Extracellular ATP stimulates volume decrease in Necturus red blood cells

Douglas B. Light, Tracy L. Capes, Rachel T. Gronau, and Matthew R. Adler

Department of Biology, Ripon College, Ripon, Wisconsin 54971


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

This study examined whether extracellular ATP stimulates regulatory volume decrease (RVD) in Necturus maculosus (mudpuppy) red blood cells (RBCs). The hemolytic index (a measure of osmotic fragility) decreased with extracellular ATP (50 µM). In contrast, the ATP scavenger hexokinase (2.5 U/ml, 1 mM glucose) increased osmotic fragility. In addition, the ATP-dependent K+ channel antagonist glibenclamide (100 µM) increased the hemolytic index, and this inhibition was reversed with ATP (50 µM). We also measured cell volume recovery in response to hypotonic shock electronically with a Coulter counter. Extracellular ATP (50 µM) enhanced cell volume decrease in a hypotonic (0.5×) Ringer solution. In contrast, hexokinase (2.5 U/ml) and apyrase (an ATP diphosphohydrolase, 2.5 U/ml) inhibited cell volume recovery. The inhibitory effect of hexokinase was reversed with the Ca2+ ionophore A-23187 (1 µM); it also was reversed with the cationophore gramicidin (5 µM in a choline-Ringer solution), indicating that ATP was linked to K+ efflux. In addition, glibenclamide (100 µM) and gadolinium (10 µM) inhibited cell volume decrease, and the effect of these agents was reversed with ATP (50 µM) and A-23187 (1 µM). Using the whole cell patch-clamp technique, we found that ATP (50 µM) stimulated a whole cell current under isosmotic conditions. In addition, apyrase (2.5 U/ml), glibenclamide (100 µM), and gadolinium (10 µM) inhibited whole cell currents that were activated during hypotonic swelling. The inhibitory effect of apyrase was reversed with the nonhydrolyzable analog adenosine 5'-O-(3-thiotriphosphate) (50 µM), and the effect of glibenclamide or gadolinium was reversed with ATP (50 µM). Finally, anionic whole cell currents were activated with hypotonic swelling when ATP was the only significant charge carrier, suggesting that increases in cell volume led to ATP efflux through a conductive pathway. Taken together, these results indicate that extracellular ATP stimulated cell volume decrease via a Ca2+-dependent step that led to K+ efflux.

volume regulation; patch clamp; potassium channel; hexokinase; calcium


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

THE ABILITY OF ANIMAL CELLS to regulate their volume is a fundamental property common to a large number of cell types (11, 20, 22, 24-26) and has been extensively reviewed (8, 12, 21, 23, 29). Volume regulation is of importance in cells exposed to anisotonic extracellular media and in cells where transport of solutes could change intracellular osmolality. Exposure of vertebrate cells to a hypotonic solution results in an initial increase in cell volume due to the relatively rapid influx of water. During continuous hypotonic stress, increases in cell volume are then followed by a slower, spontaneous recovery toward the preshock level, a process known as regulatory volume decrease (RVD). This recovery is accomplished by selectively increasing the permeability of the plasma membrane during cell swelling to allow for efflux of specific intracellular osmolytes, thereby decreasing the driving force for water influx (8, 12, 21, 23, 29). Most vertebrate cells lose K+ and Cl- during RVD (8, 12, 21, 23, 29). This may occur by electroneutral ion transport pathways (21) or by the separate activation of K+ and anion channels (8, 11, 21, 26, 35). Loss of organic anions and osmolytes also may occur during RVD (18, 29).

The cellular mechanisms that activate and regulate permeability pathways during RVD are not completely understood and appear to differ between cell types. For example, in some instances, the activation mechanism for an RVD response is Ca2+ independent (16, 20, 24). In contrast, Ca2+ appears to play a role during cell volume regulation in several cell types (4, 22, 23, 25, 38). In addition, although it has been suggested that Ca2+ directly activates ion channels during RVD (11, 23, 35), there also is evidence that several Ca2+-dependent intracellular messengers and enzymes (e.g., calmodulin, phospholipase A2, 5-lipoxygenase, and protein kinase C) are involved with cell volume regulation (12, 21-23).

It is well known that ATP is a ubiquitous intracellular source of energy. However, over 25 years ago it was proposed that ATP acts as a transmitter substance at autonomic neuromuscular junctions (3). Since then, there has been a growing body of evidence indicating that extracellular ATP plays a significant role in a number of other biological processes (6, 9, 17, 36, 37). For example, extracellular ATP has been implicated in the control of fluid secretion by salivary gland cells (28), ion and water balance of cochlear fluids (32), secretion of histamine by mast cells (9), vasodilation of coronary blood vessels (9), and production of prostacyclin (9). Extracellular ATP also has been shown to stimulate cell volume regulation (33, 36), and a number of studies have demonstrated that extracellular nucleotides are important for regulating ion channels (1, 7, 28, 31, 32). Extracellular ATP exerts its influence by acting as an autocrine and paracrine signal, binding to specific cell surface receptors termed purinoceptors (7, 27, 32, 36, 37). Purinoceptors have been subdivided into two main categories: P1 receptors, which recognize nucleosides, such as adenosine, and P2 receptors, which bind ATP and other nucleotides (28, 32, 33, 37). The P2 receptors have been further subdivided into two main groups: ATP-gated, Ca2+-permeable, nonselective channels (32, 37) and ATP-activated receptors coupled to a G protein (32, 37).

Despite recent reports concerning the physiology of extracellular ATP, there is a paucity of data on the role of ATP in RVD. Thus the potential connections between this nucleotide and cell volume regulation remain to be elucidated. In view of these uncertainties, the purpose of this study was to investigate whether extracellular ATP regulates K+ efflux during RVD in Necturus red blood cells (RBCs). The basis of this study also stemmed from our recent work in which we demonstrated that RVD in this cell type depends on a K+ conductance that is regulated during cell swelling by a Ca2+-dependent mechanism (22) and that extracellular ATP may elevate the intracellular free Ca2+ concentration by activating phospholipase C (15, 28, 33) or by directly gating Ca2+-permeable ion channels (15, 28, 33, 37). To this end, we used three different approaches: 1) hemolysis studies to examine osmotic fragility, 2) a Coulter counter to measure the volume of osmotically stressed cells, and 3) the whole cell patch-clamp technique to study membrane currents.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Animals. Mudpuppies (Necturus maculosus) were obtained from a local vendor (Lemberger, Oshkosh, WI) and kept in well-aerated, aged tap water at 5-10°C for <= 6 days before use. They were anesthetized with 3-aminobenzoic acid ethyl ester (MS-222, 1%) and killed by decapitation. Blood was obtained from a midventral incision and collected into tubes coated with heparin (10,000 U/ml). Immediately after exsanguination, the blood was spun in a centrifuge (Hermel-Z230, National Labnet, Woodbridge, NJ) at 1,000 rpm for 1 min. The supernatant was aspirated and replaced with an equal volume of amphibian Ringer solution. This process of spinning and washing the cells was repeated twice.

Osmotic fragility. Osmotic fragility was examined by determining the degree of cell lysis for a suspension of RBCs in hypotonic Ringer solution. The level of hemolysis was determined via a turbidity (cloudy-to-clear) shift that occurs when the integrity of the plasma membrane is compromised. This was detected with a spectrophotometer (Spectronic 20D, Milton Roy) 10, 15, or 20 min after blood (30-50 ml) was added to saline solutions (3 ml) of different osmolalities and compositions. Spectrophotometric experiments were conducted at 625 nm, because this wavelength provided the greatest difference in optical density (OD) between intact and lysed cells (2).

A hemolytic index (HI, percent) was determined using the following formula: HI(%) = (OD of test compound - OD of negative control)/(OD of positive control - OD of negative control) × 100, where the positive control was blood in distilled water, the negative control referred to blood in diluted Ringer solution, and the test compound was blood in diluted Ringer solution containing a specific pharmacological agent. All reported HI values were calculated using a concentration for the negative control that gave an OD reading between 0.025 and 0.030. We chose this region of OD as a standard for comparison, because approximately one-half the cells in suspension were intact at this dilution.

Coulter counter. Cell volume distribution curves were obtained by electronic sizing with use of a Coulter counter (model Z2) with Channelyzer (Coulter Electronics, Hialeah, FL). Mean cell volume was taken as the mean volume of the distribution curves. The diameter of the aperture tube orifice was 200 µm, and the metered volume was 0.5 ml. Absolute cell volumes were obtained using polystyrene latex beads (20.13 µm diameter or 4.271 × 103 fl volume) as standards (Coulter). Experiments with the latex beads showed that measured volumes were unaffected by changes in osmolality and ionic composition within the ranges used for this study. Cell suspensions were diluted 4,000-fold with amphibian Ringer solution or 2,000-fold with amphibian Ringer solution followed by a 1-fold dilution with distilled water to give a final cell density of ~5,000 cells/ml.

As described by others (16, 36), a percent volume recovery at x minutes after hypotonic exposure was calculated as follows: [(Vmax - Vx min)/(Vmax - V0)] × 100, where Vmax is the peak relative cell volume, V0 is the initial relative volume or 1, and Vx min is the relative cell volume measured x min after hypotonic exposure. We also used the peak relative volume for the control when assessing the effect of a pharmacological agent added at 0 min. A percent volume decrease was calculated as follows: (%recoveryexp /%recoverycon) × 100, where recoveryexp and recoverycon are experimental and control recovery, respectively, and maximal recovery in hypotonic Ringer solution is 100%.

Patch clamp. Patch pipettes were fabricated from Kovar sealing glass (Corning model 7052, 1.50 mm outside diameter, 1.10 mm inside diameter, Garner Glass, Claremont, CA) by means of a two-pull method (model PP-7, Narishige). Pipette tips were fire polished (model MF-9, Narishige) to give a direct-current resistance of ~5-8 MOmega in symmetrical 100 mM KCl solutions. All pipette solutions were filtered immediately before use with a 0.22-µm membrane filter (Millex-GS, Bedford, MA), and the pipettes were held in a polycarbonate holder (E. W. Wright, Guilford, CT). Membrane currents were measured with a 1010-Omega feedback resistor in a head stage (CV-201A, Axon Instruments, Foster City, CA) with a variable-gain amplifier set at 1 mV/pA (Axopatch 200A, Axon Instruments). The current signals were filtered at 1 kHz through a four-pole low-pass Bessel filter and digitized at 5 kHz with an IBM-486 computer.

Data were acquired and analyzed with P-Clamp (version 6, Axon Instruments). Data were acquired during 100-ms voltage pulses, and the command potential was set to -15 mV (close to the resting potential for RBCs) for 100 ms between each pulse. All voltage measurements refer to the cell interior.

RBCs, attached to glass coverslips (5 mm diameter, Bellco Biotechnology, Vineland, NJ) with poly-D-lysine (150,000-300,000; 1 mg/ml), were placed in a specially designed open-style chamber (250 µl volume, Warner Instruments, Hamden, CT). The bath solution could be changed by a six-way rotary valve (Rheodyne, Cotati, CA). The whole cell configuration was achieved after formation of a gigaohm seal (cell-attached configuration) by applying suction to disrupt the patch of membrane beneath the pipette or by applying a large voltage (>200 mV) to the patch. A sudden increase in the capacitance current transient accompanied disruption of the membrane.

Solutions. Amphibian Ringer solution consisted of (in mM) 110 NaCl, 2.5 KCl, 1.8 CaCl2, 0.5 MgCl2, 5 glucose, and 10 HEPES (titrated to pH 7.4 with NaOH). A low-Na+-Ringer solution was prepared by substituting choline chloride for NaCl (used for all experiments with gramicidin), and a 0.5× Ringer solution was obtained by mixing equal volumes of Ringer solution and distilled water. A stock solution of gramicidin was dissolved in methanol; stock solutions of A-23187 (Ca2+ ionophore calcimycin) and glibenclamide were prepared with DMSO. All nonaqueous stock solutions were mixed at 1,000× the final concentration and then diluted 1,000× to give an appropriate working concentration, thereby diluting the vehicle an equivalent amount. All stock aqueous solutions (e.g., ATP, hexokinase, apyrase) were diluted 100× to give an appropriate final concentration.

Patch pipettes were filled with an intracellular Ringer solution containing (in mM) 100 KCl, 3.5 NaCl, 1.0 MgCl2, 1.0 CaCl2, 2.0 EGTA, 5 glucose, 1.0 Mg-ATP, 0.5 GTP, and 5.0 HEPES (titrated to pH 7.4 with KOH). During seal formation, the extracellular solution contained (in mM) 105 NaCl, 2.5 KCl, 1.8 CaCl2, 0.5 MgCl2, 5 glucose, and 10.0 HEPES (pH 7.4). An isosmotic high-K+ bath contained (in mM) 105 KCl, 2.5 NaCl, 1.8 CaCl2, 0.5 MgCl2, 5 glucose, and 10.0 HEPES (pH 7.4). A hypotonic (0.5×) high-K+ bath contained (in mM) 2.5 NaCl, 50 KCl, 1.8 CaCl2, 0.5 MgCl2, 5 glucose, and 10.0 HEPES (pH 7.4). Currents carried by ATP at physiological concentrations are below the limits of detection (36). Accordingly, we used abnormally high ATP concentrations in the bath and pipette solutions to examine the presence of a putative ATP conductance. An isosmotic ATP solution contained (in mM) 100 Tris-ATP, 1.0 CaCl2, 1.0 MgCl2, 5 glucose, and 10.0 HEPES (pH 7.4). A hypotonic ATP solution contained (in mM) 50 or 10 Tris-ATP, 1.0 CaCl2, 1.0 MgCl2, 5 glucose, and 10 HEPES (pH 7.4).

For hemolysis experiments, cells were incubated with a pharmacological agent or its vehicle for 1-10 min before experimentation. For cell volume studies, pharmacological agents were added with hypotonic exposure (0 min) or at peak cell volume (5 min after hypotonic stress). Osmolality of solutions was measured with a vapor pressure osmometer (model 5500, Wescor, Logan, UT). Chemicals were purchased from Sigma Chemical (St. Louis, MO), Alexis Biochemicals (San Diego, CA), and ICN (Costa Mesa, CA). All experiments were conducted at room temperature (21-23°C).

Statistics. Values are means ± SE. The statistical significance of an experimental procedure was determined by a paired Student's t-test or least significant difference test with paired design of ANOVA/multivariate ANOVA, as appropriate (Data Desk software, Ithaca, NY). P < 0.05 was considered significant. Each animal served as its own control, and cell volumes at specific times were tested against each other. For patch-clamp studies, each cell served as its own control.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Osmotic fragility studies. Although osmotic fragility depends on several factors, we first examined this property as one assessment of a cell's ability to regulate volume in a hypotonic medium. The OD, measured at a concentration of amphibian Ringer solution where ~50% of the cells in suspension were intact (20.7 ± 1.3 mosmol/kgH2O), was 0.029 ± 0.002 (n = 7 experiments; Fig. 1). To determine whether osmotic fragility depended on ATP, we repeated the hemolysis assay with this nucleotide at 50 µM in the extracellular medium. In this case, the OD measured at the same concentration for the control was 0.042 ± 0.004 (n = 7, P < 0.01; Fig. 1), indicating a 45% decrease in cell lysis compared with the control.


View larger version (29K):
[in this window]
[in a new window]
 
Fig. 1.   Extracellular ATP decreased osmotic fragility. Cells were incubated for 1-10 min in isosmotic Ringer solution with ATP (50 µM, n = 7), hexokinase (hexokin, 2.5 U/ml, n = 6), or glibenclamide (glibencl, 100 µM, n = 6) before dilution. Control solution was diluted amphibian (high-Na+) Ringer solution, experimental solution was diluted Ringer solution with pharmacological agent, and experimental + ATP solution was a Ringer solution that contained antagonist and nucleotide. Values are means ± SE of optical density (OD) measurements obtained at concentrations of Ringer solution that provided an OD of 0.025-0.030 for control. * P < 0.05; ** P < 0.01.

We next looked at the effect of the ATP scavenger hexokinase on osmotic fragility. Hexokinase is an enzyme that traps ATP by transferring its gamma -phosphoryl group to a variety of C6 sugars, such as to the hydroxyl group on C-6 of glucose (27). Hexokinase (2.5 U/ml, 1 mM glucose) decreased the OD from 0.025 ± 0.003 to 0.017 ± 0.003 (n = 6, P < 0.01; Fig. 1), giving an HI of 32%. In addition, we examined the effect of glibenclamide, an antagonist of ATP-dependent K+ channels, on osmotic fragility (10). This inhibitor (100 µM) decreased the OD from 0.027 ± 0.002 to 0.021 ± 0.002 (n = 6, P < 0.05; Fig. 1), which resulted in an HI of 22%. As illustrated in Fig. 1, ATP (50 µM) reversed the inhibitory effect of glibenclamide, increasing the OD to 0.025 ± 0.004, which was not significantly different from the control (n = 6).

Cell volume studies. When RBCs were placed in a hypotonic (0.5×) Na+-Ringer solution, they quickly swelled and then slowly and spontaneously decreased in volume (Fig. 2A). As illustrated in Fig. 2A, the relative volume with ATP (50 µM) was significantly lower than the control for all measurements beyond 5 min (n = 7, P < 0.05 at >= 5 min). The percent volume decrease of the control was only 39% that of ATP at 30 min, whereas it was 73% that of ATP by 90 min. We also added ATP to the extracellular medium 5 min after hypotonic shock, when the cells were maximally swollen and when it appeared that endogenous K+ channels were activated. Even when added at this time, ATP (50 µM) still enhanced cell volume recovery (n = 6, P < 0.05 at >10 min). Interestingly, addition of ATP (50 µM) to human RBCs had no effect on the percent volume decrease for cells exposed to a hypotonic (0.67×) Ringer solution (n = 6, data not shown; human RBCs did not express a well-developed RVD response; nonetheless, on the basis of our studies with Necturus RBCs, we were interested in determining whether ATP also could influence volume in this cell type).


View larger version (20K):
[in this window]
[in a new window]
 
Fig. 2.   Effect of extracellular ATP and A-23187 on cell volume after hypotonic shock. At time 0, cells were abruptly exposed to a hypotonic (0.5×) high-Na+-Ringer solution, which caused a rapid initial increase in volume followed by a gradual recovery toward basal values, despite continued presence of hypotonic buffer (control). A: cell volume recovery was enhanced by addition of extracellular ATP (50 µM, n = 7) to extracellular medium at 0 min. Inset: percent volume recovery at 30 min; con, control. B: recovery also was enhanced by addition of A-23187 (1 µM, n = 6) 5 min after hypotonic shock (arrow). Inset: percent volume recovery at 20 min; A23, A-23187. Vehicle (DMSO, diluted 1,000×, n = 6) had no effect on cell volume recovery. Values are means ± SE.

We previously demonstrated that volume decrease in Necturus RBCs is enhanced by a Ca2+-dependent process and that RVD depends on activation of a K+ conductance (2, 22). To further examine the importance of Ca2+ in cell volume recovery, we added the Ca2+ ionophore A-23187 to the extracellular medium 5 min after hypotonic shock. As illustrated in Fig. 2B, application of A-23187 (1 µM) at this point in time enhanced cell volume recovery (n = 6, P < 0.05 at >5 min). For example, at 20 min, the percent volume decrease of the control was only 38% that of cells exposed to A-23187.

We next examined the effect of extracellular ATP, A-23187, and gramicidin (a cationophore that was used with a choline-Ringer solution to maintain a high K+ permeability) on cells bathed in an isosmotic Ringer solution. As illustrated in Fig. 3, there was a significant reduction in cell volume 2 min after the addition of ATP (50 µM) that was followed by a slower, spontaneous recovery to the original volume (n = 6, P < 0.05 at 11-70 min). The Ca2+ ionophore A-23187 produced a similar response, initially reducing cell volume, which was followed by a slower volume recovery (n = 9, P < 0.05 at 11-70 min; Fig. 3). By 80 min there was no significant difference in mean volumes between cells treated with ATP or A-23187 and the control cells (Fig. 3). Finally, gramicidin (5 µM, choline-Ringer solution) also caused a decrease in cell volume (n = 9, P < 0.001 at >4 min after gramicidin; Fig. 3). A lack of volume recovery in this instance was most likely due to the use of a choline-Ringer solution, which could inhibit a regulatory volume increase.


View larger version (24K):
[in this window]
[in a new window]
 
Fig. 3.   Effect of ATP, A-23187, and gramicidin on relative cell volume for red blood cells (RBCs) in isosmotic Ringer solution. Cells remained in isosmotic Ringer solution for 90 min (control). Addition of ATP (50 µM, n = 6) at 9 min (arrow) caused a rapid decrease in relative volume followed by a gradual recovery toward basal values. Addition of A-23187 (0.5 µM, n = 9) at 9 min (arrow) also caused a rapid initial decrease in relative volume followed by a gradual recovery toward basal values. Addition of gramicidin (5 µM, n = 9) at 11 min caused a rapid decrease in relative volume (lack of volume recovery with gramicidin was most likely due to choline-Ringer solution, which could inhibit a regulatory volume increase). Values are means ± SE.

To further examine whether extracellular ATP enhanced volume decrease, we used enzymes that dephosphorylate ATP. As shown in Fig. 4A, hexokinase (2.5 U/ml) inhibited cell volume recovery after hypotonic shock (n = 6, P < 0.05 at >10 min), reducing the percent volume decrease to 44% of control values at 90 min. Hexokinase was ineffective when glucose was omitted from the bath solution (n = 4, not shown). In addition, the inhibitory effect of hexokinase was reversed with the Ca2+ ionophore A-23187 (1 µM), such that mean values for relative cell volume with hexokinase and ionophore were significantly below control values for all measurements after hypotonic exposure (P < 0.001; Fig. 4A). For example, at 90 min cells exposed to both hexokinase and A-23187 had a 232% greater volume recovery than cells bathed in hexokinase alone and a 46% greater recovery than the control cells. Furthermore, gramicidin (5 µM) reversed the inhibitory effect of hexokinase when added 5 min after hypotonic shock (Fig. 4B). In this case, mean values for relative cell volume with both hexokinase and gramicidin were significantly below control values for all measurements after hypotonic shock (n = 6, P < 0.001 compared with control; Fig. 4B). At 90 min, cells with hexokinase and gramicidin had a 129% greater volume recovery than the control cells.


View larger version (33K):
[in this window]
[in a new window]
 
Fig. 4.   Effect of ATP scavengers on cell volume after hypotonic shock. At time 0, cells were abruptly exposed to hypotonic (0.5×) Ringer solution, which caused a rapid initial increase in relative volume (control). A and B: cell volume recovery was inhibited by addition of hexokinase (hexokin, 2.5 U/ml, n = 6) to extracellular medium at 0 min. Recovery was enhanced with hexokinase + A-23187 (1 µM, n = 6, A) or hexokinase + gramicidin (gram, 5 µM, n = 6, B). Vehicle for A-23187 (DMSO, diluted 1,000×, n = 8) or for gramicidin (methanol, diluted 1,000×, n = 8) had no effect on cell volume recovery. C: cell volume recovery also was inhibited by addition of diphosphohydrolase apyrase (2.5 U/ml, n = 6) to extracellular medium at 0 min. Recovery was enhanced with apyrase + nonhydrolyzable analog adenosine 5'-O-(3-thiotriphosphate) (ATPgamma S, 50 µM, n = 6). Insets: percent volume recovery at 90 min; gram, gramicidin; apy, apyrase; hex, hexokinase. Values are means ± SE.

We next examined the effect of apyrase on RVD. Apyrase is a diphosphohydrolase that hydrolyzes ATP into AMP and two orthophosphate anions (19). Consistent with the hexokinase results, apyrase (2.5 U/ml) reduced cell volume recovery (n = 6, P < 0.05 at >10 min; Fig. 4C). In this case, the percent volume decrease changed to 76% of the control values at 90 min. In addition, the inhibitory effect of apyrase was reversed with the nonhydrolyzable ATP analog adenosine 5'-O-(3-thiotriphosphate) (ATPgamma S, 50 µM), such that mean values for cell volume with apyrase and ATPgamma S were significantly below control values for all measurements after 0 min (P < 0.001; Fig. 4C). At 90 min, volume recovery was 126% greater in cells with apyrase and ATPgamma S than in cells with apyrase alone and 72% greater than in control cells.

Next we examined the effect of the ATP-dependent K+ channel antagonist glibenclamide on RVD. As illustrated in Fig. 5A, glibenclamide (100 µM) inhibited cell volume recovery (n = 7, P < 0.05 at >10 min), reducing the percent volume decrease to 38% of control values at 30 min. In addition, the inhibitory effect of glibenclamide was reversed with ATP (50 µM), such that mean values for glibenclamide with ATP were significantly lower than for glibenclamide alone (n = 7, P < 0.05 at >= 10 min; Fig. 5A). At 30 min, cells treated with glibenclamide and ATP had a 239% greater volume recovery than cells treated with the antagonist alone. However, cell volume recovery with a combination of glibenclamide and ATP was not significantly different from the control (Fig. 5A). Furthermore, the Ca2+ ionophore A-23187 (1 µM) reversed the inhibitory effect of glibenclamide, such that mean values for glibenclamide with A-23187 were significantly lower than for glibenclamide alone and for the control (n = 7, P < 0.05 at >0 min; Fig. 5A). At 30 min, volume recovery was 517% greater in cells treated with glibenclamide and A-23187 than in cells treated with glibenclamide alone and 136% greater than in control cells.


View larger version (26K):
[in this window]
[in a new window]
 
Fig. 5.   Effect of glibenclamide and gadolinium on cell volume after hypotonic shock. At time 0, cells were abruptly exposed to hypotonic (0.5×) Ringer solution (control). A: cell volume recovery was inhibited by addition of glibenclamide to extracellular medium at 0 min (100 µM, n = 7). Glibenclamide (gliben) + ATP (50 µM) was not significantly different from control (100 nM, n = 7); glibenclamide + A-23187 enhanced cell volume recovery (n = 7). Inset: percent volume recovery at 30 min; glb, glibenclamide. B: cell volume recovery also was inhibited by addition of gadolinium (2.5 U/ml, n = 6, triangle ) to extracellular medium at 0 min. Recovery was enhanced with gadolinium (gadol) + ATP (50 µM, n = 6). Inset: percent volume recovery at 90 min; gad, gadolinium. Values are means ± SE.

The effect of the stretch-activated channel blocker gadolinium also was examined (39). As illustrated in Fig. 5B, gadolinium (10 µM) inhibited cell volume recovery (n = 6, P < 0.05 at >10 min), such that the percent volume decrease changed to 41% of control values at 90 min. Furthermore, the inhibitory effect of gadolinium was reversed with ATP (50 µM, n = 6; Fig. 5B). In this case, the percent volume decrease with both gadolinium and ATP was not significantly different from the control at all time points after hypotonic shock (Fig. 5B).

Patch-clamp studies. We first examined whether extracellular ATP would activate whole cell currents under isosmotic conditions. After addition of ATP (50 µM) to the extracellular bath (isosmotic high-K+-Ringer solution), the conductance gradually increased until a maximum stimulation occurred by ~3-5 min (Fig. 6A). No increase in current was observed in control cells over a similar time period. With ATP, the whole cell conductance increased by 55%: from 2.9 ± 0.1 to 4.5 ± 0.8 nS (n = 4, P < 0.05; Fig. 6B). This change was not associated with a shift in the reversal potential (Erev), which was expected for cells exposed to symmetrical bath and pipette solutions.


View larger version (39K):
[in this window]
[in a new window]
 
Fig. 6.   Whole cell currents for cells in isosmotic or hypotonic KCl-Ringer solution. Cells were maintained at a holding potential of -15 mV and stepped to potentials of -100 to +100 mV in 20-mV intervals. A: whole cell currents for RBC exposed to isosmotic symmetrical KCl solution and enhancement of these currents by addition of ATP (50 µM) to bath. B: corresponding current-voltage (I-V) relationship for control (isotonic) and ATP solutions. C: whole cell currents for RBC exposed to hypotonic (0.5×) high-KCl bath solution, inhibition of these currents by addition of apyrase (2.5 U/ml), and enhancement of currents by addition of ATPgamma S (50 µM) in presence of apyrase. D: corresponding I-V relationship for control (0.5× KCl-Ringer solution, hypo), apyrase, and ATPgamma S + apyrase solutions. Insets: current levels at -100 and +100 mV; apyr, apyrase. Values are means ± SE.

We then determined whether apyrase would alter whole cell currents under hypotonic conditions with a high-KCl-Ringer solution in the pipette and a 0.5× KCl-Ringer solution in the bath. The only major ions of significance with these solutions were K+ and Cl-, and the equilibrium potentials for perfectly cation- and anion-selective conductances were -16.2 and +14.7 mV, respectively. Addition of apyrase (2.5 U/ml) to the bath reduced the whole cell conductance by 74%: from 20.7 ± 2.9 to 5.4 ± 1.3 nS (n = 9, P < 0.001; Fig. 6, C and D). Interestingly, this antagonist did not alter the Erev, which was -8.0 ± 1.0 mV for the control and -8.4 ± 0.6 mV with apyrase (n = 9; Fig. 6D). As might be anticipated, the inhibitory effect of apyrase was not affected by adding ATP (50 µM, n = 6, not shown). In contrast, it was reversed with the nonhydrolyzable analog ATPgamma S (50 µM), which increased the whole cell conductance by 170%: to 14.6 ± 3.6 nS (n = 9, P < 0.05 compared with apyrase), a value that was not significantly different from the control (Fig. 6, C and D). In addition, application of ATPgamma S to a bath containing apyrase changed Erev to -10.3 ± 1.7 mV, a value closer to the equilibrium potential for K+ (EK; n = 9).

We next looked at the effect of glibenclamide on whole cell currents after hypotonic shock. This antagonist (100 µM) decreased the whole cell conductance by 77%: from 13.7 ± 2.4 to 3.2 ± 0.6 nS, which was close to that observed for cells under isosmotic conditions (n = 4, P < 0.01; Fig. 7, A and B). Glibenclamide also caused a slight shift in Erev toward the equilibrium potential for Cl- (ECl): from -8.0 ± 1.0 to -6.8 ± 2.4 mV. Interestingly, addition of ATP (50 µM) partially reversed the inhibitory effect of glibenclamide (Fig. 7A). In this case, the conductance with ATP and glibenclamide together increased to 7.7 ± 1.1 nS, and Erev changed to -10.4 ± 1.3 mV, a displacement toward EK (n = 3; Fig. 7B).


View larger version (42K):
[in this window]
[in a new window]
 
Fig. 7.   Effect of glibenclamide and gadolinium on whole cell currents stimulated by hypotonic Ringer solution. Cells were maintained at a holding potential of -15 mV and stepped to potentials of -100 to +100 mV in 20-mV intervals. A: whole cell currents for RBC exposed to 0.5× high-KCl bath solution, inhibition of these currents by addition of glibenclamide (100 µM), and enhancement of currents by addition of ATP (50 µM) in presence of glibenclamide. B: corresponding I-V relationship for control, glibenclamide, and ATP + glibenclamide solutions. C: whole cell currents for RBC exposed to 0.5× high-KCl bath solution, inhibition of these currents by addition of gadolinium (10 µM), and enhancement of currents by addition of ATP (50 µM) in presence of gadolinium. D: corresponding I-V relationship for control, gadolinium, and ATP + gadolinium solutions. Insets: current levels at -100 and +100 mV. Values are means ± SE.

Gadolinium (10 µM) also inhibited whole cell currents in cells exposed to a 0.5× KCl-Ringer solution (Fig. 7C). This agent decreased the conductance by 95%: from 11.6 ± 5.1 to 0.6 ± 0.1 nS (n = 3, P < 0.01; Fig. 7D). It also shifted Erev toward ECl: from -5.7 ± 0.6 to -0.8 ± 0.7 mV. Addition of ATP caused a reversal of the inhibitory effect caused by gadolinium, increasing the conductance to 9.1 ± 4.1 nS (n = 3; Fig. 7, C and D), which was not significantly different from the control. In addition, Erev changed to -8.2 ± 1.4 mV, a value nearer EK.

To further examine the properties of ATP efflux during cell swelling, we used solutions in which this nucleotide was the only significant charge carrier. For this purpose, abnormally high ATP concentrations were used in the bath and pipette solutions to enhance measurements of ATP transport (see METHODS). With symmetrical 100 mM ATP solutions, whole cell currents were very small but measurable (Fig. 8A). In this case, the conductance was 1.0 ± 0.3 nS (n = 3), and Erev was -0.7 ± 0.5 mV (n = 3), which was not significantly different from zero (Fig. 8B). In contrast, cell swelling induced with a hypotonic ATP bath (50 mM) caused an increase in whole cell currents (Fig. 8A). Under these conditions, the conductance gradually increased until a maximum stimulation occurred by ~3-5 min. No increase in current was observed in control cells over a similar time period. With the hypotonic ATP bath, the whole cell conductance increased 550%: to 6.5 ± 2.2 nS (n = 6, P < 0.05; Fig. 8B). In addition, there was a shift in Erev from -0.7 ± 0.5 to +4.4 ± 2.4 mV, a value close to that predicted for currents carried by ATP (at pH 7.4, the 4th OH on ATP would be 74% ionized, giving the molecule a net charge of -3.74 or a theoretical equilibrium potential for ATP of +4.7 mV; at pH 7.4 Tris would be 83% ionized, giving an equilibrium potential for Tris of around -70 mV). Consistent with these observations, a further reduction in extracellular ATP to 10 mM resulted in a larger shift in Erev. With this 10-fold difference in the ATP concentration, Erev was +12.2 ± 2.5 mV (n = 9, the theoretical equilibrium potential for ATP for these solutions was +15.5 mV; Fig. 7, A and B). This solution also caused an increase in conductance to 9.9 ± 1.5 nS, which probably resulted from a further decrease in osmolality.


View larger version (45K):
[in this window]
[in a new window]
 
Fig. 8.   ATP conductance was activated by cell swelling. Cells were maintained at a holding potential of -15 mV and stepped to potentials of -100 to +100 mV in 20-mV intervals. A: whole cell currents for cell exposed to symmetrical 100 mM Tris-ATP solutions and enhancement of these currents by reduction of bath concentration to 50 and 10 mM Tris-ATP. B: corresponding I-V relationship for control (100 mM ATP, iso), 50 mM Tris-ATP (hypo), and 10 mM Tris-ATP (hypo) solutions. C: whole cell currents for cell with 100 mM Tris-ATP in pipette and 10 mM ATP in bath and inhibition of these currents with glibenclamide (100 µM) or gadolinium (10 µM). D: corresponding I-V relationship for control, glibenclamide, and gadolinium solutions. Insets: current levels at -100 and +100 mV. Values are means ± SE.

Having established the presence of an ATP conductance, we next tried to inhibit it with pharmacological agents. Glibenclamide (100 µM) blocked ATP currents, reducing whole cell conductance by 77%: from 9.9 ± 1.5 to 2.3 ± 0.6 nS (n = 7, P < 0.01, measured with 100 mM ATP in the pipette and 10 mM ATP in the bath; Fig. 8, C and D). In addition, this agent shifted Erev from +12.2 ± 2.5 to +4.1 ± 4.2 mV, consistent with inhibition of an anion conductance. Gadolinium (10 µM) also inhibited an ATP conductance, reducing it by 46%: to 5.3 ± 1.7 nS (n = 4, P < 0.05; Fig. 8, C and D). However, gadolinium did not significantly alter Erev. In addition, whereas glibenclamide essentially reduced inward and outward currents equally, gadolinium had a greater inhibitory effect on outward negative current, suggesting inhibition of ATP efflux.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The major finding of this study was that extracellular ATP enhanced cell volume recovery in Necturus RBCs when exposed to a hypotonic medium. Our results are most consistent with swelling-induced stimulation of ATP release through a glibenclamide- and gadolinium-sensitive conductance. This, in turn, led to a rise in intracellular Ca2+, thereby increasing K+ efflux, which contributed to solute loss and recovery of cell volume.

Key evidence for the dependence of cell volume decrease on ATP was obtained from a series of experiments in which the extracellular ATP concentration was altered. For example, endogenous extracellular ATP levels were reduced with hexokinase and apyrase, which had the effect of increasing osmotic fragility, decreasing cell volume recovery in response to hypotonic shock, and blocking whole cell currents that were activated with a hypotonic bath. It seems unlikely that these ATP scavengers acted in a nonspecific manner, because hexokinase was ineffective in the absence of glucose, and the inhibitory effect of apyrase was reversed with ATPgamma S (27). In contrast, addition of micromolar amounts of exogenous extracellular ATP had the opposite effect, reducing osmotic fragility, enhancing the percent volume decrease, and activating whole cell currents in isosmotic and hypotonic media. Furthermore, ATP caused cells to shrink under isosmotic conditions, presumably by eliciting a change that mimicked the response that occurs when cells are stimulated with hypotonic exposure. Although we did not measure the level of endogenous extracellular ATP, the concentration of exogenous ATP we added was similar to that used by others (1, 31, 32, 36). Taken together, these observations demonstrate a positive role for extracellular ATP modulation of cell volume in response to hypotonic shock.

Our experimental protocols for reducing endogenous extracellular ATP and the effect of these changes on ion efflux are consistent with a report by Schwiebert et al. (27). They also used hexokinase and apyrase to reduce the extracellular ATP concentration and found that these agents prevented cAMP and protein kinase A activation of outwardly rectifying whole cell Cl- currents in a human airway epithelial cell line. In addition, our finding that cell volume decrease was stimulated by extracellular nucleotides is consistent with reports for several other cell types. Wang et al. (36) reported that increases in cell volume lead to efflux of ATP through a conductive pathway in rat hepatoma cells. This nucleotide, in turn, acts as an autocrine that couples increases in cell volume to opening of Cl- channels through stimulation of P2 receptors. Similarly, Kim et al. (17) demonstrated a potentiation of RVD in response to extracellular UTP, which promotes Ca2+ mobilization and net K+ efflux in human submandibular salivary gland duct cells. Furthermore, Taylor et al. (33) found that hypotonic shock triggers ATP release from human airway epithelial cells and suggest that extracellular ATP plays a role in RVD. Interestingly, our initial studies with human RBCs indicated that these cells do not display a well-developed RVD response. In addition, extracellular ATP had no effect their size. Apparently, there is a fundamental difference in the way nucleated and anucleated RBCs regulate their volume, at least in response to hypotonic shock.

A logical question stemming from our observations is, What was the source of endogenous external ATP? This nucleotide was not a component of amphibian Ringer solution, nor was it normally added to the extracellular bath solution used for patch-clamp experiments. Furthermore, it has been shown by others that ATP cannot act as a blood-borne ligand, because it is subject to quick degradation in the general circulation (9). Thus, except for a few experiments where ATP was added as an agonist to the extracellular medium, the only source of this nucleotide was the RBCs themselves. In addition, our patch-clamp studies indicated the presence of an ATP-permeable conductance that was activated during cell swelling, thereby providing a pathway for ATP efflux in swollen cells. The presence of an ATP conductance is consistent with reports by Wang et al. (36) and Schwiebert et al. (27), who also showed that ATP can be released from cells via a conductive pathway.

Interestingly, the ATP conductance in mudpuppy RBCs was inhibited by glibenclamide. We originally chose this antagonist, because it has been shown to block ATP-dependent K+ channels (10) and because RVD by mudpuppy RBCs depends on a K+ conductance that is activated during cell swelling (2, 22). Although glibenclamide increased osmotic fragility, reduced cell volume recovery, and blocked whole cell currents in swollen cells, its inhibitory effects were reversed by the addition of extracellular ATP. This indicated that the site of action for glibenclamide was "upstream" to the site affected by ATP, suggesting that glibenclamide blocked ATP release from the cell. This hypothesis was supported by our patch-clamp studies, in which glibenclamide was shown to be a potent inhibitor of the ATP conductance. Glibenclamide inhibition of an ATP conductance is not unique to this cell type; it also has been reported for a human airway epithelial cell line (27).

In this study we also demonstrated that ATP enhanced RVD by stimulating a K+ permeability. This was shown pharmacologically using the cationophore gramicidin with a choline-Ringer solution. With this solution, K+ and Cl- were the only two permeable ions of significance, and addition of gramicidin ensured a continual high K+ permeability. Gramicidin consistently reversed the inhibitory effect of hexokinase. In addition, for the cell volume experiments, it did not matter whether gramicidin was added at 0 or 5 min. The effect of gramicidin was examined at 5 min, because 5 min corresponded with maximum cell swelling, indicating that several minutes were required for endogenous K+ channels to activate after hypotonic stress. Thus percent volume recovery was enhanced regardless of whether the K+ permeability was artificially enhanced with gramicidin at the time of hypotonic stress or at 5 min, even in the presence of an ATP scavenger. In addition, gramicidin caused cells to shrink under isosmotic conditions. This is consistent with these cells having a low K+ permeability under normal conditions and an elevated K+ permeability during hypotonic stress. In fact, in a previous report we showed that mudpuppy RBCs have a high basal Cl- permeability and that K+ efflux is a rate-limiting step for cell volume recovery in response to hypotonic shock (2).

Moreover, our electrophysiological studies demonstrated that the ATP-stimulated K+ permeability was a conductive pathway. For example, addition of ATP or ATPgamma S to a hypotonic KCl bath consistently changed Erev away from ECl and toward EK, indicating stimulation of a K+ conductance. Nonetheless, we cannot rule out the possibility that ATP also stimulated a Cl- permeability concomitantly with its activation of a K+ channel. However, the putative presence of voltage-sensitive, volume-sensitive, or ATP-sensitive Cl- channels does not alter our conclusion that ATP stimulated a K+ conductance during cell swelling.

Similar to gramicidin, the Ca2+ ionophore A-23187 also increased percent volume recovery whether it was added at 0 or 5 min, indicating that the rate of cell volume recovery was sensitive to the level of free Ca2+. Furthermore, A-23187 caused cells to shrink under isosmotic conditions, presumably by eliciting a change that mimicked the response that occurs when cells are stimulated by hypotonic shock. We cannot, however, rule out the possibility that A-23187 caused an RVD-type response that was fundamentally different from the swelling-induced response.

The Ca2+ ionophore also reversed the inhibitory effects of apyrase and hexokinase, indicating that the Ca2+-dependent step is "downstream" to the site of action of ATP. Furthermore, the inhibitory effects of glibenclamide and gadolinium, two agents that blocked the ATP conductance, also were reversed with A-23187. Taken together, these observations are consistent with a presumed rise in intracellular Ca2+ occurring after cell release of ATP. It is worth noting that RBCs exposed to gramicidin or A-23187 stabilized their volume at a smaller size than the control cells. In fact, under control conditions, the percent volume recovery was only ~40-50% by 90 min. Although this level of RVD was less than that expressed by several other cell types (8, 12, 13, 16), it is consistent with our previous studies on Necturus RBCs (2, 22). It is possible that, under the conditions of our study, control cells lacked a sufficient rise in intracellular Ca2+ or an adequate increase in K+ permeability to display a full RVD response. Alternatively, these cells may naturally never reach a level of RVD that is equivalent to cell volume recovery with A-23187 and gramicidin or that expressed by other cell types.

We previously reported that gadolinium increases osmotic fragility, inhibits cell volume recovery in response to hypotonic shock, and blocks whole cell currents in swollen cells (2). Furthermore, the inhibitory effects of this agent were reversed with A-23187. On the basis of the information we had at that time, we concluded that the Ca2+ influx step during cell volume decrease occurred through a Ca2+-permeable, stretch-activated channel. However, in this study we show that the inhibitory effect of gadolinium on cell volume recovery and on whole cell currents was reversed by adding micromolar amounts of extracellular ATP, suggesting that this agent blocked ATP efflux. This was further supported by our patch-clamp studies in which it was shown that gadolinium blocked the ATP conductance. In light of this new evidence, our results are most consistent with ATP efflux via a stretch-activated conductance, which in turn leads to Ca2+ influx (Fig. 9). However, we cannot rule out other effects of gadolinium or the presence of other stretch-activated conductive pathways.


View larger version (22K):
[in this window]
[in a new window]
 
Fig. 9.   Proposed model for regulatory volume decrease signaling by extracellular ATP during hypotonic swelling in Necturus RBCs. Cell swelling leads to ATP release via a glibenclamide- and gadolinium-sensitive conductance. Extracellular ATP then stimulates an increase in intracellular Ca2+, presumably by binding to a P2 receptor (P2R, nature of which remains to be determined). This, in turn, activates a K+ conductance, thereby leading to solute efflux and cell volume recovery. Dashed lines, site of inhibition by specific antagonists. Narrow arrows, effect of adding exogenous ionophores.

In some instances, there appear to be two components to the rate of volume recovery by Necturus RBCs: an initial faster component followed by a slower phase. This phenomenon may have resulted from the presence of different populations of RBCs. Unlike those in mammals and anurans, RBCs in urodeles complete their maturation in circulation, which takes ~1 mo (34). During that time, the cells change their morphology from a round contour to a more oblong shape. The circulating RBCs also have the ability to synthesize DNA, RNA, and proteins and are capable of undergoing cell division (34). Thus RBCs at different levels of maturation may respond to hypotonic shock with different rates of volume recovery. A further complicating factor is that there can be three distinct populations of RBCs in amphibians: two larval forms, one originating from the liver and the other from the mesonephros, and an adult form that appears after metamorphosis (34). Given the neotenous nature of Necturus, it is conceivable that these species possess more than one form of RBC.

Alternatively, a single RBC population could possess several transport pathways, each leading to solute efflux and subsequent cell shrinkage. For example, cell swelling in RBCs from Amphiuma, a species similar to Necturus, stimulates conductive and electroneutral K+ transport mechanisms, with the latter contributing more significantly to net K+ flux (4). It also has been reported that solute flux pathways activated with hypotonic shock may only remain active for a short period of time. For instance, Ehrlich ascites tumor cells display a Cl- transport pathway that is activated with cell swelling but inactivates within the next 10 min (13). Thus it is conceivable that the initial phase of cell volume recovery in Necturus RBCs depends on a K+ permeability pathway that no longer contributes to K+ flux during the slower phase.

Finally, on the basis of the evidence we present in this report, it is compelling to conclude that extracellular ATP regulates RVD in Necturus RBCs. We cannot, however, rule out the possibility that this nucleotide may have caused superimposed cell shrinkage that was unrelated to RVD, thereby enhancing cell volume decrease. For example, under hypotonic conditions the apyrase-sensitive current was greater than the ATPgamma S-induced current for voltages less than -25 mV. However, these two currents were not significantly different for positive voltages. Furthermore, we have not established a pharmacological potency profile for ATP and its analogs. Another factor to consider concerning the role of ATP in RVD is that whole cell currents induced by extracellular ATP under isosmotic conditions were significantly less than currents induced with hypotonic shock. This observation suggests that an additional mechanism may be involved when cells are swollen, possibly analogous to a report concerning the Ca2+ sensitivity of Amphiuma RBCs (5). With Amphiuma, Ca2+ is stimulatory to K+ loss in isosmotic and hypotonic media; however, the Ca2+ sensitivity of swollen cells is greater than that for cells at normal volume. The author concluded that cell swelling increases the Ca2+ sensitivity of the Ca2+-activated K+ transport pathway (5). By analogy, it is possible that swelling of Necturus RBCs increased their sensitivity to ATP and/or Ca2+.

In conclusion, cell volume decrease in mudpuppy RBCs was stimulated by extracellular ATP. Cell swelling activated an ATP conductance, which, in turn, stimulated a Ca2+-dependent step, thereby leading to K+ efflux and subsequent cell volume recovery. The coupling of swelling-activated ATP release and subsequent cell volume decrease represents a novel mechanism for osmotic regulation of cell function.


    ACKNOWLEDGEMENTS

We thank Lawrence Tate for conducting the experiments with human RBCs and Sharon Provinzano (Ripon College) and Erica Smith (Ripon College) for technical assistance in running the laboratory. We also thank Dr. Fiona L. Stavros (Texas Biotechnology, Houston, TX) for developing the hemolytic index and Dr. Robert L. Wallace (Ripon College) for helpful discussions and suggestions on the manuscript.


    FOOTNOTES

Research support was provided by National Science Foundation Grant MCB-9603568.

Portions of this study were presented in abstract form at Experimental Biology '99, Washington, DC, April 1999.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Address for reprint requests and other correspondence: D. B. Light, Dept. of Biology, Ripon College, 300 Seward St., Ripon, WI 54971-0248 (E-mail: LightD{at}Ripon.edu).

Received 20 January 1999; accepted in final form 11 May 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Arellano, R. O., E. Garay, and R. Miledi. Cl- currents activated via purinergic receptors in Xenopus follicles. Am. J. Physiol. 274 (Cell Physiol. 43): C333-C340, 1998[Abstract/Free Full Text].

2.   Bergeron, L. J., A. J. Stever, and D. B. Light. Potassium conductance activated during regulatory volume decrease by mudpuppy red blood cells. Am. J. Physiol. 270 (Regulatory Integrative Comp. Physiol. 39): R801-R810, 1996[Abstract/Free Full Text].

3.   Burnstock, G. Purinergic nerves. Pharmacol. Rev. 24: 509-581, 1972[Medline].

4.   Cala, P. M. Cell volume regulation by Amphiuma red blood cells: the role of Ca2+ as a modulator of alkali metal/H+ exchange. J. Gen. Physiol. 82: 761-784, 1983[Abstract].

5.   Cala, P. M. Volume regulation by Amphiuma red blood cells: characteristics of volume-sensitive K/H and Na/H exchange. Mol. Physiol. 8: 199-214, 1985.

6.   Dubyak, G. R., and C. El-Moatassim. Signal transduction via P2-purinergic receptors for extracellular ATP and other nucleotides. Am. J. Physiol. 265 (Cell Physiol. 34): C577-C606, 1993[Abstract/Free Full Text].

7.   Fitz, J. G., and A. H. Sostman. Nucleotide receptors activate cation, potassium, and chloride currents in a liver cell line. Am. J. Physiol. 266 (Gastrointest. Liver Physiol. 29): G544-G553, 1994[Abstract/Free Full Text].

8.   Grinstein, S., and J. K. Foskett. Ionic mechanisms of cell volume regulation in leukocytes. Annu. Rev. Physiol. 52: 399-414, 1990[Medline].

9.   Gordon, J. L. Extracellular ATP: effects, sources and fate. Biochem. J. 233: 309-319, 1986[Medline].

10.   Hassessian, H., P. Bodin, and G. Burnstock. Blockade by glibenclamide of the flow-evoked endothelial release of ATP that contributes to vasodilation in the pulmonary vascular bed of the rat. Br. J. Pharmacol. 109: 466-472, 1993[Abstract].

11.   Hazama, A., and Y. Okada. Ca2+ sensitivity of volume-regulatory K+ and Cl- channels in cultured human epithelial cells. J. Physiol. (Lond). 402: 687-702, 1988[Abstract].

12.   Hoffman, E. K., and P. B. Dunham. Membrane mechanisms and intracellular signaling in cell volume regulation. Int. Rev. Cytol. 161: 173-262, 1995[Medline].

13.   Hoffman, E. K., I. H. Lambert, and L. O. Simonsen. Separate, Ca2+-activated K+ and Cl- transport pathways in Ehrlich ascites tumor cells. J. Membr. Biol. 91: 227-244, 1986[Medline].

14.   Hughes, B. A., and M. Takahira. ATP-dependent regulation of inwardly rectifying K+ current in bovine retinal pigment epithelial cells. Am. J. Physiol. 275 (Cell Physiol. 44): C1372-C1383, 1998[Abstract/Free Full Text].

15.   Ikeda, K., M. Suzuki, M. Furukawa, and T. Takasaka. Calcium mobilization and entry induced by extracellular ATP in the non-sensory epithelial cell of the cochlear lateral wall. Cell Calcium 18: 89-99, 1995[Medline].

16.   Jorgensen, N. K., S. Christensen, H. Harbak, A. M. Brown, I. H. Lambert, E. K. Hofmann, and L. O. Simonsen. On the role of calcium in the regulatory volume decrease (RVD) response in Ehrlich mouse ascites tumor cells. J. Membr. Biol. 157: 281-299, 1997[Medline].

17.   Kim, H. D., J. W. Bowen, M. R. James-Kracke, L. A. Landon, J. M. Camden, J. E. Burnett, and J. T. Turner. Potentiation of regulatory volume decrease by P2U purinoceptors in HSG-PA cells. Am. J. Physiol. 270 (Cell Physiol. 39): C86-C97, 1996[Abstract/Free Full Text].

18.   Kirk, K., and K. Stange. Functional properties and physiological roles of organic solute channels. Annu. Rev. Physiol. 60: 719-739, 1998[Medline].

19.   Komoszynski, M., and A. Wojtczak. Apyrases (ATP diphosphohydrolase, EC 3.6.1.5): function and relationship to ATPases. Biochim. Biophys. Acta 1310: 233-241, 1996[Medline].

20.   Leaney, J. L., S. J. Marsh, and D. A. Brown. A swelling-activated chloride current in rat sympathetic neurones. J. Physiol. (Lond.) 501: 555-564, 1997[Abstract].

21.   Lewis, S. A., and P. Donaldson. Ion channels and cell volume regulation: chaos in an organized system. News Physiol. Sci. 5: 112-119, 1990.[Abstract/Free Full Text]

22.   Light, D. B., T. M. Mertins, J. A. Belongia, and C. A. Witt. 5-Lipoxygenase metabolites of arachidonic acid regulate volume decrease by mudpuppy red blood cells. J. Membr. Biol. 158: 229-239, 1997[Medline].

23.   McCarty, N. A., and R. G. O'Neil. Calcium signaling in cell volume regulation. Physiol. Rev. 72: 1037-1061, 1992[Abstract/Free Full Text].

24.   Montrose-Rafizadeh, C., and W. B. Guggino. Role of intracellular calcium in volume regulation by rabbit medullary thick ascending limb cells. Am. J. Physiol. 260 (Renal Fluid Electrolyte Physiol. 29): F402-F409, 1991[Abstract/Free Full Text].

25.   Moran, A., and R. Turner. Secretagogue-induced RVD in HSY cells is due to K+ channels activated by Ca2+ and protein kinase C. Am. J. Physiol. 265 (Cell Physiol. 34): C1405-C1411, 1993[Abstract/Free Full Text].

26.   Rubera, I., M. Tauc, C. Poujeol, M. T. Bohn, M. Bidet, G. De Renzis, and P. Poujeol. Cl- and K+ conductances activated by cell swelling in primary cultures of rabbit distal bright convoluted tubules. Am. J. Physiol. 273 (Renal Physiol. 42): F680-F697, 1997[Abstract/Free Full Text].

27.   Schwiebert, E. M., M. E. Egan, T.-H. Hwang, S. B. Fulmer, S. S. Allen, G. R. Cutting, and W. B. Guggino. CFTR regulates outwardly rectifying chloride channels through an autocrine mechanism involving ATP. Cell 81: 1063-1073, 1995[Medline].

28.   Soltoff, S. P., M. K. McMillian, J. D. Lechleiter, L. C. Cantley, and B. R. Talamo. Elevation of [Ca2+]i and the activation of ion channels and fluxes by extracellular ATP and phospholipase C-linked agonists in rat parotid acinar cells. Ann. NY Acad. Sci. 603: 76-92, 1990[Abstract].

29.   Strange, K. (Editor). Cellular and Molecular Physiology of Cell Volume Regulation. Boca Raton, FL: CRC, 1994

30.   Strange, K., F. Emma, and P. S. Jackson. Cellular and molecular physiology of volume-sensitive anion channels. Am. J. Physiol. 270 (Cell Physiol. 39): C711-C730, 1996[Abstract/Free Full Text].

31.   Stutts, M. J., J. G. Fitz, A. M. Paradiso, and R. C. Boucher. Multiple modes of regulation of airway epithelial chloride secretion by extracellular ATP. Am. J. Physiol. 267 (Cell Physiol. 36): C1442-C1451, 1994[Abstract/Free Full Text].

32.   Sugasawa, M., C. Erostegui, C. Blanchet, and D. Dulon. ATP activates a cation conductance and Ca2+-dependent Cl- conductance in Hensen cells of guinea pig cochlea. Am. J. Physiol. 271 (Cell Physiol. 40): C1817-C1827, 1996[Abstract/Free Full Text].

33.   Taylor, A. L., B. A. Kudlow, K. L. Marrs, D. C. Gruenert, W. B. Guggino, and E. M. Schweibert. Bioluminescence detection of ATP release mechanisms in epithelia. Am. J. Physiol. 275 (Cell Physiol. 44): C1391-C1406, 1998[Abstract/Free Full Text].

34.   Turner, R. J. Amphibians. In: Vertebrate Blood Cells, edited by A. F. Rowley, and N. A. Ratcliff. New York: Cambridge University Press, 1988, p. 129-209.

35.   Ulb, J., H. Murer, and H. A. Kolb. Hypotonic shock evokes opening of Ca2+-activated K+ channels in opossum kidney cells. Pflügers Arch. 412: 551-553, 1988[Medline].

36.   Wang, Y., R. Roman, S. D. Lidofsky, and J. G. Fitz. Autocrine signaling through ATP release represents a novel mechanism for cell volume regulation. Proc. Natl. Acad. Sci. USA 93: 12020-12025, 1996[Abstract/Free Full Text].

37.   Windscheif, U. Purinoceptors: from history to recent progress. A review. J. Pharm. Pharmacol. 48: 993-1011, 1996[Medline].

38.   Wong, S. M. E., M. C. Debell, and H. S. Chase, Jr. Cell swelling increases intracellular free [Ca] in cultured toad bladder cells. Am. J. Physiol. 258 (Renal Fluid Electrolyte Physiol. 27): F292-F296, 1990[Abstract/Free Full Text].

39.   Yang, X. C., and F. Sachs. Block of stretch-activated ion channels in Xenopus oocytes by gadolinium and calcium ions. Science 243: 1068-1071, 1989[Medline].


Am J Physiol Cell Physiol 277(3):C480-C491
0002-9513/99 $5.00 Copyright © 1999 the American Physiological Society