Department of Biology, Ripon College, Ripon, Wisconsin 54971
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ABSTRACT |
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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
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INTRODUCTION |
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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.
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METHODS |
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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 compoundCoulter 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: [(VmaxPatch 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 M 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-
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.
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.
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RESULTS |
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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.
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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).
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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.
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DISCUSSION |
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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 ATPS (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 ATPS 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.
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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 ATPS-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.
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ACKNOWLEDGEMENTS |
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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.
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FOOTNOTES |
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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.
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