§
§
§
§
From the * Department of Medicine and Department of Physiology, Cardiovascular Research Institute, and § Graduate Group in
Biophysics, University of California, San Francisco, San Francisco, California 94143-0521
A method was developed to measure the osmotic water permeability (Pf) of plasma membranes in cell layers and applied to cells and epithelia expressing molecular water channels. It was found that the integrated intensity of monochromatic light in a phase contrast or dark field microscope was dependent on relative cell volume. For cells of different size and shape (Sf9, MDCK, CHO, A549, tracheal epithelia, BHK), increased cell volume was associated with decreased signal intensity; generally the signal decreased 10-20% for a twofold increase in cell volume. A theory relating signal intensity to relative cell volume was developed based on spatial filtering and changes in optical path length associated with cell volume changes. Theory predictions were confirmed by signal measurements of cell layers bathed in solutions of various osmolarities and refractive indices. The excellent signal-to-noise ratio of the transmitted light detection permitted measurement of cell volume changes of <1%. The method was applied to characterize transfected cells and tissues that natively express water channels. Pf in control Chinese hamster ovary cells was low (0.0012 cm/s at 23°C) and increased more than fourfold upon stable transfection with aquaporins 1, 2, 4, or 5. Pf in apical and basolateral membranes in polarized epithelial cells grown on porous supports was measured. Pfbl and Pfap were 0.0011 and 0.0024 cm/s (MDCK cells), and 0.0039 and 0.0052 cm/s (human tracheal cells) at 23°C. In intact toad urinary bladder, basolateral Pf was 0.036 cm/s and apical membrane Pf after vasopressin stimulation was 0.025 cm/s at 23°C. The results establish light microscopy with spatial filtering as a technically simple and quantitative method to measure water permeability in cell layers and provide the first measurement of the apical and basolateral membrane permeabilities of several important epithelial cell types.
Key words: water transport; trachea; Fourier optics; aquaporinThe development of methods to measure the plasma
membrane osmotic water permeability of cell layers has
been motivated by the identification of a family of molecular water channels (aquaporins) widely expressed
in mammalian tissues (for review, see Nielsen and Agre,
1995; Verkman et al., 1996
). Water channels have been localized in many epithelia involved in fluid transport,
including kidney tubules, airways, and alveoli in lungs,
choroid plexus in the brain, and ciliary body in eyes. Although a considerable body of data has been obtained
on the molecular biology and expression pattern of water channels, there have been few measurements of the
water permeability of epithelial cell plasma membranes. Such functional measurements will be required
to determine whether water channels can account quantitatively for the wide variations in water permeability of
epithelial cell plasma membranes and to study the regulation of water permeability.
For example, water channels in the lung are thought
to mediate transepithelial fluid transport (Matthay et
al., 1996). Recent studies have demonstrated high water permeability in bronchial (Folkesson et al., 1996
)
and alveolar epithelia (Carter et al., 1996
). There have
been no measurements of water movement in tracheal epithelia, nor have measurements been made of water
permeability of individual apical and basolateral plasma
membranes in bronchial and alveolar epithelium. The
water channel aquaporin-4 has been localized to the
basolateral membrane of tracheal epithelia, where it is
thought to play a role in maintenance of proper mucosal hydration (Frigeri et al., 1995
). It is thus predicted that tracheal epithelial cells have high water permeability; a hypothesis to be tested in this study. Similarly, experiments with the toad urinary bladder led to
the formulation of the shuttle hypothesis to account for
vasopressin-regulated water permeability (reviewed in
Jo and Harris, 1995
). This hypothesis predicts that the
apical membrane is the rate limiting barrier and that its
permeability is modulated by the vasopressin-regulated
targeting of water channels to the apical membrane. Although the transepithelial water permeability has been
measured in many studies, there is little information about water permeability of the individual plasma membranes.
The determination of water permeability in individual cell membranes in a polarized epithelium can be
accomplished by measurement of the time course of
cell volume change in response to changes in the osmolarity of perfusate bathing one surface of the cell layer.
For epithelial cell layers with high water permeability, the rapid changes in cell volume require a method with
high temporal resolution. Several methods have been
proposed and evaluated to measure water permeability
in immobilized cell layers such as cultured epithelial
cells that are grown on glass coverslips. Light scattering
from a cell layer was found to provide a qualitative index of cell volume (Echevaria and Verkman, 1992; Fischbarg et al., 1993; McManus et al., 1993
). However, the
light scattering signal is insensitive to cell volume for
many cell types and there is no rigorous basis for predicting the sign and amplitude of volume-dependent
signal changes. Strategies to measure cell volume changes
have been proposed involving tracking of the z position of fluorescent beads at the cell surface (Crowe and Willis, 1991
; Van Driessche et al., 1993
; Kao and Verkman,
1994
) and cell shape reconstruction by imaging (Kacha-dorian et al., 1985); however, these methods are approximate, too difficult technically for routine use, and
have limited data acquisition rates. Recently, accurate determination of the relative volume of cells grown on
a transparent support was accomplished by total internal reflection (TIR)1 fluorescence microscopy (Farinas
et al., 1995
). Cell cytoplasm was stained with a fluorescent indicator such as calcein, and relative cell volume
(inversely proportional to indicator concentration) was
deduced from the TIR fluorescence from a thin (<200
nm) layer of cytoplasm near the transparent support.
Measurement of indicator concentration can also be accomplished by optical sectioning techniques using confocal (Crowe et al., 1995
) or partial confocal (Muallem et
al., 1992
) optics, as well as ion-sensitive electrodes (Alvarez-Leefmans et al., 1995
).
We recently introduced an interferometry method to
measure cell volume and water permeability in adherent and nonadherent epithelial cell layers (Farinas and
Verkman, 1996). Volume changes were shown to alter
the optical path length (the product of refractive index
and geometric path length) of light passing through a
cell layer. An interferometer was used to convert the
small changes in optical path length to measurable
changes in intensity. Cell membrane osmotic water permeability was determined from the time course of interference signal in response to osmotic gradients. Although interferometry provided the only available
method with high temporal resolution to measure water permeability in individual plasma membranes in
nonadherent cell layers, a distinct disadvantage of the
method was the strong sensitivity of the interference
signal amplitude to air currents, temperature gradients, and perfusion chamber pressure, which produced
instrument drift and reduced signal-to-noise ratio.
In the present study, a simple method to measure water permeability in cell layers has been developed based
on the cell volume-dependent changes in optical path
length demonstrated by interferometry. The method
was applied to the measurement of plasma membrane
osmotic water permeability of epithelial cell layers.
Dark field and phase contrast microscopes exploit the
technique of spatial filtering to generate images based
on optical path length differences in the object. It is
demonstrated in the theory section below that the integrated intensity of an image formed by such microscopes is sensitive to volume-dependent changes in optical path length. As shown schematically in Fig. 1, an
annulus and condenser are used to illuminate a cell
layer in the object plane of a microscope with a cone of
light. The cells diffract the light leading to a zero order
beam (thick lines emanating from the object) and higher order beams (thin lines). The zero order beam
represents the undeviated incident illumination while
the higher order beams contain the light diffracted by
the cells. The objective focuses the zero order beam at
the back focal plane while the higher order beams are
focused at the image plane. The zero order beam is localized to a distinct position in the back focal plane (a
circle) and thus can be manipulated by spatial filtering
(Hecht, 1987). In the dark field microscope, the spatial
filtering results in total attenuation of the zero order
beam, whereas in the phase contrast microscope the
zero order beam is partially attenuated and phase shifted with respect to the higher order beams. Because
cell volume affects the relative intensities of the zero
and higher order beams, attenuation of the zero order
beam leads to cell volume-dependent changes in the
light intensity at the image plane. We demonstrate here
that this optical phenomenon can be exploited to measure relative cell volume and water permeability using
phase contrast or dark field microscopes. The approach was developed theoretically, validated experimentally, and applied to measure water permeability in
cells expressing molecular water channels and intact
living tissues.
Theory
A theory was developed to relate the relative cell volume of a cell layer to the integrated intensity of an image from a dark field or phase contrast microscope.
The refractive index of a cell (ncell) depends on relative
cell volume (Vr) (Farinas and Verkman, 1996)
![]() |
(1) |
where nocell is the refractive index of the cell at Vr = 1 and nw is the refractive index of water. The optical path length (OPL(x,y)) through the cell layer of maximum thickness hmax is
OPL(x,y)=[{hmaxh(x,y)}np+h(x,y)ncell]
![]() |
(2) |
where n p is the perfusate refractive index, h(x,y) is the cell height profile, and A is the field of view. The integral term in Eq. 2 is included because only relative phase differences are important in phase contrast or dark field microscopy, so that the average OPL over the field of view must be zero
.
Assum-ing that cell shape does not change as cell volume changes (h(x,y) = Vr ho(x,y)), Eq. 2 becomes
![]() |
(3) |
where havg is the average cell layer height and ho(x,y) is the cell height profile at Vr = 1. Eq. 3 indicates that, due to volume-dependent changes in cellular refractive index and cell height, the optical path length of a cell layer depends on the relative cell volume.
Dark field and phase contrast microscopes generate
an intensity image (I(x,y)) based on the phase delay,
(x,y) = (2
/
) OPL, introduced by the phase object
(an object that alters the phase but not the amplitude
of the incident light) where
is the wavelength of light
(Goodman, 1968
). The predicted image of a thin object formed by a dark field or phase contrast microscope can be calculated using Fourier analysis. The
presence of the phase object causes the incident plane
wave of unit amplitude, with electric field Eo(x,y) = exp{
i
t} to be modified such that a position-dependent phase delay, exp{i
(x,y)}, is introduced
![]() |
(4) |
where the real part of Eo(x,y) describes the phase and
amplitude of the light at the object plane, is angular
frequency and t is time. For simplicity, coherent monochromatic light, entrance and exit apertures of infinite
extent, and a magnification of unity are assumed. Since
only relative phase differences are important, the
exp{
i
t} term can be dropped and
(x,y) was chosen
so that
.
For a thin object that introduces a small phase delay
[|(x,y)| < 1 rad], the exp{i
(x,y)} term can be approximated by the first three terms of the Taylor's series expansion
![]() |
(5) |
where a is the average value of (x,y)2. The first term in
Eq. 5 describes the light that passes through the sample
without a phase shift (the zero order beam), while the
latter terms account for the diffracted light (the higher
order beams).
At the back focal plane of the microscope, the electric field is given by the Fraunhoffer approximation
and is thus the Fourier transform of the electric field at
the object plane (Eq. 5) (Hecht, 1987)
![]() |
![]() |
(6) |
where Ebfp(u,v) is the electric field at the back focal
plane,
{} denotes the Fourier transform, and
(u,v) is
the delta function. Because both
(x,y) and {
(x,y)2
a} are real-valued functions with average values of zero,
only the first term in Eq. 6 contributes to the electric
field at the origin of the Fourier plane. Both dark field
and phase contrast microscopy generate images of
phase objects by spatial filtering of the electric field at
the back focal plane. The dark field microscope totally
attenuates the zero order beam while the phase contrast microscope phase shifts and typically (depending
on the objective) attenuates the zero order light relative to the higher order light. The spatial filtering is accomplished at the origin of the Fourier plane (located
in the back focal plane of the microscope) since this region contains all of the contribution of the zero order
beam. The phase shift and attenuation introduced by
the microscope are represented by multiplying the first
term in Eq. 6 by the factor T1/2exp(i
), where
is the
phase shift and T (transmittance) describes the attenuation of the zero order beam. For a dark field microscope,
= 0 and T = 0, while for the phase contrast microscope,
= ±
/2 and T > 0. Thus, the electric field
intensity just after passing through the back focal
plane, Ebfp(u,v) is
![]() |
![]() |
(7) |
At the image plane of the microscope, the electric field, Ei(x,y), is the Fourier transform of the electric field at the back focal plane
![]() |
![]() |
(8) |
The light intensity at the image plane, I(x,y), is the square of the modulus of the electric field at the image plane
![]() |
![]() |
(9) |
where only terms to second order in (x,y) have been
retained to be consistent with the original approximation (|
(x,y)| < 1 rad). The integrated intensity,
![]() |
![]() |
where c is an instrumental constant and the constants,
1 = cT +
2 (ncello
nw)2, and
2 = c(4
2/
2)(1
T)
do not depend on relative cell volume. Eq. 10 predicts
that the integrated intensity is approximately linearly
related to relative volume (since nw np < nocell
nw).
The sensitivity of the signal to cell volume changes depends on the difference in refractive indices of water
and the perfusate, as well as cell shape and intracellular
refractive index. Because the volume-independent
term (
1) is smaller for dark field microscopy, the sensitivity should be greater for dark field than for phase
contrast microscopy. The integration of the image intensity is accomplished either optically, by measuring the total intensity of the light emerging from the microscope, or numerically, by integrating the pixel intensity
of an image. For thick objects, |
(x,y)| > 1 rad, Eq. 4
can be used directly with Fourier analysis to numerically calculate the volume-dependent changes in transmitted light intensity (Goldstein, 1991
).
Cell Culture and Tissue Isolation
The following cell lines were grown on 18-mm diameter round
coverglasses: MDCK (CCL 34; American Type Culture Collection, Rockville, MD), CHO-K1 (Cell Culture Facility, University of
California, San Francisco), Sf9 (CRL 1711), CAKI-1 (HTB 46),
BHK (CRL 6281), astrocytes (HTB 12), and A549 (CCL 185).
Cells were grown in DME-H21, Ham's F12K, SFM, McCoy's 5a,
DME-H21, Leibovitz's L15, and Ham's F12K media, respectively.
All media except SFM were supplemented with 10% fetal calf serum. Cells were maintained at 37°C in a 95% air/5% CO2 incubator, except for Sf9 cells, which were grown at 28°C in an air incubator. Chinese hamster ovary (CHO) cell clones stably expressing rat AQP2 and AQP5 were generated by transfection with the
rat coding sequence using plasmid pcDNA3, G418 selection, and
clonal analysis as described previously for AQP1 and AQP4 (Ma
et al., 1993; Yang et al., 1996
). Primary cultures of human tracheal epithelia obtained from autopsy specimens, SK-MES cells (HTB 58; American Type Culture Collection) and MDCK cells
were grown on Cyclopore porous supports (0.45 µm; Falcon Labware, Cockeysville, MD)(Yamaya et al., 1992
). Cells were generally used when just confluent. Urinary bladders from the toad
Bufo marinus were dissected, washed, and mounted as described
previously (Farinas and Verkman, 1996
).
Instrumentation
Experiments were carried out on an inverted microscope (Diaphot; Nikon, Inc., Melville, NY) equipped with either a phase contrast condenser (LWD; Nikon, Inc.) or a dark field condenser (4029; E. Leitz Wetzlar GmbH, Wetzlar, Germany). Samples were illuminated with a 50-watt tungsten-halogen lamp powered by a stabilized DC power supply (68735; Oriel Corp., Stratford, CA). Unless otherwise specified, samples were illuminated with green light (546 nm) using a broad band interference filter and visualized using a 20× DL positive phase objective (numerical aperture 0.4; Nikon, Inc.). Transmitted light was collected and focused onto a silicon photodiode (PDA50; Thorlabs, Inc., Newton, NJ) or a cooled CCD camera (14-bit, 512 × 512 pixels; Photometrics Ltd., Tucson, AZ). The photodiode signal (0-1 V) was digitized by a 12-bit analog-to-digital converter (Computer Boards, Mansfield, MA) interfaced to a computer.
Cells grown on coverglasses were perfused in a channel-type
flow chamber (Farinas et al., 1995). Apical and basolateral surfaces of polarized cells grown on porous supports were perfused
using a dual perfusion chamber (Verkman et al., 1992
). The
same chamber was used with intact epithelial tissue layers except
that pins were used to stretch and stabilize the tissue. Solution exchange was accomplished using a 4-way valve (Hamilton Co.,
Reno, NV). Temperature was controlled using an in-line steel
coil immersed in a water bath just proximal to the flow chamber.
Perfusate flow was monitored by an in-line flow meter and temperature was monitored by a thermistor.
Solutions and Measurement Protocols
Solutions consisted of PBS (in mM): 137 NaCl, 2.7 KCl, 1.0 KH2PO4, 1.0 Na2HPO4, pH 7.4, 300 mosmol, hypotonic PBS (PBS diluted with specified amounts of distilled water), and hypertonic PBS (PBS containing specified concentrations of NaCl or glycerol). For studies in toad urinary bladder, toad Ringer's solution (in mM): 110 NaCl, 2.5 NaHCO3, 3 KCl, 2 KH2PO4, 1 CaCl2, 5 glucose, pH 7.6, 240 mosmol, was used. In some studies, solutions of specified osmolarities and refractive indices were prepared using combinations of NaCl, mannitol, and raffinose. Solution refractive indices were measured with an Abbe-3L refrac-tometer (Milton Roy, Rochester, NY). Measurements generally involved the continuous collection of transmitted light intensity during cell or tissue perfusion with solutions of specified composition.
Computation of Water Permeability for Cells on Glass Supports
Plasma membrane osmotic water permeabilities (Pf) were calculated from the time course of volume change in response to an osmotic gradient. As seen from Eq. 10, the integrated intensity is
approximately proportional to relative cell volume, Vr, [I = c1 + c2Vr, where c2 = I/(Vrf
1)]. The rate of volume change, dVr/
dt, is then related to the rate of change of intensity, dI/dt
![]() |
(11) |
where I is the amplitude of the intensity change and Vrf is the final volume. For cells grown on glass supports, and assuming that
cells act as perfect osmometers [
Vr = constant, where
is the
osmolarity], Vrf =
o/
f. The value of dI/dt|t = 0 was determined
by least squares fitting of the data (Kinfit; Olis, Inc., Jefferson,
GA). The initial rate of cell volume change (dVr/dt|t = 0) is given
by dVr/dt|t = 0 = Pf (S/Vo) vw
where S/Vo is the cell surface-to-volume ratio, and vw is the molar volume of water. Surface-to-volume ratios were measured by image reconstruction using confocal microscopy as described previously (Farinas et al., 1995
). For
cell layers grown on glass supports, it follows that the water permeability coefficient is Pf = Î/ (S/Vo vw
f) where Î = (dI/dt|t = 0 /
I) is the rate of signal change normalized to the signal amplitude.
Computation of Water Permeability for Polarized Epithelial Cells
In the case of an epithelial cell layer, both the time course of volume change and the final volume, Vrf, are dependent on both the
apical and basolateral perfusate osmolarities (ap and
bl), as
well as the apical Pfap and basolateral Pfbl water permeability coefficients
![]() |
(12) |
![]() |
(13) |
where o is the initial cell osmolarity. Because the volume
change depends on both apical and basolateral parameters, two independent measurements are required to calculate Pfap and Pfbl
from measured intensity changes. Eqs. 11-13 can be used to relate the signal to the permeabilities and relative volume. Three
distinct experimental protocols are suitable.
The first protocol uses a gradient of varying magnitude applied at any one of the two sides. Combining Eqs. 11-13 (for a change in apical membrane osmolarity)
![]() |
(14) |
A plot of Î versus perfusate osmolarity, ap, yields a straight line
whose slope depends on the permeability on the cis side and
whose intercept depends on the permeability at the trans side.
The second protocol uses a gradient applied at each of the two membranes separately. For each membrane, the normalized rate of signal change is related to the water permeabilities by Eq. 14, giving
![]() |
(15) |
![]() |
(16) |
where bl =
ap is the perfusate osmolarity applied at each side.
The third protocol uses a measurement of the sum of apical and basolateral permeabilities with a measurement of the ratio of permeabilities. The sum of permeabilities is determined by simultaneously changing the perfusate osmolarity on the apical and basolateral membranes and measuring the normalized rate of signal change. From Eqs. 11-13, the sum of the water permeabilities is related to Î
![]() |
(17) |
Since the signal intensity is proportional to the relative volume, Pfap/Pfbl is obtained using Eq. 13 and a measurement of the
signal amplitude after changing the perfusate osmolarity, i, at
the apical, and then the basolateral sides (
Iap and
Ibl, respectively):
![]() |
(18) |
where r = Iap/
Ibl. Together, Eqs. 17 and 18 yield the apical
and basolateral water permeability coefficients.
Osmotic Water Transport Measured by Spatial Filtering Microscopy
Initial studies were carried out to validate and optimize
the measurement of water permeability in various cell
types. Fig. 2 A shows the time course of transmitted
light intensity through a monolayer of MDCK cells as
the perfusate osmolarity is changed. By bright field microscopy, there was no detectable signal change; however, by either phase contrast or dark field microscopy,
there was a gradual decrease in signal intensity as cell
volume increased in response to a change in perfusate
osmolarity from 300 to 150 mosmol. The signal was reversed upon return of osmolarity to 300 mosmol and
the cycle could be repeated multiple times (not shown). The signal increased as cell volume decreased in response to a change in perfusate osmolarity from 300 to
600 mosmol. The time course of integrated intensity
(shown for phase contrast microscopy) was temperature dependent (Fig. 2 B) and in agreement with the time course measured for this cell type by TIR microfluorimetry (Farinas and Verkman, 1996). The activation
energy calculated from this data, 12 kcal/mol, is consistent with non-channel-mediated transport of water
across the plasma membrane. The time over which the
transmitted light signal changed was much slower than
the solution exchange time shown in Fig. 2 C. Taken together, these results demonstrate that the phase contrast and dark field signals are sensitive to relative cell
volume. The amplitude of the phase contrast signal was
relatively insensitive to illumination wavelength in the
range 500-700 nm (phase objective optimized for 546 nm) (data not shown).
Fig. 2 D shows that phase contrast microscopy could be used to study cells of different shapes and sizes. Signal changes for epithelial cells were generally in the range of 10-20% for a twofold change in cell volume, but somewhat smaller for cells of low height such as astrocytes. The excellent signal-to-noise ratio for the transmitted light detection (generally >100:1) permitted measurement of relative cell volume changes as small as 1%. Fig. 2 E demonstrates that solute as well as water permeability can be measured by phase contrast microscopy. After cell swelling in response to hypotonic PBS (decreased signal), addition of glycerol to the hypotonic PBS produced a rapid increase in signal due to osmotic water efflux, followed by a slower decrease in signal as glycerol and water entered the cell. A similar biphasic response was found after glycerol removal.
Images of CHO cells were acquired using the phase
contrast microscope (Fig. 3 A). Changing osmolarity
from 300 to 600 mosmol caused cell shrinking and a
corresponding increase in the brightness of the cells.
Changing perfusate osmolarity from 300 to 150 mosmol caused cell swelling and a corresponding decrease
in integrated pixel intensity. The details of the change
in integrated intensity for individual cells was investigated for Sf 9 cells since they have a simple, nearly radially symmetric shape. Images acquired at 400 mosmol
show an increase in intensity of 6% compared with images acquired at 300 mosmol (Fig. 3 B). The change in
the radial distribution of intensity was calculated for a
representative cell (Fig. 3 B, arrow) and is plotted in
Fig. 3 C. The change in intensity occurs throughout the
cell until at large radius the background intensity is
reached. It follows that the integrated intensity can be
measured either by collecting light from the full microscope field and focusing it onto a photodetector, or by
integrating the pixel intensity of a recorded image. In
the latter case, it is possible to measure the response of
individual cells to osmotic gradients.
Validation of Theory Predictions
The theory developed above explains how the phase
contrast or dark field microscope produces a volume-dependent signal. To validate the theory, testable predictions were compared with experimental results. Fig.
4 A shows the dependence of relative signal intensity on relative cell volume predicted by Eq. 10. For a
square cell (hmax = 4 µm, np = 1.334, nc = 1.366, nw = 1.3323, surface coverage = 50%) in a phase contrast
microscope ( =
/2,
= 546 nm, T = 0.09) or a dark
field microscope (T = 0), the integrated intensity is approximately a linear function of relative cell volume
with a slight upward concavity. While the transmittance
of a dark field microscope is 0 and that for a bright
field microscope is 1, phase contrast microscopes can
have any nonzero value of the transmittance. The sensitivity of the signal to cell volume changes (dI/dVr|Vr = 1), increases as the transmittance of the spatial filter decreases (Fig. 4 B). As found experimentally, maximum
sensitivity is obtained for a dark field microscope (T = 0) while a bright field microscope (T = 1) is insensitive. The model predicts a weak dependence of signal
on the wavelength of light (Fig. 4 C). For a given cell shape, the model predicts that the sensitivity is approximately proportional to the difference between refractive indices of the perfusate and water (Fig. 4 D). Practically, this indicates that for a given volume change the
signal size can be increased by increasing the perfusate
refractive index. As expected, the sensitivity increases
for objects with greater cell heights (Fig. 4 E). The sensitivity differs weakly for different cell shapes (trapezoidal, square, or elliptical) having the same degree of surface coverage (70%) and the same initial cell volume
(Fig. 4 F). Taken together, these computations indicate
that the details of the microscope (the transmittance
and wavelength), the cell type (shape and height), and
the perfusate composition (refractive index) affect the
sensitivity of the signal to cell volume. However, in all
cases a signal that is approximately linearly proportional
to relative cell volume is predicted.
Model predictions were confirmed experimentally
using a phase contrast microscope. Fig. 5 A shows a calibration of integrated intensity vs. relative cell volume
for two cell types, Sf 9 and CHO. Relative cell volume
was changed by equilibrating the cells with perfusates
of various osmolarities; assuming that cells act as ideal
osmometers, the relative cell volume equals the reciprocal relative perfusate osmolarity. The nearly linear
(with slight upward concavity) data agrees well with the
theoretical predictions (Eq. 10). As expected from Fig.
4, E and F, the taller Sf9 cells show greater sensitivity
than the CHO cells. To test the effect of perfusate osmolarity on signal size, Sf9 cells were subjected to the
same osmotic gradient (300-400 mosmol) at two different constant refractive indices (np = 1.3363 and 1.3408 for buffers containing 100 mosmol mannitol and 100 mosmol raffinose, respectively). As predicted in Fig. 4
D, Fig. 5 B shows that the signal size increases as the
perfusate refractive index increases. The signal size increased by a factor of 2.05 ± 0.2 (mean ± SEM, n = 3), which was not significantly different from the predicted
value of 1.87 (P > 0.25).
Characterization of Aquaporin-expressing Transfected Cells
Water permeability was measured in CHO cells that
were stably transfected with several mammalian aquaporins and grown on glass coverslips. Previously, water
transport in CHO cells expressing AQP1 and AQP4 was
measured by stopped-flow light scattering (Ma et al.,
1993; Yang et al., 1996
); however, it was difficult to obtain accurate permeabilities because of the cell damage
associated with the suspension procedure and cell settling during the course of the measurement. Representative phase contrast intensity data are shown in Fig. 6 A
for the response of cells to osmotic gradients (300-
150-300 mosmol) at 24°C. The rate of change of intensity was increased more than fourfold in the transfected
cells compared with the control cells. Using Eq. 11 and
an (S/V)o of 5,450 cm
1 (Ma et al., 1993
), water permeability coefficients were low in control (nontransfected)
cells (0.0012 cm/s) and significantly increased for the
CHO cells expressing AQPs 1, 2, 4, and 5 (Fig. 6 B).
Water Permeability in Epithelial Cells
As derived in methods, three protocols enable the
measurement of individual permeability coefficients in
an epithelial cell layer. Fig. 7 shows predictions of the
model to demonstrate several principles. Fig. 7 A indicates that as the size of the gradient applied at one side
of an epithelium is varied, the amplitude and rate of
volume change (and integrated intensity since signal intensity is proportional to relative cell volume) is altered (Pfap = Pfbl = 0.003 cm/s, o =
bl = 300 mosmol, vw = 0.018 l/mol, S/Vo = 2,000 cm
1,
ap as indicated). Fig. 7 B shows that the normalized rate of signal
change is linearly dependent on perfusate osmolarity,
where the slope depends on the permeability at the cis
side and the intercept depends on the permeability of
the trans side (first protocol, Eq. 14). If a gradient is applied only to one membrane, Fig. 7 C shows that the
rate of change of volume depends on the ratio of permeabilities (shown for change at apical side,
o =
bl = 300 mosmol,
ap = 200 mosmol, Pfap + Pfbl = 0.011 cm/s, Pfap/Pfbl as indicated). The half-time for volume
change is faster if the gradient is applied to the membrane of lower permeability because less water flow
must occur to reach the steady state volume. Therefore, measurement of the rate of change of volume after application of a gradient to the apical and then the basolateral membrane provides the individual membrane
water permeabilities (second protocol, Eqs. 15 and 16).
Fig. 7 C also shows that the total volume change is
strongly affected by the ratio of permeabilities. The volume change is greatest when the osmotic gradient is
applied at the membrane of higher permeability (Pfap/
Pfbl = 10). Fig. 7 D shows that simultaneous application
of a gradient to both membranes results in a change in
volume with a rate determined by the sum of the individual permeabilities (
o = 300 mosmol,
ap =
bl = 200 mosmol, S/Vo = 2,000 cm
1, Pfap+Pfbl as indicated). Together, measurements from Fig. 7, C and D
yield the individual membrane permeabilities (third
protocol, Eqs. 17 and 18).
To determine the osmotic water permeability of the
apical and basolateral membranes of an epithelial cell
layer, MDCK cells were grown on porous supports. Several porous supports were tested for their optical properties and effective solution exchange when replacing
solutions bathing the basal cell surface (through the support). It was found that the Cyclopore porous filter
(11 µm thickness, 0.45 µm pore size) did not interfere
with the quality of the phase contrast signal. Furthermore, since the Pfap and Pfbl values measured for AQP5-transfected cells grown on the Cyclopore membranes
were equal and high (as in Fig. 6 A), it was concluded that the filter did not introduce a delay in solution mixing for the dual perfusion chamber used here (data not
shown). Fig. 8 A shows the time course of phase contrast signal for swelling and shrinking of MDCK cells by
replacement of apical and basolateral perfusates. Water
permeability coefficients at 23°C were calculated using
the three protocols described in methods using measured apical and basolateral surface-to-volume ratios of
2,290 and 2,470 cm1, respectively. For the first protocol (Eq. 14), Î values were 0.0296 and 0.0857 s
1 for
changes of 300-150 and 300-600 mosmol at the apical
membrane, and 0.0299 and 0.0564 s
1 for identical
changes at the basolateral membrane. Using Eq. 14,
the average permeabilities were Pfap = 0.0024 ± 0.0007 cm/s and Pfbl = 0.0011 ± 0.0007 cm/s. The average ratio of permeabilities was Pfap/ Pfbl = 2.2. For the second
protocol, Î values were 0.0274 and 0.0286 s
1 for osmolarity changes of 300-150 mosmol at the apical and basolateral membranes, respectively, and 0.0604 and
0.0401 s
1 for osmolarity changes of 300-600 mosmol.
Using Eqs. 15 and 16, Pfap = 0.0019 ± 0.0003 cm/s and
Pfbl = 0.0009 ± 0.0004 cm/s. The third protocol described in methods was used to measure the relative
water permeability coefficients. The ratio of signal
changes for apical and basolateral changes was 1.88 for
changes in osmolarity from 300 to 150 mosmol, and 1.59 for changes in osmolarity from 300 to 600 mosmol.
Eq. 17 gives a ratio of apical-to-basolateral permeabilities Pfap/Pfbl of 1.94, in agreement with that measured
by the first method. The values obtained by the first
protocol were used to compute the predicted transepithelial water permeability for the apical and basolateral
barriers in series, PfTE = 0.00075 cm/s, where [PfTE(S/
Vo)TE]
1 = [Pfap(S/Vo)ap]
1 + [Pfbl(S/Vo)bl]
1.
Recent studies have measured the water permeability
across the alveolar and small airway epithelial barriers;
however, no information has been available for the trachea and larger airways. To measure the water permeability of the apical and basolateral membranes of tracheal epithelia, primary cultures of human trachea were grown on porous supports. Fig. 8 B shows the time
course of phase contrast signal for swelling and shrinking of tracheal cells by replacement of apical and basolateral perfusates. Apical and basolateral surface-to-volume ratios of 1,940 and 1,300 cm1 were measured. Using the first and second protocols, average Pfap and Pfbl
were computed to be 0.0052 and 0.0039 cm/s at 23°C,
respectively. The average ratio of permeabilities Pfap/
Pfbl = 1.3 agrees with that calculated from the third
protocol, Pfap/Pfbl = 1.4. These values were used to calculate the expected transepithelial water permeability
for the tracheal epithelium, PfTE = 0.0022 cm/s. The
time course of cell volume change in tracheal epithelial
cells was notably faster than that observed for the MDCK cells.
The time course of phase contrast signal for a primary culture of tracheal epithelial cells derived from a
cystic fibrosis patient is also shown in Fig. 8 B. Together
with the measured apical and basolateral surface-to-volume ratios of 1,770 and 1,480 cm1, average Pfap and
Pfbl were computed to be 0.0062 and 0.0028 cm/s, respectively. For SK-MES cells, which are a model for
lung cystic fibrosis cells, the measured Pfap and Pfbl at
23°C were 0.0018 and 0.0010 cm/s, respectively.
Vasopressin-regulated Water Permeability in Toad Bladder
The toad urinary bladder has been studied extensively
as a model system for vasopressin-regulated water permeability in kidney collecting duct. It has been assumed that water permeability is rate limited at the apical plasma membrane. To measure basolateral and apical membrane water permeability directly, toad urinary bladders were mounted in a perfusion chamber. Fig. 8
C shows the time course of integrated signal in response to a fivefold dilution of basolateral and then apical perfusate in the absence of vasopressin. A volume-dependent change in intensity was seen for changes in basolateral but not apical osmolarity due to a low value
of Pfap/Pfbl. The data can be used to set an upper limit
on Iap/
Ibl < 0.02, which, using Eq. 17, gives an upper
limit for Pfap/Pfbl < 0.07. Because the basolateral permeability is much greater than the apical permeability,
the apical membrane is essentially impermeable so that
a change in basolateral osmolarity was used with Eq. 11
to calculate Pfbl = 0.036 ± 0.007 cm/s at 23°C. To estimate apical membrane water permeability, oil was
brushed onto the basal surface to prevent volume flow
across this membrane. Under these conditions, toad bladders stimulated with 50 mU/ml of vasopressin
showed a change in signal as apical perfusate osmolarity was changed (Fig. 8 D). Eq. 11 gives a value of Pfap = 0.025 ± 0.009 cm/s at 23°C. From these results, trans-epithelial PfTE = 0.015 cm/s is computed for the vasopressin-stimulated toad bladder epithelium.
The goal of this study was to develop a simple, quantitative method to measure plasma membrane water and
solute permeabilities in cell layers for application to
cells and tissues expressing molecular water channels.
The motivation for this study was the limitation of existing methods and the need to make permeability measurements on native and transfected cell cultures of polarized epithelial cells and intact tissues. Based on the
principle that the optical path length through a cell
layer is sensitive to cell volume (Farinas and Verkman,
1996), we predicted that a phase contrast or dark field
microscope would generate an integrated intensity signal that was sensitive to cell volume. The method was
effective for permeability measurements in a wide variety of cells of different size and shape cultured on solid
or porous supports, as well as in intact epithelial sheets.
Phase contrast or dark field microscopy enabled measurement of stable signals with signal-to-noise ratios
generally exceeding 100:1. The theoretical basis of the
sensitivity of integrated phase contrast signal to cell volume was established and model predictions were validated experimentally. The technical simplicity and
high data quality afforded by the phase contrast approach should make it the method of choice for plasma
membrane water and solute permeability measurements in cell layers, as well as for studies of cell volume
regulation.
The instrumentation required to carry out the permeability measurements includes only a conventional phase contrast or dark field microscope, stabilized monochromatic light source, solid state photodetector, and cell perfusion chamber. Unlike interferometry, high quality phase contrast signals could be obtained with a conventional, incoherent illumination source, although use of a more coherent source would theoretically improve contrast. However, laser speckle pattern would present a potential difficulty. For routine use, an incandescent light source (tungsten-halogen lamp) with a stabilized power supply is recommended because of its superior stability to arc lamps. The high transmitted signal intensities permitted detection by a solid state photodiode. When imaging is not required or microscopes are not available or practical, the method can be implemented by illuminating a cell layer with a laser diode (e.g., a common red laser pointer) and passing the transmitted light through a lens with a small stop placed at its focus to attenuate the zero order light. A second lens is used to focus the higher order beams onto a photodiode. In this way, an inexpensive apparatus could be constructed that uses the principles of spatial filtering to generate volume-dependent signals. Finally, the perfusion chamber is an important component of the instrument. The chamber should permit solution exchange with rapid exchange times, little turbulence to minimize cell trauma, and clear rigid windows for quantitative microscopy. Single and dual channel flow chambers were used that were modeled after chambers designed for in vitro perfusion of isolated kidney tubules. New perfusion chamber designs with millisecond exchange times should permit measurement of fast transport processes with time resolution comparable with stop-flow mixing methods.
The phase contrast method offers considerable advantages over existing methods to measure plasma
membrane transport processes in cell layers in terms of
technical simplicity and data quality. TIR fluorescence
(Farinas et al., 1995) provides accurate relative cell volume values based on dilution of an aqueous-phase fluorescent indicator in the cytoplasm. Interference microscopy (Farinas and Verkman, 1996
) provides information about cell shape and absolute cell volume that are
not obtainable by phase contrast microscopy. However,
unlike TIR, phase contrast microscopy permits measurements on nonadherent cell layers, and unlike interference microscopy, signal detection in phase contrast
microscopy is not sensitive to small temperature gradients, air currents, or variations in the fluid layer thickness of the perfusion chamber. Light intensity signals
from phase contrast microscopy can be related to cell volume by use of a calibration procedure as in Fig. 4,
and the theory (Eq. 10) predicts the functional form of
the phase signal vs. cell volume relation. The method
shares with the light scattering method (Echevaria and
Verkman, 1992; Fischbarg et al., 1993
; McManus et al.,
1993
) the diffraction phenomena to generate an optical, volume-dependent signal. However, compared with
light scattering methods, phase contrast microscopy
has distinct advantages in terms of applicability to cells
of arbitrary shape and size, and the ability to study individual cells by image detection, as well as a rigorous
theory that predicts the magnitude and sign of the observed signal changes.
As in interferometry, the theoretical basis for the dependence of integrated intensity on cell volume involves volume-dependent changes in the optical path length through the cells. No extrinsic probe is required; instead, cell volume-dependent changes in intracellular refractive index are used. In interferometry, the optical path length changes are recorded as differences in intensity of interfering reference and sample light beams. In phase contrast and dark field microscopy, the optical path length changes are recorded as differences in integrated intensity due to spatial filtering of the electric field perturbation introduced by the object.
Model predictions concerning the dependence of the integrated phase contrast signal on perfusate refractive index were tested experimentally. It was found that the integrated signal was approximately linearly dependent on the relative cell volume. The sensitivity of the method was cell-type dependent as expected from differences in cell shape and height. Although excellent signals were obtained for most cell types with the measurement protocol used here, certain cell types, including LLC-PK1 cells, showed relatively poor sensitivity of signal to cell volume. For such cases, the sensitivity could be increased by increasing perfusate refractive index, using a shorter wavelength illumination and dark field microscopy. Since the signal amplitudes were generally large and phase contrast microscopes are more common than dark field microscopes, phase contrast microscopy was used for most of the measurements reported here. The theory developed to explain the observed intensity changes was based on diffraction by the whole cell rather than scattering from intracellular structures. Scattering from intracellular structures could also, in principle, give rise to volume-dependent intensity changes in spatially filtering microscopes. That this was not the case is shown by the fact that the sensitivity was increased by increasing the perfusate refractive index. The perfusate refractive index would not be expected to have any effect on sensitivity if scattering from intracellular structures gave rise to the observed signals.
For simplicity, the theory was developed with the assumption that cell volume change does not result in
cell shape change. Without this assumption, Eq. 10
must be modified by adding a cell volume-dependent
shape dependence to 2:
2
= f(shape,Vr). In general,
this dependence can be quite complex, but mathematically it can be treated as a Taylor's series expansion:
2
=
2 [1 + a1(Vr
1) + a2 (Vr
1)2 + . . .], where aj
are the coefficients of the Taylor's series. The form of
the dependence in Eq. 10 remains unchanged if the
higher order coefficients of the Taylor's series expansion are small (i.e., aj
0 for j > 2). As for the case of
constant cell shape, the signal is approximately linearly
dependent on cell volume and increases as the difference in the refractive index of the perfusate increases.
However, the sensitivity of the method now depends
not only on the details of cell shape but also on the details of cell shape change. Since the experimental results (Fig. 5) show a nearly linear dependence of signal
on cell volume, the assumption that aj
0 for j > 2 is
justified. Therefore, while the precise manner in which
cell shape changes affects the sensitivity of the method,
the general dependence of the signal on cell volume is
still expected to be approximately linear.
The phase contrast method was applied to measure
water permeability in CHO cells that were stably transfected with cloned mammalian aquaporins. Stably
transfected cell lines were established previously for the
erythrocyte water channel (AQP1; Ma et al., 1993) and
the mercurial insensitive water channel (AQP4; Yang et
al., 1996
). In those studies, quantitative measurement
of plasma membrane water permeability required cell
fractionation and water permeability measurement by
stop-flow light scattering because of difficulties associated with light scattering and settling of suspended intact CHO cells. The results here show very high water
permeability in the CHO cells expressing AQPs 1 and
4. Transfected CHO cells expressing the vasopressin-sensitive water channel AQP2 and the lung/gland water channel AQP5 were established here and showed a
similar increase in water permeability compared with control cells. Interestingly, CHO cells expressed AQP2
constitutively at the plasma membrane in the absence
of vasopressin, which is different from the results found
for LLC-PK1 (Katsura et al., 1995
) and CD8 epithelial
cell lines (Valenti et al., 1996
). This finding raises the
possibility that epithelial cells possess factors that participate in the targeting of AQP2-containing vesicles.
A second application of the phase-contrast method
was the analysis of water transport in epithelial cell layers. Several protocols were developed for measurement
of the water permeability of the apical and basolateral
membranes. The measurement could be made either
by applying an osmotic gradient at each side of the epithelium or by varying the magnitude of the gradient applied at a single side. A third protocol could in principle
be used to measure the permeabilities, but it is technically difficult because simultaneous application of a gradient to the basolateral and apical surfaces is required.
The third protocol is best used to measure the relative permeability values. When applied to MDCK cell layers,
low water permeabilities were obtained for the apical
and basolateral membranes, with that for the apical membrane higher than that for the basolateral membrane.
The different protocols yielded similar permeability values. Previous measurement of transepithelial water permeability for MDCK cysts gave a value of 0.00068 cm/s
(Mangoo-Karim and Grantham, 1990), which is similar
to that of 0.00075 cm/s predicted here from the measurements of the individual plasma membrane permeabilities. Recently, Timbs and Spring (1996)
have measured
plasma membrane water permeability of MDCK cells at
37°C and found Pfap and Pfbl to be 0.0011 and 0.0014 cm/s, respectively. The lower time resolution and the
presence of unstirred layers in series with the basolateral membrane in their measurement may explain the somewhat lower values than those found in the present study.
The plasma membrane water permeabilities have
been calculated using the nominal surface area of the
cells without accounting for infoldings. This is the current practice in the field (Timbs and Spring, 1996; Tripathi and Boulpaep, 1989
). The calculated permeability values thus represent an upper bound to the actual permeabilities since invaginations can increase the surface area by three- to fivefold. Cell volume and nominal
surface areas were measured by confocal microscopy of
fluorescently labeled cells. The results of this method
were in agreement with values obtained by interference
microscopy (Farinas and Verkman, 1996
).
The water permeability of individual plasma membranes in tracheal epithelia was measured using primary cultures of human tracheal epithelial cells cultured on porous supports. Immunolocalization studies
show that the molecular water channels AQP3 and
AQP4 are expressed at the basolateral membrane of
tracheal epithelial cells (Frigeri et al., 1995). Elsewhere
in lung, AQP4 is expressed throughout small airways
(Frigeri et al., 1995
) and aquaporins 1 and 5 in alveoli
(Hasegawa et al., 1994
; Raina et al., 1995
). Functional studies in intact lung (Carter et al., 1996
) and isolated
microperfused airways (Folkesson et al., 1996
) have
shown high water permeabilities and evidence of molecular water channels. No information has been available on the water permeability of tracheal epithelia.
Pfap and Pfbl values determined here were 0.0052 and
0.0039 cm/s at 23°C, respectively. The water permeability of the basolateral membrane supports the prediction of a high Pf based on the presence of water channels at this site. It will be interesting to measure Pf in
tracheal cells from transgenic knock-out mice lacking
AQP4 (Ma et al., 1997
). The high water permeability of
the apical membrane and the fact that no known water
channels have been localized to this membrane suggest
that an as yet unidentified water channel may be expressed. The expected transepithelial water permeability for the tracheal epithelium, PfTE = 0.0022 cm/s, was
threefold higher than that for the MDCK cell layer.
Previous measurements on epithelia including mammalian and amphibian proximal tubule and gallbladder gave a ratio of apical to basolateral water permeability less than one (Tripathi and Boulpaep, 1989).
Our results represent the first measurements of water
permeability of individual plasma membranes in several types of nonleaky epithelia. The results show that the
water permeability of the apical membrane is higher
than that of the basolateral membrane. This was found
for tracheal cells as well as SK-MES cells. The high apical membrane water permeability of tracheal cells may
be of physiological importance since it would facilitate the rapid equilibration of the water content of the thin
mucus layer in contact with the apical membrane of the
tracheal cells. The high water permeability of the apical
membrane couples the mucosal extracellular volume
to the cell volume, thereby slowing any changes in the
mucus water content. The calculated transepithelial water permeability of this tissue is sufficiently high so that water permeability does not limit the rate of water secretion or reabsorption (Novotny and Jakobsson, 1996
).
The water permeability of epithelial cells derived from
normal and cystic fibrosis patients was similar. The abnormal hydration of mucus in cystic fibrosis patients is
thus not related to reduced water permeability.
The method was also applied to measure water permeability in intact epithelial tissues. The basolateral
membrane osmotic water permeability of toad urinary
bladder was measured for the first time. In the unstimulated bladder, the basolateral permeability, Pfbl = 0.036 cm/s, was >14-fold higher than the apical permeability. The measured Pfap value of 0.025 cm/s for
the vasopressin-stimulated bladder is consistent with
the value of 0.04 cm/s for a forskolin-stimulated bladder measured by interferometry (Farinas and Verkman,
1996). The calculated value of the transepithelial water
permeability PfTE = 0.015 cm/s for the stimulated bladder is comparable with values measured gravimetrically: 0.023 cm/s (Levine and Kachadorian, 1981
) and
0.028 cm/s (Shi et al., 1990
). The high basolateral Pf is
consistent with the notion that the apical membrane is
the predominant barrier to water flow even under conditions of vasopressin stimulation.
Taken together, the results establish light microscopy with spatial filtering as a technically simple and accurate method for measuring osmotic water permeability in cell layers. The method provided the first measurement of apical and basolateral plasma membranes for human tracheal epithelial cell lines grown on porous supports and for the toad urinary bladder.
Address correspondence to Dr. Javier Farinas, Cardiovascular Research Institute, 1246 Health Sciences East Tower, University of California, San Francisco, San Francisco, CA 94143-0521. FAX: 415-665-3847; E-mail: javier{at}itsa.ucsf.edu
Received for publication 13 January 1997 and accepted in revised form 11 June 1997.
1 Abbreviations used in this paper: CHO, Chinese hamster ovary; Pf, osmotic water permeability; TIR, total internal reflection.We thank Dr. David Agard for useful discussions regarding phase contrast microscopy, Dr. Walt Finkbeiner for providing tracheal epithelial cultures and advice in culturing cells on porous supports, Drs. Ethan Carter and Neil Emans for help in animal tissue studies, and Drs. Baoxue Yang and B.K. Tamarappoo for AQP2 and AQP5 cell transfections.
This work was supported by grants DK-35124, HL-42368, HL-51854, and DK-43840 from the National Institutes of Health and grant R613 from the National Cystic Fibrosis Foundation. Javier Farinas was a graduate student supported by a predoctoral fellowship from the American Heart Association, California Affiliate and a Krevans Fellowship from UCSF. Megan Moore was a summer research student of the American Heart Association, California Affiliate.
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