Estrogen increases the permeability of the cultured human cervical epithelium by modulating cell deformability

George I. Gorodeski

Departments of Reproductive Biology and of Physiology and Biophysics, Case Western Reserve University School of Medicine, Cleveland, Ohio 44106

    ABSTRACT
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Abstract
Introduction
Methods
Results
Discussion
References

Estrogens increase secretion of cervical mucus in females. The objective of this research was to study the mechanisms of estrogen action. The experimental models were human CaSki (endocervical) and hECE (ectocervical) epithelial cells cultured on filters. Incubation in steroid-free medium increased transepithelial electrical resistance (RTE) and decreased epithelial permeability to the cell-impermeant acid pyranine. Estrogen treatment reversed the effects, indicating estrogen decreases epithelial paracellular resistance. The estrogen effect was time and dose related (EC50 ~1 nM) and specific (estradiol = diethylstilbestrol > estrone, estriol; no effect by progesterone, testosterone, or cortisol) and was blocked by progesterone, tamoxifen, and ICI-182780 (an estrogen receptor antagonist). Estrogen treatment did not modulate dilution potential or changes in RTE in response to diC8 or to low extracellular Ca2+ (modulators of tight junctional resistance). In contrast, estrogen augmented decreases in RTE in response to hydrostatic and hypertonic gradients [modulators of resistance of lateral intercellular space (RLIS)], suggesting estrogen decreases RLIS. Estrogen decreased cervical cell size, shortened response time relative to changes in cell size after hypertonic challenge, and augmented the decrease in cell size in response to hypertonic and hydrostatic gradients. Lowering luminal NaCl had no significant effect on RTE, and the Cl- channel blocker diphenylamine-2-carboxylate attenuated the hypertonicity-induced decrease in cell size to the same degree in control and estrogen-treated cells, suggesting estrogen effects on permeability and cell size are not mediated by modulating Na+ or Cl- transport. In contrast, estrogen increased cellular G-actin levels, suggesting estrogens shift actin steady-state toward G-actin and the cervical cell cytoskeleton toward a more flexible structure. We suggest that the mechanism by which estrogens decrease RLIS and increase permeability is by fragmenting the cytoskeleton and facilitating deformability and decreases in cervical cell size.

paracellular permeability; transepithelial transport; cervical mucus; tight junctions; lateral intercellular space; cytoskeleton; G-actin

    INTRODUCTION
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Abstract
Introduction
Methods
Results
Discussion
References

THE MAIN FUNCTION OF THE epithelium that lines the uterine cervix is to control secretion of the cervical mucus. The cervical mucus lubricates the lower genital tract, it prevents the entry of microorganisms and cells into the uterus, and it controls sperm penetration into the cervix and sperm capacitation and migration. Abnormal secretion of cervical mucus may lead to infertility and to states of disease such as mucorrhea and dryness dyspareunia (11).

The cervical mucus is a mixture of mucins and cervical plasma (11). The mucins are secreted into the cervical canal by exocytosis from endocervical cells, and their overall regulation is relatively well understood (11). The cervical plasma makes up 80-99% of the total weight of the cervical mucus, and it is believed to originate by transudation of fluid and solutes from the blood into the cervical canal (11). In contrast to the mucins, relatively little is known about the mechanisms that regulate secretion of the cervical plasma. Part of the difficulty in studying transport phenomena across the human cervical epithelium was the lack of an appropriate experimental system. Most previous studies in women used samples of cervical mucus, but studies in vivo may be inaccurate due to sampling errors and due to contamination with blood and with uterine and vaginal secretions (11). Recently, new systems to culture human cervical cells on filters were described (12, 16). These culture systems were used in the present research to study the regulation of transcervical transport.

Estrogens increase cervical secretions (11), but relatively little is known about the mechanisms of estrogen action in the cervix and about the regulation of cervical plasma. In preliminary experiments, it was found that estrogens increase the permeability of the cultured cervical epithelium. These novel data suggested that estrogens increase transcervical transport by increasing the epithelial permeability (i.e., decreasing the epithelial resistance) to flow of fluid and solutes from the blood into the cervical canal. The objective of the present work was to study the mechanisms by which estrogens regulate transcervical transport.

Human cervical cells form epithelia characterized by high permeability to solutes and fluids, and most of the transport occurs via the intercellular (paracellular) route (12, 15, 16). The theoretical model that best explains paracellular transport is the Ussing-Zerahn model (33). According to this model, movement of molecules in the intercellular space (paracellular pathway) is restricted by the resistances of the tight junctions (RTJ) and of the lateral intercellular space (RLIS). The regions of the tight junction are considered high-resistance elements, due to the occlusion of the intercellular space by the tight junctional complexes (28). In contrast, RLIS is considered a low-resistance element, and it is determined by the proximity of the plasma membranes of neighboring cells and by the length of the intercellular space from the tight junctions to the basal lamina (28). According to the Ussing-Zerahn model, RTJ and RLIS contribute to the total paracellular resistance in series, so that total paracellular resistance equals RTJ + RLIS. Furthermore, because in low-resistance epithelia such as the cultured human cervical epithelium the resistance of the paracellular pathway to passive movement of molecules determines the overall permeability properties of the epithelium (28), the transepithelial resistance (RTE) is approximately equal to RTJ + RLIS. Previous studies in cultured human cervical epithelia showed that RTJ contributes ~75%, while RLIS contributes only 25%, to the total epithelial resistance (12).

Based on these considerations, the specific aims of the present study were to determine the degree to which estrogens decrease RTJ or RLIS and the mechanisms involved. The results indicate that estrogens decrease the RLIS by a mechanism that involves alterations in cell size, enhanced cell deformability, and a shift in cytoskeletal actin toward G-actin.

    METHODS
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Abstract
Introduction
Methods
Results
Discussion
References

Cell cultures. The experiments were conducted on two cell types that represent the two types of cervical epithelia: endocervix and ectocervix. CaSki cells are a stable line of transformed cervical epithelial cells that express phenotypic markers of the endocervix (16). Human ectocervical epithelial (hECE) cells were obtained from minces of normal human ectocervix as described (14) and are a model of the stratified cervical epithelium (16). Experiments were done on passage 3 hECE cells. Cells were grown and maintained in culture dishes at 37°C in a 91% O2-9% CO2 humidified incubator. Methods for growing cells on filters were described (16). Cells were routinely tested for mycoplasma. In most experiments with estrogens, cells on filters were shifted to steroid-free medium (SFM) for 2 days. This medium was composed of phenol red-deficient DMEM-Ham's F-12 or RPMI 1640 (GIBCO, Grand Island, NY) containing 8% heat-inactivated fetal bovine serum that was previously treated with charcoal to remove steroids. Preparation of charcoal-treated serum was described (14); briefly, dextran-coated charcoal (Sigma Chemical, St. Louis, MO) was dissolved at 8% in 0.15 M NaCl, autoclaved, mixed by stirring, and spun, and the pellet was resuspended as 1 g/1.25 ml in H2O. Fetal bovine serum (HyClone, Logan, UT) was mixed with the activated charcoal-dextran at 20:1 (vol/vol) and incubated for 45 min at 55°C. At the completion of incubation, the mixture was spun twice at 800 g for 20 min and the supernatant (serum) was decanted and collected.

Before experiments, filters containing cells were washed three times and preincubated for 15 min at 37°C in a modified Ringer (test) buffer composed of (in mM) 120 NaCl, 5 KCl, 10 NaHCO3 (before saturation with 95% O2-5% CO2), 1.2 CaCl2, 1 MgSO4, 5 glucose, and 10 HEPES (pH 7.4) as well as 0.1% BSA.

Changes in paracellular permeability. Changes in paracellular permeability were determined in terms of changes in the permeability to pyranine (Ppyr) and in terms of changes in the RTE.

Determinations of Ppyr. Pyranine was chosen as a probe to assess paracellular permeability because it is a highly charged trisulfonic acid with a molecular mass of 0.51 kDa (16) and its concentration can be measured down to nanomolar levels by fluorescence techniques. Pyranine traverses epithelia via the paracellular pathway, and it does not permeate cells; cytolysis of dispersed CaSki cells that were previously incubated with 0.1 mM pyranine did not increase pyranine fluorescence significantly above background (not shown). Ppyr was determined from unidirectional (luminal to subluminal) fluxes: pyranine was added to the luminal compartment, and the amount of pyranine in the subluminal compartment was measured after 10 min (16). The transepithelial permeability coefficient (Ppyr) was calculated as previously described (13, 16).

Determinations of RTE. The electrophysiological methods, including conditions for optimal determinations of RTE across low-resistance epithelia, calibrations and controls, potential pitfalls, and the appropriate measures to prevent artifacts were previously discussed by us (15) and by others (28). Changes in RTE were determined continuously across filters mounted vertically in a modified Ussing chamber, as described (15), from successive measurements of the transepithelial potential difference (PD; lumen negative) and of the transepithelial electrical current (I; obtained by measuring the current necessary to clamp the offset potential to zero and normalized to the 0.6-cm2 surface area of the filter), switching between I (pulses of 200-1,400 µA · cm-2) and PD at a rate of 20 Hz: RTE = PD/I.

Determinations of the dilution potential. Determinations of the dilution potential (Vdil) were performed in the Ussing chamber as described (15). Transepithelial Vdil values were determined by measuring the effect of lowering NaCl in the luminal solution on changes in voltage generated across the filter (15). The Henderson diffusion equation (15) was used to interpret the Vdil in terms of ionic permeabilities as uCl/uNa, where uCl and uNa are the mobilities of Na+ and Cl- in the intercellular space. The ratio uCl/uNa ranges from ~1.52 in free solution to 0.7 in tight epithelia (15, 28).

Generation of hydrostatic gradients. Aliquots (1 ml) of test buffer were added to the subluminal compartment, thus establishing a 21-mmH2O hydrostatic gradient in the subluminal-to-luminal direction, which is in the physiological range for capillaries (12).

Generation of osmotic gradients. Extracellular osmolarity in the luminal and subluminal sides was increased by adding aliquots (~120 µl) of 2 M sucrose solution to the subluminal solution, thus establishing a hypertonic gradient in the subluminal-to-luminal direction. Hypotonicity was induced by adding aliquots of a hypotonic buffer (described in Refs. 12, 13) to both the luminal and subluminal solutions. Increasing the tonicity by adding aliquots from concentrated sucrose solution did not affect fluid resistivity (and thus RTE) significantly; in contrast, decreases in electrolyte concentration may decrease fluid resistivity (12, 13). Therefore, in experiments using hypotonic challenge, changes in fluid resistivity were determined, and levels of RTE were corrected (12, 13). All solutions were checked for osmolarity using a model 5100B vapor pressure osmometer (HyClone).

Modulation of extracellular Ca2+. Extracellular Ca2+ was lowered by adding aliquots of EGTA to the luminal and subluminal solutions (17). Concentrations of free Ca2+ were calculated from the stability constants for 37°C and pH 7.4 as described (17).

Flow cytometry. Cells grown on filters were harvested with Hanks' balanced salt solution plus 1 mM EDTA supplemented with 1% trypsin and washed twice with medium containing FCS and three times with isotonic (290 mosmol/l) Ringer buffer without FCS. Cells were brought to a final concentration of 106 cells/ml and kept on ice until assayed and not longer than 15 min. Measurements of forward angle light scatter by flow cytometry were made with an Ortho Cytofluorograph IIs system (Westwood, MA) equipped with a 5-W argon laser operating at 250 mW at 488 nm. Data were acquired with a linear amplifier and converted to digital form. The flow cytometer was calibrated for cell size before each experiment using FluoresBrite beads (0.1-100 µm diameter). Cells were aspirated into the flow cell, and forward light scatter was recorded at each osmolarity (18). Four thousand cells were collected at each osmolarity, and the mean was determined. For measurements of volume changes in response to hypertonic medium, aliquots of 2 M sucrose solution were added. For measurements of volume changes in response to hypotonic medium, aliquots of the hypotonic medium described above (Generation of hydrostatic gradients) were added.

Fluorescence of attached cells. The fluorescence experiments were conducted in a newly custom-designed fluorescence chamber, which was recently described (18). In this apparatus, a filter with cells was placed in an enclosed dark chamber maintained at a fixed temperature and under conditions that permit selective perfusion of the luminal and subluminal compartments. The cells were illuminated over the apical surface, and the intensity of the emitted light from the apical surface was measured as described (18). Cells on filters were incubated in culture medium with 7 µM fura 2-AM plus 0.25% Pluronic F127 for 45 min at 37°C. Following the incubation, cells were washed twice and reincubated with fresh culture medium for 10 min at 37°C to permit hydrolysis of the esters and to retain the polar molecules intracellularly. Measurements of fura 2 fluorescence were made at the isosbestic wavelengths [360-nm excitation/510-nm emission (F360/510) (32)]. Under these conditions, the leakage of fura 2, photobleaching, and metabolization of fura 2 are minimal (18).

The theoretical background for the changes in F360/510 was recently discussed (18). Similar to the principle of F360/510 fura 2 microfluorescence imaging of attached cells (7), changes in F360/510 are not influenced by cytosolic Ca2+ but rather reflect changes in the intracellular concentration of the fura 2 and subsequently reflect changes in cell volume. The explanation is that, in attached and confluent cells, changes in volume are the result of changes both in the cross-sectional plane (the x-y plane, parallel to basal lamina), and in height (the z-axis, from basal lamina to the apical surface). It was suggested that in the new fluorescence chamber the emitted light stems from a single (or from a few) section(s) at the x-y plane. Changes in cell volume, and in particular in cell height, will draw more fura 2 molecules into, or away from, the monitored x-y plane(s), depending on whether the cell volume decreases or increases, respectively. Therefore, if the cell volume decreases, the signal will be stronger; if the cell volume increases, the signal will be weaker.

DNase I inhibition assay. The assay was based on the method of Blikstad et al. (3), with modifications. Cells on filters were washed three times with cold PBS (4°C) and lysed in situ (104 cells/1 µl) in buffer containing 10 mM K2HPO4, 100 mM NaF, 50 mM KCl, 2 mM MgCl2, 1 mM EGTA, 0.2 mM dithiothreitol, 0.5% Triton X-100, and 1 M sucrose (pH 7.0) at 20°C. For determination of the G-actin content, 10 µl of the lysate were added to an assay mixture containing 7 µl DNase I solution [1 mg/10 ml DNase I (bovine pancreas; 600 kU/mg protein; Sigma) in 50 mM Tris · HCl, 10 mM phenylmethylsulfonyl fluoride, and 0.5 mM CaCl2 (pH 7.5)] and 1 ml DNA solution [4 mg/100 ml DNA (herring testes; Sigma) in 100 mM Tris · HCl, 4 mM MgSO4, and 1.8 mM CaCl2 (pH 7.5)]. The DNase I activity was monitored continuously in a quartz cuvette with a Spectronic 20 spectrophotometer (Fisher Scientific, Pittsburgh, PA) at 260 nm between 10 and 40 s after mixing the reagents. An increase in absorbance indicates DNase I-dependent degradation of DNA, and the slope of the increase in absorbance 10-40 s after mixing the reagents is proportional to the amount of active, uninhibited DNase I. Because the main inhibitor of DNase I under the conditions of the experiment is G-actin (3), the slope of the increase in absorbance is inversely proportional to the amount of G-actin in the homogenate. To measure total actin, aliquots of the lysates were diluted three times with lysis buffer and then incubated for 20 min with an equal volume of guanidine hydrochloride buffer [1.5 mM guanidine hydrochloride, 1 M sodium acetate, 1 mM CaCl2, 1 mM ATP, and 20 mM Tris · HCl (pH 7.5)] to depolymerize F-actin to monomeric G-actin. To express the changes in absorbance in terms of G-actin, a standard curve for 30-70% inhibition of DNase I was obtained by measuring the absorbance after adding defined amounts of rabbit skeletal muscle G-actin to the reaction mixture instead of cell lysates. G-actin and total cellular actin data are expressed per milligram of total protein. A linear relationship in the range of 30-70% inhibition of DNase I activity was obtained (not shown), as described (3).

Reagents. All reagents used for permeability/Ussing chamber experiments were added from concentrated stocks (300-1,000×) in either 1% ethanol, DMSO, or saline to both the luminal and subluminal solutions, unless stated otherwise.

Statistical analysis. Data are presented as means ± SD, and significance of differences among means was estimated by ANOVA. Trends were calculated using GB-STAT V5.3 (Dynamic Microsystems, Silver Spring, MD) and analyzed with ANOVA. Best fit of regression equations (least squares criterion) was achieved with SlideWrite Plus (Advanced Graphics Software, Carlsbad, CA), which uses the Levenberg-Marquardt algorithm, and analyzed using ANOVA.

Chemicals and supplies. Anocell (Anocell-10) filters were obtained from Anotec (Oxon, UK). Fluorescent microspheres (FluoresBrite beads, calibration grade) were obtained from Polysciences (Warrington, PA). ICI-182780 was a gift from Dr. Alan Wakeling (Zeneca Pharmaceuticals) (35). All other chemicals were obtained from Sigma.

    RESULTS
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Abstract
Introduction
Methods
Results
Discussion
References

Effects of estrogens on paracellular permeability. CaSki and hECE cells form confluent cultures on filters and express tight junctions that effectively occlude the intercellular space (12, 15-17). Table 1 shows levels of Ppyr and of RTE across blank filters (without cells) and across filters containing confluent cervical cultures. Both CaSki and hECE cells formed epithelia that significantly increased RTE and decreased Ppyr (Table 1). Preplating of 3T3 fibroblasts had no significant effect on permeability (Table 1). In EGTA-treated cells, the levels of RTE and of Ppyr were similar to those across blank filters (Table 1), indicating that lowering extracellular Ca2+ to <0.1 mM abolished the paracellular resistance, probably by disrupting the tight junctions (10, 17, 22). The net RTE that was conferred by the cervical cells was 10 ± 2 Omega  · cm2 for CaSki cells and 18 ± 2 Omega  · cm2 for hECE cells (Table 1, Fig. 1A; P < 0.01 compared with blank filters).

                              
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Table 1.   Ppyr and RTE across hECE and CaSki cultures on filters


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Fig. 1.   Means ± SD (n = 4-6 filters) of effects of steroid-free medium (SFM) and of 17beta -estradiol on levels of transepithelial electrical resistance (RTE; A) and on levels of reciprocal of pyranine permeability (1/Ppyr; B), across cultures of CaSki cells and of human ectocervical epithelial (hECE) cells. Cells were plated on filters in regular medium (RM) and after 3 days were either shifted to SFM or maintained in RM for 2 additional days. Two days before experiments, cells were also treated either with 10 nM 17beta -estradiol (Estrogen), or with vehicle (Control) for a total of 2 days. Levels of RTE were corrected for electrical resistance of filter. All differences between estrogen and control groups were significant (P < 0.02-0.01). All differences between RM and SFM for both estrogen and control groups were significant (P < 0.05-0.01).

Similar effects were obtained relative to Ppyr (Table 1); because permeability relates reciprocally to resistance, the levels of Ppyr were expressed in terms of the inverse of permeability, i.e., 1/Ppyr. Levels of 1/Ppyr were 7 ± 1 s · cm-1 · 104 for CaSki cells and 10 ± 1 s · cm-1 · 104 for hECE cells, significantly higher than the level of 1/Ppyr for blank filters (3 ± 1 s · cm-1 · 104; Table 1; P < 0.01 for both CaSki and hECE cells). These results confirm that human cervical cells form confluent epithelia on filters and restrict the free movement of solutes through the intercellular space. However, compared with other types of cells, cervical cells form epithelia with relatively low levels of RTE and 1/Ppyr (12, 15-17).

Treatment of CaSki cells with 10 nM 17beta -estradiol decreased net RTE from 10 to 6 Omega  · cm2 (Fig. 1A; P < 0.02) and 1/Ppyr from 7 to 3 s · cm-1 · 104 (Fig. 1B; P < 0.02). Treatment with estradiol also decreased net RTE across hECE cells from 18 to 9 Omega  · cm2 (Fig. 1A; P < 0.01) and 1/Ppyr from 10 to 7 s · cm-1 · 104 (Fig. 1B; P < 0.03).

To better understand the effects of estrogen on paracellular resistance, CaSki and hECE cells were incubated in SFM and then treated with 10 nM 17beta -estradiol. Incubation in SFM increased net RTE across CaSki and hECE cells by 9 and 10 Omega  · cm2, respectively, compared with cells grown in regular medium (Fig. 1A; P < 0.02 in both). It also increased 1/Ppyr compared with cells grown in regular medium by 3.5 s · cm-1 · 104 in CaSki cells and by 3 s · cm-1 · 104 in hECE cells (Fig. 1B; P < 0.05-0.02). SFM did not affect CaSki or hECE cell viability (not shown). Treatment of CaSki and hECE cells that were previously incubated in SFM with estradiol decreased net RTE by 10 and 14 Omega  · cm2, respectively, compared with cells that were not treated with the hormone (Fig. 1A; P < 0.01 in both). In cells that were previously incubated in SFM, estradiol also decreased 1/Ppyr in CaSki and hECE cells by 3.5 s · cm-1 · 104 (Fig. 1B; P < 0.02 in both). Based on these results, it is suggested that incubation in SFM increases, whereas treatment with estradiol decreases, paracellular resistance across cultured human cervical epithelia. All subsequent experiments were done on cells that were preincubated in SFM.

The effects of estrogen on RTE across CaSki and across hECE cells were not acute and required ~1 h of incubation with the hormone; maximal decreases in resistance were obtained after 6 h of incubation with estradiol (Fig. 2A). The estrogen effects were dose dependent: decreases in resistance began already with 0.1 nM and saturation was achieved with 10 nM. The EC50 of estradiol was ~1 nM for both CaSki and hECE cells (Fig. 2B). The dose-response curves for both CaSki and hECE cells could be fitted by a modified Hill equation with a Hill coefficient of ~1 for both CaSki and hECE cells (Fig. 2B), suggesting interaction of estradiol with a single class of binding sites.


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Fig. 2.   Time-response (A) and dose-response (B) effects of estradiol on RTE across cultures of CaSki and hECE cells. Cells were plated on filters and grown in regular medium for 3 days. Filters were then shifted to SFM for 2 additional days before experiments. Levels of RTE were corrected for electrical resistance of filter. A: cells were treated with 10 nM 17beta -estradiol for periods of 1-48 h before experiments. B: cells were treated with different concentrations of 17beta -estradiol ranging from 10-10 to 10-7 M for 2 days before experiments. Data of individual experiments were fitted by a modified Hill equation: R = Rmax · 1/{1 + (EC50/[E2])n} + Rmin · (1 - 1/{1 + (EC50/[E2])n}), where R is measured RTE, Rmax and Rmin are maximal and minimal RTE, EC50 is estradiol concentration that produces half-maximal effect, [E2] is concentration of estradiol, and n is Hill coefficient. Shown are means ± SD of 2 experiments, with 2-4 filters per point. Trends were significant for both A and B (P < 0.01).

Diethylstilbestrol, a potent estrogen, could mimic the effects of estradiol on RTE in both CaSki cells and hECE cells (Fig. 3, A and B). The weak estrogens estrone and estriol had significantly smaller effects on RTE in CaSki cells than did estradiol (Fig. 3A) and had no effect in hECE cells (Fig. 3B). Testosterone and cortisol affected neither the RTE nor the estrogen-induced decrease in resistance in either CaSki or hECE cells (Fig. 4, A and B). Progesterone and the estrogen receptor antagonists tamoxifen and ICI-182780 (35) also had little effect on baseline resistance in either CaSki or hECE cells, but all three agents blocked the estrogen-induced decreases in RTE (Fig. 4, A and B).


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Fig. 3.   Means ± SD (n = 3-5 filters) of effects of different estrogens on RTE across CaSki (A) and hECE (B) cultures. Cells were plated on filters and grown in regular medium for 3 days and then shifted to SFM and treated with one of estrogens (100 nM) for 2 additional days before experiments. DES, diethylstilbestrol. Levels of RTE were corrected for electrical resistance of filter. * P < 0.01 compared with cells treated with vehicle (Control).


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Fig. 4.   Means ± SD (n = 3-5 filters) of effects of steroids and estrogen receptor antagonists on RTE across CaSki (A) and hECE (B) cultures. Cells were plated on filters and grown in regular medium for 3 days and then shifted to SFM. Cells were treated with one of following agents for 2 days before experiments, in absence or presence of 10 nM 17beta -estradiol: testosterone, cortisol, tamoxifen, and ICI-182780 (100 nM) and progesterone (1 µM). Levels of RTE were corrected for electrical resistance of filter. * P < 0.01 compared with cells treated with vehicle (Control).

Estrogen decreases RLIS. Changes in paracellular resistance can be the result of changes in RTJ or RLIS (28, 33). To determine whether estrogen modulates RTJ or RLIS, three experiments were done; the experiments were conducted on CaSki cells, which are more stable than hECE cells on filters and do not detach following replacement of the luminal and subluminal solutions (16). However, some experiments were also conducted on hECE cells, with similar results (not shown).

In the first experiment, the effects of treatment with estrogen on the changes in RTE in response to hydrostatic gradients and to hypertonicity were determined. It was previously shown that hydrostatic and hypertonic gradients in the subluminal-to-luminal direction decrease RTE by decreasing RLIS (12). Because estrogen also decreases RTE across cervical cells (Figs. 1-4), it was of interest to determine the degree to which treatment with estrogen can modulate the changes in RTE induced by hydrostatic or hypertonic gradients.

Both hydrostatic gradients and hypertonic gradients decreased RTE, and the decreases in resistance were greater in estrogen-treated cells than in control cells (Fig. 5A, Table 2). The hypertonicity-induced decreases in RTE were slower than the effects of hydrostatic gradients, and they could be described by simple exponential curves (Fig. 5A). The rate of decrease in RTE in estrogen-treated cells was faster than in control cells, with a half time (t1/2) of 53 ± 11 vs. 127 ± 12 s (Fig. 5A; n = 3, P < 0.01). Based on these results, it is suggested that treatment with estrogen modulates the responses to hydrostatic and hypertonic gradients.


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Fig. 5.   Modulation by estrogen of changes in RTE across CaSki cultures induced by transepithelial hydrostatic gradients (HSG), hypertonic gradients (HTG), sn-1,2-dioctanoyl diglyceride (diC8), and low extracellular Ca2+. Cells were plated on filters and grown in regular medium for 3 days. Filters were then shifted to SFM for 2 additional days before experiments in absence (Control) or in presence (Estrogen) of 10 nM 17beta -estradiol. Levels of RTE were corrected for electrical resistance of filter. A: HSG in subluminal-to-luminal direction of 21 mmH2O were established by adding aliquots of buffer to subluminal side. HTG of 290-325 mosmol/l in subluminal-to-luminal direction were established by adding aliquots of 2 M sucrose to subluminal side. Time-related decreases in RTE in response to HTG in both control and estrogen cells were fitted by the simple exponential equation RTE = RTE,min + (RTE,max - RTE,min) · e-t/tau , where RTE is resistance at time t, RTE,max is maximal resistance, RTE,min is minimal resistance, and tau  is the time constant. Half times (t1/2) for responses were extrapolated from tau . B: diC8 was added at a final concentration of 5 µM to both luminal and subluminal solutions. C: Ca2+ in bathing solutions was lowered from 1.2 mM to ~0.6 mM by adding aliquots of 0.3 M EGTA. Experiments were repeated 3 times.

                              
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Table 2.   Modulation by estrogen of changes in RTE across CaSki cultures induced by HSG and HTG in subluminal-to-luminal direction and by diC8 and low extracellular Ca2+

Interestingly, in estrogen-treated cells, hydrostatic gradients followed by hypertonicity (or vice versa) resulted in smaller decreases in resistance than those expected from summation of the two conditions. Thus, in estrogen-treated cells, hypertonicity after a hydrostatic gradient decreased RTE only by 3.6 Omega  · cm2. This level was similar to the effect in cells not treated with estrogen and significantly smaller than the decrease in RTE induced by hydrostatic gradients in estrogen-treated cells not exposed to hypertonicity (Fig. 5A, Table 2). Similarly, a hydrostatic gradient after hypertonic challenge decreased RTE only by 3.1 Omega  · cm2. This level was similar to the decrease in resistance in cells not treated with estrogen and significantly smaller than the decrease in RTE in estrogen-treated cells exposed to hypertonic challenge before the hydrostatic gradient (Fig. 5A, Table 2). Although the resulting resistance after combined hydrostatic and hypertonic gradients was small (~2 Omega  · cm2, Fig. 5A), RTE could be further reduced to levels observed across blank filters (i.e., 0 net RTE) by adding 1.2 mM EGTA (not shown). Based on these results, it is suggested that the effects on RTE of hydrostatic and hypertonic gradients are nonadditive and that the failure of the effects of hydrostatic and hypertonic gradients to summate is not the result of loss of transepithelial resistance.

In the second experiment, we determined the effects of treatment with estrogen on the changes in RTE in response to sn-1,2-dioctanoyl diglyceride (C8:0) (diC8) and to low extracellular Ca2+. DiC8 and low Ca2+ modulate RTJ in cervical cells (15, 17), and the objective was to determine the degree to which estrogen can modulate the changes in RTE induced by these conditions. DiC8 increased RTE by 4.5 Omega  · cm2, whereas low extracellular Ca2+ decreased RTE by 2.3 Omega  · cm2, and the responses were similar in estrogen-treated cells and in control cells (Fig. 5, B and C, Table 2). A hydrostatic gradient applied after the diC8 (Fig. 5B), or after lowering extracellular Ca2+ (Fig. 5C), lowered RTE both in estrogen-treated cells and in control cells; the magnitude of the responses to hydrostatic gradients was similar to those observed in cells not pretreated with diC8 or in cells bathed in normal Ca2+, respectively (compare Fig. 5, A-C; Table 2). Based on these results, it is suggested that treatment with estrogen does not modulate the responses to diC8 or to low Ca2+.

In the third experiment, the effects of estrogen treatment on the Vdil and on uCl/uNa were determined. These parameters were chosen because the mobilities of monoions are influenced by the tight junctions and the cation selectivity reflects the degree of occlusion of the paracellular space by the tight junctions (28). Neither incubation in SFM nor treatment with estradiol affected the Vdil or the uCl/uNa (Table 3), suggesting that estrogen does not significantly modulate the RTJ.

                              
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Table 3.   Effects of SFM and of treatment with 10 nM 17beta -estradiol on Vdil, uCl/uNa, and RTE under conditions of lowered luminal NaCl across CaSki cultures

Based on the results shown in Fig. 5 and in Tables 2 and 3, it is suggested that in human cervical cells estrogen modulates paracellular resistance mainly by changing the RLIS.

Interestingly, lowering luminal NaCl did not affect the differences in RTE induced by estrogen. Changes in RTE under conditions of asymmetrical NaCl concentrations (Table 3) were determined by correcting the measured levels of RTE for fluid conductivity (15). As is shown in Table 3, levels of RTE across CaSki cultures bathed in low luminal NaCl were similar to those across cultures bathed in normal NaCl (Fig. 1A). Based on these results, it is suggested that the changes in RTE induced by SFM and by estradiol are not determined to a significant degree by changes in transcellular transport of Na+ and/or Cl-.

Estrogen modulates changes in cell size. The RLIS depends on the geometry of the intercellular space. Decreases in RLIS are usually the result of increases in the size of the intercellular space and most often are secondary to a decrease in the size of the cells that form this space. A possible mechanism for the estrogen-induced decrease in RLIS (Fig. 5, Tables 2 and 3) would be that estrogens stimulate a decrease in cell size. To test this hypothesis, the effect of estrogen treatment on CaSki cell size was studied using flow cytometry. The mean diameter (± SD) of cells that were grown in SFM was 12.3 ± 0.1 µm; because dispersed CaSki cells in solution have a homogeneous pattern (16), and, if it is assumed that they form spheres, the mean calculated volume of the cell is 974 µm3. This value is similar to that reported for other cell types in solution (e.g., Ref. 16). The diameter of cells that were grown in SFM and treated with estradiol was 11.9 ± 0.2 µm, which was significantly lower than the former value (P < 0.01, n = 12).

One of the conclusions from Fig. 5A is that estrogen augments a decrease in RLIS in response to hypertonic challenge. Decreases in RLIS in response to hypertonicity are usually secondary to loss of cellular water, cell volume decrease, and increases in the volume of the lateral intercellular space (21, 23, 28, 33). A possible explanation for the augmented decrease in RLIS in estrogen-treated cells is that the hormone facilitates a decrease in cell volume in response to hypertonic challenge. To test this hypothesis, three additional experiments were done. First, changes in cell size in response to different osmolarities between 260 and 325 mosmol/l were determined in estrogen-treated and in control cells. Brief exposure of cells to these osmolarities did not exert deleterious effects on the cells (13). Flow cytometric analysis revealed that hypertonicity decreased the mean diameter of cells in solution, whereas hypotonicity increased the mean diameter of cells in solution, in a dose-related manner (Fig. 6). More importantly, in estrogen-treated cells the decrease in cell size following hypertonic challenge in the range of 290-325 mosmol/l was greater (50 ± 9 nm · mosmol-1 · l-1) than in control cells (28 ± 6 nm · mosmol-1 · l-1) (n = 4; P < 0.01).


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Fig. 6.   Effects of osmolar changes on diameter of CaSki cells. Cells were plated on filters and grown in regular medium for 3 days. Filters were then shifted to SFM for 2 additional days before experiments in absence (Control) or presence (Estrogen) of 10 nM 17beta -estradiol. For experiments, cells were harvested from filters; 3 min before cells were loaded into flow cytometer, aliquots of 2 M sucrose solution or hypotonic buffer were added to obtain final designated osmolarities. Shown are means ± SD of 3-4 experiments. Data for changes in cell size in response to different osmolarities were fitted by straight lines (r = 0.97-0.99; P < 0.001 for all 4 lines). For +DPC (dashed lines), cells were preincubated for 10 min at 37°C with 0.5 mM diphenylamine-2-carboxylate.

Second, the experiment was repeated in cells that were pretreated with the Cl- channel blocker diphenylamine-2-carboxylate (DPC). DPC had no significant effect on the size of cells bathed in isotonic buffer or on the responses to hypotonic challenge (Fig. 6). In contrast, DPC attenuated the decrease in cell size in response to hypertonic buffer, both in control cells and in cells treated with estrogen (Fig. 6). However, despite these effects of DPC, the decreases in cell size in response to hypertonic challenge were greater in estrogen-treated cells (41 ± 11 nm · mosmol-1 · l-1) than in control cells (22 ± 9 nm · mosmol-1 · l-1) (n = 3; P < 0.02).

In the third experiment, changes in cell size in response to hypertonic gradients were determined in cells attached to filters. The experiments were conducted in a fluorescence instrument that was custom developed to house the same type of filters that were used for the permeability/Ussing chamber experiments (see METHODS and Ref. 18). Cells on filters were loaded with the fluorescent dye fura 2, and changes in cell size were determined by measuring changes in fluorescence in the Ca2+-insensitive wavelengths (F360/510).

An increase in osmolarity from 290 to 305 mosmol/l increased the F360/510 in a time-related manner both in control cells and in estrogen-treated cells (Fig. 7), and it persisted for at least 10 min (not shown). It was previously shown by us (18) and by others (7) that an increase in F360/510 correlates with an increase in cell size (18), and the responses to hypertonicity shown in Fig. 7 are interpreted as a decrease in mean cell size. Replenishment of isotonic solution resulted in a time-related decrease in F360/510 to baseline levels (Fig. 7), i.e., a return of cell size to baseline level. Lowering the osmolarity from 290 to 275 mosmol/l decreased the F360/510 in a time-related manner both in control cells and in estrogen-treated cells, indicating an increase in mean cell size. However, in contrast to hypertonic challenge, the decrease in F360/510 was transient, despite the continued hyposmolarity, and the fluorescence signal returned spontaneously to baseline within 5 min (Fig. 7).


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Fig. 7.   Effects of osmolar changes on fura 2 fluorescence in isosbestic wavelengths for Ca2+ [360-nm excitation/510-nm emission (F360/510)] in human CaSki cells attached to filters. Cells were plated on filters and grown in regular medium for 3 days. Filters were then shifted to SFM for 2 additional days before experiments in absence (Control) or presence (Estrogen) of 10 nM 17beta -estradiol. Cells were loaded with fura 2, and measurements of fluorescence were made in a fluorescence chamber designed to house filter insert. When indicated (arrows), cells were perfused with a hypertonic solution (in subluminal compartment), with fresh isotonic solution (in both luminal and subluminal compartments), or with a hypotonic solution (in both compartments). Measurements of fluorescence were expressed as arbitrary units (AU) relative to fluorescence obtained when cells were perfused with 290 mosmol/l buffer. Increases in F360/510 in response to hypertonicity were fitted by the simple exponential equation F = Fmax + (Fmin - Fmax)e-t/tau , where F is F360/510 at time t, Fmax is maximal F360/510, Fmin is minimal F360/510, and tau  is the time constant. Decreases in F360/510 in response to isotonic solution were fitted by the simple exponential equation F = Fmin + (Fmax - Fmin)e-t/tau . Values of t1/2 for responses were extrapolated from tau . Experiment was repeated 3 times.

To quantify the changes in F360/510, measurements of fluorescence were expressed relative to the fluorescence obtained when cells were perfused with the isotonic, 290 mosmol/l buffer. The rationale was that, in the range of osmolarities between 260 and 325 mosmol/l, changes in F360/510 in attached cervical cells are linear (21) and correlate with changes in size of dispersed cervical cells (Fig. 6). As is shown in Fig. 7, there were significant differences in the rate of change in F360/510 between estrogen-treated cells and control cells: the t1/2 of the increase in fluorescence in response to hypertonicity was 55 ± 12 s in estrogen-treated cells and 159 ± 28 s in control cells (n = 3, P < 0.01). The t1/2 of the decrease in fluorescence in response to normotonicity was 37 ± 6 s in estrogen-treated cells and 59 ± 10 s in control cells (P < 0.04). There were no differences in the rates of change in F360/510 between estrogen-treated and control cells following hypotonicity (Fig. 7). The fluorescence method was not sensitive enough to determine actual changes in cell size.

Based on the results shown in Figs. 6 and 7, it is suggested that treatment with estrogen decreases the size of cervical cells and facilitates changes in cell size in response to hypertonic challenge.

Estrogen increases G-actin. One of the conclusions from Figs. 5 and 7 was that hypertonic challenge decreased RLIS to a greater degree and faster in cells treated with estrogen than in control cells, suggesting that hypertonicity dilated the intercellular space in estrogen-treated cells to a greater degree than in untreated cells. Because the effect was not significantly dependent on changes in transcellular transport of Na+ and Cl- (Table 3, Fig. 6), another possible explanation is that estrogen modulates the cytoskeletal architecture of cells and reduces their rigidity; this facilitates a decrease in cell size in response to the hypertonicity. Actin filaments are the major component of the cytoskeleton in eukaryotic cells and, in most cells, the density of actin filaments depends on an equilibrium between polymerization of monomeric G-actin and depolymerization of filamentous F-actin (6, 30). Enhanced polymerization of G-actin to form F-actin is usually associated with a more dense and rigid cytoskeleton, whereas depolymerization of F-actin is associated with a more dynamic cytoskeleton. Based on these considerations, the hypothesis was that estrogens shift the equilibrium toward G-actin, thus fragmenting the cytoskeleton and rendering the cell more deformable.

To determine the effect of estrogen on G-actin content in human cervical cells, G-actin and total cellular actin were measured in control and in estradiol-treated CaSki and hECE cells, using the DNase I inhibition assay (see METHODS). In both types of cells, treatment with estradiol did not have a significant effect on total cellular actin level, which was ~150 pg/mg protein (Table 4). In contrast, levels of G-actin in estradiol-treated cells were higher than in untreated cells: the relative G-actin to total actin ratio increased in CaSki cells from 31% in control cells to 47% in estrogen-treated cells (P < 0.01) and in hECE cells from 33 to 56% (P < 0.01). Tamoxifen alone did not have a significant effect on G-actin, but it inhibited the estrogen-induced increase in G-actin in both CaSki and hECE cells (Table 4). Based on these results, it is suggested that estrogen increases G-actin in human cervical cells.

                              
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Table 4.   Effects of treatment with 10 nM 17beta -estradiol on cellular content of G-actin and total cellular actin in CaSki and hECE cells

    DISCUSSION
Top
Abstract
Introduction
Methods
Results
Discussion
References

Estrogens increase the transepithelial permeability of cultured human cervical cells by decreasing the paracellular resistance. Incubation of cells in SFM decreased the permeability, suggesting that agents present in the regular culture medium act on the cells to increase the permeability. These agents may be estrogens, present in the FCS that is used to enrich the culture medium (16), or the phenol red, which has mild estrogenic effects (2). In contrast to the effect of SFM, treatment with estrogens decreased the paracellular resistance. The changes in paracellular resistance induced by estrogen were not associated with changes in cation selectivity. Also, estrogen treatment did not affect the responses to diC8 or to low Ca2+, conditions that modulate RTJ (15, 17). Based on these results, it is suggested that estrogens do not appreciably modulate the RTJ. Because estradiol augmented the effects of hydrostatic and hypertonic gradients, conditions that modulate the RLIS (12, 13, 15), and since the combined effects of hydrostatic and hypertonic gradients were not additive, it is further suggested that estrogens and the hydrostatic and hypertonic gradients modulate a common paracellular mechanism, possibly the RLIS.

One of the conclusions from the present results is that the mechanism by which estrogens decrease the RLIS is modulation of cell size. Treatment with estrogen had three effects on the size of cervical cells: 1) it decreased the size of cells, 2) it shortened the response time relative to changes in cell size after a hypertonic challenge, and 3) it augmented the magnitude of decrease in cell size in response to hypertonicity.

The experiments utilized two methods that yielded complementary data with regard to changes in cell size: measurements of size of dispersed cells and changes of fura 2 fluorescence in the Ca2+-insensitive wavelengths in cells attached to filters. The former method yielded actual values of cell size, which allow for the direct comparison of changes in size between control cells and estrogen-treated cells. The method used dispersed cells, which can create a potential pitfall for the interpretations of the results because dispersed epithelial cells change their shape by rearrangement of the cytoskeleton and form spheres. In addition, the technique can underestimate the actual size of previously attached cells (10). Estradiol reduced cell size, and the consistent differences in cell size between estrogen-treated cells and control cells over the range of 260-325 mosmol/l suggest that the differences are not an artifact due to cell dispersion. Furthermore, hypertonicity produced a greater decrease in size of cells that were previously treated with estrogen than in control cells, and this finding correlates with the results of the fluorescence experiments in attached cells.

The fluorescence experiments utilized measurements of changes in intracellular fura 2 concentrations to determine changes in size of attached cells. One of the advantages is that the method can resolve the time course of the changes in cell size. It is, however, less sensitive than flow cytometry for determinations of actual changes in cell size. The fluorescence experiments revealed three findings. 1) Hypertonic gradients in the subluminal-to-luminal direction produced a sustained decrease in cell size, whereas hypotonic challenge produced a transient increase in cell size. This finding was similar to the effects of these perturbations on RTE (13, 18), suggesting that CaSki cells lack the ability to undergo regulatory volume increase after an acute volume decrease but accomplish regulatory volume decrease after an acute volume increase. 2) The time course of the changes in F360/510 in response to hypotonic challenge was similar in estrogen-treated cells and in control cells. 3) Estrogen treatment facilitated more rapid changes in size in response to hypertonic challenge, compared with cells deprived of estrogen.

The conclusion from the flow cytometry and the fluorescence experiments is that estrogens reduce cell size and facilitate reduction in the size of cervical cells in response to hypertonicity. Acute changes in cell size can be the result of two main mechanisms: acute water shifts, which are usually secondary to acute changes in Na+ or Cl- transport, or rearrangement of cytoskeletal proteins. Incubation in SFM or treatment with estrogen did not modulate levels of RTE in response to lowering luminal NaCl; also, treatment with estrogen did not modulate the effects of the Cl- channel blocker DPC on decreases in cell size in response to a hypertonic challenge. In the latter experiments, DPC attenuated the decreases in cell size both in control cells and in estrogen-treated cells, suggesting that loss of volume of cervical cells following a hypertonic challenge involves augmented Cl- secretion via Cl- channels (20). However, the response to DPC in estrogen-treated cells was additive to the effect of estrogen; it is therefore suggested that the mechanism by which estrogen augments the hypertonicity-induced cell volume loss in cervical cells is not by upregulating Na+ or Cl- transport and consequently not by augmenting water transport per se.

In contrast to the lack of a significant effect on Na+ or Cl- transport, estradiol modulated cellular actin homeostasis in human cervical cells by increasing the ratio of G-actin to total cellular actin, and this effect was also blocked by tamoxifen. Based on these findings, it is suggested that the estrogen-increased permeability in cervical cells is associated with increased deformability of the cells.

Previous studies showed that estrogens modulate cytoskeletal proteins in other tissues (9, 29, 31), but the present study focused on the human cervical epithelium. Actin filaments (F-actin) are made of oriented globular monomeric G-actin molecules that are maintained in a dynamic state of remodeling, which allows cells to change their shape in response to environmental and intrinsic stimuli (6, 30). In most cells, the density of actin filaments depends on an equilibrium between polymerization of monomeric G-actin and depolymerization of filamentous F-actin (6, 30). Enhanced polymerization of G-actin to form F-actin leads to a more dense and rigid cytoskeleton, whereas depolymerization of F-actin is associated with a more dynamic cytoskeleton. Because treatment with estrogen increased G-actin, it is suggested that estrogens shift actin steady-state equilibrium in cervical cells toward G-actin.

Based on this finding, the following model of estrogen regulation of permeability in the cervix is proposed: estrogen increases G-actin, thus fragmenting the cytoskeleton and rendering the epithelial cells more sensitive to a decrease in volume in response to stimuli. These stimuli are either continuous, such as the hydrostatic pressure across the epithelium, or episodic, such as increases in cytosolic Ca2+ in response to secretagogues (e.g., histamine) or neurotransmitters (e.g., ATP; Ref. 15). This new model can be summarized as follows: estrogen right-arrow increased G-actin right-arrow dynamic cytoskeleton and increased cell deformability right-arrow decreased RLIS right-arrow increased paracellular permeability right-arrow increased fluid and solute transport (blood to lumen) right-arrow increased cervical mucus.

Other possible mechanisms by which estrogens can modulate the size of cervical cells, and which have not been studied in the present paper, are changes in membrane permeability (e.g., Ref. 36), modulation of transcellular movement of water (4), and regulation of ion transport mechanisms such as the Na+-K+-ATPase (5), K+ (26) and Ca2+ channels (37), and proton transport (34). Estrogen may also modulate Ca2+ mobilization (22) and subsequently volume changes (6). Some of these mechanisms may depend on nongenomic actions of estrogen (e.g., Refs. 20, 24, 26, 37), which are not supported by the data of the present study (see below).

The pharmacological profile of the estrogen effect, including the time response, the dose response, the specificity, and the inhibition of the effect by progesterone and by estrogen receptor antagonists, is similar to that of estrogen action on the human endometrium (25). It is suggested that the effect is mediated by the estrogen receptor, a conclusion that is supported by the expression of estrogen receptors in human cervical cells (e.g., Ref. 27), including the alpha - and beta -isoforms (unpublished observation).

The experimental data on the inhibition of the estrogen effects by progesterone, tamoxifen, and ICI-182780 are novel and may be important for elucidating the mechanism of estrogen action on cervical paracellular resistance. The effects of these agents may vary according to the type of cell studied, but in the uterus their actions involve modulation of estrogen receptor-dependent regulation of gene activation (10, 19, 35). All three agents blocked the estrogen-induced increase in permeability, and tamoxifen also blocked the estrogen-induced increase in G-actin. Based on these findings, it is suggested that the molecular mechanisms of the effects of estrogen on RLIS, on cell size, and on G-actin involve gene regulation. However, more studies are needed to elucidate those mechanisms.

The present results may be important for understanding the production of cervical mucus in vivo. Our previous studies have suggested that transcervical transport occurs mainly through the paracellular pathway (12, 13, 15, 16). Estrogens increase mucus production in vivo, and until recently the main explanation was that the hormone increases blood flow into the cervix, which increases the hydraulically driven transudation of fluid from the blood through the intercellular space and into the lumen. Based on the present results, it is suggested that estrogens decrease the RLIS and therefore increase the permeability. It is predicted that during the preovulatory phase (a highly estrogenic phase), or in postmenopausal women treated with estrogen, the permeability will increase and more fluid will flux from the blood into the cervical canal. The net result will be an increase in the volume of the cervical mucus. Because progesterone blocked the estrogen effect on RLIS, it is also predicted that progesterone attenuates the effect of estrogen. These predictions are supported by clinical data in women and by experimental studies in animals (11).

The present results are also important for understanding tamoxifen-related changes in cervical mucus secretion. For instance, women with breast cancer who are being treated with tamoxifen experience frequently diminished mucus secretion; this clinical finding can be explained by the present experimental findings that tamoxifen blocked the effects of estrogen on actin homeostasis. This would lead to a more rigid cytoskeleton of cervical epithelial cells and subsequently to decreased permeability and a reduced amount of cervical mucus.

    ACKNOWLEDGEMENTS

This study was supported by National Institute of Child Health and Human Development Grants HD-00977 and HD-29924.

    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: G. I. Gorodeski, University MacDonald Women's Hospital, University Hospitals of Cleveland, 11100 Euclid Ave., Cleveland, OH 44106.

Received 21 April 1998; accepted in final form 10 June 1998.

    REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References

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Am J Physiol Cell Physiol 275(3):C888-C899
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