution (Jans et al. 2000). To gain further insight into the Mg²⁺-sensitive C_T rise, we have examined the role of Mg²⁺ in the [Ca²⁺]_i dynamics evoked by a hyposmotic shock. We have employed polarized A6 epithelia cultured on permeable supports in order to localize Ca²⁺ influx pathways to the apical and/or basolateral domains.

	METHODS

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Cell culture

All experiments were carried out on polarized monolayers of A6 cells (a kind gift from Dr J. P. Johnson, University of Pittsburgh, Pittsburgh, PA, USA), grown on permeable Anopore filters (pore size 0.2 µm; Nunc Intermed, Roskilde, Denmark) at 28 °C and 1 % CO₂ in a humidified incubator. Cells were seeded at a lowered density of 2.5 10⁵ cm^-2, which prevented verapamil-sensitive fura-2 efflux presumably through P-glycoprotein (MDR1). The growth medium was renewed twice weekly and consisted of a 1:1 mixture of Leibovitz's L-15 and Ham's F-12 media, supplemented with 10 % fetal bovine serum (Sigma, St Louis, MO, USA), 2.6 mM sodium bicarbonate, 3.8 mM L-glutamine, 95 IU ml^-1 penicillin, and 95 µg ml^-1 streptomycin. As cells held in culture beyond 9 days showed poor dye retention, we employed epithelial monolayers cultured between 5 and 9 days for all experiments.

Solutions and chemicals

Hyposmotic solutions (140 mosmol (kg H₂O)^-1) contained (mM): 70 Na⁺, 2.5 K⁺, 2.5 HCO₃^-, 1 Ca²⁺ and 72 Cl^- (pH 8.0). Hyposmotic Ca²⁺-free solutions contained (mM): 71 Na⁺, 2.5 K⁺, 68.5 Cl^-, 5 Hepes and 2 EGTA (pH 7.4). Isosmotic solutions (260 mosmol (kg H₂O)^-1) were prepared by adding 65 mM NaCl or 110 mM sucrose. In Na⁺-free solutions, N-methyl-D-glucamine (NMDG⁺) replaced Na⁺ on an equiosmolar basis. Initial experiments, as indicated below, were conducted with solutions free of Mg²⁺. In experiments with Mg²⁺, MgCl₂ was added with no osmolality correction for Mg²⁺ concentrations of up to 2 mM, but for higher concentrations an equiosmolar amount of NaCl was omitted. Suramin, U73122 and U73343 were purchased from Sigma.

Intracellular Ca²⁺ measurements

Cells were loaded apically by incubation in a 10 µM fura-2/AM (Sigma) solution containing 0.2 g l^-1 pluronic acid (F-127, Molecular Probes, Eugene, OR, USA) for 120 min at 28 °C in 1 % CO₂. After washing off the excess dye, the monolayer was placed upside-down in an Ussing-type chamber mounted on the stage of an inverted fluorescence microscope equipped with a 40 objective (Zeiss LD, Achroplan, Wetzlar, Germany). The apical and basolateral surfaces of the monolayer were perfused independently at room temperature. Fluorescence emissions to excitation at 340 nm and 380 nm were filtered through a band-pass filter centred at 510 nm (6 nm) and detected by photon counting using a photomultiplier tube (Hamamatsu H3460-04, Hamamatsu Photonics, Japan). Photon counts of the emission at each of the excitation wavelengths were corrected for autofluorescence using signals from monolayers (n = 10) not exposed to the dye. The ratio of corrected fluorescence excited at 340 nm to that excited at 380 nm (i.e. R = I₃₄₀/I₃₈₀) was then used to estimate [Ca²⁺]_i (Grynkiewicz et al. 1985):

where R_max and R_min are the corrected fluorescence ratios under saturating and Ca²⁺-free conditions in the presence of ionomycin (5 µM), respectively. K_d, the dissociation constant of fura-2 for Ca²⁺, was taken as 224 nM (Grynkiewicz et al. 1985) and R_c is the ratio of the corrected fluorescence intensities at 380 nm excitation in zero and saturating Ca²⁺.

Electrophysiological measurements

Transepithelial resistance (R_T) and short-circuit current (I_SC). Current and voltage electrodes for the Ussing-type chamber were made of Ag-AgCl electrodes in KCl and connected to the solutions with agar bridges. The monolayers were short circuited with a high speed voltage clamp. The reciprocal of R_T, or transepithelial conductance, was measured by imposing a 1 Hz, 5 mV sine wave on the tissue. Na⁺ transport was defined as the amiloride-sensitive component of I_SC.

Transepithelial capacitance (C_T). Continuous recording of C_T was based on the current elicited by five sine waves (2, 2.7, 4.1, 5.4 and 8.2 kHz), repetitively, across the monolayer, as described previously (Van Driessche et al. 1999).

Data analysis

Results are given as means ± S.E.M. along with the number (n) of epithelia investigated.

	RESULTS

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Electrophysiology

The polarized nature of the monolayers was first verified by electrophysiological studies. At day 4, R_T was rather low (0.1 k cm^-2) and Na⁺ transport was not yet established. From day 5, Na⁺ currents that were activated by hypotonicity could be recorded and C_T normalized to a value characteristic for polarized A6 epithelia (Van Driessche et al. 1999). Electrophysiological data are summarized in Table 1. Accordingly, all subsequent experiments were conducted with cells cultured for 5 to 9 days.

Biphasic increases in [Ca²⁺]_i

When cells were perfused with isosmotic solutions (260 mosmol (kg H₂O)^-1) on both sides, mean [Ca²⁺]_i was 245 ± 12 nM (n = 10). Lowering the osmolality of the apical perfusate to 140 mosmol (kg H₂O)^-1 did not result in noticeable changes in [Ca²⁺]_i for up to 30 min (data not shown). Therefore, in all subsequent experiments osmotic changes were applied simultaneously to both sides of the epithelium. Bilateral lowering of the osmolality to 140 mosmol (kg H₂O)^-1 elicited a biphasic increase of [Ca²⁺]_i ([Ca²⁺]_i) as shown in Fig. 1 (upper trace). The initial phase of [Ca²⁺]_i was immediate, and [Ca²⁺]_i peaked at 606 ± 24 nM (n = 10) approximately 40 s after applying the shock. During the decline of the first phase, a second increase in [Ca²⁺]_i was initiated. This second increase was slow, and [Ca²⁺]_i reached 628 ± 31 nM (n = 10) about 8 min after inducing the shock (Table 2). Thereafter, [Ca²⁺]_i declined slowly but still remained above the isosmotic level after 20 min of hypotonicity. Returning to isosmotic conditions rapidly restored [Ca²⁺]_i to isosmotic levels.

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Figure 1. Changes in [Ca2+]i of fura-2-loaded polarized A6 epithelia in response to a hyposmotic shock Hypotonic shock was induced by a sudden removal of 65 mM NaCl (upper trace, n = 10), 110 mM sucrose (middle trace, n = 5) or 65 mM NMDGCl (lower trace, n = 5). The continuous lines represent the mean values. S.E.M. values are shown as dotted lines. Both apical and basolateral perfusates were nominally Mg2+ free. Numerical values are shown in Table 2. The upper panel indicates the osmolarity of the solution ().

Inducing hypotonicity by lowering the NaCl concentration of the external bath alters the Na⁺ gradient across the cell membrane. This manoeuvre might cause secondary effects on Na⁺-dependent transporters. Therefore, we compared the above results with experiments in which hypotonicity was evoked by removing sucrose from the perfusion solutions. Figure 1 (middle trace) shows that in these circumstances, hypotonicity also resulted in a biphasic change of [Ca²⁺]_i. Sucrose removal elevated [Ca²⁺]_i from 220 ± 9 nM to 446 ± 14 nM and to 626 ± 21 nM (n = 5), for the first and second phase, respectively. It should be noted that the first phase of the [Ca²⁺]_i response was smaller after sucrose removal as compared to the reduction of the NaCl concentration. This might be due to impaired diffusion of released compounds as a result of the remaining sucrose sticking to the membrane thereby interfering with activation mechanisms that take place at the extracellular border. Similarly, we previously demonstrated that sucrose removal, as a tool to induce a hyposmotic shock, interfered with the basolateral efflux of K⁺ during regulatory volume decrease (Li et al. 1998).

We also performed experiments in Na⁺-free conditions to verify a possible contribution of the Na⁺-Ca²⁺ exchanger to the second phase of [Ca²⁺]_i during the hypotonic shock. In Na⁺-free solutions, reducing the concentration of NMDGCl in the perfusion solutions was used to induce the hyposmotic shock (Fig. 1, lower trace). In these conditions, the first phase of [Ca²⁺]_i was absent, whereas the second phase was still present, increasing [Ca²⁺]_i from 268 ± 26 to 617 ± 45 nM (n = 5). Therefore, in all subsequent experiments, we reduced the osmolality by NaCl removal to avoid diffusion problems with sucrose and interference of Na⁺ free conditions with Na⁺ dependent processes.

Source of the hypotonicity induced [Ca²⁺]_i increase

We investigated whether, as in other cell types, both extracellular and intracellular sources contribute to [Ca²⁺]_i. Removal of Ca²⁺ from the apical solution had no effect on [Ca²⁺]_i, induced by hypotonicity (data not shown). Maintaining Ca²⁺-free conditions at the basolateral surface was complicated. Prolonged exposure of the basolateral membrane to Ca²⁺-free solutions damaged the monolayer's integrity as found by electrophysiological studies (data not shown). However, normal values of R_T persisted longer when Ca²⁺ was removed only at the moment of osmotic shock. To prevent Ca²⁺ leakage from the apical to the basolateral surface, Ca²⁺ was removed simultaneously from both sides. The combination of a hyposmotic shock with bilateral Ca²⁺ removal led to an increase of [Ca²⁺]_i from 241 ± 7 nM in isosmotic conditions to 638 ± 60 nM (n = 5) 41 s after its application (Fig. 2, Table 2). It is noteworthy that neither the time course nor the amplitude of the first phase of [Ca²⁺]_i was affected by the removal of extracellular Ca²⁺. After this rapid transient increase, [Ca²⁺]_i returned to isosmotic levels without exhibiting the second phase of [Ca²⁺]_i normally seen in the presence of extracellular Ca²⁺. These results indicate that the second phase depends on extracellular Ca²⁺ entry, whereas Ca²⁺ release from intracellular stores accounts for the [Ca²⁺]_i increase during the first phase.

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Figure 2. [Ca2+]i changes in response to hyposmotic shock in the absence of extracellular Ca2+ In this and all subsequent experiments, epithelial monolayers were initially perfused with isosmotic NaCl Ringer solutions (260 mosmol (kg H2O)-1) on the apical and basolateral side and the hyposmotic shock (140 mosmol (kg H2O)-1) was induced by bilateral removal of 65 mM NaCl. Extracellular Ca2+ was removed simultaneously from both sides of the epithelium at the moment of the hyposmotic shock. The average value is for five tissues and S.E.M. values are shown as dotted lines. All perfusates were nominally Mg2+ free.

Characterization of the first phase of [Ca²⁺]_i increase

To identify the source of intracellular Ca²⁺ release, we explored the involvement of InsP₃-sensitive stores in mediating the first phase of [Ca²⁺]_i. We first attempted to empty the InsP₃-sensitive Ca²⁺ stores by repetitive exposure to ATP. Both apical and basolateral applications of extracellular ATP have been shown to increase [Ca²⁺]_i in A6 cells, presumably by activating PI-PLC through a Gq G protein coupled to P2Y receptors (Mori et al. 1997; Banderali et al. 1999). Thus, exposure to extracellular ATP can result in InsP₃-mediated Ca²⁺ release from intracellular stores. Hence, we examined the effect of unilateral ATP exposure on [Ca²⁺]_i, induced by hypotonicity. Cells were exposed three times, at 6 min intervals, for 2 min to 0.5 mM ATP before the hyposmotic shock was applied in the continued presence of ATP. Apical ATP (Fig. 3A) induced a rapid, but small and transient, rise in [Ca²⁺]_i from 240 ± 7 to 357 ± 21 nM, from 274 ± 12 to 333 ± 15 nM, and from 279 ± 13 to 323 ± 14 nM (n = 5), respectively, for the three consecutive exposures. This approach, however, did not alter the [Ca²⁺]_i response to the subsequent hyposmotic shock: [Ca²⁺]_i increased from 260 ± 10 to 686 ± 32 nM and 670 ± 8 nM for the first and second phase, respectively (n = 5) (Table 2). The effect of basolateral ATP on [Ca²⁺]_i was markedly different (Fig. 3B): initial exposure raised [Ca²⁺]_i rapidly from 248 ± 9 to 853 ± 91 nM (n = 5), followed by a return towards the basal level. The second and third ATP exposures caused more modest [Ca²⁺]_i increases, from 320 ± 12 to 394 ± 59 nM and from 329 ± 11 to 423 ± 57 nM, respectively. The subsequent hyposmotic shock was devoid of the first phase of [Ca²⁺]_i, whereas the second phase remained, raising [Ca²⁺]_i from 282 ± 17 to 640 ± 33 nM within 8 min (n = 5). These results suggest that the first phase of [Ca²⁺]_i is dependent on InsP₃-sensitive stores.

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Figure 3. [Ca2+]i changes in response to hyposmotic shock: effect of prior stimulation by ATP A, apical ATP. ATP (0.5 mM) was applied to the apical side for three successive 2 min periods prior to osmotic shock. The first two periods were followed by 4 min washouts; the third was followed immediately by the hyposmotic shock, during which ATP remained in the apical solution. The [Ca2+]i trace is the average of five experiments. B, basolateral ATP. Protocol identical to that in A, except that ATP was added to the basolateral solution. The [Ca2+]i time course is the mean of five preparations S.E.M. values are shown as dotted lines. All perfusates were nominally Mg2+ free.

Further support was obtained from bilateral pre-exposure of cells to U73122 (10 µM), a membrane-permeant aminosteroid that is commonly used as a specific PI-PLC inhibitor (Smith et al. 1990). Aside from its blocking effect on agonist-induced PI-PLC activation, however, U73122 has been shown to elevate [Ca²⁺]_i in Madin-Darby canine kidney (MDCK) cells by combining Ca²⁺ influx and Ca²⁺ release from intracellular stores (Jan et al. 1998). Exposure to U73122, 3 min before application of the hyposmotic shock, caused a small transient [Ca²⁺]_i rise from 222 ± 5 to 272 ± 22 nM (n = 5) (Fig. 4A). The [Ca²⁺]_i response to a subsequent hyposmotic shock again lacked the first phase. Lowering the concentration of U73122 to 5 µM only attenuated the first phase of [Ca²⁺]_i (not shown). We used the inactive enantiomer U73343 as a negative control, since it does not inhibit PLC. With U73343 (10 µM bilateral), both phases of [Ca²⁺]_i increase were fully expressed as in control conditions, increasing [Ca²⁺]_i from 227 ± 6 to 626 ± 38 nM and to 655 ± 16 nM for the first and second phase, respectively (n = 5) (Fig. 4A). Taken together, these data indicate that the initial phase of [Ca²⁺]_i reflects Ca²⁺ release from InsP₃-sensitive Ca²⁺ stores via a PI-PLC-activated pathway. These stores become depleted after repeated exposure to basolateral ATP but not apical ATP.

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Figure 4. [Ca2+]i changes in response to hyposmotic shock: inhibition of the first phase of [Ca2+] A, effect of the PLC inhibitor U73122 compared to the inactive enantiomer U73343. The PLC inhibitor U73122 (10 µM) was applied bilaterally in isosmotic conditions 3 min before applying the hyposmotic shock. In parallel experiments, the structurally related but inactive compound U73343 (10 µM) was used as a negative control. B, effect of suramin. Suramin (100 µM) was added to the basolateral solution, 4 min before the shock. For both panels, the compounds were also present in the hyposmotic solutions. Traces represent the average of five experiments with S.E.M. values shown as dotted lines. All perfusates were nominally Mg2+ free.

We further explored the mechanisms that led to the first rapid increase in [Ca²⁺]_i during the shock. Since cell swelling is known to cause release of a number of substances (Okada et al. 2001) that are able to trigger resident receptors in an autocrine activation pattern, we examined the possibility that ATP is released by applying the non-specific P2 purinoceptor antagonist suramin (100 µM) to the basolateral side of the epithelium. In the presence of suramin, the first phase of [Ca²⁺]_i was severely suppressed (Fig. 4B), whereas the second phase was present as in control conditions, increasing [Ca²⁺]_i from 248 ± 23 nM in isosmotic to 635 ± 21 nM during hypotonic conditions (Table 2). From these data, we suggest that swelling-induced ATP release underlies the rapid rise of [Ca²⁺]_i.

Effects of extracellular Mg²⁺ on Ca²⁺ entry

Given our earlier finding (Jans et al. 2000) that the C_T increase in response to hyposmotic shock is significantly attenuated by basolateral Mg²⁺, we hypothesized that extracellular Mg²⁺ might interfere with basolateral Ca²⁺ influx. It should be recalled that all experiments described so far were performed with Mg²⁺-free solutions. To assess the role of extracellular Mg²⁺, we first included the divalent cation in the apical perfusate, but found no effect on [Ca²⁺]_i (data not shown). In contrast, 2 mM Mg²⁺ in the basolateral perfusate had a marked effect on the time course of [Ca²⁺]_i in response to the hyposmotic shock (Fig. 5). Although phase 1 of [Ca²⁺]_i remained unaffected with [Ca²⁺]_i increasing from 241 ± 12 to 635 ± 25 nM (n = 5) in 50 s, the second phase was largely abolished (Table 2). This indicates that basolateral Mg²⁺ (Mg²⁺_bl) inhibits hyposmotic-induced Ca²⁺ influx at the basolateral surface.

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Figure 5. [Ca2+]i changes in response to hyposmotic shock: effect of Mg2+ at the basolateral border MgCl2 (2 mM) was included in the basolateral hyposmotic solution. The [Ca2+]i trace represents the average of five experiments with S.E.M.s shown as dotted lines.

To further investigate Mg²⁺_bl inhibition on Ca²⁺ entry, cells were acutely exposed (Fig. 6) to 2 mM Mg²⁺ during the second phase of [Ca²⁺]_i. Addition of Mg²⁺_bl caused [Ca²⁺]_i to fall rapidly towards isosmotic levels within 5 min. This fast decline of [Ca²⁺]_i in response to Mg²⁺_bl addition indicates that the mechanisms responsible for restoring normal [Ca²⁺]_i operate at a high rate. We also noticed that, at lower concentrations of Mg²⁺_bl, the rate of [Ca²⁺]_i decline was less rapid, which suggests that Mg²⁺_bl inhibits Ca²⁺ influx in a dose-dependent manner. A detailed analysis, presented in the Discussion, enabled us to determine the value (0.6 mM) of the inhibition constant, K_i.

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Figure 6. [Ca2+]i changes in response to hyposmotic shock: effect of subsequent acute addition of Mg2+ MgCl2 (2 mM) was added to the basolateral solution at time t = tA during the second rising phase of the [Ca2+]i transient, 5.5 min after the onset of the hyposmotic shock. The average and S.E.M. traces of six experiments are shown. An exponential curve (thin line) was fitted to the average trace. From its derivativ-e at t = tA the initial rate constant for [Ca2+]i decline was estimated as -1.79 ± 0.02 min-1.

	DISCUSSION

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In this study, we have examined [Ca²⁺]_i dynamics in polarized A6 epithelia in response to hyposmotic shocks. A6 cells were grown as monolayers on permeable supports and perfused independently on the apical and basolateral surfaces. The polarized nature of the monolayer was demonstrated by (1) electrophysiological characteristics, (2) the absence of changes in [Ca²⁺]_i in response to a reduction of apical osmolality alone, (3) the persistence of Ca²⁺ entry in response to a hyposmotic shock in the absence of apical Ca²⁺, (4) the failure of apical Mg²⁺ to block Ca²⁺ entry during the hyposmotic shock and (5) the differential effect of apical and basolateral ATP on [Ca²⁺]_i elevation under both isosmotic and hyposmotic conditions. Additionally, this study differs from earlier studies on A6 cells (Brochiero & Ehrenfeld, 1997; Yu & Sokabe, 1997; Urbach et al. 1999) in that monolayers were grown on permeable supports under conditions that permitted continuous measurement of [Ca²⁺]_i with fura-2 for periods of up to 60 min.

Subjecting the polarized monolayers to acute cell swelling produced profound changes in [Ca²⁺]_i characterized by a biphasic time course. The first, immediate phase of [Ca²⁺]_i was independent of extracellular Ca²⁺ and, since it was blocked by the specific PI-PLC inhibitor U73122, and not by its inactive enantiomer U73343, probably involves the InsP₃ pathway. It is possibly caused by autocrine activation after swelling-induced release of chemical mediators such as ATP, which stimulates cell surface receptors, coupled to heterotrimeric G proteins activating PLC. In this context, it has been reported that mechanical stimulation enables mammary epithelial cells to communicate with adjacent cells by InsP₃ movement via gap junctions (Boitano et al. 1992) and with more distant cells via release of ATP into the extracellular space (Osipchuk & Cahalan, 1992). Because ATP release is reported to occur in response to cell swelling (Wang et al. 1996), it is plausible that the first phase of the increase in [Ca²⁺]_i is due to ATP release. This hypothesis is supported by the observation that suramin, a purinergic receptor antagonist, also abolishes the first phase of the increase in [Ca²⁺]_i during hypotonicity. The involvement of ATP release in Ca²⁺ mobilization, however, needs further investigation. Another mechanism for the formation of InsP₃ is through activation of PLC by tyrosine kinases. Consistent with this argument, several studies have implicated tyrosine kinase activity in the response to a hyposmotic challenge (Tilly et al. 1993). Furthermore, activation of tyrosine kinases was shown to be involved in increasing C_T in A6 epithelia during hypotonicity (Osipchuk & Cahalan, 1992).

In this study, we compared three different methods of inducing a hyposmotic shock: (1) removal of NaCl, (2) withdrawal of sucrose and (3) removal of NMDGCl. Activation of the second phase of [Ca²⁺]_i was comparable in the three conditions, but the InsP₃-mediated Ca²⁺ release phase was expressed more during NaCl removal. Diffusion of ATP, released upon cell swelling, is probably impaired after sucrose removal. The suppression of the first phase of [Ca²⁺]_i when reducing the NMDGCl concentration may be the consequence of the absence of Na⁺ as a co-activator for purinergic receptors (Marsigliante et al. 2002). This view is supported by the lack of increase in [Ca²⁺]_i upon exogenous ATP application in Na⁺-free conditions (data not shown).

The second phase of [Ca²⁺]_i requires extracellular Ca²⁺. Since InsP₃-sensitive stores are emptied immediately in response to hyposmotic shock, it is conceivable that part of this Ca²⁺ influx is through capacitative Ca²⁺ entry (CCE) pathways. However, CCE does not seem to contribute significantly to the [Ca²⁺]_i changes during hypotonicity. Indeed, exogenous ATP application in isosmotic conditions only evoked a fast transient rise in [Ca²⁺]_i (data not shown, but compare Fig. 3B) without a subsequent increase due to Ca²⁺ entry, as observed in other cell types that display CCE (Putney, 1999). The lack of Ca²⁺ entry secondary to Ca²⁺ release from intracellular stores supports the notion of the absence of CCE in A6 epithelia. In contrast, our finding that the second phase is activated even after pre-exposure to U73122 or suramin suggests that pathways other than CCE are involved. Activation of the Ca²⁺ entry channels that mediate the non-capacitative pathway may be due to mechanical stress caused by the hyposmotic shock. Stretch-activated Ca²⁺ channels, sensitive to Ni²⁺ and La³⁺ have been described in A6 cells in response to shear stress (Kawahara & Matsuzaki, 1992). Similar stretch-activated Ca²⁺ channels have been observed in response to membrane hyperpolarization (Urbach et al. 1999). Voltage-dependent plasma membrane Ca²⁺ channels, related to L-type Ca²⁺ channels, were noted to mediate Ca²⁺ entry during hyposmotic treatment of A6 epithelia after membrane hyperpolarization (Brochiero & Ehrenfeld, 1997). Channels of this type have been found in the apical membranes of proximal tubule cells, but are opened upon membrane depolarization to mediate Ca²⁺ reabsorption during periods of enhanced Na⁺ reabsorption (Zhang & O'Neil, 1996). We found a hyposmotic-activated Ca²⁺ entry mechanism in A6 epithelia that is blocked by Mg²⁺_bl. Recently, extracellular Mg²⁺ was shown to block Na⁺ and Ca²⁺ currents through epithelial calcium channels expressed in human embryonic kidney (HEK 293) cells. Since inhibition was voltage dependent the Mg²⁺ binding site is presumably located within the channel pore. At -80 mV, K_i for inhibition of Ca²⁺ currents is 0.33 mM when extracellular [Ca²⁺] = 0.1 mM and as high as 9 mM when extracellular [Ca²⁺] = 2 mM (Vennekens et al. 2001). Blockage by extracellular Mg²⁺ is also a key functional property of the ionotropic N-methyl-D-aspartate (NMDA) receptor channel in the central nervous system (Mayer et al. 1984). The Mg²⁺ block and unblock of NMDA is a function of the Na⁺ concentration and the membrane potential. NMDA receptor channel is highly permeable to Ca²⁺ and is essential in synaptic transmission underlying memory, learning, and development. Mg²⁺ blocks the NMDA receptor channel within the pore at strongly negative membrane potentials. Membrane depolarization drives Mg²⁺ out of the channel pore, which then allows Ca²⁺ permeation after activation with glutamate. Additionally, PKC modulates channel activity by reducing the Mg²⁺ block of the NMDA receptor channel (Chen & Huang, 1992). PKC activity increases upon PI-PLC activation and can lead to the stimulation of non-capacitative Ca²⁺ entry (Rosado & Sage, 2000). Whether such a mechanism is applicable to A6 epithelia remains unknown.

Kinetic modelling of the Mg²⁺-sensitive Ca²⁺ entry

To systematically assess this dose dependency, we chose a fixed time (denoted by t_A) for Mg²⁺ addition to the basolateral perfusate: 5.5 min after the onset of the hyposmotic shock. (A precise definition of t_A is given in the legend of Fig. 7.) By time t_A the first phase of [Ca²⁺]_i has ended and release from intracellular stores becomes vanishingly small. For times t t_A just prior to Mg²⁺_bl addition (Fig. 7A, Period A), the [Ca²⁺]_i dynamics (Fig. 7B) are determined by (1) Ca²⁺ extrusion to the extracellular medium, (2) Ca²⁺ uptake into intracellular stores, (3) Ca²⁺ binding to intracellular buffers and (4) Ca²⁺ entry from the extracellular space. For times t t_A, after addition of a dose of Mg²⁺_bl sufficient to block further Ca²⁺ entry (Fig. 7A, Phase B), only those processes that contribute to Ca²⁺ removal from the cytosol remain (i.e. pathways 1, 2 and 3). At subsaturating Mg²⁺_bl concentrations, the [Ca²⁺]_i decline was slower because it reflects not only Ca²⁺ removal (pathways 1-3) but also the incomplete inhibition of Ca²⁺ entry (pathway 4). Thus the rate of [Ca²⁺]_i decline in the presence of various concentrations of Mg²⁺_bl is a measure of hypotonicity-induced Ca²⁺ entry. To facilitate a quantitative analysis of the rate of [Ca²⁺]_i decline, we modelled the processes of Ca²⁺ removal and Ca²⁺ entry as described below. The kinetic analysis of the inhibition of Ca²⁺ entry by Mg²⁺_bl during hyposmotic shock was performed for various concentrations of Mg²⁺ (viz. 0.1, 0.2, 0.5, 1, 2, 4, 10 and 20 mM) applied at time t_A during the rising part of the hyposmotic-induced Ca²⁺ entry phase, following the protocol of Fig. 7A. We found a K_i for inhibition by Mg²⁺_bl of 0.60 ± 0.05 mM.

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Figure 7. Model for assessing the analysis of Ca2+ entry inhibition by basolateral Mg2+ during hyposmotic shock A, basis for assessing the dose dependence of Ca2+ entry inhibition by Mg2+. The experimental curve is an expanded view of the [Ca2+]i trace around tA, the time of addition of 0.1 mM Mg2+ (tA is defined more precisely below). Period A shows the time course of [Ca2+]i before tA; it is fitted with a polynomial whose derivative at t = tA (dotted line) is labelled 'Slope A'. Period B shows the time course of [Ca2+]i after Mg2+ addition; it is fitted with an exponential function whose derivative at t = tA is labelled 'Slope B'. For simplicity, tA is defined by the intersection of slopes A and B. For comparison, the average inhibitory effect of adding 20 mM Mg2+ is also shown, scaled to the value of [Ca2+]i at the intersection of slopes A and B. An exponential was fitted to these data 'Slope C' that represents a near-maximal inhibition of Ca2+ entry and resulted in an initial rate constant for [Ca2+]i decline averaged at -3.02 ± 0.02 min-1 (n = 11). Further description is given in the text. B, processes that regulate [Ca2+]i. Rate constants used in the model are: kse for Ca2+ release from intracellular stores; ksd for ATPase-dependent Ca2+ uptake into intracellular stores; kbd and kbe for binding to and release from intracellular Ca2+ buffers; kce for Ca2+ entry from the extracellular space; and kcd for extrusion of Ca2+ from the cytosol into the extracellular medium by the Na+-Ca2+ exchanger and the plasma membrane Ca2+-ATPase. C, dose-response curve of the inhibitory effect of basolateral Mg2+ on Ca2+ entry. The inhibitory effect (Inh) of basolateral Mg2+ on Ca2+ entry, defined as: 100 (1 - ke / kemax), was calculated from the slopes of the [Ca2+]i curves at t = tA (Fig. 7A). The values were fitted with a Michaelis-Menten function constrained through the 20 mM point, yielding a Ki of 0.60 ± 0.05 mM Mg2+.

We performed a quantitative analysis of the inhibitory effect of Mg²⁺_bl on Ca²⁺ entry during the hyposmotic shock. This analysis assumes that the perfusion rate of the solutions (7.5 ml min^-1) is sufficiently high to enable rapid solution exchange adjacent to the basolateral membrane before complete replacement of the chamber content (1.5 ml). The fact that doubling the perfusion rate did not alter the kinetic feature of the Mg²⁺ block supports this assumption. As stated before, we assume that acute addition of Mg²⁺ does not affect cell volume or Ca²⁺ extrusion by either Na⁺-Ca²⁺ exchange or the plasma membrane Ca²⁺ pump. Furthermore, the model relies on the assumption that extracellular Mg²⁺ does not affect the buffering of Ca²⁺ in the cytosol or the uptake of Ca²⁺ in intracellular stores. The rate of change of [Ca²⁺]_i, i.e. the slope of a [Ca²⁺]_i versus time plot (Figs 1-7) reflects the quantity of Ca²⁺ (Q_Ca, in moles) entering or leaving the cytoplasm per unit time:

(1)

where the dimensionless term is the cytoplasm's buffering power (i.e. the number of moles of Ca²⁺ required per litre of cytoplasm to effect a [Ca²⁺]_i of 1 M) and Vol is the volume (here in litres) of cytoplasm under consideration. In general, neither nor Vol is constant. Buffering power reflects the (near-instantaneous) reactions of Ca²⁺ with cytoplasmic buffers (rate constants k^b_d and k^b_e in Fig. 7B) and may vary with [Ca²⁺]_i. Cytoplasmic volume will vary with time as cells undergo osmotic swelling and regulatory volume decrease. However, for a given [Ca²⁺]_i at a given time point such as t = t_A (Fig. 6 and Fig. 7A) and Vol are single-valued so that at or near t = t_A we define the slope as:

(2)

where flux, _net = _in + _out, is expressed in moles per litre per minute (influx is positive, outflux is negative). Furthermore, the model assumes that extracellular Mg²⁺ does not affect cytosolic Ca²⁺ removal by transport processes or buffering (rate constants k^s_d, k^c_d, and k^b_d in Fig. 7B) so that _out is identical before and after Mg²⁺ addition. This assumption is based on the invariability of the cytosolic free Mg²⁺ content (~1 mM) in relation to total cellular Mg²⁺ concentration (~20 mM; Romani, 2000).

At time t = t_A in Fig. 7A three slopes are of interest: slope A before Mg²⁺_bl addition; slope B after addition of 0.1 mM Mg²⁺_bl; and the slope C of the curve denoted '20', which summarizes 11 experiments with exposure to 20 mM Mg²⁺_bl (see Fig. 7 for the slope fitting procedure). Immediately before exposure to Mg²⁺_bl the rate of entry of Ca²⁺ is taken as maximal so that:

(3)

whereas after exposure to a subsaturating concentration of Mg²⁺_bl (which leaves some residual influx) we have:

(4)

Furthermore, at a near-maximal inhibitory concentration of Mg²⁺_bl (20 mM, as will be justified below) Ca²⁺ entry becomes negligible and:

(5)

Entering eqn (5) in eqns (3) and (4) we obtain:

(6)

and:

(7)

from which the relative inhibition of Ca²⁺ entry can be computed:

(8)

The inhibition of Ca²⁺ entry by Mg²⁺_bl during hyposmotic shock was assessed for various Mg²⁺_bl concentrations (viz. 0.1, 0.2, 0.5, 1, 2, 4, 10 and 20 mM) applied at time t_A during the rising part of the hypotonicity-induced Ca²⁺ entry phase, following the protocol of Fig. 7A. The degree of inhibition for each tested concentration of Mg²⁺_bl was then calculated according to eqn (8), and plotted as a function of the Mg²⁺_bl concentration (Fig. 7C). A preliminary fit with a Hill function yielded a Hill coefficient not statistically different from 1. We therefore fitted the data with a Michaelis-Menten function, and found a K_i for inhibition by Mg²⁺_bl of 0.60 ± 0.05 mM. Given this value, 20 mM Mg²⁺_bl will inhibit 97 % of Ca²⁺entry, which justifies the approximation implicit in eqn (5). The small second phase of [Ca²⁺]_i seen in the presence of 2 mM Mg²⁺_bl (Fig. 5) reflects a Ca²⁺ entry inhibition of approximately 80 %, as expected from the value we found for K_i.

In summary, polarized A6 epithelial cells show a biphasic increase in [Ca²⁺]_i in response to a sudden decrease in basolateral osmolality. Both phases of [Ca²⁺]_i are activated independently. The initial release of Ca²⁺ from intracellular stores depends on PI-PLC activation, whereas basolateral Mg²⁺ inhibits the subsequent non-capacitative Ca²⁺ entry in a dose-dependent manner. The latter explains the inhibitory effects of Mg²⁺_bl on the C_T increase during hypotonicity (Jans et al. 2000) The experiments in Na⁺ free conditions, i.e. NMDGCl removal, rule out any involvement of the Na⁺-Ca²⁺ exchanger in the shape of the second phase of [Ca²⁺]_i.

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Acknowledgements

This project was supported by research grants from the 'Fonds voor wetenschappelijk onderzoek-Vlaanderen' (G.0179.99), the Interuniversity Poles of Attraction Program-Belgian State, Prime Minister's Office-Federal Office for Scientific, Technical, and Cultural Affairs IUAP P4/23 and S.P. Srinivas is supported by NIH grant EY11107. P. De Weer was holder of the International Francqui Chair.

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