1 Department of Pathology and Laboratory Medicine, 2 Jules Stein Eye Institute, 4 Department of Biological Chemistry, 5 Department of Neurobiology and Brain Research Institute, University of California, Los Angeles 90095; and 3 Retina-Vitreous Associates Medical Group, Los Angeles, California 90278
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ABSTRACT |
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Na,K-ATPase regulates a variety of transport functions in epithelial cells. In cultures of human retinal pigment epithelial (RPE) cells, inhibition of Na,K-ATPase by ouabain and K+ depletion decreased transepithelial electrical resistance (TER) and increased permeability of tight junctions to mannitol and inulin. Electrophysiological studies demonstrated that the decrease in TER was due to an increase in paracellular shunt conductance. At the light microscopy level, this increased permeability was not accompanied by changes in the localization of the tight junction proteins ZO-1, occludin, and claudin-3. At the ultrastructural level, increased tight junction permeability correlated with a decrease in tight junction membrane contact points. Decreased tight junction membrane contact points and increased tight junction permeability were reversible in K+-repletion experiments. Confocal microscopy revealed that in control cells, Na,K-ATPase was localized at both apical and basolateral plasma membranes. K+ depletion resulted in a large reduction of apical Na,K-ATPase, and after K+ repletion the apical Na,K-ATPase recovered to control levels. These results suggest a functional link exists between Na,K-ATPase and tight junction function in human RPE cells.
actin stress fibers
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INTRODUCTION |
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THE RETINAL PIGMENT EPITHELIUM (RPE) forms a blood-retinal barrier at the back of the eye where it regulates the transport of fluid, ions, and metabolites between the neural retina and the choroid layer and plays an essential role in the development and normal functioning of the eye (63). Like other transporting epithelia, RPE cells contain distinct apical and basolateral plasma membrane domains separated by tight junctions that encircle the cell at the boundary between apical and lateral regions (18, 28). In transmission electron micrographs (TEM), tight junctions appear as a series of apparent fusions, also referred as "kissing points," involving the outer leaflets of the plasma membranes of adjacent cells. (2, 54). The two well-defined functions of tight junctions are the gate function regulating the passage of molecules across the paracellular space (50) and the fence function restricting diffusion of lipids and proteins between apical and basolateral membrane domains (53, 60). Although tight junctions have been characterized biochemically in cultured chicken RPE cells (5, 6, 51, 52), the mechanisms involved in the maintenance of tight junctions in mammalian RPE cells are poorly understood.
An interesting feature of RPE cells is the predominant apical
localization of Na,K-ATPase (7, 29, 40, 42, 47), which is
found in most other transporting epithelia at the basolateral plasma
membrane (33). The Na,K-ATPase, also known as the sodium pump, consists of a noncovalently linked - and
-subunit and is a
key enzyme that regulates the intracellular Na+ and
K+ homeostasis in animal cells. Recent studies indicate the
presence of a
-subunit, but its expression is restricted to certain
tissues (3, 58). Na,K-ATPase catalyses an ATP-dependent
transport of three sodium ions out and two potassium ions into the cell per pump cycle, generating a transmembrane sodium gradient that is
crucial for efficient functioning of other Na+-coupled
transport systems and provides the primary energy for uptake and
extrusion of a wide variety of solutes by epithelial cells (31,
34). Recently, we have shown that in Madin-Darby canine kidney
(MDCK) cells, inhibition of Na,K-ATPase function during epithelial
polarization prevents the formation of tight junctions
(48). This study demonstrated that the intracellular Na+ and K+ homeostasis regulated by Na,K-ATPase
plays an important role in the regulation of tight junction structure
and function.
The blood-retinal barrier (BRB), which includes retinal vascular endothelium and the RPE, excludes blood-borne proteins from the retina and regulates the ionic and metabolic gradients required for normal retinal function (16). Increased permeability of the BRB is an early complication in most retinal diseases and is thought to contribute to the development of proliferative retinopathy (13, 14). In a dystrophic Royal College of Surgeons (RCS) rat model, it has been shown that during the progression of this disease RPE cell tight junctions become leaky just as the photoreceptors begin to degenerate (10). In RCS rat RPE cells, a change in Na,K-ATPase activity from the apical to the basolateral plasma membrane domain has been observed after changes in tight junctions (11). Increased permeability of the BRB has also been suggested to be associated with diabetic macular edema (DME) (8) and in animal models for diabetes, reduced Na,K-ATPase activity in RPE cells has been shown (35). Whether a link exists between Na,K-ATPase activity and tight junction structure and function in RPE cells is not known. Our findings that Na,K-ATPase activity is necessary for the formation of tight junctions in MDCK cells (48) indicated that Na,K-ATPase function might also be involved in modulating tight junction function in RPE cells. To test this possibility, we studied the effect of Na,K-ATPase inhibition on tight junction structure and function in human RPE cells.
For our studies, we utilized cultures of human RPE cells that had been extensively characterized previously, and it has been shown that these cells have the morphological and functional properties of fresh, explanted tissues (25-27). We show that inhibition of Na,K-ATPase by ouabain and K+ depletion in tight monolayers of RPE cells results in an increased permeability to ions and nonionic molecules. This increased permeability correlated with reduced numbers of tight junction membrane contact points. Our results are consistent with a hypothesis that Na,K-ATPase function is involved in the modulation of tight junction structure and permeability in polarized monolayers of human RPE cells.
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METHODS |
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RPE cultures. Human RPE cells were established and grown in replacement Chee's essential medium (CEM), as reported earlier (25), in which cultured human RPE cells are highly differentiated and develop functional polarity appropriate for the in vitro modeling of RPE in vivo function. The cells were grown on Millicell-PCF filters 0.4-µm pore size (12 mm) (Millipore, Bedford, MA). The tenets of the Declaration of Helsinki were followed, and the patients (or their guardians) gave consent for the donation of the tissue. Institutional Human Experimentation Committee approval for the use of human eyes was obtained. For K+-free conditions, RPE cells were washed twice and incubated in a K+-free buffer (140 mM NaCl, 1.8 mM CaCl2, 1 mM MgCl2, 20 mM HEPES, 10 mM glucose, pH 7.4, and 10% FBS dialyzed against the K+-free buffer), as described (48). For K+ repletion, K+-free buffer was replaced by culture medium and the cells were incubated at 37°C, 5% CO2 for the time indicated. The control cells received a medium change at the same time. For ouabain treatment, cells were treated with 500 nM ouabain in DMSO. Control cells treated with DMSO alone were used for the experiment.
Transepithelial electrical resistance. At the indicated time points, the transepithelial electrical resistance (TER) of the RPE monolayers was determined using an EVOM Epithelial Voltohmeter (World Precision Instruments, Sarasota, FL). The values were normalized for the area of the filter after subtracting the background resistance of a filter without cells.
Paracellular diffusion studies. Paracellular flux assays were performed on confluent monolayers of RPE cells in culture medium treated with ouabain (8-h incubation) or under K+-depleted conditions (3-h incubation). For K+ repletion, the cells were incubated for 5 h in K+-free buffer before being returned to replacement CEM culture medium for another 20 h. TER was monitored. 3H-Inulin (2 µCi) or 3H-mannitol (2 µCi) (Amersham, Arlington Heights, IL) in 100 µl of culture medium with or without ouabain or K+-free medium (both at 37°C) was added to the apical side of the filter. The basal compartment medium was replaced with 200 µl of prewarmed culture medium with or without ouabain or K+-free medium, respectively, without tracer. The cells were incubated for 45 min at 37°C, and then equal-volume aliquots of apical and basal compartment media were collected. Radiolabeled tracers were quantified by counting aliquots in a liquid scintillation counter, and permeability was calculated with P = (X)B/(X/µl)i/A/T where XB is counts per minute in the basal chamber, (X/µl)i is the initial concentration in the apical chamber, A is the area of the filter in cm2, and T is the time in minutes (6). To determine maximal permeability, 5 mM EDTA was added for 1 h to the culture medium before determination of the paracellular flux.
Electrophysiological studies.
We used conventional electrophysiological techniques as previously
described (23, 27). We mounted the entire culture well in
a modified Ussing chamber in which the apical and basal surfaces of the
cultured RPE monolayer were perfused separately. The transepithelial potential (TEP) was measured differentially between two calomel electrodes that were connected to the apical and basal baths by a pair
of agar-saturated KCl bridges. The apical membrane potential (Vap) was recorded as the differential signal
between the microelectrode and the apical agar bridge. Using the
equivalent electrical circuit model for the RPE, the basal membrane
potential (Vba) was obtained from the
relationship TEP = Vba Vap. We calculated the individual resistance
parameters for the RPE cell circuit at the transition points between
normal and low K+. The transepithelial resistance
(Rt) was recorded by passing current across the
entire preparation and measuring the current-induced transepithelial
voltage change. The membrane resistance ratio (Rap/Rba) was measured as
the ratio of the current-induced voltage changes at the apical and
basal membranes. The paracellular shunt resistance
Rs is derived from the following equation:
Vap =
Eba × Rap (Rap + Rba + Rs). All
signals were amplified, collected, stored, and analyzed on a microcomputer.
Immunoblotting, immunoprecipitation, and phosphorylation of
occludin.
RPE cells were lysed in immunoprecipitation buffer (10 mM Tris, pH 7.4, 150 mM NaCl, 1% Triton X-100, 40 mM N-octylglucoside, 0.2 mM sodium vanadate, 1 mM EDTA, 1 mM EGTA, 1 mM PMSF, and 5 µg/ml each
of antipain, leupeptin, and pepstatin) on ice for 30 min. The cell
lysates were briefly sonicated and centrifuged at 4°C for 10 min at
14,000 rpm in a microfuge. The supernatants were precleared at 4°C
for 30 min with 20 µl of protein A-agarose beads (Invitrogen, Grand
Island, NY) and immunoprecipitated at 4°C for 16-18 h with 40 µl of protein A-agarose beads coated with 1 µg/ml occludin antibody
(Zymed Laboratories, S. San Francisco, CA). After immunoprecipitation,
the beads were washed four times with immunoprecipitation buffer. The
samples were digested with -phosphatase (New England Biolabs,
Beverly, MA) at 30°C for 30 min according to the manufacturer's
instructions and then subjected to SDS-PAGE and immunoblotting as
described (49).
TEM and quantification of tight junction contact points. RPE cells were fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, for 2-4 h at room temperature and processed for transmission electron microscopy as described earlier (49). The number of tight junction contact points was determined in 30-35 randomly selected cell-cell contact sites for each sample from at least two filters. Comparisons among groups (control, K+ depletion, K+ repletion) of the average number of contact points per cell were assessed by analysis of variance and t-tests, correcting for the number of comparisons. Representative results are shown.
Immunofluorescence and confocal microscopy.
Immunofluorescence was performed on cells fixed with methanol as
described previously (48, 49) with antibodies against ZO-1
and occludin (Zymed Laboratories), -catenin (Transduction Laboratories), and Pan-cadherin (Sigma Chemical). Mouse monoclonal antibodies (MAbs) raised against Na,K-ATPase
- (M7-PB-E9) and
-subunits (M17-P5-F11) have been described previously (1, 57). To detect filamentous actin, the cells were fixed in
paraformaldehyde and labeled with FITC-phalloidin (Sigma Chemical).
Epifluorescence analysis was performed using an Olympus AX 70 (Provis)
microscope. Confocal microscopy to monitor polarized distribution of
cadherin,
-catenin, and Na,K-ATPase was performed using a Fluoview
laser scanning confocal microscope (Olympus America, Melville, NY). To
detect FITC-labeled antigens, samples were excited at 488 nm with an
Argon laser and the light emitted between 525 and 540 nm was recorded.
Images were generated and analyzed using the Fluoview image analysis
software (version 2.1.39).
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RESULTS |
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To examine whether Na,K-ATPase function is involved in maintaining
established tight junction structure and function, we used well-characterized human RPE cell cultures (25-27)
that form tight monolayers and develop high TER, indicating the
presence of functional tight junctions. K+ depletion has
been used extensively to inhibit Na,K-ATPase activity in various cell
types (9, 45, 46) and, in particular, in RPE cells
(21, 41, 42). In this method, the pump can be reactivated
by repleting K+, and therefore the reversibility of the
effects of Na,K-ATPase inhibition can be studied (48). To
examine the role of Na,K-ATPase on tight junctions in RPE cells, we
first determined the effect of K+ depletion on TER, which
provides a measure of ion permeability of tight junctions. The TER of
untreated control cells was 525 ± 7 .cm2 (Fig.
1A). Within 1 h of
K+ depletion, the TER dropped to 231 ± 7
.cm2 and gradually reduced to 100 ± 4
.cm2 within 5 h. Because the change
from K+-containing medium to K+-free medium did
not result in a sudden resistance drop, the reduction was specific for
K+-free condition and not due to the medium change itself
(data not shown). The drop in resistance is not due to the increased serum levels (10%) in K+-free medium because
K+-free medium with 1% serum gave identical results (data
not shown). Transfer of cells that had been K+ depleted for
5 h to K+-containing medium resulted in a gradual
increase of the TER, reaching a value of 332 ± 9
.cm2 after ~20 h. After 48-72 h, the
TER values were similar to control cells.
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Electrophysiological studies were performed to further assess the
effects of low K+ on tight junction permeability in
cultured human RPE cells. We used conventional microelectrode
techniques to measure the changes in Rap,
Rba, and Rs in response
to low K+ (19). The Rs
is a measure of tight junction ionic permeability and reflects the
conductance of solute through the paracellular pathway formed by the
tight junctions (55). Therefore, if tight junction
permeability is increased, one would expect a decrease in
Rs. Using the equivalent electrical circuit
model for the RPE (20), we calculated the individual
resistance parameters at the transition points between normal
K+ and low K+ where K+ was
decreased from 5 to 0.5 mM outside the apical membrane of the RPE. As
shown in Table 1, we observed significant
changes in all resistance parameters, with an increase in
Rap and Rba and decrease
in paracellular Rs. Overall, the
Rt decreased. The decrease in
Rt was due to the decrease in
Rs because both the Rap
and Rba increased in low K+.
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To further confirm that the effect of K+ depletion on tight
junction permeability is due to the inhibition of Na,K-ATPase, we
utilized ouabain, a specific inhibitor of Na,K-ATPase. Ouabain treatment reduced the TER from an initial value of 443 ± 40 to 73 ± 11 .cm2 within 8 h, whereas
in control cells the TER value was 253 ± 11
.cm2 at this time point (Fig.
1B). Thus these results are consistent with a role for
Na,K-ATPase function in the regulation of tight junction ion
permeability in RPE cells.
In addition to regulating the paracellular ion flow, tight junctions
also regulate the diffusion of nonionic molecules through the
paracellular space. Tracer studies using 3H-inulin
(hydrodynamic radius ~10-14 angstroms) and
3H-mannitol (hydrodynamic radius ~4 angstroms) showed an
increase in the permeability of these tracers in
K+-depleted cells (Fig.
2A). The permeability of
mannitol in these cells was 2.6 times higher than that of inulin. In
K+-repleted cells, the permeability to these tracers was
comparable to that of control cells, indicating that Na,K-ATPase
inhibition increases the paracellular permeability of nonionic
molecules in RPE cells. Ouabain treatment increased the permeability of both inulin and mannitol (Fig. 2B). The permeability of
inulin and mannitol were three- and sixfold, respectively, higher in ouabain-treated cells compared with control cells. The permeability to
inulin and mannitol in EDTA-treated cells was 21- to 23-fold higher than in K+-depleted cells, indicating that EDTA
treatment affects tight junction permeability more drastically than
Na,K-ATPase inhibition. Taken together, these results demonstrate that
inhibition of Na,K-ATPase increases the permeability of both ionic and
nonionic molecules through the paracellular space.
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Inhibition of Na,K-ATPase during the formation of tight junctions
in MDCK cells resulted in a discontinuous staining pattern of tight
junction proteins such as ZO-1 and occludin, indicating that inhibition
of Na,K-ATPase prevented the assembly of tight junctions
(48). In contrast, in tight monolayers of RPE cells with
established tight junctions, inhibition of Na,K-ATPase by K+ depletion or ouabain did not affect the localization of
ZO-1, occludin, or claudin-3. Immunofluorescence staining for ZO-1
(Fig. 3, A-D),
occludin (Fig. 3, E-H), and claudin-3 (Fig.
3, I-L) did not reveal significant
differences in the staining patterns between control,
K+-depleted, K+-repleted, and ouabain-treated
cells. These results indicate that the increased permeability of tight
junctions in RPE cells is not accompanied by changes in the
localization of tight junction marker proteins.
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It has been suggested that occludin might contribute to the electrical
barrier function of tight junctions (4, 38) and, moreover,
occludin phosphorylation has been implicated in their permeability
function (36). In control, K+-depleted and
K+-repleted RPE cells, occludin protein levels or its
phosphorylation state did not show appreciable differences (Fig.
4), indicating that Na,K-ATPase
inhibition has a minimal effect on the levels of occludin and its
phosphorylation state in human RPE cells. We also tested the levels of
claudins 1-5 in RPE cells. The only claudin detected by immunoblot
analysis was claudin-3, and we did not detect any change in the level
of this protein between control and K+-depleted cells (data
not shown). A recent study has shown that claudin-5 is transiently
expressed during the development of chick RPE (32),
whereas we did not detect claudin-5 in human RPE cell cultures.
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Although visible differences in the localization of tight junction
proteins were not detected at the light microscopy level, a distinct
difference in tight junction structure was observed at the
ultrastructural level between control and K+-depleted
cells. Control cells showed well-developed tight junctions with
distinct, closely apposed plasma membrane contact points, a distinct
feature of tight junctions (Fig.
5A). However, in
K+-depleted cells, these contact points were not clearly
discernable and junctions appeared as an electron dense region at the
apicolateral cell-cell contact sites (Fig. 5B). After
K+ repletion, the plasma membrane contact points reappeared
and were clearly discernable (Fig. 5C). Quantification of
the TEM data (Fig. 5D) revealed that the average number of
plasma membrane contact points per tight junction in control cells of
4.30 ± 0.23 decreased to 1.60 ± 0.15 in
K+-depleted cells (P < 0.001).
Upon K+ repletion, these contact points increased to
3.22 ± 0.19, indicating that inhibition of Na,K-ATPase reversibly
affects the structure of the tight junction contact points in RPE
cells, although not quite back to control levels (P < 0.002). Another remarkable observation from the TEM was the reduction
of microvilli in K+-depleted cells compared with control
and K+-repleted cells (Fig. 5, compare asterisks in
A-C), indicating that inhibition of
Na,K-ATPase function also disrupts microvillar organization in RPE
cells.
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The apically localized Na,K-ATPase immunostaining was drastically
reduced in K+-depleted cells (Fig.
6). We used MAbs against the - and
-subunits to monitor the localization of Na,K-ATPase. Both
antibodies gave similar results, and the staining pattern of the
-subunit antibody is shown in Fig. 6. In control cells, the
Na,K-ATPase was localized at both the apical and basolateral domain
(Fig. 6, G and H). Upon K+ depletion,
the apically localized Na,K-ATPase decreased while at the same time the
basolaterally localized Na,K-ATPase increased (Fig. 6, I and
J). After K+ repletion, the apical localization
of Na,K-ATPase increased (Fig. 6, K and L),
showing a pattern similar to that in control cells. Interestingly, two basolateral markers, cadherin (Fig. 6,
A-F) and
-catenin (not shown), were
localized to the basolateral domain in K+-depleted cells,
as in control and K+-repleted cells.
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Inhibition of Na,K-ATPase in MDCK cells drastically reduced the amount
of actin stress fibers that reappeared upon K+ repletion,
indicating that Na,K-ATPase plays a role in the formation of stress
fibers (48). Inhibition of Na,K-ATPase in RPE cells drastically reduced the actin stress fibers (Fig.
7B) seen in control cells
(Fig. 7A, arrowhead), yet the cortical actin ring appeared
unaffected (arrows in Fig. 7, A-C). The
stress fibers reappeared upon K+ repletion, indicating that
Na,K-ATPase function plays a role in the formation of stress fibers in
RPE cells, as we previously observed in MDCK cells.
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DISCUSSION |
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In this study, we demonstrate that Na,K-ATPase function is involved in the maintenance of tight junction structure and permeability in human RPE cells. We show that inhibition of Na,K-ATPase by two independent methods (ouabain and K+ depletion) resulted in an increase of tight permeability to both ions and nonionic molecules. This increased permeability was accompanied by a decrease in tight junction membrane contact points rather than a change in the distribution of tight junction proteins or occludin phosphorylation.
Our electrophysiological studies support the conclusion that inhibition of the Na,K-ATPase with low K+ increases the shunt permeability. We observed a decrease in Rt, accompanied by a decrease in paracellular Rs in low K+, both of which were significant. The decrease in Rt was not due to a change in membrane (transcellular) resistance because both Rap and Rba increased in low K+. The Rs is a measure of tight junction permeability and reflects the conductance of solute through the paracellular pathway formed by the zonula occludens (55). The reduction in tight junction membrane contact points reported in this study could account for an increase in shunt permeability observed. We argue that inhibition of the Na,K-ATPase is one mechanism by which low K+ may achieve this effect on tight junction permeability. Although effects of low K+ on other transport pathways could contribute to the increase in paracellular permeability, previous electrophysiological studies in a variety of species and preparations (including human RPE) are consistent with an increase in paracellular ionic permeability caused by Na,K-ATPase inhibition. Inhibition of the Na,K-ATPase with ouabain was actually found to increase transepithelial conductance (decrease resistance) in frog (21), cat (56), and human (27). This is not consistent with the decrease in conductance expected by blocking the electrogenic Na,K-ATPase current. Blocking the pump should increase membrane resistance and produce an increase in transepithelial resistance if this was the only effect. Griff et al. (21) concluded that in frog, the observed increase in transepithelial conductance could be best explained by a decrease in Rs (increase in paracellular ionic conductance) produced by pump inhibition. They did not propose a mechanism for such an increase in paracellular permeability. The biochemical and morphological studies on tight junctions presented here explain for the first time how inhibition of Na,K-ATPase may increase paracellular shunt permeability in RPE cells.
There are several cellular mechanisms by which pump inhibition may lead to an increase in paracellular permeability. Previous studies showed that loss of tight junction structure and permeability correlated with the disruption of the circumferential actin ring (37, 59) that is located at the apical pole of polarized epithelial cells. This ring can be visualized by light microscopy by phalloidin staining for filamentous actin and colocalizes with tight junction and adherens junction proteins. We observed, however, that the increased tight junction permeability in Na,K-ATPase-inhibited RPE cells was not accompanied by an apparent change of the circumferential actin ring. Although less prominent, stress fibers projecting from the perijunctional actin ring interface at the cytoplasmic surface of tight junction membrane contact points (24, 36, 37), but their relation to tight junction permeability is unknown. We discovered that reduced stress fiber content correlated with the loss of tight junction membrane contact points and increased tight junction permeability in human RPE cells. In MDCK cells, loss of tight junctions upon Na,K-ATPase inhibition resulted in reduced stress fiber content and strongly correlated with reduced RhoA GTPase activity (48). Several studies have shown that RhoA GTPase function is also involved in the modulation of tight junction structure and function (30, 43). Because RhoA GTPase is known to be involved in the stress fiber formation (22), the reduced stress fiber content we observed in human RPE cells might be a consequence of reduced RhoA GTPase activity. Because large amounts of cells are required for such biochemical analysis, we were unable to carryout such studies in human RPE cells. However, taken together, these results indicate that stress fibers might be involved in the modulation of tight junction structure and function. We speculate that tight junction membrane contact points are maintained by a dynamic equilibrium with stress fibers. Then, it is possible that loss of stress fibers might decrease the membrane contact points and increase the permeability of tight junctions without an apparent change in the perijunctional actin ring like we observed in this study. Future studies are necessary to substantiate this hypothesis and to determine whether such regulation is unique to the RPE.
How Na,K-ATPase inhibition affects tight junction permeability is currently not known. We found that the effect on tight junctions was reversible, indicating that the mechanism is not simple degeneration of cellular function. Changes in membrane voltage alone may alter tight junction permeability if the tight junction conductance is voltage sensitive or if it creates ionic gradients across the shunt that alter its permeability. Ouabain and low K+ produced similar effects on tight junction permeability but have opposite effects on membrane voltage, arguing against a voltage-sensitive mechanism (23, 27). Pump inhibition would decrease extracellular Na+ in the unstirred layers above the cell membrane (44), and this should decrease the shunt conductance, the opposite of what we observed. Other mechanisms that cannot be ruled out include changes in signaling pathways by the intracellular Na+ increase or K+ decrease caused by the inhibition of Na,K-ATPase that might alter tight junction permeability (39, 62). Finally, although there were no obvious differences in cell size between control and Na,K-ATPase-inhibited cells (compare similar size of the XZ confocal sections in Fig. 6), we cannot rule out small cell volume changes that might contribute to changes in tight junction permeability.
Although low K+ and ouabain are both known to inhibit the Na,K-ATPase in RPE cells (21, 27, 41, 42, 56), our experiments do not rule out the possibility that cellular changes induced by low K+ independent of Na,K-ATPase inhibition might also be involved in altering tight junction permeability in RPE cells. For example, whereas both low K+ and ouabain increased tight junction permeability, the effect of low K+ on TER was more rapid compared with that of ouabain. Future experiments will determine the mechanisms involved in the alteration of tight junction permeability by low K+ and ouabain.
Our results suggest that normal Na,K-ATPase function is necessary to
localize this protein to the apical plasma membrane. Na,K-ATPase is
associated with microvilli in RPE cells (15). Although the
polarity of the basolateral markers cadherin and -catenin remained
unaltered, we found a large, reversible decrease in the amount of
Na,K-ATPase localized to the apical plasma membrane in
K+-depleted cells. This was accompanied by a loss of
microvilli in the apical membrane that was also reversible. The loss of
microvilli in low K+ that we observed could therefore
account for the change in localization of Na,K-ATPase. Microvilli in
RPE cells possess an internal core of densely packed actin filaments
(17, 61). Reduced stress fiber content in
K+-depleted cells might contribute to diminished microvilli
observed in these cells. Increased cell volume with disruption of the
actin skeleton might also alter microvilli, but we did not observe a visible change in the cell volume. Further experiments will be necessary to determine the mechanisms involved in the loss of microvilli and altered localization of Na,K-ATPase in
K+-depleted cells.
Reduced Na,K-ATPase activity might lead to increased tight junction permeability and contribute to disease progression in animal models of retinal degeneration. RPE cultures of the dystrophic RCS rats need to be maintained in a medium conditioned by normal retinas to form functional tight junctions, and replacement of this medium with unconditioned medium or medium conditioned by dystrophic RPE results in the decline of the TER (12). There are two important similarities between the current study and results reported on RCS rats regarding tight junctions and Na,K-ATPase. 1) As in human RPE cells (this study), RCS rat RPE cultures maintained in the absence of normal retina conditioned medium showed a drop in TER that was not accompanied by changes in the localization of the tight junction marker protein ZO-1 (12), and 2) as in human RPE cells (this study), after junctional breakdown in rat RPE cells, the Na,K- ATPase is localized more basolaterally compared with its apical localization in normal RPE cells. From these similarities, it is tempting to speculate that reduced Na,K-ATPase function in RPE cells of the RCS rats might be involved in the increased permeability of RPE cell tight junctions. In addition, reduced Na,K-ATPase function might be associated with other forms of retinal disease. For example, in animal models for diabetic retinopathy, RPE cells have reduced Na,K-ATPase activity (35). Also, increased permeability of the BRB is associated with DME (8). It is possible that decreased Na,K-ATPase activity in DME might increase the permeability of the BRB and play a role in the pathogenesis of diabetic retinopathy.
In conclusion, these results are the first to show that in RPE cells there is a link between Na,K-ATPase activity and tight junction structure and function. We suggest that understanding the role of Na,K-ATPase in the regulation of tight junctions should significantly improve our understanding of the functions of BRB integrity in normal and disease states of the eye.
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ACKNOWLEDGEMENTS |
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We thank Dr. Elliot Landaw for statistical analysis of the EM data.
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FOOTNOTES |
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This work is supported by National Institutes of Health Grant DK-56216 to A. K. Rajasekaran and EY-00331 and EY-00444 to D. Bok. S. A. Rajasekaran is supported by USHHS Institutional National Research Service Award T32CA09056. A. K. Rajasekaran is a member of the Jonsson Comprehensive Cancer Center. D. Bok is the Dolly Green Professor of Ophthalmology at UCLA.
Address for reprint requests and other correspondence: A. K. Rajasekaran, Dept. of Pathology and Laboratory Medicine, Rm. 13-344 CHS, Univ. of California, Los Angeles, CA 90095 (E-mail: arajasekaran{at}mednet.ucla.edu).
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.
First published February 5, 2003;10.1152/ajpcell.00355.2002
Received 1 August 2002; accepted in final form 2 February 2003.
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