Transport of fluid by lens epithelium
Jorge
Fischbarg1,2,
Friedrich
P. J.
Diecke3,
Kunyan
Kuang2,
Bin
Yu2,
Fengying
Kang2,
Pavel
Iserovich2,
Yansui
Li2,
Heinz
Rosskothen2, and
Jan P.
Koniarek2
Departments of 1 Physiology and
Cellular Biophysics and
2 Ophthalmology, Columbia
University, New York, New York 10032; and
3 Department of Physiology, New
Jersey Medical School, University of Medicine and Dentistry of New
Jersey, Newark, New Jersey 07103
 |
ABSTRACT |
We report for the first time that cultured lens epithelial cell
layers and rabbit lenses in vitro transport fluid. Layers of the
TN4
mouse cell line and bovine cell cultures were grown to confluence on
permeable membrane inserts. Fluid movement across cultured layers and
excised rabbit lenses was determined by volume clamp (37°C).
Cultured layers transported fluid from their basal to their apical
sides against a pressure head of 3 cmH2O. Rates were (in
µl · h
1 · cm
2)
3.3 ± 0.3 for
TN4 cells (n = 27) and 4.7 ± 1.0 for bovine layers (n = 6). Quinidine, a blocker of
K+ channels, and
p-chloromercuribenzenesulfonate and
HgCl2, inhibitors of aquaporins,
inhibited fluid transport. Rabbit lenses transported fluid from their
anterior to their posterior sides against a
2.5-cmH2O pressure head at 10.3 ± 0.62 µl · h
1 · lens
1
(n = 5) and along the same pressure
head at 12.5 ± 1.1 µl · h
1 · lens
1
(n = 6). We calculate that this flow
could wash the lens extracellular space by convection about once every
2 h and therefore might contribute to lens homeostasis and transparency.
cell layers; fluid circulation; lens extracellular space; lens
homeostasis; nutrient transport
 |
INTRODUCTION |
THE LENS IS AN AVASCULAR tissue thought to be nourished
solely by diffusion. However, simple diffusion of nutrients into the lens has been deemed insufficient to account for its metabolic consumption (18). This suggested to us a need to search for additional
mechanisms of nutrient transport into the lens. In this regard, the
lens is a structurally and functionally asymmetrical tissue with highly
localized transport properties. The fiber cells comprise the vast bulk
of the lens, but they have relatively low intrinsic
Na+-K+-ATPase
activity, which makes them unlikely candidates for the additional
mechanisms sought. Nutrient uptake by the epithelium followed by
transport of nutrients through cell-to-cell gap junctions may be
considered for that role, but, although the epithelium communicates
with the fiber lens cells via such junctions, it does so mostly in the
equatorial region. The lack of gap junctions was demonstrated first for
the lens of chicken by Brown et al. (7) followed by a
structure-function study (4). Subsequently, a similar lack of gap
junctions between epithelial and fiber cells was reported in the lens
of the macaque by Kuszak et al. (24). Rae et al. (38) reached a similar
conclusion using a dye distribution method. Current evidence,
therefore, suggests that such a mechanism does not appear to be
sufficient to nourish the entire lens.
A search for explanations would cover hypothetical, still undetected
transport mechanisms that would be best sought, prima facie, in the
lens epithelium. This layer covers the lens anterior surface, it has
functionally different apical and basolateral membranes, and it has
been suggested to have a central role in maintaining lens homeostasis
and integrity. Asymmetry of transport functions was first demonstrated
by Becker and Cotlier (5) and Kinsey and Reddy (23), and more recently
a detailed analysis of the transport properties of the lens has been
provided in a series of papers by Mathias, Rae, and Eisenberg (28, 30). This subject has been reviewed recently (29).
In other tissues, driving forces for transport of nutrients are usually
provided by ionic transport. In this regard, Rae, Mathias, and
colleagues (27-29) have proposed that transport of ions by the lens epithelium might lead not only to the observed ionic
currents circulating around the lens (41) but also to transepithelial
fluid entry into the lens followed by fluid circulation through the
lens and equatorial exit. However, fluid movement through the lens has
been reported only once, as a short qualitative observation for an
excised rabbit lens, and in a direction opposite to what we find (13).
Fluid transport across lens epithelium per se apparently had never been
measured directly.
From what has been studied so far, all epithelial layers involved in
water translocation (and only those) express water channels (10, 11,
32). With this background, since a recent report located the water
channel aquaporin-1 in lens epithelium but not in fiber cells (45), we
hypothesized that the lens epithelium could transport fluid. We report
here that it does, both for cultured lens epithelial cell layers and in
vitro rabbit lenses. Implications of this finding for lens homeostasis
are considered in our discussion below.
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MATERIALS AND METHODS |
Cultured mouse lens epithelial cell layers.
The simian virus 40 (SV40)-transformed
TN4 mouse cell line was used.
These cells are derived from the lens epithelium of a transgenic mouse
(26) and carry a hybrid gene of the murine
A-crystallin promoter
fused to the SV40 large T antigen. These cells maintain epithelial
morphology for many passages and do not differentiate into fiber cells.
They synthesize some of the protein markers of lens epithelial cells:
-crystallin,
-crystallin,
-insert-crystallin, and
-crystallin (this last is present in low concentrations) (42).
Sixth-generation cells were subcultured in
25-cm2 flasks filled with DMEM
with high (4.5 g/l) glucose plus 10% fetal bovine serum (FBS) and
antibiotics (penicillin and streptomycin). After confluence, the cells
were harvested with a Ca2+-free
trypsin solution, spun at 1,000 rpm (Sorvall GLC-1 centrifuge, HL4
rotor) for 5 min in DMEM, resuspended in DMEM, and seeded into
transparent, permeable, tissue culture-treated inserts (internal diameter 24 mm, no. 3492, Transwell, Costar, Cambridge, MA; cells from
1 flask were split among 4 inserts). Confluence was determined both
microscopically and by determining the specific resistance of each
insert. Cells reached confluence after 3-4 days, and experiments were performed 1-3 days after confluence.
Cultured bovine lens epithelial cell layers.
Bovine eyes were washed with physiological saline containing 50 µg/ml
garamycin. The lenses were excised under sterile conditions in a
laminar flow hood, and the epithelial cells were gently scraped off.
The dislodged cells were aspirated with a pipette and added to 5 ml of
DMEM containing 20% FBS and antibiotics. The cell suspension was
centrifuged as above, the supernatant was discarded, and the pellet was
resuspended in 2 ml of the same medium. For primary cultures, the cells
were apportioned into 75-cm3
tissue culture flasks (Falcon 3023). After 2 days and every 2-3 days thereafter, the cells were fed with fresh DMEM containing 20%
FBS. After the cells reached confluence (in 5-7 days), they were
detached and subcultured at a ratio of 1:5 using trypsin plus EDTA.
Only cells from passages
1 and
2 were used, since their morphology
remains constant during those passages (2). Within 1-3 days after
becoming confluent, the cells were harvested with trypsin plus EDTA and
plated on Costar inserts at ~106
cells/insert. Cells reached confluence in 5-7 days; they were fed
every 2-3 days and examined by microscopy throughout the culture period. Fluid transport experiments were performed 1-7 days after confluence. Confluence was determined by microscopy and was also ascertained by determining the resistance of each insert before a given
experiment (see below).
Fluid flow measurements: cultured cell layers.
The rate of fluid movement across experimental preparations was
determined by the Bourguet-Jard volume-clamp method (6) as modified in
our laboratory (12, 31). We used a specially designed chamber having a
top section and a bottom section separated by the cell layer attached
to the permeable membrane of a Costar insert and connected
hydraulically to the recording apparatus (see Fig.
1). Each chamber section had a
water jacket for temperature control (37°C). The top section of the
chamber enclosed the insert holding the cells. The bottom section of
the chamber accommodated a steel mesh disk covered by a nylon net to
prevent sagging of the flexible permeable support.

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Fig. 1.
Fluid flow apparatus used for our experiments. Cultured cell layer on a
permeable substrate (insert) separates 2 sections (upper and lower) of
experimental chamber. Chamber walls are jacketed for temperature
control; for clarity, jackets are shown on
left only. Solution in upper section
can be changed as desired. Vertical position of entire chamber can be
varied to obtain desired hydrostatic pressure difference ( P) between
it and microelectrode detector for fluid level. In most experiments,
this P was 3 cmH2O.
Inset: holder for an excised rabbit
lens, which can be clamped in chamber in place of cultured cell layer
insert.
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The nanoinjector fluid flow measurement setup was similar to that used
earlier by us (31), except for its detector. The photodetector
previously used to sense fluid flow was replaced with a glass
microelectrode (microfilament glass GCF-100-6, A-M Systems, Toledo, OH;
filled with 3 mol/l KCl) that sensed by electrical contact the level of
a water meniscus in a connecting capillary tube (Fig. 1). To avoid
microelectrode blockage, the voltage applied to it was limited to
~100 mV; an amplifier generated transistor-transistor logic voltages
to drive the nanoinjector.
This setup maintained constant volume in the bottom section of the
chamber into or out of which fluid was pumped by the experimental preparation. We recorded the rate of reinjection into or out of the
bottom section; this rate directly corresponds to the rate of transport
across the epithelial layer (Fig. 1). 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 section of
the chamber) was usually 3.0 cmH2O. Hence, when the layers
transported fluid actively, they moved fluid from the bottom chamber to
the top one against a hydrostatic pressure difference.
The analog output voltage from the nanoinjector was proportional to the
flow rate and was fed to a paper chart recorder and a computer via an
analog-to-digital interface (DI-120, DATAQ Instruments, Akron, OH; data
were analyzed with the Windaq program from DATAQ). Both sections of the
chamber were filled with identical solution (usually DMEM), except when
inhibitors of fluid transport were used. Once the preparations were
mounted, only the upper chamber was accessible for solution exchanges
or gassing. Hence, to maintain pH, we added 25 mmol/l HEPES to the
standard DMEM solutions employed.
Fluid flow measurements: in vitro rabbit lenses.
Fluid transport measurements were done in the same apparatus employed
for flow measurements across cultured cell layers, except that the
lenses were mounted in a special insert designed to fit the existing
chambers. This required a spherical cavity inside an insert and several
gaskets placed so as to separate the top and bottom compartments while
avoiding fluid leaks. A schematic diagram of the chamber and lens
insert appears in Fig. 1. Three screws join the halves of the holder
and effectively contribute to clamp the lens.
Adult male or female albino rabbits (~2 kg) were euthanized with 100 mg/kg of pentobarbital sodium solution (Butler, Columbus, OH) injected
into the marginal ear vein. Their eyes were excised and placed in a
holder; the lenses were excised and were gently transferred to the
experimental chamber (Fig. 1) using a
Teflon-coated spatula. One eye was used immediately; the second eye was
stored in a moist environment at 4°C and was used the next day.
Data obtained from both freshly dissected and stored lenses were
comparable.

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Fig. 2.
Examples of fluid transported by layers of cultured lens epithelial
cells from their basal to their apical side against a pressure head (3 cmH2O).
A: TN4 lens epithelial cells.
B: cultured bovine lens epithelial
cells. Each line represents fluid volume accumulating during a fixed
time interval (10 s), after which accumulator was reset and process was
repeated. Positive direction corresponds to fluid movement from bottom
chamber section to top chamber section, that is, from basolateral to
apical side of cells (and vice versa for negative direction). Fluid
level in top chamber section was 3 cm above that of detector
communicating with bottom chamber section. Preparations transported
fluid for several hours; rate of transport slowly decreased, and
finally flow reversed when a leak developed, driven by pressure head.
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Measurements of specific electrical resistance of cultured layers.
Electrical resistance of epithelial layers was determined using an
Endohm chamber and a resistance meter (both from WPI, Sarasota, FL);
the resistance determined with a control insert was subtracted from
that of an insert with the layer. Specific resistance was calculated by
multiplication by the area of the insert (4.7 cm2). To evaluate given layers
before an experiment, minimum acceptable values (in
· cm2)
were 80 for the cell line and 60 for the bovine cells. The average specific resistance values were, for the
TN4 cells, 130 ± 7.3
· cm2
(n = 23) and, for the bovine layers,
85.4 ± 5.9
· cm2
(n = 6).
In the experiments in which resistance was monitored as a function of
time, the Endohm chamber was placed inside a 37°C incubator. To
determine the effect of
p-chloromercuribenzenesulfonate
(PCMBS), the solution used was DMEM plus HEPES, as above. To determine the effect of Ca2+- and
Mg2+-free exposure, the solution
used for the initial control period was a bicarbonate-HEPES medium (in
mM: 81.4 NaCl, 5.4 KCl, 44.0 NaHCO3, 0.8 NaH2PO4,
0.4 MgSO4, 1.8 CaCl2, 25 glucose, and 25 HEPES).
The test solution had the CaCl2
and the MgSO4 replaced with NaCl
on a molar basis and also included 2 mM
Na2EDTA.
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RESULTS |
Fluid transport by cultured cell layers.
We examined the behavior of 68 cultured layers. In 35 of them, fluid
leaked through the layer, which we attribute to either the trauma of
mounting in the chamber or lack of complete confluence. However, the
other 33 preparations transported fluid from their basal to their
apical sides against a head of pressure; in vivo, that direction
corresponds to transport from the anterior chamber of the eye to the
interior of the lens. A number of experiments were conducted with
varying pressure heads applied to the preparations. Preparations could
transport against pressure heads as large as 10 cmH2O without apparent change in
the rate of transport. However, because such high pressure differences
are unlikely across an in vivo layer, we chose to standardize on a
pressure head of 3 cmH2O. Our
experimental device requires some pressure head to keep the tissue and
its support relatively immobile, so lower pressure heads were deemed
inappropriate for this reason.
The preparations transported fluid spontaneously and continuously for
several hours (Fig. 2); the longest time monitored was 6.5 h (not
shown). Afterwards, a leak ensued, driven by the pressure head (Fig.
2). For the bovine cell layers, the average rate of fluid transport was
4.69 ± 1.0 µl · h
1 · cm
2
(n = 6); for the
TN4 mouse cell
line layers, that rate was 3.31 ± 0.31 µl · h
1 · cm
2
(n = 27).
Because the experimental setup is based on keeping the volume of the
bottom compartment constant, this compartment is inaccessible during
the experiment. For this reason, pharmacological agents could only be
added to the top compartment (in contact with the apical cell
membranes). Quinidine sulfate, a blocker of
K+ channels, inhibited fluid
transport in a dose-dependent manner in four experiments. After
addition of this inhibitor (50 µmol/l), a slight decrease of fluid
transport was observed. After increase of the inhibitor concentration
to 150 µmol/l, a further decrease of fluid pumping was observed,
which decreased to almost zero within 60 min. Increasing the inhibitor
further (to 300 µmol/l) resulted in fluid leak, as fluid movement
reversed, driven by the existing pressure head. Figure
3A
illustrates a typical experiment.

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Fig. 3.
Experiments demonstrating effects of inhibitors on fluid transport by
TN4 cultured cells. Inhibitors were added to apical side of layer
(top chamber section). P = 3 cmH2O.
A: quinidine sulfate inhibited fluid
transport in dose-dependent fashion. When quinidine concentration
reached 300 µmol/l, fluid transport ceased.
B:
p-chloromercuribenzenesulfonate
(PCMBS; 1 mmol/l) inhibited fluid transport after an ~30-min delay.
C:
HgCl2 (1 mmol/l) inhibited fluid
transport after a short delay. In all 3 cases, after fluid transport
ceased, a leak rapidly developed.
Inset: specific resistance of cultured
cell layers (relative to its initial value;
R/Ro)
is plotted so that its time course is superimposed on that of changes
in fluid transport before and after PCMBS exposure
(B). Each point is an average of 4 determinations. Average
R/Ro
for cultured layers were 69.5 ± 1.7 and 98.3 ± 7.7 · cm2 for
PCMBS treated and Ca2+ free,
respectively.
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PCMBS is an unspecific but potent blocker of Hg-sensitive aquaporins.
This inhibitor was used at a concentration of 1 mmol/l in seven
experiments. After a 30-min delay, fluid pumping began to decrease
slowly, and it stopped 50 min later. The direction of fluid flow
reversed, indicating a leaky preparation (Fig.
3B). Similar results were obtained
with 0.1 mmol/l HgCl2 (Fig.
3C). In another series of
experiments, we attempted to determine whether the effect of mercurials
can be attributed to aquaporin blockade or to a nonspecific effect by
measuring the electrical resistance across the cell layer during
application of PCMBS or substitution of a
Ca2+- and
Mg2+-free solution with 2 mM EDTA.
As shown in Fig. 3B,
inset, after exposure to PCMBS and
after a delay, both electrical resistance and fluid transport
decreased; the decrease in fluid transport seems to precede slightly
the decrease in resistance. The replacement of ambient
Ca2+ and
Mg2+ (Fig.
3B,
inset), as may be expected, resulted
in a precipitous and nearly instantaneous drop in resistance. The
effect of HgCl2, although fast,
appears more gradual by comparison.
Last, we found no effects from addition of ouabain (1 mmol/l), Prozac
(1 mmol/l), 5-nitro-2-(3-phenylpropylamino)benzoic acid, or amiloride
to the apical side. This result is consistent with the existence of an
intercellular junction of intermediate resistance, which would diminish
the permeation of such inhibitors.
Fluid transport across in vitro rabbit lens.
We detected a spontaneous movement of fluid across lenses; the
direction of such movement was consistent with that observed in
cultured lens cell layers.
1) Lenses were mounted
"inverted," that is, with their posterior pole facing the top
compartment, and with a pressure head of 3 cmH2O applied to the posterior
side of the lens. In this case, fluid moved from the bottom to the top
compartment, against the pressure head (fluid transport), at a rate of
10.3 ± 0.62 µl · h
1 · lens
1
(n = 5). One of these experiments
is shown in Fig.
4A. Fluid transport was generally observed for some 3 h after mounting, during
which time it decreased slowly and continuously. Given the
inaccessibility of the epithelium (facing the bottom chamber, cf. Figs.
1 and 4A), no inhibitors were used
in this case.

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Fig. 4.
Fluid transported across excised rabbit lenses. Rates shown in boxes
represent average for initial 0.5 h of experiment.
A: lens inverted, with its anterior
side facing bottom chamber section. Fluid moves against pressure head.
B: lens upright, with its anterior
side facing top chamber section. Fluid moves along pressure head. In
both A and
B, fluid moved from anterior side to
posterior side of lens.
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2) Lenses were mounted
"upright," that is, with their anterior pole facing the top
chamber, and with a pressure head of 3 cmH2O applied to the anterior side
of the lens. In this case, fluid moved from the top to the bottom
chamber (from the anterior side to the posterior side of the lens,
following the pressure head) at a rate of 12.5 ± 1.1 µl · h
1 · lens
1
(n = 6) and for 1-3 h after
mounting. As in the inverted preparation above, the fluid movement
decreased continuously and slowly after mounting. A representative
experiment is shown in Fig. 4B. In four experiments, after ouabain (1 mmol/l) was added to the top chamber, fluid movement ceased (not shown). However, given the relatively short duration of fluid movements in control experiments, no
reversal of the inhibition was explored.
Because the fluid movements took place along the pressure head, these
flows could therefore reflect transport across the epithelium or a
passive leak driven by the pressure head. However, the magnitude of the
flow was very similar to that of the fluid transport (against a
pressure head) observed in the section above and was much lower than
the rate of leak through decapsulated lenses (see below), so such
movements are consistent with fluid transport by the lens epithelium.
In addition, the slow decrease with time of the fluid movements in both
cases is consistent with an active process that decays after mounting
rather than with a passive leak or an artifactual phenomenon.
So far, we have expressed flows as "per lens" because it is
somewhat uncertain which area of lens epithelium would be traversed by
these flows. When the epithelium faced upwards, a gasket pressed against it, and, when it faced downwards, the lens capsule rested on a
plastic surface with perforations. Hence, in both cases, the epithelial
area giving rise to the fluid movements detected could have been less
than the total epithelial area. Still, the flow is approximately the
same for both cases, so perhaps these uncertain factors played a minor
role. Using published dimensions for a rabbit lens epithelium (37),
namely, equatorial diameter of 10 mm and anteroposterior length (lens
thickness) of 7.0 mm, we calculate an epithelial area of 1.2 cm2 using an equation to compute
the segment of a sphere. With this correction, the in vivo lens
epithelium would be transporting fluid at rates of 8.8-10.4
µl · h
1 · cm
2.
Experimental hydraulic conductance of the lens.
The results above suggested that the epithelium was transporting fluid
across itself and into the lens. This posed the question of whether the
lens mass could exhibit a hydraulic conductance large enough to admit
such flow, since otherwise the fluid transported would be either
leaking back through intercellular junctions or be driven out of the
lens via some other paraepithelial route. To explore this matter, after
excising a lens, we dissected off a circular piece consisting of most
of its anterior capsule with the epithelium attached. In four
experiments, we verified that a hydraulic conductance across the lens
exists. Figure 5 exemplifies such data;
Fig. 5A shows a representative record
of the time course of flow vs. pressure difference, and
Fig. 5B shows the data points collected from all the experiments. The slope of the fitted line (24.1 ± 3.0 µl · h
1 · lens
1 · cmH2O
1)
represents the measured hydraulic conductance of the lens
(Lpm). Using
the cross-sectional area of the chamber (0.9 cm2),
Lpm
was
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(1)
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Fig. 5.
Passive flow of fluid leaking through excised decapsulated (hence
deepithelialized) rabbit lenses. Lenses were mounted with their
anterior sides facing top chamber section.
A: an individual recording showing
leak driven by 2 pressure heads in a single experiment.
B: rate of leak vs. pressure head;
each point represents average leak at a set pressure head during an
individual experiment. Points are from 4 experiments. Line represents
least squares fit (forced through origin;
r = 0.78, P = 0.02); its slope is hydraulic
conductance (24.1 ± 3.0 µl · h 1 · lens 1 · cmH2O 1).
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Theoretical calculation of lens hydraulic conductance.
Given this experimentally determined lens hydraulic conductance, we
asked ourselves whether the geometry of the lens was consistent with a
paracellular hydraulic conductance of such magnitude. For the ratio of
lens extracellular volume to total volume, the literature describes
values of 5-12% determined with cell-impermeant radiolabels (8,
33, 46) and 0.5% as determined from electron microscopic images (33).
Further work will be required to reconcile this discrepancy; here we
chose a mean estimate of 7%, representing the work with impermeant
tracers. From this value, given the histology of lens fibers, the
intercellular distances can be calculated. We assumed the simple
geometry exemplified in Fig. 6, with cell dimensions comparable to those reported for rabbit lens (34). Values
and formulas were
To
simplify, we neglect the z-axis, and
we assume that the proportion of extracellular to total volume will
correspond to that of the extracellular to total area in a tangential
section. With these assumptions, we can calculate the intercellular
distance that will result in a 7% ratio of extracellular area to total area. The extracellular area to be apportioned to each cell (hAs) is
the product of the cell perimeter times the half-width of the intercellular spaces (hws)
Because
hAs/(hAs + Ac) = 0.07, it follows that hws = 950 Å. Further parameters
required are
To
calculate the lens hydraulic conductance
(Lpc), we
assumed that the average anteroposterior length of the interfiber
clefts (lifc) was 5 mm. Using Poiseuille's equation for parallel
plates (
= 0.007 poise at 37°C),
Lpc
is
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(2)
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This value falls within the order of magnitude of the
experimentally determined value
(Lpm/Lpc = 6.2), which reinforces the validity of our measurements.

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Fig. 6.
Schematic cross section of a lens depicting fiber cells and interfiber
paracellular clefts on an arbitrary scale. Labels signify dimensions of
fiber cells and paracellular clefts (dashed lines); lcnf, lens fiber
cell narrow face; lcbf, lens fiber cell broad face; lcw, lens fiber
cell width; lct, lens fiber cell thickness; hws, half-width of
intercellular spaces. Solid lines and arrows highlight paracellular
diffusional pathways.
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The issue of whether transport of fluid by the epithelium might lead to
an undesirable subepithelial pressure buildup also deserves some
mention. In this regard, lens conductance values of the order found
appear physiologically compatible with the observed rates of fluid
transport. Given our
Lpm of 26.7 µl · h
1 · cm
2 · cmH2O
1,
a pressure difference of only 0.3-0.4
cmH2O would suffice to induce the
translenticular rates of fluid flow we report here (8.8-10.4
µl · h
1 · cm
2).
Thus the epithelial transport mechanism could induce fluid flow through
the lens with only minimal subepithelial pressure buildup.
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DISCUSSION |
These experiments demonstrate for the first time that the lens
epithelium actively transports a sizable amount of fluid from its basal
to its apical side. In cultured cell layers, the rates of transport
observed (3.3 and 4.7 µl · h
1 · cm
2
for the
TN4 cell line and the bovine lens epithelial cells, respectively; Figs. 2 and 3) were of the order of those in other fluid-transporting layers (e.g., 4-6
µl · h
1 · cm
2
for the rabbit corneal endothelium). The rate of fluid transport across
in vivo lens epithelium was somewhat larger, ~11
µl · h
1 · lens
1
(Fig. 4) or 8.8-10.4
µl · h
1 · cm
2.
In both cultured layers and in vitro lens, fluid movement took place
against a hydrostatic pressure difference, which is characteristic of
energy-requiring mechanisms for the transport of fluid. Moreover, in
the cultured layers, toward the end of the experiments (or in response
to inhibitors), when transport stopped, the fluid movement reversed its
direction and a leak ensued across these preparations, driven by the
hydrostatic pressure difference. This observation is important because
it confirms that the layers were intact and functional before cessation
of transport.
The blocking of fluid transport by quinidine added to the apical side
of the cultured layers (Fig. 3) was consistent with its known
inhibitory effect on K+ channels
(1, 40). Although this observation is consistent with
quinidine-sensitive K+ channels
playing a direct role in fluid transport, this cannot be affirmed
unambiguously. The blocking of fluid transport by the mercurials PCMBS
and HgCl2 added to the apical side
is consistent with the water channels (45) of the lens epithelium being
central to fluid transport. The delay for PCMBS to inhibit fluid
transport was of the order of the incubation time required for this
agent to act in other systems, such as Xenopus
laevis oocytes expressing water channels. However,
PCMBS administration also resulted in a decrease of the electrical
resistance of the cell layer, which progressed with a time course
somewhat delayed but otherwise similar to that of the decrease of fluid
transport. A Ca2+- and
Mg2+-free solution expected to
cause junctional opening practically eliminated electrical resistance
almost instantly (Fig. 3B,
inset), so the gradual decrease in
electrical resistance after PCMBS could be due to changes in cell shape
and enlargement of the paracellular space. Still, given such resistance
changes, our current experiments with mercurials do not permit a
definitive conclusion regarding the path of water movement across the
cell layer. It should be noted, however, that recent evidence obtained
with aquaporin-1 knockout mice (44) strongly suggests a transcellular
route for transepithelial water transport.
The rate of fluid transport across in vitro lenses is comparatively
large, ~10
µl · h
1 · cm
2.
For comparison, the rabbit kidney proximal tubule transports ~150
µl · h
1 · cm
2,
based on area of luminal epithelial surface excluding microvilli (19,
43); if the area of microvilli is considered, that number decreases to
3.75 µl · h
1 · cm
2.
Fluid transport through the lenticular surface thus would be larger
than that through the kidney tubule. However, if one considers that the
basolateral area of the lens epithelium is several times larger than
the lenticular surface, given the infoldings of its membrane (17), the
transcellular flow per unit area falls to the same order of magnitude
as that seen in the kidney. The surface of the apical membrane of the
lens epithelium is relatively smooth; however, the apicolateral
membranes show infoldings (24) that may contribute to an enlargement of
the apical membrane area. In addition, the apical membrane is in close
proximity to the underlying cell fiber membranes [a 0.1-µm
separation can be estimated from electron micrographs (17)],
which might lead to more efficient solute-solvent coupling and rates of
fluid transport in excess of those seen in other epithelia with more
conventional geometry.
Possible physiological significance of lens epithelial fluid
transport.
From the evidence in this paper, the direction of fluid transport in
the lens would be from the anterior surface to the interior of the
lens, which may result in fluid circulation within the lens. Given that
the lens is an avascular tissue, such circulation might greatly aid in
making nutrients available to the lens fibers and removing their waste
products, possibly playing an important role in maintaining lens
homeostasis and transparency.
In this regard, nutrients could reach the interior of the lens via two
conceivable pathways. Some electron microscopic evidence has led to the
proposal that, after the uptake of glucose and other nutrients by the
epithelial cells, diffusion of nutrients could take place via gap
junctions between the epithelial and fiber cells (16). However, more
recent studies place this possibility in doubt. Brown et al. (7),
Bassnett et al. (4), and Kuszak et al. (24) reported a virtual absence
of gap junctions between epithelium and fiber cells in the central area
of lens epithelium of chicken and macaque. They found junctions, but
rather sparse ones and located only toward the equatorial region of the
epithelium. In addition, from dye-transfer studies (38), only 10% of
the epithelial cells would be coupled with fiber cells.
The second pathway would be similar to that existing in most other
tissues, namely, nutrients reaching cells via diffusion along the
extracellular space. For the lens, several authors have discussed their
evidence in those terms (9, 35). Such a view is apparently consistent
with the ubiquitous presence of facilitative glucose transporters in
lens fiber membranes (22, 25). Metabolites would leave the lens by the
same path. However, the extracellular space of the lens is tortuous,
and the effective diffusion coefficient for cell-impermeant solutes in
the radial direction is only some 8.5% of that in free solution (34).
This poses the question of whether diffusion alone can really provide
an adequate supply of nutrients to the lens and clear its waste products.
Faced with this enigma, we performed calculations to explore it. If
glucose diffuses in and is being consumed, consumption might outstrip
diffusion, depending on their relative rates. In that case, glucose
concentration could fall to zero, and some regions of the tissue would
go unnourished. A similar question gave rise to a classical treatment
of diffusion-consumption in muscle by Hill (20), extended to spherical
geometry by Gerard (14). Gerard's equation is
|
(3)
|
where
r is the radial distance from the
center of the lens, Cl is the
glucose concentration in the extracellular space of the lens,
Ce is the glucose concentration in
the external medium surrounding the lens, U is the glucose consumption
in
mg · s
1 · g
lens tissue
1,
D is a diffusion
coefficient, and A is the
lens radius.
The limiting radial distance at which glucose concentration would
become zero
(rG0) is
|
(4)
|
We
have calculated limiting distances for the lenses of rat, rabbit, and
calf. Table 1 shows the results of our
calculations, along with values obtained from the literature (3, 15,
21, 47, 48) for the geometry and glucose consumption of lenses of known
weight. D for glucose was 8.5% of
that in free solution, or 7.7 × 10
7
cm2/s (34). As Table 1 shows,
regardless of species, size, or rate of consumption, extracellular
diffusion seems clearly insufficient to meet the metabolic demand, as
glucose would penetrate only into the outer 8-10% of the lens
radius before being completely depleted. From this, extracellular
glucose would be found only in the superficial 25% of the lens mass;
in a rabbit lens, this outer layer would correspond to approximately
the 400-µm thickness of the layer of differentiating cells and the
anterior epithelium (29). The remaining 75% of lens mass would remain
without glucose; similar conclusions would apply to other nutrients.
View this table:
[in this window]
[in a new window]
|
Table 1.
Glucose consumption of lenses of different species and calculated
distance to which glucose will penetrate by diffusion
|
|
Against this background, we describe a mechanism of fluid transport by
the epithelium and postulate a resulting circulation of fluid through
the lens extracellular space that would exchange the contents of that
space by convection about once every 2 h. Fluid circulation of such
magnitude could be very important for lens homeostasis. In the rest of
the body, a cell cannot survive unless it has a blood capillary within
some 20-100 µm (39). Lens fiber cells exist at distances from
aqueous humor many times larger than that physiological limit; clearly
some special mechanisms must be involved in their homeostasis. In fact,
it has been noted that an active process is required to bring glucose
into the lens (18). Apparently no follow-up studies to explore this
issue have been conducted in the intervening time, but this isolated observation fits very well with the ideas we present here.
Last, several authors (27-29, 41, 49) have described a circulation
of electrical current flow around the lens, originating in
Na+ extrusion by the epithelial
cells of the equatorial region and reentering the lens passively at the
anterior and posterior poles. The current loop would then be completed
by Na+ entry into fiber cells and
diffusion back toward the equatorial epithelial cells. It has been
postulated that such current flow might be the driving force for a
circulatory flow of fluid along the same circular paths through the
lens (27-29, 49). Such hypothetical fluid movement would require
fluid to 1) be transported out of the equatorial epithelial cells in the apical-to-basolateral direction and 2) passively enter the lens
through the epithelium at the anterior pole and the paracellular spaces
of the posterior pole. However, contrary to
requirement 1, we observe that
cultured lens epithelial cells transport fluid in the
basolateral-to-apical direction, so they would be expected to drive
fluid into the lens (rather than out of it) through the equatorial
region. Contrary to requirement 2, the
fluid transport across the anterior epithelium is active, not passive,
and we observe that it exits the lens through the posterior pole,
rather than entering through it. Of course, out of necessity, we
clamped the lens in such a way that possible fluid movements across
part of its equatorial region were excluded from detection (Fig. 1), so
equatorial fluid exit cannot be ruled out without further work. It has
been reported (36) that the magnitude of the short-circuit current in
rabbit lens is ~22.6 µA/cm2.
If this short-circuit current is due to the recirculating
Na+ current, and if we assume
isosmotic coupling, such a current would result in a fluid flow of 5.7 µl · h
1 · cm
2,
which is about one-half of the rates we measure for the in vitro rabbit
lens. Moreover, this short-circuit current is inhibited nearly 40% in
HCO
3-free solutions and 20% in Cl
-free solutions,
suggesting that the current flow is not just due to passive flow of
Na+ through the anterior pole of
the epithelium. Hence, the circulatory model cannot completely account
for our observations. The arguments above suggest that our results are
due to a mechanism different from and perhaps in parallel with the
measured currents and the postulated coupled circulatory fluid flow.
Our results clearly suggest that the epithelium transports fluid into
the lens. Taking into account the known presence of membrane channels,
transporters, and an aquaporin in lens epithelium, we believe that the
simplest explanation for our observations is the heretofore unknown
presence of a classical epithelial fluid transport mechanism in this
layer. The sheer magnitude of the phenomenon we report suggests that it
may be of great importance for lens homeostasis.
 |
ACKNOWLEDGEMENTS |
This work was supported by National Eye Institute Grant EY-06178
and by Research to Prevent Blindness, Inc.
 |
FOOTNOTES |
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. §1734 solely to indicate this fact.
Address for reprint requests: J. Fischbarg, Dept. of Physiology and
Cellular Biophysics, College of Physicians and Surgeons, Columbia
University, 630 West 168th St., New York, NY 10032.
Received 30 June 1998; accepted in final form 2 December 1998.
 |
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