Ovine male genital duct epithelial cells differentiate in vitro and express functional CFTR and ENaC

Marko Bertog2,*, David J. Smith1,*, Andreas Bielfeld-Ackermann2, John Bassett3, David J. P. Ferguson4, Christoph Korbmacher2, and Ann Harris1

1 Paediatric Molecular Genetics, Institute of Molecular Medicine, Oxford University, John Radcliffe Hospital, Oxford OX3 9DS; 2 University Laboratory of Physiology, Oxford OX1 3PT; 3 Growth and Development Unit, Field Laboratory, Oxford University, Oxford OX2 8QJ; and 4 Department of Pathology, Oxford University, John Radcliffe Hospital, Oxford OX3 9DU, United Kingdom


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

To investigate the biology of the male genital duct epithelium, we have established cell cultures from the ovine vas deferens and epididymis epithelium. These cells develop tight junctions, high transepithelial electrical resistance, and a lumen-negative transepithelial potential difference as a sign of active transepithelial ion transport. In epididymis cultures the equivalent short-circuit current (Isc) averaged 20.8 ± 0.7 µA/cm2 (n = 150) and was partially inhibited by apical application of amiloride with an inhibitor concentration of 0.64 µM. In vas deferens cultures, Isc averaged 14.4 ± 1.1 µA/cm2 (n = 18) and was also inhibited by apical application of amiloride with a half-maximal inhibitor concentration (Ki) of 0.68 µM. The remaining amiloride-insensitive Isc component in epididymis and vas deferens cells was partially inhibited by apical application of the Cl- channel blocker diphenylamine-2-carboxylic acid (1 mM). It was largely dependent on extracellular Cl- and, to a lesser extent, on extracellular HCO-3. It was further stimulated by basolateral application of forskolin (10-5 M), which increased Isc by 3.1 ± 0.3 µA/cm2 (n =65) in epididymis and 0.9 ± 0.1 µA/cm2 (n = 11) in vas deferens. These findings suggest that cultured ovine vas deferens and epididymis cells absorb Na+ via amiloride-sensitive epithelial Na+ channels (ENaC) and secrete Cl- and HCO-3 via apical cystic fibrosis transmembrane conductance regulator (CFTR) Cl- channels. This interpretation is supported by RT-PCR data showing that vas deferens and epididymis cells express CFTR and ENaC mRNA.

ovine genital ducts; cystic fibrosis transmembrane conductance regulator; epithelial sodium channel


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

THE EPIDIDYMIS AND VAS DEFERENS form the proximal and distal parts, respectively, of the secretory duct system of the male reproductive tract. The epithelium lining these ducts secretes and absorbs ions, organic solutes, and water to provide an appropriate luminal environment for normal sperm maturation. Male infertility can result from a number of factors involving the genital ducts. These may be physical, as in the case of congenital absence of the genital ducts, or functional, for example, inadequate fluid secretion. The electrophysiology of genital duct function has been evaluated in a number of model systems, including rodent and human intact ducts and cultured cells. It has been proposed that the epididymis primarily secretes Cl- (27; see Refs. 21 and 44 for review) following a mechanism common to many secretory epithelia. This involves an anion accumulation step at the basolateral side of the epithelium (possibly mediated by an Na+-K+-2Cl- cotransporter, or an Na+/H+ exchanger in parallel with a Cl-/HCO-3 exchanger) with apical anion secretion through ion channels. In addition, the genital ducts have a high luminal K+ concentration, which seems to be important for normal sperm function. In the vas deferens this may be is achieved by a characteristic maxi-K+ channel (34, 35), and the epididymis by an ATP-activated K+ conductance (4). Different species show important variation in the bioelectrical properties and anion secretion along the length of the genital duct system (6).

Male infertility resulting from absence of the vas deferens or epididymal obstruction is one of the characteristic features of cystic fibrosis (CF). It is not clear whether this disruption of the genital ducts is due to malformation during development or to their obstruction by secreted material after the midtrimester of gestation (17, 18). The latter explanation seems most likely but is difficult to investigate in the human fetus in utero. Attempts to investigate the cause of genital duct obstruction in CF with use of CF mouse models have been disappointing, as these animals have intact genital ducts and are not infertile (22).

We previously showed that the CF transmembrane conductance regulator (CFTR) is expressed in the epithelium of human genital ducts from the midtrimester of human gestation onward (9, 16, 38). The CFTR cAMP-activated Cl- conductance is thought to make a significant contribution to Cl- secretion by the genital duct epithelium (27). The absence of CFTR or the presence of a mutant CFTR protein is thought to underlie the genital duct obstruction. Interestingly, the CF mouse genital duct expresses a predominant Ca2+-mediated Cl- channel that may compensate for mutation in CFTR (22). Because the causes of male genital duct abnormalities are impossible to investigate in humans in vivo, the aim of this study was to evaluate whether the ovine genital duct would provide a suitable model to investigate the role of CFTR in genital duct epithelial function.

We already established cell cultures of human fetal genital duct cells and showed these to express CFTR mRNA and CFTR Cl- channels in vitro (16, 17, 27). Epididymis cells have also been cultured from adult human epididymis epithelium (10). In this report we show that cultured ovine epididymis and vas deferens epithelial cells express CFTR mRNA and Cl- channels and also the epithelial Na+ channel (ENaC). These cells provide an excellent culture system for examining the function and regulation of the ovine CFTR gene and the CFTR protein and its role in genital duct abnormalities associated with CF.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. Testicles and genital ducts were removed from 10 male lambs that died perinatally for nonpathological reasons. The lambs were the progeny of Mule ewes mated with Poll Dorset or Charolais rams.

Genital ducts were dissected from the testicular capsule and divided into vas deferens and epididymis; each was minced into 1-mm3 explants and placed in a minimal volume of CMRL 1066 medium with 20% fetal bovine serum, insulin (0.2 U/ml), hydrocortisone (1 µg/ml), cholera toxin (10-10 M), penicillin (100 U/ml), and streptomycin (100 µg/ml) on Primaria flasks (Falcon, Becton Dickinson). Once explant cultures were established, fibroblasts contaminating the epithelial cell cultures were removed before each passage by exposure of the cultures to a short trypsin prewash or by physical disruption with a cell scraper. Cells were passaged by standard trypsinization (0.25% trypsin-1 mM EDTA).

We were not successful in establishing cultures from two sets of genital ducts harvested >12 h postmortem.

Electron microscopy. Cells for transmission electron microscopy were grown to confluence on permeable collagen membranes (Cellagen disks, ICN Biomedical, Aurora, OH), fixed in gluteraldehyde, postfixed in osmium tetroxide, and dehydrated. Samples were then treated with propylene oxide and embedded in epoxy resin. Thin sections were cut and stained with uranyl acetate and lead citrate and then examined with a transmission electron microscope (model 1200EX, Jeol).

Transepithelial electrical measurements. For transepithelial studies, cells were seeded onto 12-mm-diameter Millicell-HA culture plate inserts (Millipore, Bedford, MA). Cells were allowed to grow to confluence, and transepithelial voltage (Vte) and resistance (Rte) were routinely checked using a commercially available epithelial volt-ohm-meter (EVOM) with "chopsticks" electrodes (World Precision Instruments, Sarasota, FL). For the resistance measurements, the EVOM uses an alternating square-wave current of ±20 µA at 12.5 Hz. The chopsticks electrodes were immersed in the cell culture medium bathing the apical and basolateral sides of the cultures under sterile conditions so that cultures could be monitored repeatedly for several days. EVOM measurements were also performed on monolayers grown on Cellagen disks for electron microscopy, and data obtained from Millicell inserts and Cellagen disks were pooled. Some of the Millicell inserts were subsequently transferred to modified Ussing chambers for continuous equivalent short-circuit current (Isc) measurements. Ussing experiments were performed using a computer-controlled clamp device (model CVC 6, Fiebig, Berlin, Germany), as previously described (1). In these Ussing experiments, Rte was evaluated every 20 s by measuring the voltage deflections induced by 200-ms symmetrical square-current pulses of ±25 µA. Open-circuit Vte was also measured, and the equivalent Isc was calculated according to Ohm's law. Conventionally, a lumen-negative Vte corresponds to a positive Isc, which may be due to electrogenic cation absorption and/or electrogenic anion secretion. The bath solution was identical on the apical and basolateral sides of the epithelial monolayer and contained (in mM) 140 Na+, 4 K+, 1 Ca2+, 1 Mg2+, 124 Cl-, 24 HCO-3, and 5 glucose. A reservoir connected to each half-chamber contained bath solution, which was kept at 37°C by a temperature-controlled water jacket. The solution was continuously recirculated from the reservoir through the half-chamber by use of a bubble lift, which gassed the solution with 95% O2-5% CO2, maintaining pH at 7.4. The volume of each half-chamber was 1.8 ml, and the total volume of circulating solution was 10 ml. Drugs were added from stock solutions to the apical or basolateral side of the Ussing chamber. Drugs were washed out by repeated simultaneous removal of 5 ml of solution from both half-chamber reservoirs and readdition of the same volume of fresh prewarmed and pregassed bath solution. This procedure was repeated >= 10 times, which corresponded to a >1,000-fold dilution of the added drug and usually appeared to be an efficient washout, as shown by the reversibility of the amiloride effect (see RESULTS). However, in experiments using basolateral forskolin (10-5 M), washout appeared to be delayed, possibly because of an "unstirred" layer phenomenon within the filter membrane. In some experiments, cholera toxin was omitted from the tissue culture medium 24 h before the cultures were used in Ussing experiments to avoid downregulation of cAMP-mediated pathways by chronic stimulation with cholera toxin.

Amiloride hydrochloride, bumetanide, DIDS, and forskolin were obtained from Sigma-Aldrich (Steinheim, Germany), 5-nitro-2(3-phenylpropylamino)-benzoate (NPPB) from RBI (Natick, MA), and diphenylamine-2-carboxylic acid (DPC) from Fluka (Neu-Ulm, Germany). Amiloride was made up as a 1 mM stock solution in standard bath solution. DMSO stock solutions were prepared from forskolin (1 mM), NPPB (100 mM), bumetanide (20 mM), and DPC (100 mM). Final DMSO concentration in the bath solution was maximally 1%. In control experiments, DMSO at this concentration did not inhibit Isc (n = 3). All stock solutions were freshly prepared on the day of the experiment.

Expression of CFTR and ENaC mRNA. Cells were harvested with a cell scraper, and total RNA was prepared with an RNeasy kit (Quiagen). RNA was extracted from intact genital ducts by guanidinium isothiocyanate extraction (8). RT-PCR was performed as described previously (except RNA was transcribed using Superscript RT), with amplification of the ovine C fragment (bases 1806-2597) of the CFTR cDNA (37). RNA semiquantitation was provided by simultaneous RT-PCR of the housekeeping gene, subunit c of sheep mitochondrial ovine ATP synthase (25).

For the detection of ENaC transcripts, RNA was transcribed using Superscript RT (Life Technologies, Paisley, UK) and random hexamer primers (Pharmacia, Freiburg, Germany). Sequence information for the sheep ENaC is not available. Therefore, primers were chosen that correspond to regions highly conserved between alpha -, beta -, and gamma -subunits of ENaC (3) and were kindly provided by Prof. B. C. Rossier and Dr. L. Schild (Lausanne, Switzerland). The sense 5' primer was 5'-CGCGAATTCGG(C/G)AACTGCT(A/T)CACITT(C/T)AA-3'; the antisense 3'-primer was 5'-CGCGGATCCAT(C/G)T(C/T)(C/G)TC(C/T)TGGAA(A/G)CA-3'. PCR parameters were 95°C for 4 min 15 s, 40 cycles at 95°C for 1 min 15 s, 50°C for 1 min, and 72°C for 2 min, followed by 72°C for 3 min.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell cultures. Cultures of vas deferens and epididymis epithelium were established from 10 pairs of lamb genital ducts. Epithelial cells migrated from explants 48-72 h after establishment of cultures on Primaria flasks (Becton Dickinson). Figure 1 shows the morphology of epididymis (A and B) and vas deferens (C and D) cells in culture. The morphology is similar to that of human genital duct epithelial cells (17), with at least two predominant cell types in culture, one showing a more cuboidal appearance. The cells were routinely passaged with trypsin-EDTA and maintained their differentiated morphology for at least four passages at split ratios of 1:2-3. This yielded 6-8 × 75 cm2 flasks from each genital duct. It was essential to monitor cultures closely for fibroblast contamination at the initial stages. Fibroblasts were removed by a wash in 0.25% trypsin-1 mM EDTA or by physical disruption with a cell scraper. Cells were successfully cryopreserved in 95% fetal bovine serum and 5% DMSO.


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Fig. 1.   Cultures of ovine primary epididymis (A and B) and vas deferens (C and D) epithelial cells. Scale bars, 100 µm (A and C) and 50 µm (B and D).

Further analysis of epididymis and vas deferens epithelial cells grown on permeable collagen supports was carried out by electron microscopy (Fig. 2). When grown on these supports, epididymis (A and B) and vas deferens (C and D) cells were observed to generate a polarized monolayer and to have tight junctions and surface microvilli. In some cultures, myofibroblast contamination was seen below the epithelial monolayer (A).


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Fig. 2.   Electron microscopy of genital duct epithelial cells. A: cross section through epididymis cells cultured on a permeable support, showing monolayer of tightly packed epithelial cells (E) with a number of underlying myofibroblasts (MF). Scale bar, 5 µm. B: detail showing fully formed junctional complex between adjacent epididymis cells. T, tight junction; I, intermediate junction; D, desmosome. Scale bar, 200 nm. C: cross section through vas deferens epithelial cells cultured on a permeable support. Epithelial cells form a polarized monolayer with apical attachments but with dilations along lateral margins (arrows). Scale bar, 5 µm. D: detail of apical junctional complex between adjacent vas deferens epithelial cells. Scale bar, 200 nm.

Figure 2, B and D, shows details of the apical junctional complexes between adjacent epididymis and vas deferens cells, respectively, including tight and intermediate junctions and desmosomes.

Electrophysiological characterization of the epididymis and vas deferens epithelial cells. For these experiments, epididymis cells were derived from different primary cultures and used in passages 2-6. Rte and Vte of cells grown on permeable supports were repeatedly monitored using chopstick electrodes connected to an EVOM. At 4-5 days after seeding, the epididymis cells had developed an average Rte of 1,325 ± 34 Omega  · cm2 and a lumen-negative Vte of -21.1 ± 0.8 mV (mean ± SE, n = 179). In vas deferens monolayers derived from three different primary cultures and seeded onto permeable supports at passage 3, similar Rte and Vte measurements averaged 889 ± 104 Omega  · cm2 and -10.6 ± 1.2 mV (n = 22), respectively, 4 days after seeding. Thus, when grown on a permeable support, epididymis and vas deferens cells formed epithelial monolayers with a high Rte consistent with the presence of tight junctions as detected by electron microscopy (see above). Moreover, the lumen-negative Vte indicated the presence of active transepithelial ion transport, which could be electrogenic absorption of cations and/or electrogenic secretion of anions.

To further characterize the underlying transport mechanisms, we performed continuous Isc measurements on some of these monolayers using modified Ussing chambers that enabled us to test the effect of various drugs. In epididymis cultures mounted in modified Ussing chambers, the initial Rte, Vte, and Isc averaged 774 ± 26 Omega  · cm2, -17.1 ± 0.9 mV, and 20.8 ± 0.7 µA/cm2, respectively (n = 150). The corresponding values in the vas deferens cultures were 625 ± 60 Omega  · cm2, -8.4 ± 0.7 mV, and 14.4 ± 1.1 µA/cm2 (n = 18), respectively. Thus the Rte and Vte values obtained in the Ussing chambers are similar to those measured in tissue culture medium with the EVOM.

To test the possibility that a component of the Isc was due to electrogenic Na+ absorption via amiloride-sensitive Na+ channels, we applied amiloride to the apical side of the monolayers. A typical recording obtained from an epididymis culture is shown in Fig. 3A, where increasing concentrations of amiloride reduced the Isc in a concentration-dependent way. Figure 3B summarizes the results of 19 similar experiments, and a Michaelis-Menten fit of the data reveals a half-maximal inhibitor concentration (Ki) of 0.64 µM. Amiloride was similarly effective in vas deferens epithelial cells with a Ki of 0.68 µM (n = 5; Fig. 3B). This value indicates that the channel responsible for apical Na+ entry into sheep male genital duct cells belongs to a group of highly amiloride-sensitive Na+ channels.



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Fig. 3.   Effect of amiloride (Ami) on short-circuit current (Isc) of cultured ovine epididymis epithelial cells. A: continuous Isc recording demonstrating effect of apical (ap) amiloride application in increasing cumulative concentrations. B: results of experiments similar to those in A are summarized for 19 epididymis cultures () and 5 vas deferens cultures (open circle ). Difference between Isc measured under steady-state conditions and Isc measured in presence of each amiloride concentration was calculated. For each experiment, differences were normalized to difference observed with 1 µM amiloride. Normalized average Isc difference (normalized Delta IAmi) is plotted vs. amiloride concentration. Vertical bars, SE. A Michaelis-Menten fit of data reveals half-maximal inhibitor concentrations (Ki) of 0.64 µM for epididymis and 0.68 µM for vas deferens.

Interestingly, Isc was usually not completely inhibited by amiloride, and an amiloride-insensitive Isc of variable magnitude remained. In epididymis cells grown continuously in the presence of cholera toxin (10-10 M), application of 10-5 M amiloride reduced Isc by 43 ± 8% from 27.0 ± 5.5 to 11.5 ± 1.1 µA/cm2 (n = 11). Thus a rather large portion of the observed Isc was amiloride insensitive. As shown in Fig. 4, apical application of the Cl- channel blocker DPC (10-3 M) to these monolayers inhibited a sizable component of Isc, suggesting that this component is due to anion secretion via apical DPC-sensitive anion channels. In six similar experiments, DPC reduced Isc by 40 ± 1% from 13.3 ± 0.7 to 8.0 ± 0.6 µA/cm2. Sequential application of amiloride (10-4 M) and DPC (10-3 M) largely abolished Isc (Fig. 4).


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Fig. 4.   Expression of an amiloride-sensitive Isc component and one that is sensitive to Cl- channel blocker diphenylamine-2-carboxylic acid (DPC) in cultured ovine epididymis epithelial cells. A continuous Isc recording is shown, and apical application of 10-4 M amiloride and/or DPC is indicated by horizontal bars. In this experiment a large component of Isc remained in presence of a high concentration of apical amiloride. Apical application of 10-3 M DPC inhibited a component of Isc. Application of DPC (10-3 M) in presence of amiloride (10-4 M) almost completely abolished Isc.

In subsequent experiments, epididymis and vas deferens cells were routinely preincubated overnight in culture medium devoid of cholera toxin to minimize chronic stimulation of the cAMP pathway. In this series of experiments, application of 10-4 M amiloride reduced Isc by 39 ± 2% from 20.7 ± 0.7 to 13.4 ± 0.7 µA/cm2 (n = 129) in epididymis cultures and by 57 ± 2% from 14.4 ± 1.1 to 5.9 ± 0.2 µA/cm2 in vas deferens cultures (n = 18). Thus, even in cultures incubated overnight in the absence of cholera toxin, a substantial amiloride-insensitive current component remained. In epididymis cultures, DPC (10-3 M) reduced this spontaneous amiloride-insensitive Isc component by 6.6 ± 1.1 µA/cm2 (n = 24). The relative magnitude of the spontaneous amiloride-insensitive Isc component was rather variable, ranging from 10 to 97% of the total Isc in epididymis cells and from 25 to 58% in vas deferens cells.

In the example shown in Fig. 5, the spontaneous amiloride-insensitive current was relatively small. After washout of amiloride, application of basolateral forskolin (10-5 M) elicited an increase of Isc, with a transient peak and a sustained plateau phase. During the plateau phase, the stimulated Isc component averaged 4.3 ± 1.0 µA/cm2 (n = 3), which corresponded to a 56 ± 19% increase above the baseline level. Sequential application of DPC (10-3 M) and amiloride (10-4 M) had a similar effect, as shown in Fig. 4, largely abolishing Isc. This indicates that forskolin stimulates a DPC-sensitive Isc component.


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Fig. 5.   Identification of a DPC-sensitive Isc component stimulated by basolateral application of forskolin in cultured ovine epididymis epithelial cells. In presence of 10-4 M apical amiloride, Isc was very small and increased after washout of amiloride to a steady-state level. Application of 10-5 M forskolin (FSK) to basolateral (bl) solution caused an increase of Isc with a transient peak and a sustained plateau phase. Forskolin effect persisted after washout; this suggests that washout was incomplete possibly because of forskolin trapped within filter membrane on which cells were grown. Apical DPC and amiloride were applied as indicated.

The experiments shown in Fig. 6 demonstrate that the stimulatory effect of forskolin on Isc is not prevented by amiloride. In the continuous presence of 10-4 M amiloride, application of forskolin elicited a sustained increase of Isc that averaged 3.1 ± 0.3 µA/cm2 (n = 65) in epididymis monolayers and 0.9 ± 0.1 µA/cm2 in vas deferens monolayers (n = 11). In ~50% of the experiments the sustained increase was preceded by a transient peak increase of variable size. The lack of amiloride effect demonstrates that the forskolin-induced Isc increase is not due to stimulation of the amiloride-sensitive Na+ component.


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Fig. 6.   Amiloride does not prevent forskolin-induced stimulation of Isc. Amiloride (10-4 M), 5-nitro-2(3-phenylpropylamino)-benzoate (NPPB, 10-4 M), and DPC (10-3 M) were added to apical side (ap) of cultured ovine epididymis epithelial cells, and forskolin (10-5 M) was added to basolateral side (bl).

After stimulation with forskolin, Isc was inhibited by DPC. In the presence of basolateral forskolin (10-5 M) and apical amiloride (10-4 M), application of apical DPC (10-3 M) reduced Isc by 5.5 ± 0.7 µA/cm2 (n = 35) in epididymis cultures and by 0.9 ± 0.04 µA/cm2 (n = 10) in vas deferens cultures. The Cl- channel blocker NPPB (10-4 M) also had an inhibitory effect when applied before (n = 2; Fig. 6) or after (n = 5) DPC. However, apical application of DIDS (300 µM) had no significant inhibitory effect in vas deferens cultures (n = 6) or epididymis cultures (n = 21; data not shown).

Taken together, these findings demonstrate that sheep male genital duct epithelia express an amiloride-insensitive Isc component that is spontaneously present to a variable degree and may be further stimulated by forskolin. The involvement of cAMP in its activation, its DPC sensitivity, and its insensitivity to DIDS suggests that this Isc component is due to electrogenic anion secretion, possibly via apical CFTR Cl- channels.

Cl- secretion via apical Cl- channels may only occur if Cl- is accumulated intracellularly above electrochemical equilibrium. To this end, Cl- secretory epithelia typically possess a basolateral Na+-K+-2Cl- cotransporter that utilizes the inwardly directed Na+ gradient to transport Cl- into the cell. To test whether in epididymis cultures Na+-K+-2Cl- cotransport may serve as a Cl- entry pathway, bumetanide, a known inhibitor of the Na+-K+-2Cl- cotransport, was used and applied basolaterally at 20 µM. As shown in Fig. 7, application of bumetanide significantly reduced the amiloride-insensitive Isc component by 24 ± 3% from 12.3 ± 1.0 to 8.9 ± 0.5 µA/cm2 (n = 19). This finding suggests that the Na+-K+-2Cl- cotransporter contributes to intracellular Cl- accumulation and, hence, to Cl- secretion. The incomplete inhibitory effect of bumetanide suggests that additional Cl- uptake mechanisms may exist or, alternatively, that Cl- is not the only secreted anion contributing to the amiloride-insensitive Isc component.


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Fig. 7.   Basolateral bumetanide (Bum) reduces amiloride-insensitive Isc component. Amiloride (10-4 M) was added to apical side (ap) and bumetanide (20 µM) to basolateral side (bl) of cultured ovine epididymis epithelial cells.

To further evaluate the ionic basis of the amiloride-insensitive Isc component, anion-substitution experiments were carried out using matched monolayers seeded from the same batch of epididymis cells. Figure 8A shows a control experiment performed in standard bath solution containing Cl- and HCO-3. In this set of control experiments, application of amiloride revealed a spontaneous amiloride-insensitive Isc component that averaged 13.0 ± 1.1 µA/cm2 (n = 16). The amiloride-insensitive current was further increased by forskolin, with a sustained increase averaging 2.8 ± 0.4 µA/cm2 (n = 16). Subsequent application of DPC reduced Isc by 5.1 ± 1.5 µA/cm2 (n = 7; Fig. 8A). The amiloride-insensitive Isc was significantly smaller in cultures maintained in the continuous absence of extracellular Cl- than under control conditions, averaging 3.3 ± 0.4 µA/cm2 (n = 17, P < 0.001; Fig. 8B). This is consistent with a substantial Cl- dependence of the amiloride-insensitive Isc. Interestingly, the stimulatory effect of forskolin was preserved and possibly even enhanced in the absence of extracellular Cl-, with an average sustained Isc increase of 3.9 ± 0.4 µA/cm2 from 3.3 ± 0.4 to 7.2 ± 0.3 µA/cm2 (n = 17). Figure 8B also demonstrates that the effect of forskolin is reversible on washout and that forskolin can elicit a second response in the same experiment. The experiment also shows that DPC inhibits the forskolin-stimulated amiloride-insensitive Isc component, even in the absence of extracellular Cl-, causing an average decrease of Isc by 2.1 ± 0.7 µA/cm2 from 6.9 ± 0.5 to 4.8 ± 0.6 µA/cm2 (n = 7). This suggests that the forskolin-stimulated, Cl--independent and amiloride-insensitive Isc component is also mediated via a DPC-sensitive apical conductance.




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Fig. 8.   Anion dependence of amiloride-insensitive Isc. Representative Isc traces are shown using matched epididymis cultures from same batch of cells. Amiloride and DPC were applied to apical side, and forskolin was applied to basolateral side. A: control experiment performed in standard bath solution containing Cl- (124 mM) and HCO-3 (24 mM). B: experiment performed in continuous absence of apical and basolateral extracellular Cl- (Cl- free). C: experiment performed in absence of extracellular HCO-3 (HCO-3 free) and extracellular Cl- (Cl- free) with subsequent readdition of HCO-3.

To investigate the possibility that the Cl--independent amiloride-insensitive Isc component is due to HCO-3 secretion, additional experiments were performed in the absence of extracellular Cl- and HCO-3 (Fig. 8C). In these experiments the remaining amiloride-insensitive Isc component was minimal, averaging 0.9 ± 0.4 µA/cm2 (n = 17; Fig. 8C). Furthermore, in the absence of extracellular HCO-3 and extracellular Cl-, the stimulatory effect of forskolin was largely abolished, averaging 0.8 ± 0.2 µA/cm2 (n = 17). However, after readdition of HCO-3 to the bath, a forskolin response reappeared (Fig. 8C), with an average Isc increase of 2.3 ± 0.6 µA/cm2 from 2.8 ± 0.7 to 5.1 ± 0.7 µA/cm2 (n = 8). These anion-substitution experiments confirm that in epididymis cultures the amiloride-insensitive Isc component is anion dependent and is likely to represent combined Cl- and HCO-3 secretion.

Taken together, our transepithelial measurements suggest that ovine genital duct epithelial cells are capable of absorbing Na+ via amiloride-sensitive ENaC and of secreting Cl- and HCO-3 via apical anion channels. These latter channels may be activated by an increase in intracellular cAMP, which suggests that they are CFTR Cl- channels.

Expression of CFTR and ENaC. To investigate whether cultured ovine epididymis and vas deferens cells express the CFTR gene and subunits of ENaC, we performed RT-PCR analysis. Figure 9 shows the amplification of the C fragment (bases 1806-2597) of the ovine CFTR cDNA in RNA extracted from three sets of genital ducts established in culture and from intact genital duct tissue. Although the RT-PCR is not quantitative, comparison of the CFTR-derived product with the ovine ATP synthase product amplified in the same reaction gives a measure of the variation in the CFTR levels in different genital duct cell cultures. Figure 10 shows a 370-bp RT-PCR product generated by ENaC-specific primers in RNA from epididymis (lane 4) and vas deferens (lane 5).


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Fig. 9.   Agarose gel showing results of RT-PCR with ovine cystic fibrosis transmembrane conductance regulator-specific primers (C set), producing a 792-bp fragment, and ovine ATP synthase, producing a 530-bp fragment. Lanes 1 and 15, 1-kb ladder; lane 2, no RNA; lane 3, no RT; lane 4, cultured primary fibroblasts from ovine genital ducts (may contain some epithelial cells); lane 5, human fetal primary genital duct cells; lane 6, lamb trachea; lane 7, lamb cecum; lanes 8-10, genital duct RNA from 3 different lambs; lanes 11 and 12, ovine primary epididymis epithelial cell cultures; lanes 13 and 14, ovine primary vas deferens epithelial cell cultures.



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Fig. 10.   Agarose gel electrophoresis of RT-PCR products obtained using epithelial Na+ channel-specific primers producing a 370-bp fragment. Lanes 1 and 6, 1-kb ladder (MBI-Fermentas, St. Leon-Rot, Germany); lane 2, no RNA; lane 3, no RT; lane 4, ovine primary epididymis epithelial cell culture (same sample as Fig. 9, lane 12); lane 5, ovine primary vas deferens epithelial cell culture (same sample as Fig. 9, lane 14).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Male infertility due to absence of the vas deferens or epididymal abnormalities is characteristic of CF. This has generated a research interest in the biology of the male genital duct epithelium in this disease. It is known that CFTR mRNA is expressed from the midtrimester of gestation and that at this gestational age CF genital ducts are generally morphologically normal (13, 16). It is not clear whether the ducts fail to continue normal development during later gestation in CF or whether they become obstructed by secretory deposits, as happens in the pancreatic ducts (16, 29). Epithelial cell cultures that we established from human fetal male genital ducts (17) express the CFTR gene and the CFTR cAMP-activated Cl- channel in addition to other Cl- conductances and a significant maxi-K+ channel (27, 35, 44).

Many aspects of the pathology of CF are difficult to investigate in humans and so necessitate the evaluation of alternative animal models. CF knockout mice have been useful in investigating certain aspects of CF pathology. However, they are inadequate for the study of CF disease in the airway, pancreas, and genital ducts, as they do not exhibit pathology in these organs. The limitations of CF mouse models have led to a search for alternative animal models of CF. Ovine and human development and respiratory physiology have been shown to be similar (11, 26, 39, 40). Furthermore, we have isolated the ovine CFTR cDNA and evaluated its expression through development into adult life (37). The ovine and human CFTR genes show close parallels at the sequence level and in their expression patterns. These similarities, together with recent advances in cloning technology (2, 41), have resulted in the sheep being promoted as a suitable large animal model of CF (15). To investigate the physiology of ovine genital ducts and their similarity to the human vas deferens and epididymis, we have established primary cultures of ovine genital duct epithelial cells.

The cultured cells were morphologically similar to those derived from human genital duct epithelium (10, 17). When cultured on permeable supports, vas deferens and epididymis cells formed monolayers with a high Rte, indicating the presence of tight junctions, as confirmed by electron microscopy. Moreover, our transepithelial measurements provided evidence for the presence of active transepithelial ion transport. We observed an Isc component sensitive to amiloride and another component sensitive to the Cl- channel blockers DPC and NPPB. These findings indicate that the observed basal Isc can be attributed to electrogenic Na+ absorption and Cl- secretion in sheep male genital duct epithelial monolayers.

The epididymis epithelium can function as a Cl- and HCO-3 secretory epithelium (5, 21, 27), and this secretion may be stimulated by a variety of agents known to increase intracellular cAMP (5, 27, 44). The stimulatory effect of forskolin on Isc in ovine epididymis cultures described here is consistent with these observations. The underlying secretory channels are believed to be CFTR Cl- channels, since they are known to be activated by cAMP, and CFTR transcripts have been shown to be expressed in the genital duct epithelium. CFTR Cl- channels are known to be permeable to various polyatomic anions, including HCO-3 (14, 31), with an anion permeability sequence NO-3 > Cl- > HCO-3 > formate > acetate (23). Thus it is conceivable that the HCO-3-dependent secretory Isc component observed in the present study may also be mediated via CFTR Cl- channels. Indeed, it has been estimated that in rat epididymis CFTR may account for 70% of HCO-3 secretion and the Na+-HCO-3 cotransporter may account for 30% (5). Furthermore, single-channel patch-clamp recordings on cultured human fetal epididymis cells have demonstrated the presence of small-conductance forskolin-activated Cl- channels with biophysical properties similar to CFTR Cl- channels (27).

The inhibitory effect of millimolar DPC that we have described is consistent with the inhibition of CFTR Cl- channels and is in agreement with previous findings in rat epididymis (19). The Cl- channel blocker NPPB (100 µM) also partially inhibited Isc in ovine epididymis cells, a finding that is similar to observations on cultured rat epididymis epithelium, where apical NPPB was shown to inhibit Isc with a Ki of ~50 µM (43). NPPB is a potent blocker of large-conductance outwardly rectifying Cl- channels that have been described in excised patches of numerous epithelial preparations, including cultured human fetal epididymis cells (27). Ca2+-activated Cl- channels have been demonstrated in whole cell patch-clamp recordings in rat epididymis cells (7) and may also be inhibited by NPPB. Hence, it is possible that CFTR Cl- channels are not the only channels contributing to Cl- secretion in genital duct epithelia (21). However, the molecular nature of these other Cl- channels is not clear. In this study, the functional evidence for expression of CFTR-associated Cl- channels is supported by the detection of CFTR mRNA in ovine epididymis and vas deferens cells by RT-PCR.

Amiloride has been reported to have no effect on the Isc of cultured rat epididymis epithelial cells (19). In contrast, we detected a large amiloride-sensitive Isc component in cultured ovine epididymis and vas deferens epithelia. Our findings appear to be consistent with the in vivo situation, as studies in isolated and microperfused rat cauda epididymis have demonstrated the presence of amiloride-sensitive Na+ and fluid reabsorption in this tissue with a Ki for amiloride of 1.6 µM (45, 46). Moreover, our data are consistent with recent findings in mouse epididymis cultures, in which apical application of 10-4 M amiloride also inhibited a component of the equivalent Isc. Interestingly, the mouse epididymis cells also showed a cAMP-stimulated Cl- secretory response in addition to the amiloride-sensitive Isc component (22). In sheep genital duct epithelia we have demonstrated a high sensitivity of the Isc to amiloride with a Ki of 0.6-0.7 µM. This suggests that the rate-limiting Na+ entry step occurs via highly amiloride-sensitive apical Na+ channels that most likely belong to the ENaC family (3). This conclusion is further supported by our RT-PCR evidence, which demonstrates the presence of ENaC transcripts in sheep epididymis and vas deferens epithelial cells. Thus our data suggest that ENaC is the channel responsible for Na+ absorption in genital duct epithelia. To our knowledge, similar data are not available for other species, and it will be interesting to see whether ENaC is also expressed in human genital duct epithelia.

Epithelial Na+ channels may be inhibited during activation of Cl- secretion in cells expressing both CFTR and ENaC, which suggests that there is a regulatory relationship between these two conductances (12, 20, 30, 36). However, in the genital duct epithelium, it is not clear whether CFTR and ENaC are separately expressed in different cell types or coexpressed in the same cells where they may interact. This question warrants further evaluation.

In conclusion, we have established cultured ovine epididymis and vas deferens cells, which differentiate into transporting epithelia in vitro and will have multiple applications.

First, they will provide an excellent tool for examination of the transepithelial transport properties of genital duct epithelia, which appear to absorb Na+ via an amiloride-sensitive ENaC and secrete Cl- via apical CFTR Cl- channels. The coexpression of these two channels in genital duct epithelia may be interesting in the light of recent observations that CFTR and ENaC may have a regulatory relationship and that this interaction may be central to CF airway disease. Indeed, it has been suggested that, in normal airway epithelium, CFTR constitutively suppresses Na+ channel activity, whereas in CF airway epithelial cells, i.e., in the absence of normal CFTR, Na+ channels are hyperactive (24, 36). Such a regulatory mechanism may also be important for the pathophysiology of male infertility in CF, where lack of Cl- secretion may cause hyperabsorption of Na+ by the genital duct epithelia.

Second, these cells will provide an additional useful tool in studies to elucidate the mechanisms of regulation of expression of the CFTR gene. One approach to finding the elements conferring tight tissue-specific and temporal regulation of CFTR expression has been to identify DNase I hypersensitive sites associated with the locus (32, 33). This type of approach is rendered much more useful if chromatin is available from primary cell populations that express the CFTR gene endogenously, rather than relying on transformed cell lines. Similarities in the patterns of expression of CFTR in sheep and humans suggest that these two species may share common regulatory elements (37; unpublished data). Evaluation of the ovine CFTR gene control elements in the cultured primary genital duct cells will enable the future in vivo analysis of developmental expression of the CFTR gene.

Third, the similarities between the ovine and human cultured male genital duct cells suggest that once an ovine model of CF becomes available (15), it should be suitable for evaluating the pathology of the genital duct associated with CF and its progression through gestation.


    ACKNOWLEDGEMENTS

We are grateful to Cliff Hanson and Dr. John Cuffe for assistance and Dr. Mike Gray for helpful discussions.


    FOOTNOTES

* M. Bertog and D. J. Smith contributed equally to this work.

This work was supported by the Cystic Fibrosis Research Trust, the Edward Penley Abraham (EPA) Research Fund, the Association Française de Lutte Contre La Mucoviscidose and the Wellcome Trust (D. J. Smith and A. Harris), the Wellcome Trust (A. Bielfeld-Ackermann, C. Korbmacher, and D. J. P. Ferguson), and the Deutscher Akademischer Austauschdienst and the EPA Cephalosporin Fund (M. Bertog).

Part of this work was presented at the North American Cystic Fibrosis Conference, Montreal, in October 1998 and The Physiological Society, Southampton, in September 1998, and published in abstract form [Paediatr Pulmonol Suppl 17: 216-217, 1998, and J Physiol (Lond) 513: 60P, 1998].

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 and other correspondence: A. Harris, Paediatric Molecular Genetics, Institute of Molecular Medicine, Oxford University, John Radcliffe Hospital, Oxford OX32 9DS, UK (E-mail:aharris{at}molbiol.ox.ac.uk).

Received 20 October 1998; accepted in final form 17 December 1999.


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