IOVS
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

(Investigative Ophthalmology and Visual Science. 2004;45:4423-4432.)
© 2004 by The Association for Research in Vision and Ophthalmology, Inc.
DOI:  10.1167/iovs.04-0816
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (9)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Levin, M. H.
Right arrow Articles by Verkman, A. S.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Levin, M. H.
Right arrow Articles by Verkman, A. S.

Aquaporin-Dependent Water Permeation at the Mouse Ocular Surface: In Vivo Microfluorimetric Measurements in Cornea and Conjunctiva

Marc H. Levin and A. S. Verkman

From the Departments of Medicine and Physiology, Cardiovascular Research Institute, University of California San Francisco, San Francisco, California.


   Abstract
Top
 Abstract
Methods
 Results
 Discussion
 Appendix
 References
 
PURPOSE. Fluorescence methods were developed to quantify membrane and tissue water permeabilities at the ocular surface and to compare water transport in wild-type mice versus transgenic mice lacking each of the water channels, aquaporin (AQP)-1, -3, and -5, normally expressed in cornea or conjunctiva.

METHODS. Membrane water permeabilities (Pfmem) of calcein-stained surface epithelial cells were measured from the kinetics of fluorescence quenching in response to rapid (<0.2 seconds) changes in extraocular fluid osmolarity. Tissue water permeabilities (Pftiss) across intact cornea and conjunctiva—the relevant parameters describing water movement into the hyperosmolar tear film in vivo—were determined by a dye-dilution method from the fluorescence of Texas red-dextran in an anisosmolar solution in a microchamber at the ocular surface.

RESULTS. Osmotic equilibration occurred with an exponential time constant ({tau}) of 1.3 ± 0.2 seconds (Pfmem = 0.045 cm/s) in calcein-loaded corneal epithelial cells of wild-type mice, slowing 2.1 ± 0.4-fold in AQP5-deficient mice; {tau} was 2.4 ± 0.1 seconds in conjunctiva (Pfmem = 0.025 cm/s), slowing 3.6 ± 0.7-fold in AQP3-deficient mice. In dye-dilution experiments, Pftiss of cornea was 0.0017 cm/s and decreased by greater than fivefold in AQP5-deficient mice. Pftiss in AQP5-null mice was restored to 0.0015 cm/s after removal of the epithelium. Pftiss of conjunctiva was 0.0011 cm/s and was not sensitive to AQP3 deletion.

CONCLUSIONS. These results define for the first time the water-transporting properties of the two principal ocular surface barriers in vivo. The permeability data were incorporated into a mathematical model of tear film osmolarity, providing insights into the pathophysiology of dry eye disorders.

The ocular surface consists of the cornea and conjunctiva, which make contact with the tear film. The corneal stroma is covered externally by a stratified squamous epithelium and internally by a single-layered endothelium. Maintenance of stromal transparency requires precise regulation of extracellular water content. The conjunctiva, which covers more of the ocular surface than the cornea (17 times more area in humans),1 consists of a superficial stratified epithelium interspersed with goblet cells and a deeper stroma with arterial and lymphatic networks. The palpebral and bulbar conjunctivae cover the posterior lid and the anterior scleral surfaces, respectively.

The ocular surface plays a role in regulation of tear film volume and composition. Near-isosmolar tear fluid is secreted actively by lacrimal and ocular surface tissues, dispersed by blinking, and drained through the nasolacrimal duct. Fluid secretion measurements across the cornea and conjunctiva in several species provide evidence that ocular surface epithelia contribute significantly to tear fluid under basal conditions and may be particularly important in lacrimal gland dysfunction.2 3 4 Indeed, new therapies for dry eye, or keratoconjunctivitis sicca (KCS), target signaling pathways involved in corneal and conjunctival chloride–driven fluid secretion.5 6 7

Ocular surface water transport plays an additional role in maintenance of tear volume and osmolarity by replacing evaporative water losses. Although under normal conditions, evaporation is retarded by a thin outer lipid tear layer secreted by meibomian glands,8 9 10 it is still responsible for at least 10% to 25% of tear turnover,11 driving transcellular osmotic water transport across ocular surface tissues. Evaporative water loss is an important determinant of tear film ionic content and is primarily responsible for the elevated tear film osmolarity and ocular surface changes common to all forms of dry eye disease.12 Transepithelial osmosis is also an important determinant of corneal hydration. Hypoxia from contact lens wear generates lactate osmoles in the cornea,13 inducing stromal swelling, which is reversed mainly by evaporation from tears rather than endothelium pump function.14 Transepithelial osmosis is responsible for the diurnal 4% thinning of corneas during the first few waking hours of each day.15 Though these phenomena are well described, little information is available about water permeability properties of ocular surface tissues, particularly in vivo. High water permeability at the ocular surface is thought to facilitate fluid secretion into the tear film in response to active ion secretion and to minimize tear film hyperosmolarity during evaporation.

Aquaporin (AQP)-type water channels make up a family of small transmembrane proteins that facilitate osmotically driven water transport across cell plasma membranes, providing the major molecular pathway for water movement. Three AQP proteins are strongly expressed at plasma membranes in cornea and conjunctiva, including AQP1 in corneal endothelium and keratocytes, AQP5 in corneal epithelium, and AQP3 in conjunctival epithelium.16 17 18 19 Transgenic knockout mice have been informative in elucidating the role of AQPs in mammalian physiology, manifesting a variety of phenotypes such as defective urinary concentrating ability in mice lacking AQP1 or AQP3,20 21 altered cerebral fluid balance and neural signal transduction in mice lacking AQP4,22 23 and defective salivary and submucosal gland secretion in mice lacking AQP5.24 25 In the eye, mice lacking AQP1 and AQP4 (expressed in ciliary epithelium) have reduced intraocular pressure and aqueous fluid production,26 mice lacking AQP4 (expressed in retinal Müller cells) have reduced flash-induced retinal potentials,27 and mice lacking AQP1 show delayed restoration of corneal transparency after experimental swelling.28 The presence of AQPs in ocular surface tissues suggests that they have a role in fluid transport and tear film homeostasis.

The goals of this study were to measure osmotic water permeability of the two principal ocular surface barriers—the cornea and conjunctiva—in mice in vivo and to investigate the role of AQPs as the molecular pathway of water permeation. Mice were chosen because of the availability of transgenic mice lacking each of the major ocular surface AQPs, permitting quantitative analysis of the role of each AQP by comparative permeability measurements in wild-type versus AQP knockout mice. Measurements were made in vivo, with vascular supply intact, to minimize concerns about altered tissue architecture, regulatory mechanisms, and AQP expression in isolated or cultured ocular tissues. In addition, in vivo measurements have direct relevance in relating tear-stromal osmotic driving forces to induced water flow into the tear film layer. Novel microfluorimetric methods were developed to measure water permeability in ocular surface cells and intact ocular tissues. Several experimental challenges required the design of custom microperfusion chambers for in vivo optical measurements, including the rapid osmotic equilibration of surface cells, slow net water transport across intact ocular surfaces, and physical constraints imposed by the mouse orbit. The new methods were applied to characterize water permeation mechanisms and the roles of AQPs in cornea and conjunctiva. The results were incorporated into a quantitative model of tear film osmolarity with relevance to clinical dry eye syndromes.


   Methods
Top
 Abstract
 Methods
Results
 Discussion
 Appendix
 References
 
Transgenic Mice
Transgenic mice deficient in AQP1, -3, and -5 in a CD1 genetic background were generated by targeted gene disruption, as described previously.20 21 24 Wild-type and knockout mice were matched by age and weight (ages 6–8 weeks, 22–25 g). Investigators were blinded to mouse genotype in all functional studies until completion of data analysis. Protocols were approved by the University of California at San Francisco Committee on Animal Research and are in compliance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research.

Fluorescence Microscopy
Fluorescence was measured with a stereo epifluorescence microscope (SMZ1500; Nikon, Tokyo, Japan) equipped with a 1.6x objective lens (working distance, 24 mm; numerical aperture, 0.21), adjustable zoom set to 6x, and custom filter sets for Texas red and FITC (Chroma, Rockingham, VT). The illumination source was a 100-W tungsten-halogen lamp powered by a stabilized direct current source. Fluorescence was detected with a 14-dynode photomultiplier, amplifier, and analog-to-digital converter, and recorded with custom-written software (written in LabView; National Instruments, Austin, TX). In some studies, fluorescence was imaged with a cooled charge-coupled device (CCD) camera (CoolSnap HQ; Photometrics, Tucson, AZ).

Ocular Surface Perfusion
Two stainless-steel perfusion microchambers (volumes, ~4 and 33 µL) were constructed to measure cell and tissue water permeabilities at the ocular surface in vivo. Scaled drawings with chamber dimensions are provided in Figure 1A . The upper (lens-facing) surfaces were bounded by sapphire windows (Edmund Optics; Barrington, NJ) fixed to the chambers with optical cement. The bottom (ocular surface-facing) surfaces were beveled to the contour of the mouse globe (~3.2-mm diameter sphere). The 2-mm diameter bottom dimension was chosen to isolate corneal tissue from the limbus and conjunctiva. The same beveled contact surface achieved an airtight seal with the conjunctiva. Inlets and outlets were made from 21-gauge steel needles. Solutions were perfused using PE-90 tubing and a gravity pinch valve system (ALA Scientific Instruments, Westbury, NY). Chambers were positioned with an x-y-z micromanipulator (World Precision Instruments, Sarasota, FL) as depicted in Figure 1B .



View larger version (56K):
[in this window]
[in a new window]
  		FIGURE 1. Methods for ocular surface perfusion. (A) Microchambers for measurement of rapid changes in ocular surface cell volume (top) and steady state water flux (bottom) across the cornea and conjunctiva. Chambers are drawn to scale. (B) Schematic of experimental setup for measurement of cell volume changes with the 33-µL chamber positioned on the mouse cornea by using a three-axis micromanipulator. Optical elements for fluorescence measurements are depicted, along with the perfusion system and stereotaxic platform. (C) Photographs of the 33-µL chamber in contact with the cornea of a proptosed globe (left), the 4-µL chamber forming a seal with the exposed conjunctival surface (middle), and an area of conjunctiva exposed by retracting the upper lid and depressing the globe into the orbit (right).

 
Mouse Preparation
Mice were anesthetized intraperitoneally with ketamine (40 mg/kg) and xylazine (20 mg/kg). Corneas were treated with topical proparacaine (0.5%) and kept hydrated with PBS supplemented with NaCl to 320 mOsM to match mouse serum osmolarity. Mice were immobilized for optical measurements with a custom-built stereotaxic device with a rotating jaw clamp, and the eye under study facing upward (Fig. 1C) . Body temperature was maintained at 37 ± 1°C with a heating pad and rectal temperature probe. A wire clip applied gentle upward pressure to present the globe to the chamber for corneal permeability measurements without inducing trauma or corneal edema. In some experiments, much of the corneal epithelium was mechanically removed without trauma to the stroma using a Beaver blade (BD Biosciences, Mountain View, CA) and a standard scraping procedure.29 For measurements on conjunctiva, a flat area of tissue consisting of portions of the upper tarsus and fornix was exposed with a pair of hook retractors threaded through the upper eyelid with sutures and fixed to posts with elastic bands. The globe was depressed into the orbit with surgical sponges (Medtronics, Jacksonville, FL; Fig. 1C ).

Cell Membrane Water Permeability In Vivo
A calcein-quenching method developed previously for water permeability measurement in brain astrocyte cell cultures30 was adapted to the mouse eye. Surface epithelial cells were loaded with calcein by exposure of the cornea or conjunctiva for 30 minutes to 25 µL PBS containing 10 µM calcein-AM (Molecular Probes, Eugene, OR). The solution remained in place by surface tension, and, if necessary, solution was added during the incubation. After 30 minutes, the loading solution was rinsed from the eye surface with PBS. The 33-µL microchamber was then positioned over the cornea or conjunctiva for continuous measurement of cell calcein fluorescence during perfusion with solutions of specified osmolarities. Complete solution exchange time was <50 ms at a perfusion rate of 55 mL/min achieved with the outlet under vacuum (Fig. 3A , bottom). The chamber was designed with its inlet close to the ocular surface, to establish a flow pattern with even faster exchange at the surface of the eye. For water permeability measurements, superficial cell loss was minimized by slowing perfusion approximately threefold, with an adjustable valve on the outlet tubing. Solutions were exchanged between PBS (290 mOsM) and hypoosmolar (145 mOsM, PBS diluted with distilled water) or hyperosmolar (590 mOsM, PBS with added D-mannitol) saline. Solution osmolarities were measured with a freezing point–depression osmometer (Precision Systems; Natick, MA). The time course of fluorescence in response to solution osmolarity changes, F(t), was fitted to a single exponential time constant, {tau}: F(t) = A + Be–t/{tau}, where A and B are related to system sensitivity and background signal. Pfmem was computed with a mathematical model of osmosis through a multilayer tissue, as described in the Appendix .



View larger version (24K):
[in this window]
[in a new window]
  		FIGURE 3. AQP-dependent water permeability in ocular surface cells measured by a calcein fluorescence-quenching method. (A) Top: fluorescence micrograph of calcein-stained corneal epithelial cells. Bottom: solution exchange time measurement in 33-µL chamber perfused with saline and fluorescein-containing saline. (B) Representative calcein fluorescence in cornea of wild-type mouse showing changes in response to hypoosmolar (top) and hyperosmolar (bottom) challenge. (C) Time course of corneal (left) and conjunctival (right) calcein fluorescence in response to hypoosmolar osmotic gradient in wild-type and indicated AQP-deficient mice. (D) Summary of exponential time constants for swelling response. Each point represents average data for 8 to 15 curves in individual mice, with means ± SE for each genotype shown. * P < 0.05, ** P < 0.01.

 


View larger version (25K):
[in this window]
[in a new window]
  		FIGURE 7. Model for computation of corneal epithelial cell plasma membrane water permeability from calcein fluorescence quenching experiments. (A) Schematic of multilayered corneal epithelium. An osmotic gradient is imposed at the most superficial cell layer (apical osmolarity, {Phi}0). Depicted are layer thicknesses, hi(t); osmolarities, {Phi}i(t); and interlayer water fluxes Ji. Tissue osmolarity, {Phi}6, is fixed at serum osmolarity. Pf represents the intrinsic osmotic water permeability of the epithelial cell plasma membrane (Pfmem), assumed to be the same in each layer. (B) Time course of relative corneal epithelial cell thicknesses, hi(t)/hi(0), in response to a sudden decrease in {Phi}0 from 300 to 150 mOsM. Parameters: Pfmem = 0.045 cm/s, {Phi}i(0) = 300 mOsM, h1(0) = 2 µm, h2(0) = 3 µm, h3(0) = 7 µm, h4(0) = 7 µm, h5(0) = 21 µm. Experimental data are shown overlying h1(t). The same model was implemented for the thinner conjunctival epithelium, except that four layers were included, with h1(0) = 2 µm, h2(0) = 3 µm, h3(0) = 5 µm, h4(0) = 10 µm, giving a Pfmem of 0.025 cm/s for conjunctiva of wild-type mice (not shown).

 
Transcorneal and Transconjunctival Water Permeabilities In Vivo
Water permeability across intact cornea and conjunctiva was measured by a dye-dilution method in which a cell-impermeant, photostable dye (Texas red-dextran, 3 kDa; Molecular Probes) was used as an inert marker of water flux. Texas red-dextran (0.05 mg/mL) was dissolved in hypoosmolar (~150 mOsM), isosmolar (~310 mOsM), or hyperosmolar (~580 mOsM) saline and infused into the microchamber positioned over the cornea or conjunctiva. Flow was stopped after solution exchange, and Texas red fluorescence was monitored continuously over 4 minutes. In some experiments, the solution was supplemented with 10% (wt/vol) 500-kDa dextran to increase viscosity 10-fold. Solute-free water movement across the corneal surface produced linear changes in Texas-red concentration and measured fluorescence. Water flux, Jv (in cubic centimeters per second), was computed from the product of chamber volume (Vc), and the rate of fluorescence change (after background subtraction), d(F/F0)/dt. Vc was determined to be 3.8 µL when in contact with the curved corneal surface and 5.4 µL when in contact with the flat conjunctiva. With the assumption that unstirred layer effects are negligible, the osmotic water permeability coefficient, Pftiss (in centimeters per second), is defined from the relation: Jv = PftissSvw ({Phi}1 – {Phi}2), where S is the tissue surface area assuming a smooth surface, vw is the partial molar volume of water (18 cm3/mol), and ({Phi}1 – {Phi}2) is the osmotic gradient, giving: Pftiss = Vc [d(F/F0)/dt]/[Svw({Phi}1 – {Phi}2)].

Corneal Thickness Measurement by Confocal Microscopy
Corneal thickness changes were measured after removal of the epithelium and exposure to solutions of various osmolarities, according to a method described previously.28 After removal of the epithelium, corneas in living mice were exposed to hypoosmolar, isosmolar, or hyperosmolar solutions for specified times. Corneas were dried rapidly by blotting, and thickness was determined by scanning bright-field confocal microscopy, with an upright Nipkow wheel-type confocal microscope with a confocal/coaxial module (Technical Instrument Co., San Francisco, CA) and 20x air objective (working distance, 20.5 mm; numerical aperture, 0.35; Nikon). Images were acquired in the reflectance mode every 5 µm with a cooled CCD camera.

Immunocytochemistry
After mouse sacrifice, globes were enucleated and embedded in Tissue-Tek OCT compound, and 7-µm cryostat sections were cut. Immunocytochemistry was performed on acetone-fixed sections by using polyclonal anti-AQP1, -3, and -5 antibodies (Chemicon, Temecula, CA) and secondary Cy3-conjugated anti-rabbit IgG (Sigma-Aldrich, St. Louis, MO). Fluorescence micrographs were obtained using a Leica upright fluorescence microscope with 20x air objective and 3-color cooled Spot CCD camera (Diagnostic Instruments, Sterling Heights, MI).


   Results
Top
 Abstract
 Methods
 Results
Discussion
 Appendix
 References
 
AQP Expression in Ocular Surface Tissues in Mice
Immunocytochemistry showed AQP1 in corneal endothelium and stromal keratocytes, and AQP5 in corneal epithelium, with no specific staining in corresponding knockout mice (Fig. 2) . AQP3 was expressed in conjunctival and corneal epithelia. RT-PCR analysis of whole mouse corneas with AQP-specific primers confirmed expression of transcripts encoding AQP1, -3, and -5, with no detectable transcripts encoding AQP0, -2, -4, -6, -7, -8, and -9 (not shown).



View larger version (39K):
[in this window]
[in a new window]
  		FIGURE 2. AQP expression at the mouse ocular surface. Immunofluorescence of cornea (top) and bulbar conjunctiva (bottom) from wild-type (left) and indicated AQP-null (right) mice, stained for AQPs 1, 3, or 5.

 
Water Permeability of Cells Lining the Ocular Surface
Measurement of cell membrane water permeability involved loading ocular surface cells of anesthetized mice with a membrane-permeant, nonfluorescent calcein derivative, which when de-esterified in the cytoplasm becomes fluorescent and membrane impermeant. Figure 3A (top) shows calcein staining of most superficial epithelial cells across the entire cornea, as visualized by epifluorescence stereomicroscopy through the 33-µL microperfusion chamber (Fig. 1A , top). Initial calcein-quenching experiments at the maximum perfusion rate indicated osmotic equilibration in ~1 to 2 seconds in cornea and conjunctiva of wild-type mice. Because cell exfoliation often occurred, in most studies flow was slowed approximately threefold (exchange time, <150 ms) to minimize cell loss. A slight downward force was applied to the chamber with a three-axis micromanipulator to achieve an airtight seal over corneal and conjunctival tissues.

A calcein fluorescence-quenching method was adapted to the mouse eye to compare in vivo osmotic water permeability in wild-type versus AQP-deficient ocular surface cells. As found in other cell types,30 calcein fluorescence in ocular surface cells was sensitive to the size and direction of the imposed osmotic gradient (Fig. 3B) . Nearly twofold changes in relative superficial cell volume produced by changing perfusate osmolarity from 290 mOsM to 145 or 590 mOsM yielded a 5% to 8% change in fluorescence signal. Signal changes were independent of magnification, as expected with a cytoplasmic quenching mechanism (not shown). Figure 3C shows representative kinetics of reversible corneal (left) and conjunctival (right) cell swelling in response to serial perfusion with solutions of osmolarities 290, 145, and 290 mOsM. AQP5 deletion in cornea slowed osmotic equilibration, as did AQP3 deletion in conjunctiva. Figure 3D summarizes exponential time constants ({tau}) for a series of mice, which are inversely proportional to swelling rates. Osmotic equilibration was slowed 2.1 ± 0.4-fold in AQP5-deficient corneal epithelium (P < 0.05) and 3.6 ± 0.7-fold in AQP3-deficient conjunctiva (P < 0.01). Osmotic equilibration was not slowed in corneas of AQP3-deficient mice ({tau} =1.2 ± 0.3 seconds, not shown in the Figure), in agreement with the qualitatively lower AQP3 than AQP5 protein expression in cornea. Also, corneal epithelial cell membrane water permeability in AQP1-deficient mice was similar to that in wild-type mice (Fig. 3D) , as expected, since AQP1 is expressed in corneal endothelium and keratocytes but not in corneal epithelium.

The osmotic water permeability coefficients (Pfmem) of corneal and conjunctival epithelial cell membranes were computed from the time course of fluorescence change in response to osmotic challenge, assuming multilayered epithelia containing five (cornea) or four (conjunctiva) cell layers of increasing thickness from superficial to basal layer, as estimated from histologic sections (see the Appendix ).31 Deduced Pfmem (in centimeters per second) were: 0.045 (wild-type) and 0.020 (AQP5-null) for cornea, and 0.025 (wild-type) and 0.007 (AQP3-null) for conjunctiva. The high Pfmem of corneal epithelium in wild-type mice may be important in facilitating rapid water transport across the multilayered corneal epithelium, which contains ~10 membranous barriers in series.

Water Permeability across Intact Cornea and Conjunctiva
A different fluorescence approach was developed to measure steady state, osmotically driven water transport across intact cornea and conjunctiva. A very small 4-µL chamber (Fig. 1A , bottom) was constructed to measure changes in dye concentration in anisosmolar solutions contacting ocular surface epithelia, where submicroliter net osmotic water fluxes are predicted over several minutes. Texas red-dextran (3 kDa) was found to be a useful volume marker because of its photostability, membrane impermeability, and, compared with other dyes tested, minimal binding to the microchamber and ocular surfaces. The experimental procedure involved measurement of dye fluorescence in isosmolar saline for 4 minutes to establish a baseline signal. In some experiments there was slow signal loss (generally <1%/min) over this time, possibly due to reflex tearing from eye irritation and/or dye binding to the chamber walls, which was corrected for in osmotic water permeability studies. The isosmolar solution was then changed to a hyperosmolar (~580 mOsM) or hypoosmolar (~150 mOsM) solution for 4 minutes, returned to the control isosmolar solution, and then changed to hypoosmolar or hyperosmolar solutions, respectively. Figure 4A shows approximately linear kinetics of the increase in fluorescence with hypoosmolar solutions (water entering tissue resulting in increased dye concentration) and decrease in fluorescence with hyperosmolar solutions (water exiting tissue resulting in dye dilution). The fluorescence signal changed by only a few percent over 4 minutes in the 4-µL chamber because of the small ocular surface area available for steady state osmosis. In control experiments, there was no appreciable signal change when the 33-µL chamber was used, as expected, because of the correspondingly smaller fractional changes in dye concentration caused by osmotic water transport. Figure 4A also shows reduced changes in fluorescence for water transport across corneas from AQP5- but not AQP1-null mice. Similar slopes were produced when conjunctivae of wild-type were compared with AQP3-null mice.



View larger version (21K):
[in this window]
[in a new window]
  		FIGURE 4. Osmotic water transport across intact cornea and conjunctiva. The ocular surface was covered by a 4-µL microchamber containing Texas-red dextran in hypo-, iso-, or hyperosmolar solutions. (A) Representative kinetics of fluorescence intensities for measurements on cornea (top) or conjunctiva (bottom) of mice of indicated genotype. Osmotic gradients induced water movement into (hyperosmolar) or out of (hypoosmolar) the chamber, producing dye dilution (decreasing fluorescence) or concentration (increasing fluorescence), respectively. (B) Averaged rates of relative fluorescence signal change, d(F/F0)/dt (±SE, 4–6 eyes per condition, multiple measurements done on each eye). Where indicated (open bars), the corneal epithelium was removed by scraping. (C) Whole-tissue osmotic water permeability coefficients Pftiss (±SE) computed from data in (B). *P < 0.05, **P < 0.01.

 
One concern in this approach was the potential for error in deduced Pftiss due to dye diffusion between the small chamber and the 21-gauge inlet and outlet ports over the 4-minute measurement time. To rule out significant diffusional effects, experiments were done in corneas of wild-type mice as in Figure 4A (top), except that the viscosity of bathing solutions was increased 10-fold by addition of an inert, high-molecular-weight dextran. There were no significant differences in water permeability with the viscous solutions (for example, d(F/F0)/dt x 10–4 s–1 = –1.7 ± 0.3 for nonviscous versus –1.4 ± 0.2 for viscous hyperosmolar solutions in same eyes), indicating the absence of significant diffusional effects.

Figure 4B summarizes averaged slopes for many experiments as in 4A, including data from corneas in which the epithelium was removed by a scraping procedure. Figure 4C summarizes the corresponding Pftiss values. AQP5 deletion reduced Pftiss significantly in intact cornea, as measured with both hypo- and hyperosmolar solutions. Removal of the epithelium had no significant effect on Pftiss in cornea from wild-type mice, though it restored Pftiss in cornea from AQP5-null mice to the level in wild-type mice. These results implicate the involvement of AQP5 in water movement across the intact corneal epithelium and show that the epithelium can be the rate-limiting barrier in osmosis across the intact cornea in the absence of AQP5. Corneal Pftiss was reduced to a lesser extent by AQP1 deletion and only for hypoosmolar challenge after removal of the epithelium. Pftiss was lower in conjunctiva than cornea. Despite the reduced water permeability of surface conjunctival cells in AQP3-null mice, as shown in Figure 3D , AQP3 deletion did not affect osmosis across intact conjunctiva, indicating that AQP3 is not the rate-limiting barrier for osmosis across intact conjunctiva.

Corneal Thickness Measurements
Measurements of corneal thickness were made after removal of the corneal epithelium by scraping to investigate possible differences in stromal properties in AQP deficiency and to determine whether an osmotically sensitive, semipermeable barrier remained on the stromal surface. Baseline and serial corneal thickness measurements were made by z-scanning bright-field confocal microscopy as described previously.28 Figure 5A shows an example of serial confocal images used to determine corneal thickness, with outer and inner corneal surfaces producing characteristic morphologic features. Figure 5A (bottom, right) summarizes baseline thicknesses for undisturbed corneas (in micrometers): 130 ± 1 (wild-type), 139 ± 2 (AQP5-null), and 100 ± 1 (AQP1-null), in agreement with previous results.



View larger version (51K):
[in this window]
[in a new window]
  		FIGURE 5. Corneal thickness changes after removal of the epithelium measured by z-scanning bright-field confocal microscopy. (A) Top: bright-field confocal images of cornea from wild-type mouse taken at various z-positions (a–e), where corneal epithelial and endothelial surfaces are identified by characteristic morphometric features as depicted (bottom, left). Averaged baseline corneal thickness with epithelium intact (bottom, right; ±SE, 10–12 eyes per group). **P < 0.01 compared with wild-type. Scale bar = 1 mm. (B) The time course of corneal thickness (Tc) after removal of the epithelium and exposure of stromal surface to either hypoosmolar (left), isosmolar (middle), or hyperosmolar (right) saline (±SE, 3–4 corneas per condition per genotype) *P < 0.05 compared with wild-type.

 
As expected, exposure of intact corneas to isosmolar solution produced no change in thickness (not shown). After removal of the epithelium, corneal thickness increased when the surface was bathed in hypoosmolar (150 mOsM) or isosmolar solutions, and decreased in hyperosmolar (580 mOsM) solution (Fig. 5B) . Swelling of de-epithelialized corneas in hypoosmolar solution was markedly greater in corneas from AQP1-null mice than in wild-type and AQP5-null mice. Swelling also occurred when de-epithelialized corneas were bathed in isosmolar solution, reflecting the imbibition pressure of the dehydrated stroma. For all mice, hyperosmolar challenge produced corneal thinning that was lower in absolute magnitude than was the swelling measured under hypoosmolar challenge. Hyperosmolar-induced corneal thinning was slowest in AQP1-deficient mice. In each case, results were similar for wild-type and AQP5-null mice, suggesting that AQP5 deletion does not affect stromal properties.

Model of Tear Film Osmolarity
Based on the ocular surface water permeabilities measured in this study, a simple model was constructed to predict tear film osmolarity under normal physiological conditions and for two traditionally distinct classes of dry eye syndrome—secretory dysfunction and excessive evaporation. As depicted in Figure 6 (top), tear fluid is generated by active secretion (Js) and osmotic flux (Jv), and removed by evaporation (Je) and drainage (Jd). In the steady state

(1)



View larger version (27K):
[in this window]
[in a new window]
  		FIGURE 6. Theoretical dependence of tear film osmolarity on rates of evaporation (Je) and tear fluid secretion (Js). Top: schematic of ocular surface geometry. Jv, osmotic volume flow across the corneal and conjunctival surfaces; Jd, tear fluid removal by nasolacrimal drainage; {Phi}t and {Phi}s, osmolarities of tear film and surface tissue, respectively. Bottom: {Phi}t – {Phi}s computed from the model described in the text (equation 3) . See text for explanations.

 
Near-isotonic fluid (serum osmolarity, {Phi}s) is actively secreted (Js) by various ocular tissues, including the main and accessory lacrimal glands, conjunctiva, and cornea. Evaporation from the air-exposed surface of the eye (primarily from cornea in mouse, with surface area Scornea ~0.09 cm2) concentrates the preocular film uniformly and rapidly. Osmosis (Jv) across the entire corneal and conjunctival surfaces (Scornea + Sconjunctiva) drives water into the tears, partially reducing tear film osmolarity ({Phi}t). Jv is defined by the relation

(2)

where, for simplicity, Pf denotes Pftiss. Pf (cornea) = 0.0017 cm/s, and Pf (conjunctiva) = 0.0011 cm/s as measured in this case, and Pf' represents a single weighted value of ocular surface whole-tissue permeability. Because nasolacrimal drainage is assumed to be the only route for solute removal, Jd = Js({Phi}s/{Phi}t). Combining this relation with equations 1 and 2 , and solving for {Phi}t

(3)

{Phi}t – {Phi}s is plotted in Figure 6 (bottom) over the range of Je measured in rabbits and humans at 30% to 40% humidity,32 and a range of plausible Js based on measured mouse tear production rates.33 {Phi}s for mice was taken as 320 mOsM and Sconjunctiva/Scornea as 5 based on microdissection measurements, somewhat lower than Sconjunctiva/Scornea of 17 in humans and 9 in rabbits.1 Figure 6 (bottom) shows that for a physiological evaporative rate of 1.5 x 10–6 g/cm2 per second and Js of 3 x 10–6 mL/s, tear film osmolarity is predicted to be mildly elevated over serum osmolarity ({Phi}t – {Phi}s is ~6.5 mOsM). If Js is zero at a normal evaporative rate, {Phi}t – {Phi}s increases to 12 mOsM. In contrast, if Je is elevated fourfold as in meibomian gland dysfunction,10 {Phi}t – {Phi}s becomes 26 mOsM. If Js is zero and Je is increased fourfold, {Phi}t – {Phi}s becomes 46 mOsM. The relationship between {Phi}t and Je and Js has relevance to understanding dry eye syndromes (see the Discussion section).


   Discussion
Top
 Abstract
 Methods
 Results
 Discussion
Appendix
 References
 
Novel microfluorimetric methods were applied to measure water permeability of the two principal ocular surface tissues, the cornea and conjunctiva, at the cell membrane and whole tissue levels. Measurements were made in living mice with selective AQP deletions to assess the contributions of AQP1, -3, and -5 to osmotic fluid transport across intact surface tissues. As discussed in the introduction, the motivation for this work was the recognition of the role of the ocular surface in tear film homeostasis and as a promising target for development of therapies for dry eye syndromes. Approximately one half of middle-aged women report dry eye symptoms, with KCS affecting 10% to 15% of the elderly population.32 Dry eye symptoms are also a common side effect of contact lens wear and laser refractive surgery in all age groups.

Water permeability of ocular surface cells was measured by a calcein fluorescence-quenching method, which is based on rapid changes in cytoplasmic calcein fluorescence in response to changes in concentration of cytoplasmic anionic proteins and hence to changes in cell volume.30 The wide-field fluorescence detection method used for calcein fluorescence measurement is insensitive to small pulsatile and perfusion-related eye movements, which precludes water permeability measurement by confocal detection methods. Osmotic equilibration in corneal and conjunctival epithelial cells was very fast in wild-type mice, ~1 to 2 seconds, requiring the development of an ocular surface perfusion chamber capable of fluid exchange on a much faster time scale. A model relating induced cell volume changes to osmotic water permeability (Pfmem) was developed for a stratified epithelium, in which the kinetics of thickness of calcein-containing surface cells was computed after a change in osmolarity of solution bathing the ocular surface. Pfmem is the single most informative parameter characterizing the water-transporting capacity of a membrane barrier, particularly in assessing the potential role of water transport by molecular pores. Mishima and Hedbys34 and Fischbarg and Motoreano35 estimated water permeability of ~0.01 cm/s in full thickness corneal epithelium from osmotically induced changes in corneal thickness in living rabbits and mounted tissues. This value is lower than the Pfmem of 0.045 cm/s determined in the present study for plasma membrane in mouse cornea, probably because of the presence of multiple barriers and possible unstirred layers in full-thickness rabbit corneal epithelium. A Pfmem of greater than ~0.01 cm/s provides evidence of the presence of molecular water channels.

Water permeability across the intact corneal and conjunctival barriers was measured by a steady state dye-dilution method in which the concentration of an inert, membrane-impermeant fluorescent dye was measured at the external ocular surface in response to a sustained osmotic gradient. The detection of appreciable changes in dye concentration required the design of a microchamber with <4-µL solution volume and the selection of a fluorescent dye with minimal surface binding and excellent photostability. Dye fluorescence signal changed by ~1%/min for a ~200 mOsM osmotic gradient in cornea and conjunctiva. The direction of signal changes depended on the direction of the osmotic gradient, as expected, with linearly decreasing signal for a hyperosmolar solution driving water into the external solution. The deduced Pftiss for cornea and conjunctiva was independent of the direction of the osmotic gradient, and substantially lower than Pfmem because of the multiple layers of epithelial cell membranes and other barriers in series.

Mechanical removal of the entire stratified epithelial layer of the cornea did not enhance osmotically driven water transport in corneas of wild-type mice. Of the corneal tissues, the epithelium has been considered rate limiting to water movement. However, in this study the normal epithelium did not impede osmotically induced transcorneal fluid movement in vivo. Indeed, the Pftiss of ~0.002 cm/s for whole cornea measured was approximately five times lower than water permeability of full-thickness corneal epithelium of ~0.01 cm/s, as mentioned earlier. Although the corneal epithelium in wild-type mice was not rate-limiting for transcorneal osmosis, reducing epithelial water permeability by AQP5 deletion caused a marked ~5-fold slowing of transcorneal water flux in intact cornea, which was restored by removal of the epithelium to the level in wild-type mice. These results indicate that AQP5 provides the principal route for osmotically driven water flux across the intact corneal epithelium and that most water moves from corneal stroma to the tear film layer by a transcellular route.

Osmotic water movement across intact corneas of AQP1-deficient mice was similar to that in wild-type mice under both hypoosmolar and hyperosmolar conditions, suggesting that AQP1 is not a rate-limiting barrier for transcorneal osmosis. Unexpectedly, there was apparent asymmetry in water transport in corneas of AQP1-null mice after removal of the epithelium, where transcorneal osmosis was reduced ~2.5-fold only after hypoosmolar challenge. Measurements of stromal thickness by bright-field confocal microscopy were obtained to investigate possible differences in stromal properties in corneas from AQP1-null mice that might account for this result. Thickness measurements indicated corneal swelling and thinning after exposure of the denuded corneal surface to hypoosmolar and hyperosmolar solutions, respectively. Because salt gradients are not thought to induce osmosis across exposed stromal tissue, it is likely that one or more membranes remained at the stromal surface after the denudation procedure. Grossly, corneas from AQP1-null mice scattered more light throughout the swelling process than did corneas from wild-type or AQP5-null mice. Both hypoosmolar and isosmolar swelling was increased in AQP1-null mice compared with wild-type and AQP5-null mice, which may be related to impaired endothelial fluid pump function and/or differential stromal properties of the relative thin corneas in AQP1-null mice. AQP1 thus plays an important role in the removal of excess fluid from the corneal stroma after experimental corneal edema, rather than in osmotically driven water movement from the aqueous to tear film compartments.

Immunostaining revealed AQP3 expression at the plasma membranes of conjunctival epithelia, in agreement with data in rat.18 Cell water permeability in AQP3-deficient mice was substantially reduced (3.6-fold) compared with that in wild-type mice, although osmotically induced water movement across the intact conjunctiva was not AQP3 dependent. Therefore, the AQP3-containing conjunctival epithelial cell layer is not the rate-limiting barrier for osmosis in full-thickness conjunctiva, and thus AQP3-facilitated water transport does not play a role in transconjunctival fluid movement. Water movement across complex tissues such as conjunctiva could be impeded by rate-limiting obstacles other than plasma membranes. A similar conclusion was reported for AQP3 in skin, which has a structure similar to that of conjunctiva and other mucous membranes, except for the presence in skin of a superficial watertight layer of stratum corneum consisting of lipidic, cornified cell envelopes. Although water permeability was reduced after AQP3 deletion in epidermal cells, measurements of skin hydration in response to altered rates of evaporative water loss indicated that the AQP3 water transport function was not responsible for the reduced hydration and other abnormalities in AQP3-deficient mice.36 37 Decreased epidermal and stratum corneum glycerol content was found in the AQP3-null mice, which, when normalized by systemic glycerol administration, resulted in correction of the skin phenotype abnormalities.38 It was concluded that the glycerol-, rather than the water-transporting function of AQP3, was responsible for the abnormal skin phenotype. As in skin, we speculate that AQP3-facilitated glycerol transport in conjunctiva and other mucous membranes plays a functional, but at present unknown, physiological role.

Osmotic water permeation across the corneal and conjunctival barriers is important for replacement of evaporative water loss in the tear film when the ocular surface is exposed to ambient humidity, and for preventing significant tear film hyperosmolarity. The model of tear film osmolarity presented in Figure 6 provides a quantitative prediction of tear fluid hyperosmolarity under physiological and pathologic conditions. Measurements on human and rabbit tear samples by freezing-point depression osmometry have shown normal tears to be mildly hyperosmolar ({Phi}t – {Phi}s ~10–14 mOsM),10 39 40 41 slightly higher than the ~7 mOsM predicted in our model for the mouse tear film. In our model, tear secretions are assumed to be isosmolar, whereas the lacrimal component of tear fluid secretions may be slightly hyperosmolar.42

Tear film osmolarity is increased in both evaporative (caused by excessive tear evaporation rates) and tear-deficient (caused by inadequate tear fluid secretion) dry eye. Tear fluid osmolarity has been established as an objective, quantitative index of KCS severity.39 Mathers et al.43 found both diminished tear flow and increased evaporative rates to correlate with osmolarity. The model presented in this study makes predictions for tear film osmolarity in evaporative and tear-deficient states of varying severities alone and in combination. The {Phi}t – {Phi}s of ~26 mOsM predicted for pure meibomian gland dysfunction recapitulates the {Phi}t – {Phi}s of 20 to 30 mOsM measured in a rabbit model.44 In pure tear deficiency, decreased tear turnover and consequent greater evaporation time accounts qualitatively for the hyperosmolarity in KCS, even at normal evaporation rate. However, this effect ({Phi}t – {Phi}s ~12 mOsM) is smaller than that measured in tear-deficient KCS ({Phi}t – {Phi}s >22 mOsM).41 45 Possible explanations for this quantitative difference include changes in tear film evaporation at low secretion rates (due to abnormal tear composition),41 46 and/or increased osmolarity of lacrimal secretions at very low secretion rates.42 However, the latter mechanism probably would not substantially increase tear film osmolarity because of the low lacrimal secretion rate compared with secretion from ocular surface tissues. The quantitative difference in predicted versus measured tear film osmolarity in tear-deficient dry eye syndrome emphasizes the need for further investigation into the complex interplay of aqueous and meibomian secretions in retarding tear film evaporation in dry eye syndromes.


   Appendix
Top
 Abstract
 Methods
 Results
 Discussion
 Appendix
References
 
Computation of Corneal Epithelial Pfmem
Plasma membrane permeability in corneal epithelial cells was computed from the kinetics of cell volume change in response to changing the osmolarity of fluid bathing the cornea according to the model shown in Figure A1 , panel A. An approximately linear dependence of calcein fluorescence on cell volume was assumed, as reported previously.30 Only the outermost corneal cell layer was loaded with calcein-AM under the conditions of our experiments as seen by confocal microscopy, consistent with ex vivo experiments using esterified dyes,47 48 dye injection experiments,49 and ultrastructural studies,50 providing evidence against gap junctional coupling among superficial cells and between superficial and deeper wing cell layers. Pf is defined, for simplicity, as the intrinsic plasma membrane water permeability (Pfmem), assumed to be the same throughout the corneal epithelium. For model computations, apparent water permeability at the outermost surface was taken as 2Pf, to account for microvillar surface convolutions51 52 and apparent water permeability of interfaces between two layers (containing two membrane barriers) was taken as Pf/2. Unstirred layer effects were considered insignificant.

Water flux (Ji) across each membrane barrier is: Ji = Pfi Svw({Phi}i+1 – {Phi}i), where dVi(t)/dt = Ji–1 – Ji. Because osmotic equilibration occurs much faster than ionic equilibration, the product of cell volume (Vi) and osmolarity ({Phi}i) remains constant over the time course of the measurement: {Phi}i(0)Vi(0) = {Phi}i(t)Vi(t). Combining these equations and expressing as the ratio of volume-to-surface area, hi(t)

(4)

The five coupled differential equations were numerically integrated by using the forward Euler method (time-step, 0.01 ms) to obtain hi(t). Computed hi(t)/hi(0) are shown in Figure 7B for the best fitted Pf, along with a comparison of an experimental curve with computed h1(t).


   Acknowledgements
 
The authors thank Liman Qian for mouse breeding and genotype analysis; Marios Papadopoulos, Yaunlin Song, and Tong Da for help with microsurgical technique; and Greg Friedland for assistance with computer modeling.


   Footnotes
 
Supported by National Eye Institute Grant EY13574, National Institute of Diabetes and Digestive and Kidney Diseases Grant DK35124, National Heart, Lung, and Blood Institute Grant HL59198, and Grant EB00415; and Drug Discovery and Research Development Program grants from the Cystic Fibrosis Foundation.

Submitted for publication July 12, 2004; revised August 25, 2004; accepted September 10, 2004.

Disclosure: M.H. Levin, None; A.S. Verkman, None

The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Corresponding author: Alan S. Verkman, 1246 Health Sciences East Tower, Cardiovascular Research Institute, University of California San Francisco, San Francisco, CA 94143-0521; verkman{at}itsa.ucsf.edu.


   References
Top
 Abstract
 Methods
 Results
 Discussion
 Appendix
 References
 

  1. Watsky MA, Jablonski MM, Edelhauser HF. Comparison of conjunctival and corneal surface area in rabbit and human. Curr Eye Res. 1988;7:483–486.[ISI][Medline][Order article via Infotrieve]
  2. Candia OA, Zamudio AD. Chloride-activated water permeability in the frog corneal epithelium. J Membr Biol. 1995;143:259–266.[ISI][Medline][Order article via Infotrieve]
  3. Murakami T, Fujihara T, Nakamura M, Nakata K. P2Y(2) receptor stimulation increases tear fluid secretion in rabbits. Curr Eye Res. 2000;21:782–787.[CrossRef][ISI][Medline][Order article via Infotrieve]
  4. Yang H, Reinach PS, Koniarek JP, Wang Z, Iserovich P, Fischbarg J. Fluid transport by cultured corneal epithelial cell layers. Br J Ophthalmol. 2000;84:199–204.[Abstract/Free Full Text]
  5. Li Y, Kuang K, Yerxa B, Wen Q, Rosskothen H, Fischbarg J. Rabbit conjunctival epithelium transports fluid and P2Y2(2) receptor agonists stimulate Cl– and fluid secretion. Am J Physiol. 2001;281:C595–C602.
  6. Murakami T, Fujihara T, Horibe Y, Nakamura M. Diquafosol elicits increases in net Cl– transport through P2Y2 receptor stimulation in rabbit conjunctiva. Ophthalmic Res. 2004;36:89–93.[CrossRef][ISI][Medline][Order article via Infotrieve]
  7. Fujihara T, Murakami T, Fujita H, Nakamura M, Nakata K. Improvement of corneal barrier function by the P2Y(2) agonist INS365 in a rat dry eye model. Invest Ophthalmol Vis Sci. 2001;42:96–100.[Abstract/Free Full Text]
  8. Mishima S, Maurice DM. The oily layer of the tear film and evaporation from the corneal surface. Exp Eye Res. 1961;1:39–45.[ISI][Medline][Order article via Infotrieve]
  9. Iwata S, Lemp M, Holly FJ, Dohlman CH. Evaporation rate of water from the precorneal tear film and cornea in the rabbit. Invest Ophthalmol. 1969;8:613–619.[Abstract/Free Full Text]
  10. Mathers WD. Ocular evaporation in meibomian gland dysfunction and dry eye. Ophthalmol. 1993;100:347–351.
  11. Mishima S. Some physiologic aspects of the precorneal tear film. Arch Ophthalmol. 1965;73:233–241.
  12. Lemp MA. Report of the National Eye Institute/Industry Workshop on clinical trials for dry eyes. CLAO J. 1995;21:221–232.[Medline][Order article via Infotrieve]
  13. Klyce SD. Stromal lactate accumulation can account for corneal oedema osmotically following epithelial hypoxia in the rabbit. J Physiol. 1981;321:49–64.[Abstract/Free Full Text]
  14. O’Neal MR, Polse KA. In vivo assessment of mechanisms controlling corneal hydration. Invest Ophthalmol Vis Sci. 1985;26:849–856.[Abstract/Free Full Text]
  15. Mandell RB, Fatt I. Thinning of the human cornea on awakening. Nature. 1965;208:292–293.[CrossRef][Medline][Order article via Infotrieve]
  16. Frigeri A, Gropper M, Turck CW, Verkman AS. Immunolocalization of the mercurial-insensitive water channel and glycerol intrinsic protein in epithelial cell plasma membranes. Proc Natl Acad Sci USA. 1995;92:4328–4331.[Abstract/Free Full Text]
  17. Patil RV, Saito I, Yang X, Wax MB. Expression of aquaporins in the rat ocular tissue. Exp Eye Res. 1997;64:203–209.[CrossRef][ISI][Medline][Order article via Infotrieve]
  18. Hamman S, Zeuthen T, La Cour M, et al. Aquaporins in complex tissues: distribution of aquaporins 1–5 in human and rat eye. Am J Physiol. 1998;274:C1332–C1345.
  19. Wen Q, Dieche FPJ, Iserovich P, Kuang K, Sparrow J, Fischbarg J. Immunocytochemical localization of aquaporin-1 in bovine corneal endothelial cells and keratocytes. Exp Biol Med (Maywood). 2001;226:463–467.[Abstract/Free Full Text]
  20. Ma T, Song Y, Yang B, et al. Nephrogenic diabetes insipidus in mice deficient in aquaporin-3 water channels. Proc Natl Acad Sci USA. 2000;97:4386–4391.[Abstract/Free Full Text]
  21. Ma T, Yang B, Gillespie A, Carlson EJ, Epstein CJ, Verkman AS. Severely impaired urinary concentrating ability in transgenic mice lacing aquaporin-1 water channels. J Biol Chem. 1998;273:4296–4299.[Abstract/Free Full Text]
  22. Manley GT, Fujimura M, Ma T, et al. Aquaporin-4 deletion in mice reduces brain edema following acute water intoxication and ischemic stroke. Nat Med. 2000;6:159–163.[CrossRef][ISI][Medline][Order article via Infotrieve]
  23. Li J, Verkman AS. Impaired hearing in mice lacking aquaporin-4 water channels. J Biol Chem. 2001;276:31233–31237.[Abstract/Free Full Text]
  24. Ma T, Song Y, Gillespie A, Carlson EJ, Epstein CJ, Verkman AS. Defective secretion of saliva in transgenic mice lacking aquaporin-5 water channels. J Biol Chem. 1999;274:20071–20074.[Abstract/Free Full Text]
  25. Song Y, Verkman AS. Aquaporin-5 dependent fluid secretion in airway submucosal glands. J Biol Chem. 2001;276:41288–41292.[Abstract/Free Full Text]
  26. Zhang D, Vetrivel L, Verkman AS. Aquaporin deletion in mice reduces intraocular pressure and aqueous fluid production. J Gen Physiol. 2002;119:561–569.[Abstract/Free Full Text]
  27. Li J, Patil RV, Verkman AS. Mildly abnormal retinal function in transgenic mice without Müller cell aquaporin-4 water channels. Invest Ophthalmol Vis Sci. 2002;43:573–579.[Abstract/Free Full Text]
  28. Thiagarajah JR, Verkman AS. Aquaporin deletion in mice reduces corneal water permeability and delays restoration of transparency after swelling. J Biol Chem. 2002;277:19139–19144.[Abstract/Free Full Text]
  29. Kao WW, Kao CW, Kaufman AH, et al. Healing of corneal epithelial defects in plasminogen- and fibrinogen-deficient mice. Invest Ophthalmol Vis Sci. 1999;39:502–508.
  30. Solenov E, Watanabe H, Manley GT, Verkman AS. Sevenfold-reduced osmotic water permeability in primary astrocyte culture from AQP-4-deficient mice, measured by a fluorescence quenching method. Am J Physiol. 2004;286:C426–C432.
  31. Smith RS, Sundberg JP, John SWM. The anterior segment and ocular adnexae. Smith RS eds. Systemetic Evaluation of the Mouse Eye: Anatomy, Pathology, and Biomethods. 2002;3–23. CRC Press Boca Raton, FL.
  32. Mathers WD. Evaporation from the ocular surface. Exp Eye Res. 2004;78:389–394.[CrossRef][ISI][Medline][Order article via Infotrieve]
  33. Moore M, Ma T, Yang B, Verkman AS. Tear secretion by lacrimal glands in transgenic mice lacking water channels AQP1, AQP3, AQP4, and AQP5. Exp Eye Res. 2000;70:557–562.[CrossRef][ISI][Medline][Order article via Infotrieve]
  34. Mishima S, Hedbys BO. The permeability of the corneal epithelium and endothelium to water. Exp Eye Res. 1967;6:10–32.[CrossRef][ISI][Medline][Order article via Infotrieve]
  35. Fischbarg J, Motoreano R. Osmotic permeabilities across corneal endothelium and antidiuretic hormone-stimulated toad urinary bladder structure. Biochim Biophys Acta. 1982;690:207–214.[Medline][Order article via Infotrieve]
  36. Ma T, Hara M, Sougrat R, Verbavatz JM, Verkman AS. Impaired stratum corneum hydration in mice lacking epidermal water channel aquaporin-3. J Biol Chem. 2002;277:17147–17153.[Abstract/Free Full Text]
  37. Hara M, Ma T, Verkman AS. Selectively reduced glycerol in skin of aquaporin-3-deficient mice may account for impaired skin hydration, elasticity, and barrier recovery. J Biol Chem. 2002;277:46616–46621.[Abstract/Free Full Text]
  38. Hara M, Verkman AS. Glycerol replacement corrects defective skin hydration, elasticity, and barrier function in aquaporin-3-deficient mice. Proc Natl Acad Sci USA. 2003;100:7360–7365.[Abstract/Free Full Text]
  39. Gilbard JP, Farris RL, Santamaria J. Osmolarity of tear microvolumes in keratoconjunctivitis sicca. Arch Ophthalmol. 1978;96:677–681.[Abstract]
  40. Gilbard JP, Rossi SR, Gray KL. A new rabbit model for keratoconjunctivitis sicca. Invest Ophthalmol Vis Sci. 1987;28:225–228.[Abstract/Free Full Text]
  41. Craig JP, Signh I, Tomlinson A, Morgan PB, Efron N. The role of tear physiology in ocular surface temperature. Eye. 2000;14:635–641.
  42. Gilbard JP, Dartt DA. Changes in rabbit lacrimal gland fluid osmolarity with flow rate. Invest Ophthalmol Vis Sci. 1982;23:804–806.[Abstract/Free Full Text]
  43. Mathers WD, Lane JA, Sutphin JE, Zimmerman MB. Model for ocular tear film function. Cornea. 1996;15:110–119.[ISI][Medline][Order article via Infotrieve]
  44. Gilbard JP, Rossi SR, Gray KL. Tear film and ocular surface changes after closure of the Meibomian gland orifices in the rabbit. Ophthalmology. 1989;96:1180–1186.[ISI][Medline][Order article via Infotrieve]
  45. Gilbard JP, Gray KL, Rossi SR. A proposed mechanism for increased tear-film osmolarity in contact lens wearers. Am J Ophthalmol. 1986;102:505–507.[ISI][Medline][Order article via Infotrieve]
  46. Rolando M, Refojo MF, Kenyon KR. Increased tear evaporation in eyes with keratoconjunctivitis sicca. Arch Ophthalmol. 1983;101:557–558.[Abstract]
  47. Wolosin JM. Gap junctions in rabbit corneal epithelium: limited permeability and inhibition by cAMP. Am J Physiol. 1991;261:C857–C864.
  48. Jester JV, Li H, Petroll WM, R, et al. Area and depth of surfactant-induced corneal injury correlates with cell death. Invest Ophthalmol Vis Sci. 1998;39:922–936.[Abstract/Free Full Text]
  49. Williams KK, Watsky MA. Dye spread through gap junctions in the corneal epithelium of the rabbit. Curr Eye Res. 1997;16:445–452.[CrossRef][ISI][Medline][Order article via Infotrieve]
  50. McLaughlin BJ, Caldwell RB, Sasaki Y, Wood TO. Freeze-fracture quantitative comparison of rabbit corneal epithelial and endothelial membranes. Curr Eye Res. 1985;9:951–961.
  51. Nichols B, Dawson CR, Togni B. Surface features of the conjunctiva and cornea. Invest Ophthalmol Vis Sci. 1983;24:570–576.[Abstract/Free Full Text]
  52. Gipson IK. Distribution of mucins at the ocular surface. Exp Eye Res. 2004;78:379–388.[CrossRef][ISI][Medline][Order article via Infotrieve]



This article has been cited by other articles:


Home page
 IOVSHome page
S. K. Swamynathan, J. Davis, and J. Piatigorsky
Identification of Candidate Klf4 Target Genes Reveals the Molecular Basis of the Diverse Regulatory Roles of Klf4 in the Mouse Cornea
Invest. Ophthalmol. Vis. Sci., August 1, 2008; 49(8): 3360 - 3370.
[Abstract] [Full Text] [PDF]