1 Laboratory of Physiology, Limburgs Universitair Centrum, Universitaire Campus Gebouw D, B-3590 Diepenbeek; and 2 Laboratory of Physiology, K. U. Leuven, Campus Gasthuisberg, B-3000 Leuven, Belgium
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ABSTRACT |
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In renal ischemia, tubular obstruction induced by swelling of epithelial cells might be an important mechanism for reduction of the glomerular filtration rate. We investigated ischemic cell swelling by examining volume regulation of A6 cells during metabolic inhibition (MI) induced by cyanide and 2-deoxyglucose. Changes in cell volume were monitored by recording cell thickness (Tc). Intracellular pH (pHc) measurements were performed with the pH-sensitive probe 5-chloromethyl-fluoresceine diacetate. Tc measurements showed that MI increases cell volume. Cell swelling during MI is proportional to the rate of Na+ transport and is not followed by a volume regulatory response. Furthermore, MI prevents the regulatory volume decrease (RVD) elicited by a hyposmotic shock. MI induces a pronounced intracellular acidification that is conserved during a subsequent hypotonic shock. A transient acidification induced by a NH4Cl prepulse causes a marked delay of the RVD in response to a hypotonic shock. On the other hand, acute lowering of external pH to 5, simultaneously with the hypotonic shock, allowed the onset of RVD. However, this RVD was completely arrested ~10 min after the initiation of the hyposmotic challenge. The inhibition of RVD appears to be related to the pronounced acidification that occurred within this time period. In contrast, when external pH was lowered 20 min before the hyposmotic shock, RVD was absent. These data suggest that internal acidification inhibits cellular volume regulation in A6 cells. Therefore, the intracellular acidification associated with MI might at least partly account for the failure of volume regulation in swollen epithelial cells.
distal tubular epithelial cell line; chemical ischemia; cyanide; 2-deoxyglucose; pH-sensitive fluorescent probe 5-chloromethylfluoresceine diacetate; hyposmotic shock; regulatory volume decrease
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INTRODUCTION |
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A FUNDAMENTAL PROPERTY of animal cells is the ability to maintain cell volume constant. Cellular volume is determined ultimately by two factors: the total cell content of osmotically active particles and the osmolality of the bathing medium. The principal threat to cell volume in vivo is a change in the amount of osmotically active molecules in cells bathing in isosmotic solutions. A change in total solute content of cells can occur if the balance is altered between solute entry and extrusion from the cells. A dramatic example of such isosmotic swelling in vivo occurs during ischemia. In ischemic renal injury, cellular ATP depletion inhibits the basolateral Na+ pump that normally maintains constant cell volume by offsetting the tendency of cells to swell because of impermeant cellular solutes. The resulting swelling of tubular cells is an early, important step in the development of tubular necrosis. It has also been proposed that cell swelling contributes to renal dysfunction by leading to obstruction of the tubular lumen and, thus, to a decrease of glomerular filtration rate (GFR) (2). Furthermore, cell swelling occurs at the expense of vascular space, and the compression of vessels may impede reflow through injured tissue (18). The understanding of epithelial cell volume regulatory mechanisms and their modulation by ischemia could be extremely valuable for designing novel therapeutic strategies to improve organ function, e.g., in transplants.
In this study, polarized A6 epithelia derived from the distal part of
the nephron of the kidney of Xenopus laevis were
used as model cells. The A6 cell line is a well-established cell line with transport properties specific for mammalian collecting duct cells
(30). The inhibition of cellular energy metabolism was used as an experimental model to simulate ischemic cell injury and was realized by inhibiting both cellular glycolysis (with 2-deoxyglucose) and oxidative phosphorylation (with cyanide). The aim
of the present study was to examine whether metabolic inhibition (MI)
causes swelling of these distal epithelial cells and, if so, whether
the cells can readjust their volume by a mechanism known as regulatory
volume decrease (RVD). Furthermore, we investigated whether and how MI
interferes with cell volume control mechanisms in anisotonic solutions.
In most cell types, including A6 cells (8), exposure to a
hypotonic solution elicits a RVD that is accomplished mainly by KCl
efflux through specific volume-activated K+ and
Cl channels, different from the native K+ and
Cl
channels. A fall in intracellular pH (pHc)
occurs in most cells exposed to ischemic conditions
(2). Recently, it was reported that internal protons are
able to inactivate apical epithelial Na+ channels (ENaCs)
(14, 41) and the native basolateral K+
channels (14) in A6 cells. Although activation and
regulation of volume-activated channels have been studied in detail,
data on proton modulation are rather scarce. In freshly isolated S2 segments of renal proximal tubules of rabbit kidney (34),
it was shown that a rise in PCO2 of the
perfusate reduces the rate of volume regulation in response to a
hypotonic shock without affecting the extent of the RVD response.
Moreover, it was shown that lowering of the external perfusate pH
exhibits an RVD-inhibitory effect in A6 cells (21).
Neither study distinguished between the effects of extracellular
acidification and the intracellular acidosis associated with the
acidifying maneuver on the observed RVD retardation. Some very recent
reports indicate pHc-sensitive volume-activated
K+ channels in primary cultures of seawater fish gill cells
(11), in villus epithelial cells (22), and in
Ehrlich ascites tumor cells (15). Therefore, we verified
whether intracellular protons might inhibit RVD, possibly via
inhibition of volume-activated ion channels.
This report shows that cell swelling during MI depends on the salt transport rate of the epithelial cell. We demonstrate that MI inhibits the RVD in response to isosmotic cell swelling (due to the MI itself) as well as to anisosmotic cell swelling (elicited via a reduction of the extracellular tonicity). Furthermore, our findings suggest that the intracellular acidification that accompanies MI might play a role in RVD inhibition.
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MATERIALS AND METHODS |
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Cell Culture
A6 cells (passages 105-111) obtained from Dr. J. Johnson (University of Pittsburgh, PA) were cultured at 28°C in a humidified incubator in the presence of 1% CO2 in the air (for details, see Ref. 36). A6 cells were seeded on a permeable tissue culture support (25 mm in diameter; Nunc Anopore; 1.5 million cells/support). For volume measurements, Anopore membranes were coated with fluorescent microspheres 1 µm in diameter (L-5081; Molecular Probes, Eugene, OR) embedded in a thin gelatin layer before cells were seeded. After 10-21 days of culture, confluent and polarized monolayers were used to perform volume experiments, while for fluorimetric pHc measurements, polarized monolayers 4-6 days old were used to obtain higher signal-to-noise ratios.Cell Volume Measurements
This method has been described previously in detail (37). Briefly, tissues were mounted in an Ussing-type chamber enabling solution exchange on both sides. Cell thickness (Tc) was used as an index for cell volume of confluent monolayers. The apical (upper) side of the monolayer was labeled with fluorescent biotin-coated microbeads. Focusing of the microbeads was automatically performed with a piezoelectric focusing device (PIFOC; Physik Instrumente, Waldbronn, Germany). Tc is defined as the vertical distance between the basolateral and apical beads. Measured Tc values were corrected for the diameter of the fluorescent microbeads by subtracting 1 µm. Changes in cell height are expressed as a percentage of the value recorded just before the hyposmotic challenge or metabolic inhibition was imposed. Average values of Tc were calculated from the recordings from a number of beads (nB) that remained attached to the monolayer during the entire experiment.The tissues were short-circuited during the entire course of the experiment by using Ag-AgCl voltage and current electrodes that were connected to the bath solutions with agar bridges containing 3% agar in 1 M KCl medium. Transepithelial conductance (Gt) and short-circuit current (Isc) were recorded. Isc mainly reflects transepithelial Na+ absorption in A6 cells because, in general, Isc disappears in the presence of 0.1 mM apical amiloride.
Fluorescence Imaging Microscopy to Measure pHc With CMFDA
Use of CMFDA as a pH-sensitive probe. CMFDA (5-chloromethylfluorescein diacetate; Molecular Probes, Eugene, OR) is an analog of 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF). Except for one abstract in which the use of CMFDA as a pH probe is mentioned without details (28), this is, to our knowledge, the first full report illustrating the suitability of CMFDA for pHc measurements. CMFDA shows pH-dependent spectral shifts in a somewhat more acidic cytosolic pH range than BCECF because the pKa is ~6.4 [determined for the unconjugated hydrolyzed product in buffer (after conjugation to an intracellular thiol or amine, the pKa value may be different)] compared with a pKa of ~7 for BCECF. The advantage of CMFDA is that it accumulates and retains far better into A6 cells than BCECF. Once the membrane-permeant form of the probe enters a cell, esterase-mediated hydrolysis converts nonfluorescent CMFDA to fluorescent 5-chloromethylfluoresceine, which can then react with intracellular thiols to yield well-retained products (16). Cytotoxic effects caused by this intracellular reaction are unlikely, because many cell types loaded with CMFDA remain viable for at least 24 h after loading and often through several cell divisions (1).
At first, we checked whether this fluorescein derivative was a suitable pH probe for A6 cells. CMFDA calibrations were performed as shown for one typical experiment in Fig. 1A. CMFDA was used in the dual-excitation (495 and 440 nm) mode (for details, see Use of fluorescence imaging microscopy to measure pHc). Intracellular pH was forced to equilibrate with the external pH according to the method of Thomas et al. (35). Cells were exposed to solutions containing 137 mM KCl, 1 mM CaCl2, 10 mM HEPES, and 13 µM nigericin. The K+ concentration used approximates the reported cytosolic K+ concentration in A6 cells (25). Different pH values of the solution (pHsol) were obtained with Tris. The CMFDA fluorescence (F) ratio R = F(495)/F(440) vs. the different imposed pHc values is depicted in Fig. 1B.
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Use of fluorescence imaging microscopy to measure
pHc.
A6 tissues were mounted in an Ussing-type chamber (chamber opening 0.7 cm2). After the background signal was measured, cells were
loaded from the apical side with the probe CMFDA. Cells were then
exposed to a final concentration of 10 µM of the acetoxymethyl ester
form of the dye (stock solution 5 mM in DMSO). Loading was performed for 60 min at room temperature in apical control Ringer solution (Table
2). After loading, excessive dye was
removed by replacing the apical bath solution several times. During the
experiment, monolayers were continuously superfused (0.75 ml/min) on
both sides. The fluorescence was measured with an inverted
epifluorescence microscope Zeiss Axiovert 100 (Jena, Germany).
Excitation light of a 75-W xenon lamp (Osram, Berlin-München,
Germany) was filtered at 440 and 495 nm with excitation filters
(bandwidth of 10 nm; Chroma Technology, Brattleboro, VT), which were
inserted in a computer-controlled filter wheel (Lambda 10-2;
Sutter Instruments, Novato, CA). The fluorescence collected by the
objective (Zeiss LD Achroplan ×20/0.4 corr.) was transmitted through a
>500-nm long-pass dichroic mirror and a 535/50-nm band-pass emission
filter (Chroma Technology) to a Quantix charge-coupled device (CCD)
camera (Photometrix, Tucson, AZ). The camera was equipped with a Kodak KAF 1400 CCD (grade 2, MPP) with 1317 × 1035 pixels and cooled to
25°C by a thermoelectric cooler. The acquisition of pairs of images
for this dual-excitation radiometric dye, CMFDA, was controlled by a
homemade program that uses V for Windows software (Digital Optics,
Auckland, New Zealand). Camera exposure time for 1 image was 1 s.
Signals were obtained by spatially integrating pixels over the field of
view. The background image, because of reading noise and the dark
current of the CCD camera and autofluorescence of tissue and tissue
support, was automatically subtracted pixel by pixel from the image of
the loaded cells. At the end of each experiment, a calibration was
performed by using the nigericin-high-K+ technique
(35) as described earlier. Heterogeneity in the cellular response of different regions in the monolayer was checked after each
experiment with a homemade program that uses V for Windows software.
This analysis revealed that no significant differences existed in the
cellular responses of different regions in one monolayer to
pHc-changing manipulations (Fig. 1A).
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Statistics
Values from n experiments (different monolayers) are given as means ± SE.Solutions and Chemicals
The compositions of the solutions used in this study are given in Table 2. MI was induced by bilateral sodium cyanide (CN, 2.5 mmol/l) and 2-deoxy-D-glucose (2-DG, 20 mmol/l). The mitochondrial substrate pyruvate was substituted for glucose in the recovery solution after MI because it has been reported that 2-DG has an irreversible action on cellular glycolysis (20).Isosmotic solutions had an osmolality of 260 ± 4 mosmol/kg H2O, which is the osmolality of the growth medium for the cells. Hyposmotic solutions had an osmolality of 140 ± 4 mosmol/kgH2O. All apical solutions in Figs. 3-8 were hyposmotic to avoid an osmotic gradient from apical to basolateral side during the basolateral hyposmotic challenge. Reduction of apical osmolality does not alter the volume of A6 cells (9). The osmolality of the solutions was verified with a cryoscopic osmometer (Osmomat 030; Gonotec, Berlin, Germany).
Ethylisopropylamiloride (EIPA) was purchased from Research Biochemicals (Natick, MA). CN was from UCB (Brussels, Belgium). 2-DG, nigericin, omeprazole, magnesium nitrate, and the sodium salt of pyruvic acid were from Sigma (St. Louis, MO). HOE-642 and S-3226 were from Hoechst Marion Roussel (Frankfurt am Main, Germany). All experiments were carried out at room temperature.
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RESULTS |
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Influence of Metabolic Inhibition on Cell Volume in High- and Low-Rate Salt-Transporting Epithelia
To investigate whether metabolic inhibition had an influence on cell volume, we monitored epithelial cell thickness, Tc, during a 45-min incubation period with CN (2.5 mM) and 2-DG (20 mM) and during a subsequent 45-min recovery period when CN and 2-DG were removed from the Ringer solution and pyruvate was provided as a substrate. We found a striking difference between high- and low-rate Na+-transporting A6 cells. As reported by Wills et al. (40), high rates of Na+ transport (Isc = 22.3 ± 0.8 µA/cm2, n = 6) in A6 cells are obtained by perfusion of the monolayers with a hypotonic (200 mosmol/kgH2O) Ringer solution. Metabolic inhibition in these epithelia resulted in a progressive increase of cell volume, reaching a plateau value of 120% within 25 min (Fig. 2A). Perfusion with recovery Ringer for 30 min allowed the cells to partially recover to 107%. Figure 2B illustrates the behavior of low-rate Na+-transporting epithelia (Isc = 2.9 ± 0.3 µA/cm2, n = 4) that were perfused with isotonic Ringer solution (260 mosmol/kgH2O). Interestingly, under these conditions, cells were able to maintain a cell volume constant during metabolic inhibition and the recovery phase.
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Because no volume regulation was observed after the pronounced isosmotic volume increase during MI in high-rate salt-transporting epithelia (200 mosmol/kgH2O Ringer), we checked whether MI had an influence on cellular mechanisms to regulate cell volume in anisosmotic conditions.
Tc Changes after a Basolateral Hypotonic Shock in Metabolically Inhibited Cells with Low-Rate Na+ Transport
Figure 3 illustrates the control behavior of Tc during a basolateral hyposmotic shock from 260 to 140 mosmol/kgH2O and subsequent perfusion with isotonic Ringer. When basolateral osmolality was lowered, A6 cells swelled quickly due to basolateral water influx. Subsequently, this volume increase was rapidly counteracted by a RVD, accomplished by the loss of K+ and Cl
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Effect of Metabolic Inhibition With Minimized Na+ Influx on RVD
One possible explanation for the absence of RVD seen in ischemic cells might be that the inhibition of active Na+ extrusion from the cell leads to an accumulation of Na+ in exchange for K+. Because the swelling-activated cation channels in A6 cells are impermeable to Na+ (21), cellular K+ depletion will impede cell volume regulation. To test the role of K+ depletion in the inhibition of RVD after MI, we designed a protocol to minimize Na+ influx into the cells. First, apical Na+ was replaced with N-methyl-D-glucamine (NMDG+). This apical Na+ uptake-abolishing manipulation did not affect cell volume in isotonic conditions (9). Second, the basolateral Na+ concentration was lowered to 10 mM to reduce the chemical driving force for Na+ influx. In this protocol, metabolic inhibition was realized with 2-DG (20 mM) at both sides of the monolayer and CN (2.5 mM) exclusively at the basolateral side to maintain Na+-free conditions at the apical side. The solid line in Fig. 4 illustrates the behavior of cells that were exposed to CN and 2-DG 45 min before and during the hyposmotic challenge. Although Na+ accumulation and K+ depletion were prevented, these ischemic cells still failed to downregulate their volume after a hypotonic shock. Because the reduction of the basolateral Na+ concentration had a pronounced volume decreasing effect in isotonic conditions, we checked to see whether this manipulation as such had an influence on RVD evoked by a hypotonic shock. However, RVD was still present in cells that were superfused for 75 min with a basolateral Ringer that contained only 10 mM Na+. These results suggest that another MI-associated phenomenon must play a role in RVD inhibition.
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Intracellular Acidification During Metabolic Inhibition in A6 Cells
Recently, we showed that intracellular protons are able to close Na+ channels in A6 cells (41). As a consequence, the hypothesis arose that intracellular protons might also inhibit volume-activated K+ or Cl
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Influence of Intracellular Acidification on Volume Recovery After a Hyposmotic Challenge
Because pHc changes affect transepithelial Na+ transport (INa) changes in A6 cells (41) and even a small INa has an inhibiting effect on RVD (9), the effects of pHc on RVD were examined in nontransporting conditions in which NaCl was replaced with NMDGCl in the apical saline. As mentioned earlier, this apical Na+ uptake-abolishing manipulation does not affect cell volume in isotonic conditions (9).Intracellular acidification via the NH4Cl prepulse
method in combination with EIPA.
First, the NH4Cl prepulse technique was used to acidify the
cells without changing external pH. The basolateral NH4Cl
pulse was performed under strict and volume-controlled experimental conditions (41): 40 mM sucrose in the basolateral control
saline was isotonically replaced with 20 mM NH4Cl for 25 min. Subsequently, cells were bilaterally superfused for 15 min with
control saline containing 50 µM EIPA that arrests the basolateral
Na+/H+ exchanger (3). As shown in
Fig. 6A, this method evoked an intracellular acidification from pHc 7.00 ± 0.08 to
6.4 ± 0.1 (n = 4). Although EIPA was used, the
cells gradually recovered from this pronounced acidification during the
subsequent basolateral hypotonic shock (pHc = 6.8 ± 0.1 after 20 min). Further attempts, including the use of sodium
propionate (30 mM), the Na+/H+ exchanger
blockers HOE-642 (10 µM) and S-3226 (10 µM), and the combination of
the inhibitors EIPA (50 µM), omeprazole (0.1 mM), and magnesium
nitrate (10 mM) to block, respectively, the
Na+/H+ exchanger and the putative
K+/H+-ATPase and H+-ATPase, to keep
the cells acidic for a longer period failed (results not shown). In
general, cell swelling leads to cytosolic acidification (19). Nevertheless, we investigated in control experiments
whether the basolateral hyposmotic treatment as such was not a
pHc-increasing manipulation in A6 cells. If this were the
case, it would neutralize previous acid-inducing maneuvers. Replacement
of basolateral isotonic Ringer with a hypotonic Ringer solution for
1 h induced no significant pHc change in 6 different
monolayers (Fig. 6A). Tc experiments (Fig. 6B) reveal that a hyposmotic challenge still elicits a
complete RVD in cells that were previously acidified to pHc
6.4 with the NH4Cl prepulse technique. However, RVD was
profoundly slowed down during the first 20 min compared with control
experiments. This delay in RVD might be attributed to the presence of
intracellular protons that gradually leave the cells during the
hypotonic shock.
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Intracellular acidification via lowering of pHsol.
Second, pHc was modified by lowering pHsol at
both sides of the epithelium. As indicated in Fig.
7A, a 30-min exposure period to pHsol 6 reduced the intracellular pH from 7.06 ± 0.02 to 6.59 ± 0.03 (n = 6). A subsequent
basolateral hypotonic shock of 60 min acidified the cells further to
pHc = 6.3 ± 0.2 (n = 3). These acidified cells still elicited an RVD that was, however, partially inhibited as shown in the Tc measurements of
Fig. 7B. To obtain a more pronounced pHc drop
(from 6.99 ± 0.06 to 6.15 ± 0.07, n = 6),
other monolayers were superfused for 20 min with a Ringer of
pHsol 5 (Fig. 7A). The acidifying process continued during the hypotonic shock. Figure 7B shows that, under these
conditions, RVD was completely inhibited. The fact that the shrinking
of cells still occurred when isotonic Ringer was applied demonstrates
that this very low pHsol did not damage cell membranes.
Moreover, the mean transepithelial conductance for these acidic
monolayers did not exceed a value of 0.85 mS/cm2 during
this experimental protocol.
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DISCUSSION |
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In the present study, MI with CN and 2-DG was used as an experimental model to simulate ischemic cell injury. We investigated the effects of MI on cell volume and cell volume regulation of a tight epithelium, A6.
Metabolic Inhibition Induces Cell Swelling in High-Rate Na+-Transporting A6 Epithelia
Under steady-state conditions, epithelial cell volume is maintained by balancing the rates of apical and basolateral ion transport in such a way that intracellular solute content remains constant. When MI is applied, cellular ATP depletion will hinder the active basolateral Na+ extrusion from the cells. In A6 epithelia with high rates of Na+ transport (Fig. 2A), the reduced pump rates during MI were insufficient to match massive apical Na+ influx. During the first 25 min of ischemia, cell volume gradually increased up to a plateau value of 120%. A further rise of cell volume was probably prevented by a reduction of apical Na+ entry by the action of intracellular protons (41) elicited during MI (Fig. 5). Hence, it seems that the plateau phase in Fig. 2A is comparable with the situation in low-rate salt-transporting epithelia (Fig. 2B). There, the hindered Na+ pump was still able to offset apical Na+ influx, and, therefore, accumulation of intracellular osmolytes and cell swelling could be avoided during MI.The cell swelling during MI in high-rate salt-transporting epithelia could also be a consequence of basolateral Na+ influx rather than apical Na+ influx as described above. A candidate for basolateral Na+ influx is the basolateral Na+/H+ exchanger (3) that is probably activated in these acidic cells (Table 1). However, this explanation is less plausible because low-rate salt-transporting epithelia (Fig. 2B) displayed no change in cell volume during MI, although these epithelia acidified to the same extent (Fig. 5), and, therefore, at least a similar activation of the basolateral Na+/H+ exchanger could be expected.
Absence of Isosmotic Volume Regulation in Metabolically Inhibited A6 Cells
High-rate salt-transporting A6 cells displayed no volume regulatory event after cell swelling due to MI. The similar behavior of Madin-Darby canine kidney cells during MI (preliminary experiments) indicates that the absence of cell volume regulation in metabolically inhibited cells, as described in this study, is not limited to amphibian epithelial cells.Absence of Anisosmotic Volume Regulation in Metabolically Inhibited Cells
This study further explored the capacity of distal epithelial cells (A6) to regulate cellular volume during MI by challenging them with a basolateral hyposmotic shock. After swelling, nonischemic A6 cells exhibit a RVD by K+ and ClIn addition, cellular ATP depletion during MI leads to reduced activity of the Na+ pump. Experiments that were designed to minimize Na+ influx and thus to avoid K+ depletion during MI revealed that RVD was still impaired (Fig. 4). Hence, RVD inhibition in metabolically inhibited A6 cells cannot be ascribed solely to intracellular K+ depletion.
Role of Intracellular pH on RVD Inhibition
This study indicates that MI is associated with a pronounced intracellular acidification in A6 cells (Fig. 5). A fall in intracellular pH occurs in most cells exposed to ischemic conditions (for review, see Ref. 2). Because we used 2-DG to block cellular glycolysis, the fall in pHc was not due to increased lactate production. Acidosis resulted probably from the imbalance between proton accumulation due to ATP hydrolysis (13, 32, 38, 39) and the constant passive influx of protons on one hand, and the suppression of H+ extruding transport processes in ATP depleted cells on the other hand (10).Li et al. (21) have shown that lowering the external pHsol exhibits an RVD-inhibitory effect in A6 cells. Our data suggest that this inhibitory action is exerted by internal rather than external protons. As shown in Fig. 6B, RVD was clearly delayed in A6 cells, acidified by means of the NH4Cl prepulse technique, with normal external pHsol. Similarly, Sullivan et al. (34) found that the rate of volume regulation in response to hypotonic media was reduced in rabbit proximal tubules that were acidified by increasing the PCO2 in the perfusate from 5 to 15%. Because the high PCO2 treatment in those experiments lowered both the external pHsol (from 7.44 to 6.97) as well as pHc (from 7.39 to 7.08 in isotonic conditions), no distinction was possible between modulation of RVD by external and internal protons. Moreover, pHc was not measured during hypotonic conditions. Thus it is unknown whether pHc remained acidic in hypotonic media with high PCO2 pressure. In our study, the extra internal protons resulting from the NH4Cl prepulse were extruded out of the cells in the first 20 min of the hypotonic shock, so their inhibitory action on RVD was limited in time. Further experiments are needed to elucidate the identity of the H+-extruding mechanism(s). In the work of Sullivan et al. (34), RVD was not complete except to the same extent as in control experiments. In contrast, we found in the present study that the RVD in acidified A6 cells was complete in both control and experimental conditions. It has been shown in A6 cells that the extent of RVD is dependent on the Na+ transport rate (9). Hence, the apparent discrepancy concerning the completeness of RVD in A6 cells and proximal tubular cells might be explained by the difference in their transport capacity.
Our results indicate that a complete and immediate inhibition of RVD after a hyposmotic challenge is achieved only when the following conditions are fulfilled. First, the extra amount of internal protons needs to be trapped inside the cell. This was the case in ATP-depleted cells (Fig. 5), and in cells not ATP depleted, protons remained inside by lowering the external pHsol. Second, a profound acidification is necessary. This was evidenced by the fact that RVD was only retarded (Fig. 7B) in cells that were acidified to some extent (pHc 6.59). In contrast, an immediate and complete blockage of RVD was seen in cells that were profoundly acidified to pHc values as low as 6.15. Acute addition of external protons (pHsol 5), simultaneously with the hypotonic shock, could not inhibit RVD immediately (Fig. 8A). The clear delay in RVD blockage seen in acutely treated cells is accompanied with a rapid intracellular acidification (Fig. 8B). Although it is conceivable that external protons could also interact with the volume-regulated ion channels either directly or indirectly at a site unrelated to the pore, the slow time course of the effect of lowering external pHsol acutely on RVD gives evidence against this explanation. Recently, it has been shown that inactivation of cloned amphibian (derived from A6 cells) and mammalian (human) renal chloride channels (ClC-5) after exposure to external acidic solutions occurred very rapidly (within 1 min) (24). Therefore, the involvement of external protons in RVD inhibition is less likely.
In the present study, the mechanism of RVD inhibition by internal
protons in A6 cells was not identified. Further experiments are needed
to resolve this issue. Native basolateral K+ channels are
pHc sensitive in A6 cells (14). The closure of these channels in acidified cells does not imply RVD inhibition because, in many cell types including A6 cells, Cl and
K+ efflux occurs through a separate set of volume-activated
ion channels (6, 8). Although the underlying molecular
mechanism of the inhibitory effect of protons on some types of
K+ channels is well described, e.g., on ROMK1 (4, 12,
33), the inhibition of volume-activated K+ channels
by protons is still unclear. However, some recent studies reported that
volume-activated channels might indeed be pHc sensitive. In
primary cultures of seawater fish gill cells, the swelling-activated K+ channel is impaired in acidic cells (11).
In addition, a role for intracellular acidification in the inactivation
of volume-regulated K+ channels is described recently for
Ehrlich ascites tumor cells (15).
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ACKNOWLEDGEMENTS |
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We acknowledge E. Lariviere for excellent technical assistance with volume measurements. We also thank J. Simaels, A. Roosen, J. Janssen, R. Van Werde, G. Raskin, P. Pirotte, and W. Leyssens for technical help. We are grateful to Dr. W. Zeiske for critical comments and helpful suggestions.
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FOOTNOTES |
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Address for correspondence: P. Steels, MBW-Laboratory of Physiology, Biomedisch Onderzoeksinstituut, Limburgs Universitair Centrum, Universitaire Campus Gebouw D, B-3590 Diepenbeek, Belgium (E-mail: paul.steels{at}luc.ac.be).
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. Section 1734 solely to indicate this fact.
April 3, 2002;10.1152/ajpcell.00371.2001
Received 3 August 2001; accepted in final form 26 March 2002.
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