Fluid transport by human nonpigmented ciliary epithelial layers in culture: a homeostatic role for aquaporin-1

Rajkumar V. Patil1,2, Zhiqiang Han1, Maimaiti Yiming3, Junjie Yang1, Pavel Iserovich3, Martin B. Wax1, and Jorge Fischbarg3,4

Departments of 1 Ophthalmology and Visual Sciences and 2 Molecular Biology and Pharmacology, Washington University School of Medicine, St. Louis, Missouri 63110; and Departments of 4 Physiology and Cellular Biophysics and 3 Ophthalmology, Columbia University, New York, New York 10032


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We report for the first time that cultured nonpigmented human ciliary epithelial (NPE) cell layers transport fluid. Cells were grown to confluence on permeable membrane inserts, and fluid transport across the resulting cell layers was determined by volume clamp at 37°C. These cell layers translocated fluid from the apical to the basal side at a steady rate of 3.6 µl · h-1 · cm-2 (n = 4) for 8 h. This fluid movement was independent of hydrostatic pressure and was completely inhibited by 1 mM ouabain, suggesting it arose from fluid transport. Mercuric chloride, a nonspecific but potent blocker of Hg2+-sensitive aquaporins, and aquaporin-1 antisense oligonucleotides both partially inhibited fluid transport across the cell layers, which suggests that water channels have a role in NPE cell homeostasis. In addition, these results suggest that of the two ciliary epithelial layers in tandem, the NPE layer by itself can transport fluid. This cultured layer, therefore, constitutes an interesting model that may be useful for physiological and pharmacological characterization of ciliary epithelial fluid secretion.

aquaporins; ciliary epithelium; aqueous humor


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

CILIARY EPITHELIUM IS RESPONSIBLE for the secretion of aqueous humor into the posterior chamber of the eye. Most of the aqueous humor secretion is driven by the active transport of electrolytes from the plasma into the posterior chamber followed by rapid movement of water and solutes (17). The ciliary epithelium is composed of two layers juxtaposed, the proximal (or outer) pigmented epithelium (PE) and the distal (or inner) nonpigmented epithelium (NPE). These two layers exhibit characteristics found in transporting epithelia (16, 43). Ciliary epithelial cells have been cultured for several years and have been characterized for the presence of several ion transporters and receptors (24, 29, 41). In addition, ex vivo preparation of rabbit ciliary epithelium has been established and utilized for studying the ion transport across this bilayer (7, 22, 36). For the NPE layer (both in vivo and in cultured cells), the Na+-K+-ATPase, which is located on its basolateral membranes, provides the gradient required by a host of Na+-dependent cotransporters (2, 12, 13, 28).

In contrast to the rich information on electrolyte movements, very little is known about water movements across ciliary epithelial cell membranes. Presumably, part of the reason for that is the lack of convenient in vitro models for studying fluid transport across ciliary epithelial layers. We now report that the NPE cells, when grown on permeable supports, can be advantageously utilized to study translayer fluid movements. It is not clear whether or not ciliary fluid secretion in vivo requires the simultaneous presence of both the PE and NPE layers transporting in tandem. Our findings now strongly suggest that the cultured NPE layer alone transports fluid actively. The implications of these findings for the mechanism of fluid secretion in vivo remain to be determined.

Last, using molecular, immunological, and biochemical techniques, we have recently demonstrated that functional water channels (aquaporin-1 or AQP1) are expressed in simian virus 40 (SV40)-transformed human NPE cells (15). Our present results suggest that AQP1 contributes to the homeostasis of these cells.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cultured human NPE cell layers. The SV40-transformed and well characterized clone (ODM2) of human NPE cells (3, 6, 41) was used in this study. NPE cells (passage 8) were grown on 24.5-mm-diameter transparent, permeable, collagen-treated supports (Transwell, Costar) at a density of 1 × 106 per support in Dulbecco's modified Eagle's medium (DMEM), supplemented with 10% fetal bovine serum (FBS), in a 5% CO2 atmosphere at 37°C. Experiments were performed within 1-7 days after confluence. Confluence was determined by microscopy and was also ascertained by determining the resistance of each insert before the experiment. Electrical resistance of NPE layers was determined using an Endohm chamber and a resistance meter (WPI, Sarasota, FL). The resistance determined with a control insert was subtracted from that of an insert with the cell layer. Specific resistance was calculated by multiplication by the area of the insert (4.7 cm2); the usual value for a control insert was 56.4 Omega  · cm2. For a set of five NPE cultured layers representative of those utilized, the (corrected) specific resistance was 73.32 ± 15.6 Omega  · cm2. These values are of the order of those seen for other cell layers grown on the same support by us (alpha TN4 cells: 130 ± 7.3 Omega  · cm2; bovine lens epithelium: 85.4 ± 5.9 Omega  · cm2) (8). For 2 h before beginning and throughout the experiments, the cell layers were bathed by DMEM containing FBS without antibiotics. The basal side of the confluent NPE layer grown in this fashion is closer to the collagen support, and the apical side is facing away from the support.

Antisense oligonucleotides. A phosphorothioated antisense oligonucleotide targeted toward the region encompassing the ATG start codon of AQP1 mRNA (from base -7 to +11) (39) was synthesized. The oligonucleotide had natural nucleotides flanked by four phosphorothioate-modified deoxynucleic acids at the 5' and 3' ends (TCGCTGGCCATGCTGGCA). Bold letters in the oligonucleotide refer to phosphorothioated bases, and the underlined region corresponds to the location of the start codon. Phosphorothioated oligos are stable (exonuclease resistant) and are transported into cells more efficiently than unmodified oligos (35). A second oligonucleotide, with the same base composition but with a scrambled (nonsense) sequence (TGGGCTCACACCGTCGGT), was used as control for nonspecific or potential toxic effects of the oligonucleotide. Scrambling was done using DNA analysis software (MacVector). Cells were grown without oligonucleotides and incubated with antisense or nonsense oligonucleotide (5 µM) for 16-18 h in serum-free media before measurements.

Fluid flow measurements. The chamber utilized has been described in prior publications (8). Briefly, it consists of two Lucite halves with water jackets for temperature control that are kept saturated with water when not in use. The top half of the chamber holds the Costar Transwell insert, and a rubber gasket separates the plastic bottom of the insert from a seat in the bottom chamber. The clamping force to seal the chamber is exerted on the plastic surfaces and not on the cells. The bottom chamber seat for the insert accommodates a stainless steel wire mesh to prevent sagging of the flexible permeable support. The bottom chamber is sealed by a plug and an O-ring and is pierced by a 16-gauge stainless steel tube. The top chamber is also sealed by a plug and an O-ring and pierced by several tubes; it was gassed continuously with moist 95% air and 5% CO2. The steel tube coming from the bottom plug is connected to the automatic fluid level detector. The current nanoinjector method uses the signal from the detector to provide negative feedback and to keep the volume of the bottom compartment constant. Volume flow recordings correspond to the rate of injection (or withdrawal) of fluid from the bottom chamber. The relative positions of the chamber and the detector were such that the hydrostatic pressure difference applied to the apical side of the cell layer (top chamber) was 3 cmH2O. To be noted, the configuration of the setup precludes gassing of the bottom compartment.

Immunoblotting of AQP1 in NPE cell membranes. Immunoblot analysis was carried out on membrane preparations of the cultured cells. Cells were grown without oligonucleotides and were incubated for 18 h with sense and antisense AQP1 oligonucleotides (5 µM). For membrane preparations, cells were first lysed in the lysis buffer (2 mM HEPES and 2 mM EDTA, pH 7.4) on ice for 15 min, vortexed briefly, and centrifuged at 37,000 g for 30 min. The membrane pellet was resuspended in Tris · HCl (50 mM, pH 7.5) buffer, solubilized in the homogenizer, 25 µg of membrane protein was mixed with the sample buffer and resolved by 13% SDS-PAGE, and electrotransferred to a nitrocellulose membrane. The membrane was blocked with 1% bovine serum albumin in Tris-buffered saline (TBS; 20 mM Tris · HCl, pH 7.5, 150 mM NaCl, and 0.05% Tween 20) for 1 h at room temperature and was incubated with AQP1-specific polyclonal antibody (Alomone Labs, Jerusalem, Israel). The membranes were washed four times with TBS, incubated with an anti-rabbit IgG-alkaline phosphatase conjugate antibody (Promega, Madison, WI), and immunoreactive bands were detected by enhanced chemiluminescence (Amersham, Arlington Heights, IL).

Northern blotting of AQP1 in NPE cells. Cells were grown without oligonucleotides and were incubated for 18 h with sense and antisense AQP1 oligonucleotides (5 µM) for 18 h. Northern blot analysis was performed with the poly(A)+ RNA isolated using a poly(A)+ RNA isolation kit from Qiagen (Chatsworth, CA). Denatured poly(A)+ RNA (1.0 µg) was separated on 1.5% agarose gel containing formaldehyde. After electrophoresis, the RNA was electrotransferred to a nitrocellulose membrane using the Bio-Rad Transblot apparatus. RNA was cross-linked to the nitrocellulose membrane by an ultraviolet crosslinker. The blot was prehybridized at 42°C for 2 h with 100 µg/ml of denatured salmon sperm DNA in 50% formaldehyde, 0.04% polyvinylpyrrolidone, 0.04% bovine serum albumin, 0.04% Ficoll, and 1% SDS in 5× saline sodium citrate (SSC) buffer. PCR-amplified DNA labeled with 32P was used as a probe. Hybridization was carried out at 42°C for 16 h. Filters were washed twice in 2× SSC at room temperature for 15 min and once in 1× SSC for 15 min containing 1% SDS. The blots were exposed to Kodak X-Omat AR film with intensifying screen at -80°C for 3 days to detect the AQP1 mRNA expressed in NPE cells.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Fluid transport by NPE cell layers. The introduction of permeable tissue culture supports has made it possible to measure the ability of cells to transport fluid in monolayer cultures as well as to measure the electrical resistance and potential differences across the monolayers. We used a modified Bourguet-Jard technique (10, 30) to measure fluid transport across cultured monolayers of NPE cells. This method has been successfully used to measure water and fluid transport across lens epithelium and corneal limiting layers (8, 9, 11, 26, 45), gives resolutions as high as 1-3 nl, and can be applied to any layer of cultured cells. From the direction of fluid transport across the NPE cell layer in vivo, if the cultured cells would behave likewise, we would expect fluid movement from the top chamber to the bottom chamber. Figure 1 shows that fluid movement across an NPE cell layer indeed occurs in the expected direction, from the apical to the basal surface, corresponding to the activity of an absorptive epithelium. This movement of fluid took place spontaneously and continuously for several hours; the longest time monitored was 8 h. In other experiments, varying pressure heads were applied to the NPE cell layer. Hydrostatic pressure differences (up to 7 cmH2O) had no effect on fluid movement, as exemplified in Fig. 2. Because such high pressure differences are unlikely across an in vivo layer, we chose a pressure head of 3 cmH2O as a standard. This pressure is required to keep the cells and their support relatively immobile; at the same time, it lessens capillarity artifacts in the sensor tube. After ~8 h in the chamber, the rate of fluid movement increased progressively, which is consistent with an increasing leak across the deteriorating cell layers (driven by the existing pressure head). The average rate of fluid movement was 3.6 + 0.3 µl · h-1 · cm-2 (n = 4).


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Fig. 1.   Chart recording of fluid movement by cultured nonpigmented ciliary epithelial layer (NPE). Recording begins after an ~15-min stabilization period. Individual deflections represent fluid accumulating during a fixed time interval (10 s) in the downward direction, after which the accumulator is reset and the process restarts. The downward direction corresponds to fluid movement from the top chamber to the bottom chamber, i.e., from the apical to the basal side of cells. The fluid level in the top chamber section was 3 cm above that of the detector communicating with the bottom chamber. The cultured monolayer transported fluid for several hours; the longest time monitored was 8 h (not shown). Similar results were obtained in 4 separate experiments with different batches of NPE cells.



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Fig. 2.   Chart recording of fluid movement by cultured NPE layer under varying pressure heads. Arrows indicate the hydrostatic pressure (in cmH2O) applied to the NPE monolayer. Hydrostatic pressure differences up to 7 cmH2O had no effect on fluid movement. Similar results were obtained in 3 separate experiments with different batches of NPE cells.

We further investigated the nature of this absorptive fluid movement by exposing the apical side of the cells to ouabain (1 mM), which blocks the Na+-K+-activated ATPase. Figure 3 shows a representative experiment (n = 4); after an ~20-min delay, fluid movement began to decrease and was eventually abolished over a 45-min period.


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Fig. 3.   Effect of ouabain on fluid movement across cultured NPE cells. Recording begins after an ~15-min stabilization period. Ouabain (1 mM) was added to the apical side of the layer (top chamber) and inhibited fluid movement after an ~20-min delay. Similar results were obtained in 4 separate experiments with different batches of NPE cells.

Mercuric chloride is an unspecific but potent blocker of Hg2+-sensitive aquaporins. This inhibitor at a concentration of 300 µM was added to the top chamber (in contact with the apical cell membranes). As Fig. 4 exemplifies, after an ~30-min delay, fluid movement began to decrease slowly over a 30-min period. The average rates of fluid movement in control and 30 min after mercuric chloride addition were 3.9 ± 0.3 µl · h-1 · cm-2 (n = 4) and 2.1 ± 0.2 µl · h-1 · cm-2 (n = 4), respectively. This effect is consistent with inhibition of water channels; the delay for mercuric chloride action is in the order of the incubation time required for this agent to act in other systems, such as fluid transport by lens epithelium (8) and oocyte-expressing water channels (34). Subsequently, the rate of fluid movement increased continuously, perhaps due to the toxicity of mercuric chloride resulting in an increasing leak across the cell layer.


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Fig. 4.   Effect of mercuric chloride on fluid movement across cultured NPE. Recording begins after an ~15-min stabilization period. Mercuric chloride (300 µM) was added to the apical side of the layer (top chamber) and inhibited fluid movement after an ~30-min delay. Similar results were obtained in 4 separate experiments with different batches of NPE cells.

Figure 5 shows the effect of AQP1 antisense oligonucleotide on fluid movement across NPE cells. We incubated NPE cells (on their permeable support) with AQP1 nonsense (control) and antisense (treated) oligonucleotides at a final concentration of 5 µM added to the cell culture media 16-18 h before measurements of fluid flow. The initial rates of fluid movement in both control and treated layers were similar. As the experiment progressed, however, the steady-state rates of fluid movement measured over 6-8 h were different, namely, 3.25 + 0.2 µl · h-1 · cm-2 (n = 3) across cell layers pretreated with nonsense oligonucleotide and 1.8 + 0.2 µl · h-1 · cm-2 (n = 4) across the layers treated with antisense oligonucleotide, for a reduction of 54%. Interestingly, such reduction (Fig. 5) was about the same as that for cell layers treated with mercuric chloride (Fig. 4).


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Fig. 5.   Effect of aquaporin-1 (AQP1) antisense oligonucleotides on fluid movement by NPE. A phosphorothioated antisense oligonucleotide targeted toward the region encompassing the ATG start codon of AQP1 mRNA was used. Cells were grown without oligos and incubated with antisense oligonucleotide (5 µM) for 16-18 h before measurement of fluid movement. The initial transient change in rate of fluid movement lasting 0.5 h represents the adjustment of the cell layer and support after the mounting procedure. The steady-state rate of fluid flow was much lower than in the control series exemplified in Fig. 1. Similar results were obtained in 4 separate experiments with different batches of NPE cells.

The partial inhibitions in fluid transport with HgCl2 and antisense oligonucleotides, together, are consistent with these manipulations, resulting in blockage of water channels. However, the initial rate of fluid transport by the cells incubated with antisense oligonucleotides (Fig. 5) appears to be as large as the rates seen in Figs. 1 and 4; it is only later, during those experiments, that the inhibition gradually appears. To determine whether the effect by AQP1 antisense oligonucleotide was specific, we performed Western and Northern blots on the protein extracts and mRNA preparations, respectively, from NPE cells after oligonucleotide treatment. In Fig. 6, AQP1 antisense oligonucleotide treatment with NPE cells significantly decreased the protein concentrations (lane 2) and moderately reduced mRNA levels (lane 4) compared with the AQP1 nonsense oligonucleotide treatment (lanes 1 and 3), which had no effect on the AQP1 protein and mRNA levels. The inhibitory action of antisense oligonucleotide at the transcriptional level may be due to the degradation of antisense oligonucleotide-mRNA heteroduplex by RNase H, which destroys the RNA but leaves the antisense oligonucleotide intact to hybridize with yet another mRNA target, due to the inhibition of specific RNA processing steps such as 5' capping, splicing, and nuclear export. There were no differences in AQP1 protein or mRNA levels in cells treated with or without nonsense oligonucleotide treatment (data not shown).


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Fig. 6.   Effect of AQP1 antisense oligonucleotides on protein and mRNA expression levels determined by Western and Northern blots, respectively. Cells were grown without oligonucleotides and were incubated for 18 h with sense (lanes 1 and 3) and antisense (lanes 2 and 4) AQP1 oligonucleotides (5 µM) for 18 h. Western and Northern blots were performed on the protein extracts (lanes 1 and 2) and mRNA preparations (lanes 3 and 4), respectively, from NPE cells after oligonucleotide treatment. Lanes 1 and 2 were loaded with 25 µg of membrane proteins, and lanes 3 and 4 were loaded with 1.0 µg of mRNA preparation. Similar results were obtained in 3 separate experiments with different batches of NPE cells.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

In prior experimental work that examined fluid secretion by ciliary epithelium in vivo (4, 18, 32), it has not been possible to ascertain whether the simultaneous presence of both layers is required for fluid secretion because its two layers (PE and NPE) are always together. In addition, there is no prior instance in which fluid movements have been determined across an isolated or cultured preparation of this important tissue. Here we report for the first time that the NPE layer by itself (albeit cultured) is capable of translocating fluid in a direction (from apical to basolateral; Fig. 1) and at a rate (3.6 ± 0.3 µl · h-1 · cm-2; cf. Fig. 1) consistent with fluid transport across it. The direction of fluid movement observed is the same as that for fluid transport across this absorptive layer in vivo. In addition, the blockage of fluid movement by ouabain further suggests that fluid movement across NPE cells indeed represents solute-coupled fluid transport.

Whether these observations across cultured cells are totally applicable to the ciliary epithelium in vivo remains to be determined. Still, there is that presumption a priori, since the cultures transport fluid actively, and they do so in the same direction as the in vivo process. Anatomically, it is well documented in the literature that NPE cells in vivo establish zonular structures at the intersections of their apical-lateral membrane domains (27, 42). Correspondingly, NPE cells in culture also form zonulae in primary culture (1, 38).

As mentioned earlier in this paper, varying hydrostatic pressures progressively between 3 and 7 cm of H2O had no effect on the rate of fluid movement (Fig. 2). This suggests that the fluid movement observed is due to fluid transport and not to a leak or an artifact. To further characterize this finding, we chose to study the possible role of the AQP1 water channel in fluid transport. The presence of AQP1 has been demonstrated by others and ourselves in human and rat NPE (14, 31, 37), and, more recently (15), in the same cultured human NPE cells with which this study was done. As detailed in RESULTS, we found that fluid movement across the NPE layer was significantly reduced in cells treated with AQP1 antisense oligonucleotide (Fig. 5). Nonsense oligonucleotide that had the same base composition as the antisense oligonucleotide, but scrambled, had no effect on fluid movement. This, again, suggests that the fluid movement observed is due to an active cellular mechanism (rather than a leak) and that AQP1 plays a role in the cellular processes, resulting in fluid transport by NPE cells.

The observed partial inhibition of fluid movement by HgCl2 is telling in at least two senses. It also militates against the existence of a fluid leak or an artifact, since under the applied hydrostatic pressure, a nonspecific or toxic effect of HgCl2 would have presumably increased the leak, and, therefore, increased the rate of fluid movement. Instead, the opposite happened. However, an explanation for this effect must also take into account the fact that the inhibition was partial (~50%). If the water flow observed traversed membrane water channels, one might argue that perhaps half of the flow would traverse other water-permeable membrane routes. Against this, however, is the observation that upon mounting, cells treated with antisense AQP1 oligonucleotide displayed an initial rate of fluid transport quite comparable to those of untreated cells. As mentioned earlier in this paper, the AQP1 protein content was already much decreased in the antisense oligonucleotide-treated cells, which points away from the fluid movement traversing AQP1 water channels.

An alternative explanation for the inhibitions seen with HgCl2 and antisense oligonucleotide is that AQP1 could have a central role in maintaining cell homeostasis. To be noted, as mentioned previously, despite downregulation of AQP1, the initial rate of transepithelial water transport is not reduced but the later water flow is lowered. This suggests that the downregulation may have a primary role on transepithelial solute flux in some unidentified way and is not limited to a unique effect on the water conduits. Consistent with this, clearly, cells in which AQP1 is diminished or inhibited show a functional deficit some time after mounting. In going from the incubator to the chamber, the surrounding fluid is the same but the lower chamber cannot be gassed with air-CO2, and its pH increases with time. There are reports that AQP1 serves as a conduit for CO2 gas (5, 25, 33), although there is also evidence against that possibility (44). Hence, although the precise reason remains unclear, lack of proper water or gas permeation might conceivably underlie the functional deficit seen in our case when AQP1 channels are inhibited or diminished in number.

As mentioned earlier in this paper, the rate of fluid movement observed was 3.6 ± 0.3 µl · h-1 · cm-2, which is of the order of those in other fluid-transporting layers (8, 26). The ciliary epithelium comprises ~70 major ciliary processes that are ~2 mm long and 0.5 mm wide (23), which results in, roughly, a 7-cm2 surface area for the folded ciliary epithelium. If a factor of 6.7 is used, as described by Krupin et al. (20, 21), to relate the real area to the projected area for unfolded rabbit ciliary epithelium, the 7-cm2 surface area will result in a 46.9-cm2 surface area. Because the height of the ciliary processes is not constant, the real area for unfolded ciliary epithelium is an estimate. If such a surface area is used, the rate of fluid transport that we observe yields a putative rate of aqueous humor secretion of 2.8 µl/min-1, which is in agreement with the in vivo values reported previously in the literature (19, 20).

Our observation of fluid transport by NPE alone leads naturally to a discussion of the presumed role of the gap junctions known to exist between PE and NPE. Their role and the modulation of their open and closed states is unclear, but they have been mentioned in the literature as a possible route for fluid movement from PE to NPE working in tandem (20). For example, Walker et al. (40) recently observed sequential cell regulatory volume changes first in PE and then in NPE, consistent with cyclic movement of fluid from PE to NPE. From their observations, one might argue that fluid could have crossed from PE to NPE across timely opening gap junctions. However, in our own results, NPE cells transport fluid without PE present. Our findings, therefore, pose the questions of whether the tandem model can explain the results in vivo or whether an alternative explanation needs to be sought. In this connection, we note that fluid production by the ciliary epithelium is considerable. Given a ciliary epithelial volume of 8 µl in vivo (20) and a rate of secretion of aqueous humor of 3-4 µl/min, the ciliary epithelium transports its own volume approximately every 2.5 to 3 min (20). Given our current observation that the NPE layer by itself transports fluid, it might be argued that transport by the PE and NPE layers juxtaposed could add up, as it would with two impellent pumps in series. This, of course, would require the PE to be able to transport fluid on its own, the capability of which is unknown.

In summary, we present evidence that a human cultured NPE layer actively transports a sizable amount of fluid from its apical to its basal side and that AQP1 has a role to allow these cells an optimal rate of transport. To be noted, there are aquaporins in all the tissues involved in the major route of aqueous humor flow. The possibility exists that NPE in vivo could transport fluid on its own, and improvements in our understanding of the role of aquaporins in connection with this transport could provide crucial new insights into physiological and disease mechanisms. Our results also indicate that cultures of the individual cell layers, like the one utilized here, could constitute useful in vitro models to study the mechanisms of aqueous humor secretion.


    ACKNOWLEDGEMENTS

We thank Drs. Kunyan Kuang and Quan Wen for technical expertise.


    FOOTNOTES

This work was supported by National Eye Institute Grants EY-10423 (to R. V. Patil), EY-06178 (to J. Fischbarg), Core Grant EY-02687, and, in part, by unrestricted grants from Research to Prevent Blindness, Inc. R. V. Patil is a Research to Prevent Blindness Olga Keith Wiess Scholar.

Address for reprint requests and other correspondence: R. Patil, Dept. of Ophthalmology and Visual Sciences, Washington Univ. School of Medicine, 660 S. Euclid, St. Louis, MO 63110 (E-mail: patil{at}vision.wustl.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.

Received 22 May 2000; accepted in final form 16 May 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Am J Physiol Cell Physiol 281(4):C1139-C1145
0363-6143/01 $5.00 Copyright © 2001 the American Physiological Society




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