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Department of Pediatrics (C.R.), University of Kansas Medical Center, Kansas City, Kansas 66160; Department of Pediatrics (J.G.), HuaXi Medical Center, Western China University of Medical Sciences, Chengdu, China 610041; and Department of Pediatrics (N.B., J.R.), Vanderbilt University Medical Center, Nashville, Tennessee 37232-2370
Address all correspondence and requests for reprints to: Jeff Reese, D4106 Medical Center North, Department of Pediatrics, Vanderbilt University Medical Center, Nashville, Tennessee 37232-2370. E-mail: jeff.reese{at}vanderbilt.edu.
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Abstract |
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Introduction |
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The movement of fluids across cell membranes is an important aspect of reproduction. In the female reproductive tract, tissue fluid shifts result in uterine edema at several different stages. The cycling endometrium undergoes recurrent uterine stromal edema in response to hormonal stimuli. The uterus also manifests an inflammatory-like response at the time of insemination that is marked by generalized edema and hyperemia. Most importantly, a dramatic increase in uterine vascular permeability occurs with the onset of implantation and is restricted to the implantation site (9, 10). In rodents, this response is easily detected by uterine uptake of protein-bound dyes or radioisotopes from the intravascular compartment (10). This localized uterine edema coincides with increased expression of vasoactive mediators around the implanting blastocyst (11, 12). However, tissue edema, glandular secretions, and fluid shifts across endothelial and epithelial compartments also occur in the preimplantation period (13, 14) and help prepare the uterus for the onset of implantation. Morphologic assessment of the uterine edema that precedes embryo implantation suggests a "clasping" effect that results in immobilization of the free-floating blastocyst and close approximation of the blastocyst trophectoderm with the uterine luminal epithelium (13, 15). The human uterine lumen is also considered to be closed at the time of implantation, and there are accompanying edematous changes (16, 17). In all mammals, the uterus contains endometrial glands that synthesize or secrete substances that are essential for survival of the developing embryo (18). Absorption and secretion of uterine luminal fluids also facilitate the movement of the free-floating zygote to the eventual site of implantation. Despite extensive studies to resolve the basis of uterine receptivity for embryo implantation (19, 20), there is little information available on specific water and ion transport channels during implantation.
Numerous factors that effect vascular tone and permeability are active during the periimplantation period, including histamine, kinins, platelet-activating factor, prostaglandins and other eicosanoids, and vascular permeability factor/vascular endothelial growth factor (VEGF; Refs. 12 and 21, 22, 23, 24). In addition, structural changes in the receptive endometrium such as the appearance of pinopodes and alterations in apical microvilli may also contribute to tissue fluid shifts during implantation (25, 26). Ovarian estrogen and progesterone influence the expression of many of these factors. Estrogen, in particular, has a marked impact on uterine edema, characterized initially by fluid imbibition and enhanced uptake of macromolecules, followed by a period of cell growth and proliferation. Fluid imbibition may be the result of increased synthesis of vasoactive mediators (11, 12), an increase in pinopodes (26), or structural changes in vascular endothelium (27, 28, 29). Alternatively, estrogen may stimulate fluid movement via specific water-transporting pathways.
At the cellular level, hydrostatic and osmotic forces regulate water balance across membrane barriers via specific ion pumps, ion channels, and solute transport or exchange proteins (30). Recently, the extensive water permeability of renal collecting ducts, erythrocyte membranes, and other epithelia suggested the presence of a specific bulk water transport mechanism. The fortuitous discovery of a red blood cell protein with membrane channel characteristics led to the discovery of the aquaporin (AQP) family of water channels (31, 32). Aquaporins are a conserved group of small hydrophobic integral membrane proteins whose six transmembrane domains are predicted to form barrel-like channels that function as pores for water transport. AQPs exist in plants, bacteria, insects, and diverse members of the animal kingdom. Of the currently known AQPs, AQP3, AQP7, and AQP9 are also permeable to glycerol and other solutes. The presence of uterine edema before implantation and the rapid onset of highly localized vascular permeability surrounding the implanting mouse blastocyst suggest that AQPs might regulate tissue fluid balance during implantation. However, a role for AQPs in the female reproductive tract has not been established. To determine their contribution to uterine fluid homeostasis, we examined the expression and steroid regulation of the 10 known murine AQPs in the periimplantation period.
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Materials and Methods |
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Hormone treatment regimen
To evaluate the effects of estrogen and progesterone on AQP gene expression, adult virgin females were ovariectomized; allowed to recover (7 d); and treated with progesterone, 17ß-estradiol, or a combination of these. One group of ovariectomized females received oil or progesterone (2 mg/mouse, sc, daily) for 3 d and was killed 6 h later. A second group of mice received daily progesterone for 3 d (2 mg/mouse) plus a single dose of estrogen (25 ng/mouse, sc) and was killed 1, 6, and 24 h later. A third group of mice received a single dose of estrogen (25 ng/mouse, sc) and was killed 1, 6, and 24 h later. Uteri were collected and stored at -80 C for later analysis.
RT-PCR
Total RNA was extracted from uteri on d 4 and 5 of pregnancy and from adult eye, lung, testis, kidney, liver, and brain to serve as control tissues (TRIzol, Life Technologies, Inc., Gaithersburg, MD). Reverse transcription was performed by a standard protocol (Superscript II, Life Technologies, Inc.). Two microliters of reverse transcription product were amplified by PCR with primers specific for each AQP transcript or the housekeeping gene, ribosomal protein L7 (rpL7). General thermocycling conditions included denaturation at 95 C for 3 min and 40 cycles of amplification with 94 C for 30 sec, 55 C for 20 sec, and 72 C for 45 sec. AQP-specific primers and cycling conditions were adapted from published literature (33, 34) or generated from known sequence (AQP0 sense 5'-GAA ACC TAG CGC TCA ACA CG-3'; AQP0 antisense 5'-ATT GGA GTC ACT GGG TCT GG-3'). Internally positioned oligonucleotides were generated for each predicted cDNA product. RT-PCR products were visualized in 2% agarose gels and transferred onto nylon membranes. Southern hybridization was performed with 32P-labeled internal oligonucleotides to verify the identity of the amplified cDNAs.
In situ hybridization
To complement the RT-PCR results, AQP0, AQP1, and AQP3–9 amplification products were cloned into a suitable vector (TOPOII, Invitrogen, Carlsbad, CA) and their identity and orientation were determined by T7 or M13-primed sequencing at a core sequencing facility at our institution. These cDNAs were used to generate 35S-labeled sense and antisense cRNA probes for in situ hybridization.
Replicate uterine sections from each time point were mounted with the appropriate controls on the same glass slide. Positive control tissues included adult eye (AQP0), kidney (AQP1, AQP3, AQP6), lung (AQP5), liver (AQP8, AQP9), testis (AQP7), and brain (AQP4). In situ hybridization was performed as previously described (11). Briefly, 10-µm sections of frozen tissues were mounted onto poly-L-lysine-coated slides and fixed in 4% paraformaldehyde/PBS for 10 min at 4 C. Tissue sections were then acetylated, prehybridized, and hybridized at 45 C for 4 h in buffer containing an 35S-labeled antisense cRNA probe. After hybridization and washing, the slides were incubated with RNase A (20 µg/ml) at 37 C for 20 min. RNase A-resistant hybrids were detected by autoradiography using NTB2 emulsion (Eastman Kodak Co., Rochester, NY). Slides were developed after 2- to 10-d exposure periods. Parallel tissue sections hybridized with 35S-labeled sense probes served as negative controls. Sections were briefly poststained with hematoxylin and eosin.
Immunohistochemistry
A goat polyclonal antibody raised against the carboxy terminus of human AQP5 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) was used to examine the cellular localization of AQP5 by immunohistochemistry. Mouse and human epitopes differ by three amino acids in this region and antibody cross-reactivity has been demonstrated with rat and mouse tissues. Frozen sections of mouse uteri from d 4 and 5 were mounted onto poly-L-lysine-coated slides and fixed in Bouin’s solution for 10 min at 4 C. Immunolocalization was performed by serial washing steps, blocking nonspecific staining with 10% nonimmune serum for 10 min, and overnight incubation with 1:500 dilution of primary antibody at 4 C according to the manufacturer’s recommendations (Zymed Laboratories, Inc., San Francisco, CA). Slides were then washed and incubated with secondary antibody for 10 min, briefly exposed to 0.23% periodic acid to block endogenous peroxidase activity, washed, and exposed to peroxidase substrate under direct visualization to determine maturity of the reaction. Immunoreactive protein was detected as red-brown deposits. Sections were lightly counterstained with hematoxylin.
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Results |
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). Faint bands were also visible for AQP0, AQP3, AQP6, AQP7, and AQP9 after longer exposures of Southern blots (data not shown), whereas there was no evidence of AQP2 mRNA expression. Uterine RT-PCR products for AQP0, AQP1, and AQPs 3–9 were gel purified and cloned. Sequence analysis confirmed the correct amplification of each target gene.
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). AQP1 mRNA was expressed in the myometrial smooth muscle on d 1, 4, 4.5, and 5 of pregnancy. There was no AQP1 localization in the smooth muscle layers of the large mesometrial blood vessels or smaller branch vessels that penetrate the layers of the myometrium. AQP1 expression in the uterine myometrium was more prominent in the inner circular layers, although less prominent signals were noted in longitudinal layers. There was also faint accumulation of AQP1 signals in stromal regions around the luminal epithelium on d 1, 4.5, and 5. It was unclear whether these regions represent the microvascular bed within the stroma or other specific structures. There was no localization of AQP1 in stromal or luminal epithelial cells in the immediate vicinity of the implanting blastocyst. AQP4 signals were localized only in the luminal epithelium on d 1 and 4 of pregnancy, although less hybridization was noted on d 4. There were no specific AQP4 signals in the glandular epithelium or near the implanting blastocyst. Diffuse signals were noted in the uterine stroma on d 5, but AQP4 sense probes showed a similar pattern, suggesting nonspecific hybridization. AQP5 mRNA expression was restricted to the glandular epithelium on d 1, 4, 4.5, and 5 of pregnancy. In comparison with the modest localization on d 1, intense accumulation of AQP5 signals was noted in the glands on d 5. Longitudinal sections that included implantation and interimplantation regions (data not shown) revealed the same pattern of AQP5 expression in the glandular epithelium along the length of the uterus and demonstrated that AQP5 transcripts were not restricted to the site of implantation.
RT-PCR also detected the expression of AQP8 and AQP9 mRNAs. However, autoradiographic signals were not localized in d 5 uteri by in situ hybridization. Instead, AQP8 and AQP9 mRNAs were expressed in the implanting blastocyst (Fig. 2). In the embryo, AQP8 was localized to the inner cell mass but not the surrounding mural or polar trophectoderm. AQP9 was localized to the mural trophectoderm, with sparing of the polar trophectoderm and inner cell mass. Serial sections and evaluation of implantation sites from different females confirmed this pattern of expression. Sections through d 8 implantation sites revealed highly restricted expression of AQP8 mRNA in embryonic endoderm and in the mesometrial decidualizing stroma at the site of future placenta formation. AQP8 was also expressed in the receding uterine glands at the periphery of the antimesometrial decidua (Fig. 2). Low levels of AQP9 accumulation were localized in the mesometrial decidua at this time but were not present in the developing embryo.
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) but not on d 4 (data not shown). There was minimal AQP5 protein in the myometrium, luminal epithelium, or stroma. The surface of the blastocoele cavity also appeared to contain low levels of AQP5 in comparison with the accumulation of AQP5 in the nearby glands. In all sections taken, high-power magnification suggested preferential localization of AQP5 protein in the basal region of the glandular epithelium (Fig. 3).
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), similar to that observed in pregnant females (Fig. 1B). Among those animals that received estrogen alone, a biphasic response was observed. After 1 h of estrogen exposure, a general increase in the intensity of AQP1 signals was observed in the inner circular myometrial layers. Animals that were killed after 6 h of estrogen treatment showed an overall reduction in myometrial AQP1 signals but the presence of scattered AQP1 expression in the uterine stroma. After 24 h of estrogen exposure, this pattern was accentuated, and the distribution of AQP1 expression localized to the small blood vessels throughout the uterine stroma. In the group of animals that received both a priming dose of progesterone and varying lengths of estrogen exposure, a similar biphasic response was observed, with initial AQP1 expression in the myometrium followed by a shift to the uterine stromal vascular bed. There was no AQP1 expression in the luminal or glandular epithelium.
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). Animals that received progesterone or estrogen treatment alone did not show any uterine AQP5 expression. The most intense accumulation of AQP5 signals appeared in the glandular epithelium of progesterone-primed uteri after 6 h of estrogen exposure. After 24 h of estrogen, APQ5 signals were less prominent but remained restricted to the glandular epithelium. The influence of estrogen was dependent on prior exposure to progesterone because animals that were treated with estrogen alone did not express AQP5, even after 6–24 h of exposure. There was no evidence of AQP5 expression in the myometrium or stromal vasculature.
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Discussion |
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Uterine edema is a general feature of implantation in rodents, nonhuman primates, and humans (14, 16, 35). An initial phase of generalized uterine swelling facilitates closure of the uterine lumen around the free-floating blastocyst (13, 14, 36). A second phase of localized, prominent edema occurs at implantation sites that coincide with the onset of increased capillary permeability around the implanting blastocyst (10). There are several molecular mechanisms that are associated with each edematous response. Prostaglandins produced by cyclooxygenase (COX) are implicated in the production of periimplantation edema because they are inflammatory in nature, they are increased at implantation sites, and uterine edema and implantation are suppressed by inhibitors of prostaglandin synthesis (24). In addition, the constitutively expressed COX-1 isoform is localized to the uterine luminal epithelium before attachment, whereas the inducible COX-2 isoform is expressed at the time of implantation but restricted to the implantation site (11), corresponding to the two different stages of uterine edema. In support of this, COX-1- and COX-2-deficient mice have reduced uterine weight and vascular permeability (37, 38). Cox-1-/- females maintain normal implantation by compensatory up-regulation of COX-2 (37). However, a single dose of a selective COX-2 inhibitor can completely block implantation in these mice (39), suggesting that prostaglandin signaling is indispensable and that uterine edema may be an important aspect of the implantation process. It is unclear whether prostaglandin-induced fluid shifts during the periimplantation period are mediated by the currently known AQPs because there is no colocalization of AQPs with cellular sites of COX-1 or COX-2 expression. However, the prostaglandin E2 receptor EP3 has the same distribution as AQP1 in the uterine myometrium (40), suggesting that AQP and prostaglandin signaling pathways may interact to produce uterine edema before the onset of implantation. In this respect, prostaglandin E2 affects AQP2 activation in the kidney (41) and mice lacking the initial enzyme in prostaglandin synthesis, cPLA2, have abnormal AQP1 expression and function in the kidney (42). Thus, close examination of uterine phenotypes in AQP or prostaglandin-deficient mice may provide insight into the function of AQPs during the establishment of pregnancy.
We previously detected uterine AQP1 expression by microarray analysis (8). The function of AQP1 in the inner circular myometrium is unknown because AQP1-deficient mice are fertile (43), although subtle reproductive deficits in Aqp1-/- females cannot be ruled out until rigorous evaluation of uterine edema and vascular permeability have been performed. The influence of estrogen and progesterone on AQP1 expression by microarray analysis suggested that AQP1 is a hormonally regulated gene. Our current findings extend these results to show that AQP1 has a biphasic expression pattern in response to ovarian steroids. The shift of AQP1 expression to the stromal vasculature in response to estrogen suggests a role for AQP1 in uterine vascular permeability. AQP1 has been detected in a human uterine cDNA library (44) and mediates rapid transport of water across human vascular smooth muscle barriers in vitro (45). AQP1 is also present in uterine vascular smooth muscle cells of the rat (45) and is up-regulated after several hours of estrogen exposure (46). The two classical phases of estrogen response include uterine edema and increased vascular permeability followed by a second phase of cellular proliferation. In the first phase, estrogen induces fenestration of uterine capillary vessels and gaps between endothelial cells (28, 29), but the molecular pathways that govern these changes are unknown.
Phase I estrogenic responses are partially mediated by COX-derived prostaglandins but may also be carried out by other uterine genes that affect vascular permeability. VEGF/vascular permeability factor is a potent etiologic factor in uterine edema and is predominantly up-regulated by estrogen (12). VEGF receptors are expressed in the endothelium of uterine stromal blood vessels and are also influenced by ovarian steroids (47, 48). The estrogen-induced expression of AQP1 in the stromal vasculature suggests that AQPs may interact with the VEGF signaling pathway to regulate uterine fluid balance. Extensive uterine edema also occurs in response to the hormone relaxin. Unlike prostaglandin and VEGF actions, relaxin-induced uterine edema appears to be upstream of estrogen receptor signaling (49). Histamine is another potent mediator of uterine vascular permeability, but its synthesis is primarily regulated by progesterone (21). Similar to the pattern of COX-1-derived prostaglandins, histamine synthesis is restricted to the uterine luminal epithelium on d 4 of pregnancy and may be more important for the initial stages of generalized uterine edema and luminal closure during preparation for implantation. Nitric oxide, inflammatory cells, cytokines, and other factors also contribute to uterine edema (17), but their potential interactions with AQP water channels await further study.
We also observed unique expression patterns for AQP4, AQP8, and AQP9, although their distribution does not imply a role in uterine fluid balance at the time of implantation. The mouse uterus is markedly edematous on the morning after mating. At this time, the uterus is highly estrogenized and sustains an inflammatory-like cellular and vascular response to insemination. This edema is partially mediated by prostaglandins because COX-2 is up-regulated in the uterine luminal epithelium on d 1 of pregnancy (11). We observed that AQP4 is unresponsive to estrogen or progesterone alone, but its expression is superimposed on the distribution of COX-2, suggesting that this water channel may facilitate fluid exudation or reabsorption in a prostaglandin-dependent manner. Fluid balance in the blastocyst may also be AQP mediated because previous reports have identified the expression of AQPs 1–9 in preimplantation embryos by RT-PCR (33, 34). In those studies, AQP8 and AQP9 expression were present only at the morula-blastocyst stage of development. Our results showing differential localization in the implanting blastocyst and at sites of placental development suggest that AQP8 and AQP9 have distinct functions in cellular turgor or fluid/solute transport during embryo-placental development and that gene targeting studies may result in embryonic compromise.
AQP5 expression in the uterine glandular epithelium corresponds to epithelial localization of AQP5 in the cornea, types I and II pulmonary alveolar cells, airway epithelial and submucosal glands, and salivary and lacrimal glands (31, 32). Apical-basal polarity appears to be important because AQP5 localizes to the apical surface in most of these tissues, and transfected kidney cells preferentially sort AQP5 to the apical surface (50). Aberrant trafficking of AQP5 has been implicated as a cause of abnormal salivary and lacrimal secretions in Sjögren’s syndrome (51, 52). AQP5 expression after blastocyst attachment and in estrogen-treated, progesterone-primed uteri suggests that AQP5 is not specifically involved in uterine preparation for implantation but may be important for general fluid homeostasis following implantation and during placentation. In contrast to other epithelia, AQP5 immunolocalization was more intense in the basolateral region of the uterine glands. Although this location is less common, AQP5 has been reported in the basolateral region of bronchiolar epithelia and type I pneumocytes (53) and in the secretory portion of sweat glands (54). Basolateral expression in d 5 uterine glands suggests that the volume or content of uterine secretions may be regulated via an AQP5-specific mechanism.
All mammalian uteri contain endometrial glands that release histotrophic substances that promote uterine receptivity and embryonic development (18). These factors can be locally synthesized or transported from the vascular system or underlying stroma. There is a net influx of fluid, proteins, and tracer molecules from the maternal vasculature to the glandular epithelium at this time (55, 56), suggesting that basolateral AQP5 may facilitate transcellular fluid movement into the uterine lumen. AQP5-deficient mice have pulmonary and salivary abnormalities but no overt reproductive deficits (43). Despite this, more thorough investigation of reproductive phenotypes may reveal uterine abnormalities that are subtle or masked by compensatory expression of other water transport mechanisms. Indeed, fluid transport during implantation might also occur via alterations in cell-cell junctional complex proteins of the paracellular pathway, by passive transfer with ion cotransporters or by unique pore-forming mechanisms such as the CPE-1 receptor for bacterial toxins (6, 30).
Overall, our results suggest that AQPs are selectively expressed in the uterus and participate in uterine edema and estrogen-induced water imbibition during uterine preparation for implantation but are not responsible for the discrete region of increased vascular permeability surrounding the implanting blastocyst. The final mechanism of uterine edema in response to prostaglandin signals and other vasoactive substances is still unknown. Whether uterine edema is required for embryo implantation remains unclear. Pharmacologic antagonists of AQP channels are either unavailable or too toxic for in vivo studies. The development of specific AQP inhibitors and definitive analysis of AQP knockout uteri may identify a specific function for AQPs in reproduction and help to address these questions.
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Acknowledgments |
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Footnotes |
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Abbreviations: AQP, Aquaporin; COX, cyclooxygenase; VEGF, vascular endothelial growth factor.
Received November 12, 2002.
Accepted for publication December 18, 2002.
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