1Unidad de Reproducción y Desarrollo, Departamento de Ciencias Fisiológicas, Pontificia Universidad Católica, and 2Departamento de Biología, Facultad de Química y Biología, Universidad de Santiago, Santiago, Chile
Submitted 2 October 2003 ; accepted in final form 10 January 2005
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ABSTRACT |
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water channels; epithelial cells; estradiol; progesterone
In many tissues, water channel proteins known as aquaporins (AQPs) have been implicated in transmembrane water transport (2). Eleven mammalian AQPs have been cloned, several of which have been related to physiological processes (2, 52). Moreover, specific mutations of AQPs are responsible of some human and rodent inherited diseases (27).
AQPs have been documented in the reproductive tract of the male and female. In male rats, several AQPs have been detected (3, 5, 8, 40, 46, 48), and their expression has been related to the formation of the seminiferous fluid, processes under steroid hormone control (9, 10, 21, 22, 41). In the female, the presence of AQP1 mRNA has been demonstrated in the human uterus (31) and in the frog oviduct (1). AQP9 mRNA is present in the rat oocyte only throughout proestrus (13), AQP-7, -8, and -9 have been detected in rat granullosa cells (34), and AQP1 has been detected in smooth muscle cells of the rat vagina and uterine tube (16). Recently, the differential expression of various AQPs and AQP mRNAs in the mouse uterus has been detected after ovariectomy and in response to hormonal replacement (24, 43). To our knowledge, the expression of AQPs by oviductal epithelial cells has not been reported; thus elements of the molecular mechanism that control the volume of oviductal fluid in different physiological states remain to be determined.
The work presented herein demonstrates that the oviduct of cycling rats expresses AQP5, -8, and -9 in epithelial cells with differing subcellular localization. Moreover, the expression of AQP5, -8, and -9 is differentially regulated by ovarian hormones. Finally, we present evidence suggesting that the different levels of AQP9 detected in the epithelium of cycling rats correspond to 17-estradiol (E2) and P4 regulation at the mRNA and translation levels, respectively.
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MATERIALS AND METHODS |
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RNA preparation. The rat oviductal epithelium was obtained mechanically following the procedure described by Morales et al. (37). Briefly, oviducts were removed and placed in Hanks' solution at pH 7.4. Oviducts were cut into 4- to 8-mm2 pieces and placed in MEM solution containing 5 mM EDTA for 3040 min at 37°C. Tissue pieces were transferred to sterile Hanks' solution, and the epithelium of each tissue piece was mechanically removed from the rest of the tissue. Total RNA was isolated using acid guanidinium thiocyanate-phenol-chloroform extraction as described by Chomczynski and Sacchi (6). Total RNA was quantified by spectrophotometry, and its quality was checked by electrophoresis in agarose gels. mRNAs were isolated from total RNA by standard procedures using an oligo(dT) cellulose affinity column (Sigma Chemical; St. Louis, MO).
Osmotic water permeability assay. The method used is based on real-time quantitative imaging described by Zhang et al. (54). Oocytes at stages V and VI were harvested from Xenopus laevis and defolliculated by incubation for 13 h in collagenase (1.5 mg/ml type 1, Worthington Biochemicals; Freehold, NJ) in the presence of trypsin inhibitor (0.5 mg/ml type III-0, Sigma Chemical). Oocytes were injected using an automatic Eppendorf microinjection system on the day after isolation with either 100 nl of water or mRNA (2 ng/nl) and incubated at 18°C for 72 h in modified Barth's buffer solution [in mM: 88 NaCl, 1 KCl, 0.82 MgSO4, 0.33 Ca(NO3)2, 0.41 CaCl2, 2.4 NaHCO3, and 10 HEPES; pH 7.4, 200 mosM]. Oocytes were transferred from 200 to 10 mosM of the modified Barth's buffer solution at 25°C, and oocyte swelling was monitored by videomicroscopy using a Nikon inverted phase contrast microscope equipped with a Sony video camera operating at a fixed gain. The images were recorded in a computer through Data Translation hardware boards for analysis. The time course of oocytes swelling was obtained by plotting the relative oocyte volume as a function of time. Relative oocyte volume (V/Vo) was calculated in 250-ms intervals from the relative oocyte area (A/Ao) in the focal plane using the equation V/Vo = (A/Ao)3/2. Oocyte area was determined using Imagepro software. The coefficient of osmotic water permeability (Pf, in cm/s x 104) was calculated from the initial 15- to 30-s response of oocyte swelling using the equation Pf = [Vo x d(V/Vo)/dt]/[S x Vw x (osmin osmout)], where Vo is the initial oocyte volume and equaled 9 x 104 cm3; S is the initial oocyte surface area and equaled 0.045 cm2, Vw is the molar ratio of water and equaled 18 cm3/mol, and osmin is 200 mosM and osmout is 40 mosM. The inhibition of water channel function was determined by incubation of oocytes in Barth's buffer solution containing HgCl2 (0.3 mM) for 5 min before the swelling assay was performed in the presence of HgCl2. The reversibility of water channel function inhibition was determined by incubation of oocytes with 2-mercaptoethanol (5 mM) for 15 min after the HgCl2 treatment.
RT-PCR detection of AQP mRNA.
RT was performed using total RNA (2 µg) previously treated with DNase I amplification grade for 15 min at 25°C. DNase activity was stopped by the addition of 1 µl EDTA (25 mM) and heating to 70°C for 15 min. One microliter of oligo-dT (10 µM) was added, and the mixture was heated for 10 min at 70°C. Seven microliters of a solution containing 1 µl dNTPs (10 mM), 2 µl first-strand buffer (x5), and DTT (100 mM) were added. The reaction mixture was heated for 2 min at 42°C, followed by the addition of 100 units of SuperScript II enzyme. RT was achieved by heating the reaction mixture for 50 min at 42°C and then for 15 min at 70°C. The final volume of the RT reaction was 20 µl. An extra reaction mixture without SuperScript II enzyme was used as a control for DNA contamination. PCR experiments for each AQP and -actin were performed using 2 µl of the RT product plus 23 µl of PCR mix containing specific primers (0.4 µM). Primer sequences were obtained from Ford et al. (13). The PCR amplification cycle consisted of 30 s for denaturing at 95°C, followed by 30 s for annealing with varying temperatures according to the primers used and 45 s for the extension reaction at 72°C. After 40 cycles, final products were extended for 5 min at 72°C. The annealing temperature and the expected product sizes for each cDNA were as follows: AQP2, 56°C and 277 bp; AQP3, 65°C and 645 bp; AQP5, 65°C and 441 bp; AQP8, 60°C and 433 bp; AQP9, 65°C and 374 bp; and
-actin, 56°C and 281 bp. All reagents for RT-PCR were from GIBCO-BRL Life Technologies (Gaithersburg, MD). AQPs and
-actin amplified cDNA fragments were resolved in agarose gels (1.5%) and revealed by ethidium bromide staining, and their electrophoretic migration was compared against a 100-bp DNA ladder (Winkler; Santiago, Chile). Total RNA isolated from the rat kidney, lung, colon, and liver was used as a positive control for the detection of AQP PCR products. PCR products were isolated from a low melting agarose gel and purified using the Wizard PCR Preps DNA purification System (Promega; Madison, WI). Their identity was confirmed to correspond to fragments from rat AQPs by automated sequencing using an ABI Prism310 sequencer (Perkin-Elmer) as described by Muscillo et al. (38).
Relative levels of AQP9 mRNA were measured by semiquantitative RT-PCR using 25 and 24 cycles for AQP9 and -actin, respectively. For an improved quantitative estimation, PCR products were electrophoresed in 12% polyacrylamide gels (47), revealed by silver staining (Winkler), and scanned with a Bio-Rad model GS-700 imaging densitometer (Bio-Rad; Hercules, CA). The optical density of bands was quantified using NIH Image 1.61 software. Values were normalized against those of
-actin.
Animal castration and hormonal treatment.
Animals were anesthetized using a mixture of ketamine-xylacine (80/10 mg/kg im), after which they underwent surgical removal of the ovaries. One week after surgery, ovariectomized rats were injected subcutaneously with either 5 µg E2 (Sigma Chemical) in 100 µl propylene glycol, 5 mg P4 (Sigma Chemical) in 500 µl olive oil, 5 µg E2 followed 6 h later by an administration of 5 mg P4, or 5 mg P4 followed 6 h later by another administration of 5 mg P4. Control animals were injected with the respective vehicles. Twenty-one hours after single hormone administration or 15 h after the second hormone administration, anesthetized rats were killed by cervical dislocation, and their oviducts were removed. The dose of 5 µg E2 was selected as it reflected the physiological estrogen plasma levels (11, 44). Because the ratio of the maximal plasma levels of estrogen with respect to that of P4 during proestrus is 1:1,000 (44), we used 5 mg P4. Furthermore, these doses have been previously reported to produce estrogenic and progestogenic responses in the reproductive tract (20) and other organs (29).
Tissue fixation, immunohistochemistry, and immunofluorescence. To obtain the organs used as positive controls, anesthetized rats were perfused via the left cardiac ventricle with 1x PBS (pH 7.4), followed by 3% paraformaldehyde, 75 mM lysine, and 10 mM periodate (PLP) fixation for 15 min. Organs were removed, minced into small pieces, and further fixed with PLP for 1 h at room temperature. Oviducts were removed from anesthetized rats and separated into ampullar and isthmic sections under a dissection microscope. Each segment was fixed in PLP for 1 h at room temperature. Organ pieces were mounted in tissue freezing medium (Electron Microscopy Science), and cryosections (5 µm thick) were obtained. Endogenous peroxidase activity was blocked by incubation for 15 min at room temperature with 5% H2O2 in methanol. Sections were incubated for 10 min at 80°C in 1 mM Tris·HCl and 0.5 mM EDTA (pH 9.5) for antigen recovery. Sections were blocked for 1 h at room temperature with 1% immunoglobulin-free BSA (Sigma Chemical) dissolved in a mixture of 25% rat serum, 25% goat serum, and 50% TCT [Tris·HCl (pH 7.6), 0.7% carragenan, and 0.25% Triton X-100]. Sections were incubated overnight at 4°C with primary antibody dilution (Alpha Diagnostic) in blocking solution. Tissues were washed three times, 10 min each, with 0.02% Triton X-100 in Tris-buffered saline (TTBS) and then incubated for 1 h at room temperature with secondary goat anti-rabbit IgG biotynilated antibody (DAKO). Tissues were washed three times, 10 min each, with TTBS, followed by incubation for 1 h at room temperature with streptoavidin-peroxidase complex (DAKO). Tissues were further washed with TTBS, and color development was achieved by incubation for 15 min in 0.5 mg/ml diaminobenzidine-0.1% H2O2 in TBS. The reaction was stopped by washing tissue in tap water, followed by distilled water. Tissues were counterstained with hematoxylin (Merck; Darmstadt, Germany), mounted on glass slides with Permount (Merck) and observed under a Nikon labophot-2 microscope with white light illumination. The specificity of the immunoreactivity was assessed by preabsorbing each antibody dilution with its corresponding antigenic peptide.
For colocalization studies, double immunofluorescence assays were done using serum A (see below) to detect AQP9 and a polyclonal goat anti-rat Mucin 1 antibody (Santa Cruz Biothecnology; Santa Cruz, CA) to label the apical plasma membrane or the oviduct. Immunofluorescence was revealed using cy2-labeled and rhodamine-labeled F(ab')2 fragments of goat anti-rabbit and donkey anti-goat IgGs, respectively (Jackson Immuno Research Laboratories;West Grove, PA). Tissue sections were mounted in glycerol containing 1 mg/ml DABCO (1,4-diazabicyclo[2,2,2]octane, Sigma Chemical) dissolved in PBS. Samples were observed under a Zeiss LSM 510 confocal microscope equipped with argon and helio/neon lasers. Excitation wavelengths of 488 and 543 nm, and beam path controls BP 505530 and LP 560 were used for detecting green and red fluorophores, respectively.
Western blot analysis. Total membrane preparations from the rat testis and liver were obtained by tissue homogenization in 7.5 mM phosphate buffer containing 0.3 M sucrose, 1 mM EDTA, 3 mM PMSF, 10 µM leupeptin, 0.7 µM aprotinin, and 7 µM pepstatin. Homogenates were centrifuged at 1,000 g for 10 min at 4°C, and the supernatant was then centrifuged at 200,000 g for 1 h at 4°C. The pellet was dissolved in buffer A containing 20 mM Tris·HCl (pH 8), 5 mM EDTA, and 2% SDS by passing it through a 21-gauge syringe several times. Oviductal epithelial cells were collected in the homogenization phosphate buffer at 4°C and centrifuged for 1 min at maximal speed in an Eppendorf centrifuge. The pellet was suspended in buffer A, and proteins were sonicated in position 1 (Microson ultrasonic cell disrupter, Heat Systems; Farmingdale, NY). Proteins were measured in aliquots by the Lowry method (Bio-Rad). After protein samples (200 µg) were mixed with Laemmli buffer, they were boiled immediately for 4 min and resolved by 12% SDS-4 M urea PAGE. Low-range prestained molecular weight standards were used (Bio-Rad). Gels were then blotted onto nitrocellulose membranes at 300 mA for 80 min. Blots were then incubated in 5% nonfat milk in 0.02 M Tris·HCl (pH 7.5) and 0.5 M NaCl (TBS) and then incubated overnight at 4°C either with an affinity purified rabbit anti-AQP9 polyclonal antibody (Chemicon; Temecula, CA) diluted 1:200 in TBS with 0.05% Tween (TTBS) or with the diluted antibody preabsorbed with a 70 M excess of the antigenic peptide. Blots were rinsed repeatedly with TBS-Tween and then incubated for 1 h at room temperature in goat anti-rabbit IgG antibody conjugated to horseradish peroxidase (AP132P, Chemicon) diluted 1:5,000 in TTBS. Blots were revealed by incubation with a chemiluminescence reagent (Perkin-Elmer). Bands were scanned with an Epson model Expression 636 imaging densitometer. The optical density of bands was quantified using NIH Image 1.61 software. Values of AQP9 levels were normalized against those of total proteins in the respective lane stained with Coomassie blue.
Characterization of the polyclonal serum. Rabbit serum A containing poyclonal antibodies was generated against the synthetic peptide MKAEPSENNLEKHELSVIM corresponding to the COOH-terminal amino acids 277295 of rat AQP9 coupled to hemocyanin (BiosChile Ingeniería Genética; Santiago, Chile). Serum A was applicable for immuhistochemistry but not for Western blot analysis. When tested in sections of the rat testis, this antibody labeled mainly Leydig cells, as described by Elkjaer et al. (8). After antibody preabsortion with a 20 M excess of the antigenic peptide, this label was lost (Fig. 1).
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RESULTS |
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DISCUSSION |
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Herein, we report, for the first time in the mammalian oviduct, the expression of AQP5, -8, and -9 in epithelial cells. The presence of AQP9 in the membrane suggests that it may be, at least, one of the responsible components of water movement toward the lumen of the oviduct. Thus its detection unravels a possible mechanism to the still-unexplained phenomenon of oviductal fluid formation. A similar role in water transport has been suggested for epithelial AQPs in the uterine fluid secretion (24) and for apical AQPs in exocrine glands and respiratory epithelial cells (2). Although AQP2 and -3 have been proposed as responsible for water movement in the uterus (24), our results indicate that this role in the oviduct may rely mainly on AQP9 and probably on AQP5 and -8. This differential pattern of expression of AQPs between the epithelium of the oviduct and uterus is similar to the heterogeneity observed within different segments of the renal (39), respiratory (27), and gastrointestinal epithelium (32). Other epithelial proteins, such as MUC-1, have a varying pattern of expression along the reproductive tract epithelium and during the menstrual cycle, suggesting a correlation with its putative antiadhesive function (25). In a similar manner, AQP5 and -8 are present or absent in the oviduct at different stages of the estrous cycle, and AQP9, although always present, has varied expression levels also within the cycle. As demonstrated in other organs (39, 18, 53), epithelial oviductal cells express more than one AQP. This is possibly related to their differential permeability as AQP5 and -8 are water-selective channels, whereas AQP9 is also permeable to glycerol and other small neutral solutes (50, 2). Apical localization of AQP9 channels and their permeability properties may indicate that their function is related not only to water transport but also to metabolite secretion or absorption from the luminal fluid to maintain the appropriate environment for reproductive functions (26). Alternatively, each AQP may be sorted to different cell membrane compartments to accomplish transcellular water transport, as has been demonstrated in kidney epithelial cells (39). In oviductal epithelial cells, AQP9 localization to luminal plasma membrane is equivalent to that found in epithelial cells along the male reproductive tract (41). Microtubule disruption in the epididymus does not affect its cell surface localization, suggesting that AQP9 membrane insertion is constitutive and does not involve a regulated vesicle trafficking mechanism (41). Currently, AQP5 has been detected only in the apical membrane of epithelial cells in various organs (27, 4). On the other hand, AQP8 has been detected in the basolateral and apical membranes (53, 4) and also in cytoplasmic vesicles in different tissues (15, 7). Recently, it has been demonstrated that in hepatocytes, cytoplasmic AQP8 may redistribute to the plasma membrane after stimulation with glucagon or cAMP (33, 17), suggesting that regulated trafficking to the plasma membrane might also occur in oviductal epithelium. In our system, although it remains to be determined the precise localization of AQP5 and -8 within the epithelial cell, it is possible according to previous reports that AQP5 might localize mainly in the apical membrane (27, 4) and that AQP8 may localize either in the basolateral or apical membrane (4, 33, 53). Furthermore, the variable presence of AQP5 and -8 compared with the permanent expression of AQP9 along the oviductal epithelium during the estrous cycle (Table 1), together with their distinctive permeability properties, suggest a different role for each AQP channel in fluid formation in the oviduct.
Differential levels of AQP9 were detected throughout the estrous cycle with the highest levels detected in estrus and proestrus. The high AQP9 levels correlate with those stages where water content in the oviductal fluid is elevated (30, 23). Cyclic changes in AQP9 levels could result in variations of water permeability of the apical membrane of the oviductal epithelium. This could explain the higher rate of fluid secretion into the lumen of the oviduct detected in estrous compared with the luteal phase in many mammalian species (23). Furthermore, fluid accumulation reaches its maximum around the estrus, but it declines after ovariectomy (23). However, the latter effect can be counteracted by a systemic administration of E2 (35). In our study, we detected that the expression of epithelial AQP5, -8, and -9 was lost in ovariectomized rats. Only AQP9 expression was restored after a specific combination and timing of E2 and/or P4 administrations, further supporting a role for AQP9 in fluid formation. The latter suggests that sex-steroid hormones control AQP9 expression and that other molecules produced by the ovary or by other organ in an ovary-dependent manner could control AQP5 and -8 expression. Similar to our results, castration produces downregulation of AQP9 in rat epididymal epithelium that is restored by androgen replacement (41). Other authors have also reported that not all AQP proteins or mRNAs lost after ovariectomy are restored (43) and that different AQPs are expressed after ovarian hormone replacement (24). These observations indicate that each AQP may need a specific combination or sequence of ovarian hormones to reproduce the physiological conditions required for their expression. However, it remains to be addressed whether water channels formed by de novo synthesis of AQP9 after E2, P4, or combined hormone replacements are functional channels. Interestingly, hormonal regulation of AQPs only occurs in epithelial cells of the uterus but not in other cell types of this organ also expressing AQPs (24). This suggests that the control of water transport relies on cells lining the tubal lumen. Thus further clarification of the contribution of each AQP channel to the transcellular water transport through the genital tract epithelium is needed to understand the variations in water volume at different stages of the reproductive cycle.
The increased AQP9 levels detected under hormonal replacement treatments following ovariectomy did not always parallel changes in mRNA levels, suggesting that E2 and P4 treatments triggered different mechanisms for AQP9 upregulation. We observed that, whereas similar mRNA levels were detected after the administration of E2 alone or of E2 followed by P4, protein levels were further elevated in the latter treatment. Moreover, two consecutive administrations of P4 increased protein levels but slightly decreased AQP9 mRNA levels. Therefore, our results suggest that E2 may trigger a mechanism that increases mRNA levels by activating its transcription rate or inhibiting its degradation. On the other hand, P4 administered after priming with E2 or P4 may trigger a mechanism to activate AQP translation. Similar posttranscriptional mechanisms have been proposed to occur in the estrous cycle-dependent expression of c-Fos in the rat uterine epithelium (36). Also, a posttranscriptional regulation of the E2- and P4-dependent expression of microtubule-associated protein-2 in hippocampal neurons has been described in ovariectomized rats (42). Although a single P4 administration to ovariectomized rats did not induce AQP9 expression, priming with P4 induced AQP9 synthesis to similar levels compared with those detected after a priming injection of E2. This effect on protein expression was not preceded by mRNA increased levels. We discarded the possibility of missing a previous and transient expression, after two consecutive P4 injections, by analyzing AQP9 mRNA levels at smaller time intervals. In accordance with our results, P4 administered alone reduces water channel function in assays utilizing mRNAs isolated from Bufo arenarum oocytes and urinary bladder (12), possibly through a mechanism that reduces the transcription of yet unidentified AQP mRNAs. This is in agreement with previous observations showing that in ovariectomized ewes, fluid formation was not restored after a single progesterone administration (35). However, it has been established the importance of hormonal priming in protein expression and cellular responses to different hormonal treatments. In fact, similar to our observations on the priming effect of P4, it has been reported that E2 administration induces AQP expression in uteri of ovariectomized mice only after P4 priming (43). Furthermore, P4 priming differentially affects P4 single treatment on the proliferative/differentiative response in breast cancer cells (19). These data support our results that indicate a differential response of epithelial cells of the oviduct after P4 administration alone or after a priming hormone dose.
No sex hormone response elements in the AQP9 gene have been described; however, a binding motif for glucocorticoid receptor has been found (51). Therefore, the possibility that sexual hormones directly regulate the transcription of AQPs needs further analysis. Besides a direct effect of hormones on the promoter activity of AQPs, their effect could be mediated through an indirect pathway that involves the expression of other intracellular mediators. Permeability regulators that vary along the reproductive cycle, such as VEGF receptor transcripts and platelet-activating factor receptor expressed by epithelial cells of the oviduct, are possible candidates to explain this mechanism (14, 49).
In this study, we demonstrated, for the first time, the expression of AQP5, -8, and -9 in the oviductal epithelium and that their presence depends on ovarian signals. Moreover, we demonstrate that steroid hormones control AQP9 expression most probably by regulating mRNA levels and protein translation. Taken together, our results provide new evidence to suggest the involvement of AQP water channels in water transport in the oviductal epithelium. Thus the hormonal fine tuning of oviductal fluid characteristics, mediated in part by the regulation of water channels expression, will determine the success of fertilization and early embryonic development.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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. Section 1734 solely to indicate this fact.
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