Basolateral Na⁺-K⁺-2Cl⁻ Cotransport in Cultured and Fresh Bovine Corneal Endothelium

Abstract

purpose. To examine whether Na⁺-K⁺-2Cl⁻ cotransport has the potential to contribute to corneal endothelial ion and fluid transport in cultured and fresh bovine corneal endothelial cells.

methods. Cl⁻ and Na⁺ sensitive fluorescent dyes were used to measure furosemide-dependent ion fluxes in cultured and fresh endothelial cells. Immunoblot analysis and immunofluorescence were used to determine expression and location of the Na⁺-K⁺-2Cl⁻ cotransporter (NKCC1).

results. Application of furosemide (50–100 μM) reduced Cl⁻ and Na⁺ influx in approximately 50% of trials using cultured cells and only 10% of trials with fresh cells; however, in all cases pretreatment with furosemide slowed Cl⁻ efflux when cells were bathed in Cl⁻-free Ringer’s. Double-sided perfusion of cultured cells indicated that furosemide-sensitive Cl⁻ fluxes were located on the basolateral side. Immunoblot analysis revealed 174-kDa bands in both fresh and cultured cells, but the bands were denser in fresh endothelial cells. Immunofluorescence showed distinct lateral membrane staining in addition to significant amounts of perinuclear staining.

conclusions. The Na⁺-K⁺-2Cl⁻ cotransporter is present in both fresh and cultured bovine corneal endothelium, and the expression is apparently higher in the fresh cells. The cotransporter is present on the lateral membrane consistent with a role in loading endothelial cells with Cl⁻, thereby possibly contributing to a transendothelial Cl⁻ flux. However, in the resting cell, net flux through the transporter is often not apparent.

Maintenance of corneal hydration is provided by the ion and fluid secretion properties of the corneal endothelium. The endothelium is a thin monolayer of cells that are well coupled to each other through gap junctions but show discontinuous bands of tight junctions leading to very low transendothelial electrical resistance (20–40Ω /cm²). Being so leaky, the endothelium is not readily amenable to the study of transport by the usual measurements of short circuit current or net fluxes of radioactive tracers in the Ussing-type chamber. Nevertheless, early attempts have shown that short circuit current is dependent on the presence of bicarbonate and reduced by carbonic anhydrase inhibitors, ^{1 2} which is consistent with the reduction in fluid transport by these same manipulations. ^{3 4} More recently, it has been shown that uptake of HCO₃ ⁻ by the endothelium is provided by an electrogenic Na⁺-nHCO₃ ⁻ cotransporter (n ≤ 2), which is most likely positioned at the basolateral membrane. ⁵ Furthermore, it has been shown that Cl⁻ is necessary for fluid transport. ⁶ Intracellular [Cl⁻] is above electrochemical equilibrium in cultured endothelial cells, ⁷ indicating that a transporter uptake mechanism is likely to be present. One possibility for Cl⁻ uptake is coupling of Na⁺/H⁺ exchange with Cl⁻/HCO₃ ⁻ exchange. However, recently we have shown that the anion exchanger is not expressed in cultured endothelium, and expression in freshly isolated tissue is very low. ⁵ Another possible uptake mechanism, used by many secretory epithelia, is through the loop diuretic-sensitive Na⁺-K⁺-2Cl⁻ cotransporter. In fact, bumetanide-sensitive Rb⁺ uptake, further activated by cell shrinkage, has been demonstrated in cultured bovine corneal endothelial cells (BCECs), ⁸ indicating the presence of Na⁺-K⁺-2Cl⁻ cotransport, a role in regulatory volume increase and possible role in fluid transport. On the other hand, Rb⁺ uptake studies using fresh rabbit corneas have failed to show bumetanide-sensitive uptake in endothelium bathed in isotonic or hypertonic media. ⁹ Further, fluid transport studies using freshly isolated rabbit corneas have so far failed to show bumetanide sensitivity of fluid transport. ^{4 9} Taken together, these studies indicate that the Na⁺-K⁺-2Cl⁻ cotransporter may not be present in fresh endothelium, and its expression could be upregulated by culturing. 

Given this uncertainty about the role of Na⁺-K⁺-2Cl⁻ cotransport in endothelial fluid transport, we used immunoblot analysis and immunofluorescence to determine whether the Na⁺-K⁺-2Cl⁻ cotransporter is present in cultured and fresh bovine corneal endothelium. Further, fluorescent probes for intracellular [Cl⁻ _i] and [Na⁺ _i] were used to determine whether the cotransporter provides net influx under isosmotic conditions in cultured BCECs. We also tested whether the cotransporter is segregated to the basolateral membrane. Together with an apical efflux mechanism (e.g., anion channels) a basolateral location for the cotransporter would be consistent with a role in transendothelial Cl⁻ and fluid transport. Lastly, we attempted to extend the measurements for intracellular [Cl⁻] to fresh endothelial cells to determine whether Na⁺-K⁺-2Cl⁻ cotransport-dependent Cl⁻ fluxes could be demonstrated. 

Materials and Methods

Cell Culture

BCECs were cultured to confluence on 25-mm round coverslips or 13-mm filters (AnoDisc; Whatman, Clifton, NJ), as previously described. ⁵ Briefly, primary cultures from fresh cows’ eyes were established in T-25 flasks with 3 ml Dulbecco’s modified Eagle’s medium (DMEM), 10% bovine calf serum, and antibiotic-antimycotic (100 U/ml penicillin, 100 μg/ml streptomycin, and 0.25 μg/ml Fungizone [Bristol–Myers Squibb, Princeton, NJ]), gassed with 5% CO₂-95% air at 37°C and fed every 2 to 3 days. These were subcultured to three T-25 flasks and grown to confluence in 5 to 7 days. The resultant second-passage cultures were then further subcultured onto filters or coverslips and allowed to reach confluence within 5 to 7 days. Cells were transferred to 1% serum-DMEM for at least 24 hours before the experiments. 

Solutions and Chemicals

The composition of the HCO₃ ⁻-free Ringer’s solution used throughout this study was (in mM) 150 Na⁺, 4 K⁺, 0.6 Mg²⁺, 1.4 Ca²⁺, 118 Cl⁻, 1 HPO₄ ⁻, 10 HEPES⁻, 30 gluconate⁻, and 5 glucose. Ringer’s was equilibrated with air and pH adjusted to 7.5 at 37°C. Cl⁻-free Ringer’s was prepared by equimolar substitution of NaCl with sodium nitrate. Osmolarity was adjusted to 300 ± 5 mOsM with sucrose. The chloride-sensitive fluorescent dyes SPQ and MEQ were obtained from Molecular Probes (Eugene, OR). All other chemicals were obtained from Sigma (St. Louis, MO). Cell culture supplies were obtained from Gibco (Grand Island, NY). 

Perfusion

A coverslip perfusion system for fluorescence measurements with an inverted microscope was used as described previously. ¹⁰ Briefly, the coverslip with the monolayer of cells formed the bottom of a perfusion channel (volume, 80 μl), the top of which was formed by another permanently sealed coverslip. Each end of the channel was fitted with 23-gauge stainless steel tubing and connected to perfusion syringes by gas-impermeable tubing (Phar-Med, Fisher Scientific, Fairlawn, NJ). The perfusion chamber was seated on a water-jacketed (37°C) brass collar, held on the stage of an inverted microscope (Diaphot; Nikon, Melville, NY). The cells were viewed with a ×40 oil-immersion objective (Fluor, 1.3 numerical aperature; Nikon). Ringer’s solutions were placed in syringes held in a Plexiglas warming box maintained at 37°C. The flow of the perfusate (∼0.5 ml/min) was achieved by gravity. The desired solution was selected by means of an eight-way valve. 

For independent perfusion of the apical and basolateral sides, a double-sided perfusion chamber was used (see Reference 5 for details). Cell-coated filters (AnoDisc) were sandwiched between two thin (1-mm) plastic (Kel-F) plates, both of which had a perfusion slot cut out at the center. Each perfusion slot (7 mm long × 3.1 mm wide) was connected to 23-gauge stainless steel tubing. The filter was placed in a 40-μm recess in the bottom plate with the cells facing downward. Thus, only the cells facing the open slot were perfused. Round glass coverslips were seated on the outer surface of each plate with a thin layer of vacuum grease to form compartments (∼22 μl) for independent apical and basolateral perfusion. Stainless steel clamps on the outer surface of the plastic plates were screwed together sandwiching the filter firmly. The assembled chamber was placed on a water-jacketed (37°C) brass collar held on the microscope stage. The apical compartment faced the microscope objective. The apical and basolateral compartments were connected to separate tubes (Phar-Med; Fisher), which in turn were connected to syringes in the Plexiglas warming box, as described. Two independent eight-way valves were used to select the desired perfusate for the apical and basolateral chambers. A long-working-distance, water-immersion ×40 objective (1.2-mm working distance, 0.75 numerical aperture; Carl Zeiss, Thornwood, NY) was used for fluorescence measurements. 

Measurement of Cellular Fluorescence

Cellular fluorescence was measured with a microscope spot fluorometer (Photon Technology, Monmouth Junction, NJ). Fluorescence excitation was provided by a 75-W xenon arc. Excitation wavelengths were obtained by passing the light through a DeltaRam monochromator (Photon Technology). The excitation light was directed to the objective by a dichroic mirror. The fluorescence emission collected by the objective passed through the dichroic mirror and a barrier filter and was led to a photomultiplier for detection. Neutral density filters (1–2 optical density) were included in the excitation path to minimize photobleaching. Synchronization of excitation with emission measurement and data collection was controlled by software (Felix; Photon Technology). Fluorescence ratios were obtained at one per second. 

Measurement of SPQ Fluorescence and Intracellular[ Cl⁻]

Cultured endothelial cells were loaded with the halide-sensitive dye SPQ at room temperature by a 6-minute exposure to hyposmotic Ringer’s (180 mOsM NO₃ ⁻ Ringer’s) containing 20 mM SPQ. ^{7 11} After loading, the cells were allowed to recover from the hyposmotic shock for 30 to 40 minutes in isosmotic HCO₃ ⁻-free Ringer’s also containing 20 mM SPQ. Subsequently, the adhering dye was washed with the same solution, and the coverslip was mounted into the perfusion chamber. The excitation for SPQ fluorescence was 365 ± 10 nm, the dichroic mirror was centered at 400 nm, and the barrier filter was a 420- to 450-nm band-pass. Relative differences in Cl⁻ flux between control and experimental conditions were determined by comparing the percentage change in SPQ fluorescence (ΔF/F) after removal or addition of Cl⁻. In some experiments, absolute intracellular[ Cl⁻] was calculated by the Stern–Volmer equation after determination of fluorescence at 0[ Cl⁻ _i] and halide-insensitive fluorescence (i.e., background fluorescence by perfusion with 150 mM SCN⁻). ^{7 11} Because these dyes are quenched by chloride, fluorescence increases as cytoplasmic[ Cl⁻] decreases. 

Fresh corneal endothelial cells were prepared by dissecting small strips (3 × 7 mm) of Descemet’s membrane-endothelium. Relative[ Cl⁻ _i] changes in fresh cells were assessed with the halide-sensitive fluorescent dye MEQ, which was also excited at 365 nm. Fresh endothelial cells were exposed to the nonfluorescent reduced cell-permeant quinoline derivative of MEQ (diH-MEQ), ^{12 13} which is oxidized to MEQ within the cytoplasm. diH-MEQ was usually synthesized on the day of the experiment and saved for no more than 4 days in chloroform at −20°C. Fresh cells were exposed to 10 μM diH-MEQ for 15 minutes at 37°C, washed twice with Ringer’s solution and kept at 37°C for another 10 minutes. Endothelial strips were placed in the open coverslip perfusion chamber, and glass fibers were laid on top of the strips to restrict movement during perfusion. The chamber was then sealed with a blank coverslip. Strips were viewed with the ×40 water-immersion long-working-distance objective. 

Measurement of Intracellular [Na⁺]

Cultured cells were loaded with the Na⁺-sensitive fluorescent dye SBFI. SBFI-AM stock was first mixed 1:1 with 25% weight–volume solution of Pluronic F-127 and then added to Ringer’s at a final concentration of 10 μM. Cells were incubated for 30 to 60 minutes at room temperature, washed, and placed in the coverslip perfusion chamber. The dye was alternately excited at 340 and 380 nm. The dichroic mirror was centered at 400 nm, and the barrier filter was a 420- to 500-nm band-pass. Calibration of the fluorescence ratio against intracellular[ Na⁺] was performed as previously described. ¹⁴  

Immunoblot Analysis

Fresh BCECs were scraped from dissected corneas, placed into ice-cold phosphate-buffered saline (PBS) containing a protease inhibitor cocktail (Complete; Boehringer–Mannheim, Indianapolis, IN) and centrifuged at low speed for approximately 5 minutes. Cell pellets was resuspended in 2% sodium dodecyl sulfate (SDS) sample buffer containing protease inhibitors. Cultured cells were dissolved directly in sample buffer. Both preparations were sonicated (model 250; Branson, Danbury, CT) briefly on ice and then centrifuged at 6000g for 5 to 10 minutes. An aliquot of the supernatant was taken for protein assay using the Bradford method (Bio-Rad, Hercules, CA).β -Mercaptoethanol (5%) and bromphenol blue were added to the remainder of the supernatant and heated at 80°C for 4 minutes. The samples were applied to a 7.5% polyacrylamide gel with 4.5% stacking gel (60 μg/lane). After electrophoresis at 20 mA, proteins were transferred to a polyvinylidene difluoride membrane overnight at 4°C. Membranes were incubated in PBS containing 5% nonfat dry milk for 1 hour at room temperature and washed in PBS containing 0.05% Tween two to three times for 5 minutes. The blots were then incubated with antibodies against the Na⁺-K⁺-2Cl⁻ cotransporter. T4 anti-NKCC1 mouse monoclonal antibody, (Developmental Studies Hybridoma Bank, Iowa University, Iowa City) or N1 anti-NKCC1 rabbit polyclonal antibodies (1:100; a generous gift from C. Lytle, University of California, Riverside) were used. Next, the blots were washed four times with PBS-Tween, incubated with secondary antibody coupled to horseradish peroxidase (Sigma), and finally developed by enhanced chemiluminescence (Dupont, Wilmington, DE). Films were scanned to produce digital images that were then assembled and labeled (PowerPoint; Microsoft, Redmond, WA). 

Immunofluorescence

Coverslips with cultured cells in 6-well plates were washed three to four times with warmed (37°C) PBS, and fixed for 30 minutes in fixation solution (PLP; containing 2% paraformaldehyde, 75 mM lysine, 10 mM sodium periodate, 45 mM sodium phosphate, [pH 7.4]) at 37°C on a rocker. After fixation, the cells were washed three to four times with PBS. Coverslips were then kept in PBS for 5 minutes containing 1% SDS to unmask epitopes ¹⁵ (and Christian Lytle, personal communication, March 1998) and washed three times in PBS. Cells were then washed with 0.01% saponin in PBS for 15 minutes and blocked for 1 hour in PBS containing 0.2% bovine serum albumin and 5% goat serum, 0.01% saponin, and 50 mM NH₄Cl. T4 or N1 antibodies (1:100 in PBS), were applied at room temperature for 1 hour. Coverslips were washed three times for 15 minutes in PBS containing 0.01% saponin. Texas red or fluorescein-conjugated secondary antibody (1:500 dilution) was applied for 1 hour at room temperature. Coverslips were then washed and mounted with medium (Prolong Antifade; Molecular Probes), according to the manufacturer’s instructions. 

To prepare fresh endothelial cells for immunofluorescence staining, corneas were dissected within 15 minutes of death, washed with warmed PBS, and immediately fixed with warmed PLP fixative buffer at 37°C for 10 minutes. Corneas were rinsed with PBS and endothelium-Descemet’s strips were peeled off the corneas and flattened onto microscope slides (Superfrost; Fisher). Strips were fixed again at room temperature for 20 minutes and washed with PBS. The remainder of the procedure was the same as for staining cultured cells. 

Results

If Na⁺-K⁺-2Cl⁻ plays a role in fluid transport, it should be a contributor to the salt uptake in the resting unstimulated endothelial cell. In all the experiments that are to be described, the loop diuretic furosemide was used as the Na⁺-K⁺-2Cl⁻ cotransport inhibitor, because the more potent inhibitor bumetanide is highly fluorescent at the excitation wavelengths used for SPQ and SBFI. Figure 1 shows the effect of 50 μM furosemide on steady state[ Cl⁻ _i] in cultured cells, as measured by SPQ. Furosemide caused a small increase in fluorescence indicating a decrease in[ Cl⁻ _i]. To determine the actual change in concentration of intracellular Cl⁻, the steady state fluorescence at 0[ Cl⁻] and the halide-insensitive fluorescence were determined. The lower portion of Figure 1 shows that the baseline[ Cl⁻ _i] was approximately 37 mM. Furosemide caused a small decrease to approximately 26 mM or a decrease of 11 mM. In 6 of 11 experiments furosemide caused a decrease in [Cl⁻ _i], and the average decrease was 7 ± 4 mM. In the other five experiments, no change in SPQ fluorescence was elicited. If these five experiments are included in the calculation of[ Cl⁻ _i] change, the average decrease was only 3.8 mM. Therefore, in approximately 50% of coverslips tested, furosemide inhibited chloride influx consistent with its blocking net Na⁺-K⁺-2Cl⁻ uptake; however, in some instances net uptake by the cotransporter was not apparent. 

Figure 2A shows the effect of furosemide on baseline[ Na⁺ _i], as measured by SBFI. Steady state [Na⁺ _i] was approximately 16 mM, consistent with previous estimates in the absence of HCO₃ ⁻. ¹⁴ In Figure 2 , Furosemide caused a slow decrease in[ Na⁺ _i] to approximately 11 mM, a decrease of 5 mM. In six of nine experiments, furosemide decreased[ Na⁺ _i] (average decrease, 4 ± 3 mM). In three of the nine trials, furosemide had no effect on steady state [Na⁺ _i], and if these experiments are included, the average decrease in[ Na⁺ _i] was only 2.7 mM. Figure 2B shows the effect of furosemide on the rate of Na⁺ leakage into endothelial cells after the application of 100 μM ouabain. Furosemide slowed the rate of Na⁺ entry by 19 ± 3% (n = 4). These results are consistent with modest net uptake by the Na⁺-K⁺-2Cl⁻ cotransporter in cultured endothelial cells. 

If Na⁺-K⁺-2Cl⁻ cotransport is to contribute to transendothelial ion and fluid transport, it should be located on the basolateral side of the endothelium to load cells with chloride and a Cl⁻ efflux pathway (e.g., anion channels) would be located apically. Figure 3 shows the effect of chloride removal from apical or basolateral sides, separately or both sides simultaneously, on[ Cl⁻ _i] of endothelial cells grown on a permeable substrate. When chloride was removed from the basolateral side, SPQ ΔF/F was approximately twice the ΔF/F when chloride was removed from the apical side but only 25% of the ΔF/F when chloride was removed from both sides together. Figure 3B shows that in the presence of 100 μM furosemide the relative changes in fluorescence when chloride was removed from both sides was reduced. Further, in the presence of furosemide, ΔF/F was almost eliminated when chloride was removed from the basolateral side; however, ΔF/F was unaffected when chloride was removed from the apical side. Table 1 summarizes these results and suggests that furosemide slowed Cl⁻ efflux from both sides by approximately 32% and from the basolateral side by 75%, and that it had no effect when Cl⁻ was removed from the apical side. These results indicate that furosemide-sensitive Cl⁻ flux was located basolaterally in cultured cells. 

To test whether furosemide-sensitive fluxes were present in fresh tissue, endothelium-Descemet’s membrane explants were dissected and loaded with the chloride-sensitive fluorescent dye MEQ. Figure 4 shows changes in MEQ fluorescence on chloride removal in the absence and presence of 100 μM furosemide. In this experiment when Cl⁻ free Ringer’s was removed, there was a small, quick decrease in fluorescence followed by the expected increase. The cause of this initial sharp transition is not known and did not always occur. Addition of furosemide caused a small increase in fluorescence, suggesting inhibition of net Cl⁻ uptake. However, in 10 trials with fresh tissue this was the only experiment that showed an increase in MEQ fluorescence caused by furosemide. The other 9 experiments showed no change in fluorescence. When Cl⁻ was removed in the presence of furosemide, the initial slope of ΔF/F was reduced by 30% (mean reduction, 50%; n = 10; P < 0.05). After 5 minutes, Cl⁻ was again added and in the presence of furosemide, the rate of decrease was reduced by 63% (mean reduction, 67%; n = 10; P < 0.05). 

To confirm the presence of the Na⁺-K⁺-2Cl⁻ cotransporter in corneal endothelium, we used immunoblot analysis with two antibodies to NKCC1. Figure 5 shows that the T4 and N1 antibodies produced positive bands at approximately 174 kDa, which is in the expected range for mammalian Na⁺-K⁺-2Cl⁻ cotransporters. ¹⁶ Fresh corneal epithelium was included as a positive control. ¹⁷ Figure 5 also indicates that the apparent density of T4 and N1 bands were considerably higher with the fresh cell preparations than with the cultured cells. 

Figure 6 shows immunofluorescence localization of the Na⁺-K⁺-2Cl⁻ cotransporter in fresh and cultured bovine corneal endothelium and fresh rabbit endothelium. We included rabbit endothelium because the endothelial fluid transport experiments that concluded that Na⁺-K⁺-2Cl⁻ cotransport may not be expressed in fresh endothelium were performed with rabbit. ⁹ Both T4 monoclonal antibodies and N1 polyclonal antibodies revealed lateral membrane staining in fresh bovine and rabbit endothelium. In addition, in bovine there was a significant amount of cytoplasmic staining with the nucleus prominently excluded. Cultured cells also showed lateral membrane staining with N1, but it was weaker than with fresh cells. We were unable to reveal any lateral membrane staining in cultured cells with the T4 antibody. 

Discussion

Previous evidence for Na⁺-K⁺-2Cl⁻ cotransport in cultured corneal endothelial cells was based on bumetanide-sensitive Rb⁺ uptake and bumetanide sensitive regulatory volume increase. ⁸ In the present study, we confirmed these physiological findings by demonstrating furosemide-sensitive Cl⁻ and Na⁺ fluxes in the cultured cells. Furthermore, we showed that furosemide-sensitive Cl⁻ flux was present across the basolateral membrane, but not the apical membrane. 

Similar types of experiments using freshly isolated endothelial tissue are technically more difficult and fraught with more artifacts. The chloride-sensitive dye SPQ can be used with long-working-distance objectives; however, SPQ binds to the basement membrane of dissected endothelium and can be taken up into the stroma of intact corneas. MEQ is a halide-sensitive fluorescent dye with spectral properties similar to SPQ. MEQ is loaded in the form of the nonfluorescent compound diH-MEQ, which is membrane permeable and converted to MEQ intracellularly by an oxidative mechanism. Initially, we tried MEQ experiments with intact corneal buttons, but the responses to removing chloride were very slow. This was probably because the stroma acts as a large reservoir of chloride, so that removing chloride from the endothelial side causes stromal chloride to enter the cells leading to slow and incomplete depletion of cellular chloride. To avoid the effects of this stromal chloride, we used dissected Descemet’s–endothelial explants loaded with MEQ. As in the cultured cells, furosemide-sensitive Cl⁻ fluxes were observed in MEQ-loaded fresh endothelial cells. 

Western blot analyses showed that cultured cells expressed NKCC1 and the lateral membrane N1 antibody staining seen in the cultured cells was consistent with the furosemide-sensitive Cl⁻ flux being on the basolateral side. It is not clear why we were unable to demonstrate lateral staining in cultured cells using the T4 antibody. Although not a general requirement, the NKCC1 epitopes recognized by T4 and N1 required unmasking by SDS treatment in the corneal endothelial cells. Five minutes of 1% SDS exposure was sufficient to unmask both epitopes in fresh cells and the N1 epitope in cultured cells. In an attempt to unmask the T4 epitope in cultured cells, we tried longer exposure to SDS; however, this treatment caused the fixed cells to separate from one another and in some cases to come off the coverslip. Although there was a question about whether Na⁺-K⁺-2Cl⁻ cotransport was even present in fresh cells, ⁹ Western blot analysis indicated that expression was apparently greater in the fresh cells than in the cultured cells. This agrees with preliminary work showing greater transcriptional production of the Na⁺-K⁺-2Cl⁻ cotransporter message in fresh cells as well. ¹⁷ Immunofluorescence staining with N1 and T4 antibodies was most apparent at the lateral membranes in the fresh bovine and rabbit endothelial cells; however, in the absence of higher spatial resolution we cannot exclude an apical or basal location as well. 

These results indicate that NKCC1 is expressed in both cultured and fresh bovine corneal endothelium, it is located in the lateral membrane, and it is functional, inasmuch as furosemide slows induced Cl⁻ fluxes. Loop diuretics such as furosemide have also been shown to block Cl⁻ channels. ¹⁸ Thus, it is possible that the inhibition of Cl⁻ fluxes seen in these experiments could have been due to this effect. To block channels, however, 1 mM furosemide was needed, which is 10 times more than used currently. Further, if furosemide had a significant effect on anion channel conductance, then we would expect the membrane voltage to hyperpolarize and[ Na⁺ _i] could increase. Previously, we have shown that furosemide had no effect on corneal endothelial membrane potential, ⁷ and in these experiments we showed that furosemide decreased[ Na⁺ _i]. Thus, it is unlikely that furosemide-sensitive anion channel fluxes exist in corneal endothelial cells. 

Another possible source of Cl⁻ fluxes is K⁺-Cl⁻ cotransport, which can be measured as bumetanide-insensitive–furosemide-sensitive Rb⁺ fluxes. ¹⁹ There have not been any studies that have specifically examined whether K⁺-Cl⁻ cotransport is present in corneal endothelium. However, high concentrations of furosemide are used to block K⁺-Cl⁻ cotransport (1 mM vs. 50–100 μM used in the present study), and, furthermore, it has been shown that Rb⁺ fluxes are reduced to less than 10% of control in the presence of bumetanide and ouabain, ⁸ arguing against a significant role of K⁺-Cl⁻ cotransport in corneal endothelial cells. Taken together, these results are consistent with inhibition of coupled electroneutral Na⁺-K⁺-2Cl⁻ entry and not with furosemide-sensitive Cl⁻ fluxes due to Cl⁻ channel blockade or K⁺-Cl⁻ cotransport. 

In addition to the lateral membrane staining obtained using the Na⁺-K⁺-2Cl⁻ cotransporter antibodies, there was significant cytosolic staining. This may indicate that there is a significant amount of Na⁺-K⁺-2Cl⁻ cotransporter in reserve that may respond to upregulation in chloride transport (e.g., adenosine exposure ⁷ or cell shrinkage ⁸ ). Further studies are needed to explore this possibility. 

Does Na⁺-K⁺-2Cl⁻ cotransport contribute to salt and water secretion across the corneal endothelium? The lateral membrane location of the Na⁺-K⁺-2Cl⁻ cotransporter indicates potential contribution to vectorial salt and water transport. In many secretory epithelia, for example, basolateral Na⁺-K⁺-2Cl⁻ cotransport provides Cl⁻ influx, and Cl⁻ channels serve as an apical efflux mechanism. When Cl⁻ channels are closed,[ Cl⁻ _i] is relatively high, which acts as a negative regulator of cotransporter activity (see Reference 20 for review). When apical Cl⁻ channels are open, Cl⁻ efflux occurs as long as[ Cl⁻ _i] is above electrochemical equilibrium. This requires continual influx from the basolateral Na⁺-K⁺-2Cl⁻ cotransporter, the negative regulation of which is possibly released by an initial decrease in[ Cl⁻ _i]. Otherwise, the electrochemical gradient for Cl⁻ is depleted. In fact, we have previously shown in cultured corneal endothelium that[ Cl⁻ _i] is unaffected by Cl⁻ secretagogues (e.g., forskolin) but is depleted if Cl⁻ efflux is stimulated by forskolin and cotransporter uptake is blocked simultaneously by furosemide. ⁷  

In the present study we asked whether the cotransporter brings net salt into the cell in the unstimulated condition. If it does, then furosemide should reduce[ Cl⁻ _i], even transiently, albeit at a much slower rate than in the stimulated cell. This occurred in approximately 50% of cultured cells and in only 1 of 10 trials in fresh cells. These results indicate that transendothelial Cl⁻ flux that has the Na⁺-K⁺-2Cl⁻ cotransporter as a component is possible in corneal endothelial cells, but that in the unstimulated condition, especially in fresh cells, it is close to equilibrium. 

This contrasts with cotransporter activity assayed as bumetanide-inhibitable Rb⁺ uptake in the presence of ouabain, which indicates significant activity under isotonic unstimulated conditions in cultured cells. ⁸ Rb⁺ uptake could still occur if the cotransporter were at equilibrium (i.e., if influx and efflux through the cotransporter are equal or if the cotransporter affected a net outward flux). In addition, evidence from other cell types indicates that at equilibrium the cotransporter can “run in neutral,” allowing the partial reaction (K⁺[Rb⁺]/K⁺ exchange) to occur without any Cl⁻ flux due to inhibition by intracellular Cl⁻. ²¹ Thus ⁸⁶Rb uptake, which measures unidirectional fluxes, does not provide information about the existence or direction of a net chloride flux. 

Our results, together with the negative effects of bumetanide on rabbit corneal endothelial fluid transport, ^{4 9} suggest that the cotransporter does not have a major direct role in ion-coupled fluid transport in the unstimulated corneal endothelium. Thus it may be that Na⁺-K⁺-2Cl⁻ cotransporter blockade would have very little effect on endothelial fluid transport, as shown in rabbit corneas, ^{4 9} unless fluid transport is stimulated by a Cl⁻-dependent process as can occur with adenosine or cyclic adenosine monophosphate. ^{7 22}  

 Supported by Grant R01 EY08834 from the National Eye Institute.

 Submitted for publication June 16, 1999; revised August 18, 1999; accepted September 14, 1999.

 Commercial relationships policy: N.

 

Figure 1.

 

Effect of furosemide (50 μM) on the steady state intracellular chloride concentration in cultured corneal endothelial cells[ Cl⁻ _i]. dF/F, relative change in SPQ fluorescence. Arrows: solution changes.

Figure 2.

 

Effect of furosemide (50 μM) on steady state[ Na⁺ _i] and Na⁺ leak. (A) Furosemide caused a slow decrease in intracellular[ Na⁺ _i], and (B) slowed the rate of Na⁺ leakage into endothelial cells exposed to 100 μM ouabain.

Figure 3.

 

Effects of chloride withdrawal from apical or basolateral sides of cultured endothelial cells on intracellular[ Cl⁻ _i] in the absence (A) and presence (B) of 100 μM furosemide. Boxes indicate the time when cells were exposed to Cl⁻-free solutions.

Table 1.

 

Percentage Increase in SPQ Fluorescence (ΔF/F) after 5 Minutes of Cl⁻-Free Perfusion in the Presence and Absence of Furosemide

	Control	Furosemide*
Both sides	52 ± 0.10	34 ± 0.13, †
Apical	6 ± 0.02	6 ± 0.02
Basolateral	13 ± 0.01	3 ± 0.01, †

  n = 5 for all conditions.

*  100 μM.

†  Significantly different from control (P < 0.05). Data from unpaired experiments.

Figure 4.

 

Effect of furosemide on MEQ fluorescence in freshly isolated BCECs. Raw photomultiplier output in counts per second (cps) is plotted on the y-axis. Rate of fluorescence change, ΔF/F [cps × min⁻¹ × (baseline cps)⁻¹ × 10³] when chloride was removed or added again are indicated adjacent to straight lines drawn through the data. Furosemide (100 μM) slowed Cl⁻ efflux by 30% and influx by 67%. Boxes indicate the time when cells were exposed to Cl⁻-free solutions.

Figure 5.

 

Immunoblots of cultured and fresh bovine corneal endothelium (Endo Cult) and fresh corneal epithelium (Endo Fresh) using the T4 and N1 antibodies to the Na⁺-K⁺-2Cl⁻ cotransporter. Each lane contained 60 μg of protein.

Figure 6.

 

Immunofluorescence localization of Na⁺-K⁺-2Cl⁻ cotransporter in fresh rabbit (A, B), and bovine (C, D) corneal endothelium, and in cultured BCECs (E, F). (A, C, and E) T4 monoclonal antibodies; (B, D, and F) N1 polyclonal antibodies. Scale bars, 50 μm.