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. Author manuscript; available in PMC: 2015 May 15.
Published in final edited form as: Dev Biol. 2014 Feb 21;389(2):124–136. doi: 10.1016/j.ydbio.2014.01.025

Lhx1 is required in Müllerian duct epithelium for uterine development

Cheng-Chiu Huang 1,2, Grant D Orvis 2, Kin Ming Kwan 2,3, Richard R Behringer 1,2,4
PMCID: PMC3988469  NIHMSID: NIHMS569747  PMID: 24560999

Abstract

The female reproductive tract organs of mammals, including the oviducts, uterus, cervix and upper vagina, are derived from the Müllerian ducts, a pair of epithelial tubes that form within the mesonephroi. The Müllerian ducts form in a rostral to caudal manner, guided by and dependent on the Wolffian ducts that have already formed. Experimental embryological studies indicate that caudal elongation of the Müllerian duct towards the urogenital sinus occurs in part by proliferation at the ductal tip. The molecular mechanisms that regulate the elongation of the Müllerian duct are currently unclear. Lhx1 encodes a LIM-homeodomain transcription factor that is essential for male and female reproductive tract development. Lhx1 is expressed in both the Wolffian and Müllerian ducts. Wolffian duct-specific knockout of Lhx1 results in degeneration of the Wolffian duct and consequently the non cell-autonomous loss of the Müllerian duct. To determine the role of Lhx1 specifically in the Müllerian duct epithelium, we performed a Müllerian duct-specific knockout study using Wnt7a-Cre mice. Loss of Lhx1 in the Müllerian duct epithelium led to a block in Müllerian duct elongation and uterine hypoplasia characterized by loss of the entire endometrium (luminal and glandular epithelium and stroma) and inner circular but not the outer longitudinal muscle layer. Time-lapse imaging and molecular analyses indicate that Lhx1 acts cell autonomously to maintain ductal progenitor cells for Müllerian duct elongation. These studies identify LHX1 as the first transcription factor that is essential in the Müllerian duct epithelial progenitor cells for female reproductive tract development. Furthermore, these genetic studies demonstrate the requirement of epithelial-mesenchymal interactions for uterine tissue compartment differentiation.

Introduction

Two pairs of urogenital ducts, the Wolffian and Müllerian ducts, form during embryonic development in mammals, reptiles and birds (Massé et al., 2009). In male mammals, the Wolffian duct differentiates under the influence of testosterone into the vas deferens, epididymis and seminal vesicle of the reproductive system. In females, due to the lack of androgen support, the Wolffian duct degenerates during late embryonic development (Josso, 2008; Welsh et al., 2009). In female mammals, the Müllerian duct differentiates into the oviducts, uterus, cervix and upper portion of the vagina of the reproductive tract. In males, the Müllerian ducts are actively eliminated by the anti-Müllerian hormone signaling pathway (Behringer et al., 1994; Mishina et al., 1996; Jamin et al., 2003; Orvis et al., 2008). Thus, amniotes develop through an ambisexual stage relative to the urogenital ducts that subsequently requires differentiation toward the male or female reproductive tract phenotypes.

The Wolffian duct develops from the intermediate mesoderm (Jacob et al., 1991; Obara-Ishihara et al., 1999) at embryonic day (E) 9 in the mouse and its formation is complete by E10.5. Müllerian duct epithelial cells are initially specified in the rostral mesonephric epithelium around E11.5 in the mouse and subsequently invaginate and elongate caudally along the Wolffian duct (Guioli et al., 2007; Orvis and Behringer, 2007). Elongation of the Müllerian duct is complete when its caudal tip connects to the urogenital sinus at E13.5. Previous studies have demonstrated that the elongation of the Müllerian duct requires the presence of the Wolffian duct (Gruenwald, 1941, Kobayashi et al., 2005; Orvis and Behringer, 2007). The Wolffian duct expresses Wnt9b and genetic studies suggest that WNT9B secreted by the Wolffian duct is a trophic factor required for Müllerian duct elongation (Carroll et al., 2005). Hnf1b encodes a POU homeodomain transcription factor that is expressed in the Wolffian duct and directly activates Wnt9b transcription. Hnf1b knockout mice have defects in Wolffian duct development and consequently a lack of Müllerian duct formation (Lokmane et al., 2010).

Genetic studies in mice suggest that Wnt4 in mesonephric mesenchyme is essential for Müllerian duct invagination and elongation (Vainio et al., 1999). As the Müllerian duct elongates mesenchyme cells surround the Müllerian duct epithelial cells and separate the epithelium from the Wolffian duct and mesonephric epithelium in a rostral to caudal manner. Studies suggest that the Müllerian duct mesenchyme that initially surrounds the Müllerian duct epithelium is derived from the mesonephric mesenchyme and lateral mesonephric epithelium (Zhan et al., 2006; Guioli et al., 2007). In females, differentiation of the Müllerian duct into the adult female reproductive tract is highly dependent on the interactions between the Müllerian duct epithelium and mesenchyme (Kurita et al., 2001). Signaling molecules such as WNT4 (Bernard and Harley, 2007), WNT5A (Mericskay et al., 2004), WNT7A (Miller and Sassoon, 1998; Carta and Sassoon, 2004) and several HOX transcription factors (Hsieh-Li et al., 1995; Warot et al., 1997; Du and Taylor, 2004) all participate in the differentiation of the female reproductive tract during late gestation and early postnatal stages.

Lhx1 (also known as Lim1) encodes a LIM-homeodomain transcription factor (Jurata et al., 1998). Lhx1 is essential for the formation of the head organizer, kidneys, and retinal layers (Shawlot and Behringer 1995; Kobayashi et al., 2005; Poché et al, 2007). It is also required redundantly with Lhx5 for cerebellar cell survival (Zhao et al., 2007). Previous studies demonstrated that Lhx1 is expressed in the Wolffian and Müllerian ducts and is critical for female reproductive tract formation and/or maintenance (Kobayashi et al., 2004; Pederson et al., 2005; Orvis and Behringer, 2007). In addition, heterozygous LHX1 missense mutations have been found in patients with Müllerian aplasia, the congenital loss of the uterus and vagina (Sandbacka et al., 2013). Wolffian duct-specific knockout of Lhx1 results in Wolffian duct degeneration but also secondary loss of the Müllerian duct (Kobayashi et al., 2005). Thus, the role of Lhx1 specifically in the Müllerian duct is currently unknown. Emx2 and Pax2 also encode transcription factors that are expressed in both the Wolffian and Müllerian ducts that are required for female reproductive tract development (Torres et al., 1995; Miyamoto et al., 1997). However, because mutants for these genes also have Wolffian duct defects these studies need to be interpreted with caution because of the potential non cell-autonomous effect of defective Wolffian duct development on subsequent Müllerian duct development.

In the present study, we generated Müllerian duct epithelium-specific Lhx1 conditional knockout mice (Lhx1 cKO), using a Wnt7a-Cre transgene (Winuthayanon et al., 2010). Unlike Lhx1 null mutant mice (Shawlot and Behringer, 1995), Lhx1 conditional knockouts are viable. Lhx1 cKO males are healthy and fertile, however Lhx1 cKO females are sterile because they have a shortened oviduct and lack a uterus, cervix and upper vagina. Interestingly, residual uterine tissue in the Lhx1 cKO females appears to be the longitudinal muscle layer. Developmental studies demonstrate that Müllerian duct elongation is blocked, resulting in an absence of caudal Müllerian duct. Increased cell death and decreased cell proliferation in the Müllerian duct epithelium provide two cellular mechanisms responsible for the block in ductal elongation. Time-lapse imaging in organ culture revealed high motility and cell protrusions of the Müllerian duct leading tip during elongation. This phenotype is compromised upon conditional loss of Lhx1. These data suggest a cell-autonomous requirement of Lhx1 for Müllerian duct cell survival, proliferation and elongation. In addition, these genetic studies indicate complex epithelial-mesenchymal interactions for the development of distinct uterine tissue compartments.

Results

Wnt7a-Cre activity is specific to the Müllerian duct epithelium during urogenital organogenesis

WNT7A is a member of the WNT glycoprotein family. Wnt7a is expressed in the Müllerian duct epithelium and is required for sexual dimorphic differentiation of the reproductive tract (Parr and McMahon, 1998). Wnt7a-Cre transgenic mice were generated by modifying a bacterial artificial chromosome that contains the Wnt7a locus (Winuthayanon et al., 2010). Matings between Wnt7a-Cre mice and Rosa26 lacZ Cre reporter (R26R-lacZ) mice resulted in β-gal expression in the Müllerian duct. The β-gal expression pattern in these mice was similar to the endogenous expression pattern of Wnt7a in the Müllerian duct epithelium (Huang et al., 2012). β-gal activity was also detected in regions of the developing central nervous system, limb buds and hair follicles (data not shown).

Further characterization of β-gal expression mediated by the Wnt7a-Cre transgene during urogenital development showed that transgene activity was detected in the Müllerian duct beginning at tail somite stage 22 (TS 22) (Fig. 1A and B). Subsequently, the Wnt7a-Cre transgene faithfully recombines the R26R-lacZ transgene in the Müllerian duct during its elongation. β-gal activity was specifically detected throughout the Müllerian duct epithelium but was not detected in the Wolffian ducts. The Wnt7a-Cre transgene also labels the caudal tips of the elongating Müllerian ducts (Fig. 1C–F). These results demonstrate that the Wnt7a-Cre transgenic mouse line can be used for Müllerian duct epithelium-specific gene modifications.

Fig. 1. Temporal and spatial characterization of Wnt7a-Cre-mediated lacZ expression in developing urogenital tissues.

Fig. 1

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Wnt7a-Cre males were crossed with Rosa26 lacZ Cre reporter (R26R-lacZ) females. Bigenic embryos were dissected and stained at various embryonic stages determined by tail-somite (TS) counts. A, At E11.75 (TS 20), no urogenital tissues were β-gal positive. B, At E12.0 (TS 22), the tip of the Müllerian duct shows β-gal expression (red arrows). Both the Wolffian ducts and kidneys are negative for β-gal. C, At E12.25 (TS 26), the entire forming Müllerian duct is stained positive for β-gal. D, At E12.5 (TS 28), the entire Müllerian duct is stained positive for β-gal. Wolffian ducts and kidneys are negative for β-gal signal. E, At E13.0 (TS 32), the elongating Müllerian duct starts to migrate toward the midline. F, At E13.5 (TS 34), The Müllerian duct completes its formation and connects to the urogenital sinus. The black scale bar represents 200 μm in A and B. The white scale bar represents 500 μm in C–F.

Generation and characterization of Müllerian duct-specific Lhx1 conditional knockout mice

Given the overlapping temporal expression pattern of Lhx1 and Wnt7a (Parr and McMahon, 1998; Kobayashi et al., 2004) in the developing Müllerian duct, it should be feasible to generate viable, Müllerian duct-specific Lhx1 conditional knockout (Lhx1 cKO) mice using the Wnt7a-Cre transgene. Lhx1lacZ/+; Wnt7a-Cre Tg/+ bigenic males were bred with females homozygous for an Lhx1 flox allele (Lhx1fx/fx). The Lhx1lacZ allele serves as a null allele but also marks Lhx1-expressing cells by simple X-gal staining (Kania et al., 2000). We found that both control and Lhx1 cKO neonates were born at the expected Mendelian ratio. Analysis of the female reproductive tract from control and Lhx1 cKO neonates revealed several structural differences. Control female neonates (postnatal day 0, P0) developed normal female reproductive tracts (Fig. 2A), including oviducts, uterus, cervix and vagina. However, Lhx1 cKO female neonates were found to have short oviducts relative to controls but their uterus, cervix and upper vagina were replaced by a thin membrane-like tissue (Fig. 2B). Lhx1 cKO females can survive to adulthood but are infertile. In contrast, Lhx1 cKO adult males were normal and fertile. When compared to control animals (Fig. 2C), it was apparent that the uterine horns, cervix and upper vagina did not develop in Lhx1 cKO female adults (Fig. 2D). Oviducts from control adult females were highly coiled (Fig. 2E) but in the Lhx1 cKO adult females the oviducts were shorter, although they did form a coiled structure and had normal cytoarchitecture (Fig. 2E, F).

Fig. 2. Ablation of Lhx1 results in uterine aplasia in neonatal and adult females.

Fig. 2

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A, B, Comparisons of the female reproductive tracts between neonatal control and Lhx1 cKO mice. In Lhx1 cKO mice, the oviducts are shorter and the uterus is replaced by a thin membranous tissue. C–F, Comparisons of the female reproductive tract between adult control and Lhx1cKO mice. The controls have coiled oviducts of normal length (white dashed line). In the mutants, there is no uterus, cervix or upper vagina. The mutants do possess coiled but shortened oviducts (white dashed line) with normal histology (F, inset). Follicles and corpus lutea are found in control and mutant ovaries, indicating their normal function in both groups. G, H, Histological analysis of the neonatal uterus of control and Lhx1 cKO mice. Sections are as indicated by dashed lines in A and B. While the luminal epithelium is present in control mice (white arrow in G), it is absent in the thin membranous tissue of mutant mice. Mesenchymal cells surrounding the luminal epithelium are condensed and well-developed in the control group, whereas they are less condensed and show signs of cell death in Lhx1 cKO mice (H and inset). I, J, Sections are as indicated by dashed lines in C and D. H & E staining shows that components of the uterus, including the endometrium (luminal epithelium, LE; glandular epithelium, GE; and stroma, Str), two smooth muscle layers (Myo), and serosa (Ser) were present as expected in control mice (I). Apparently, only the outer longitudinal muscle layer is present in the Lhx1 cKO mice (J). K, L, The identity of the “muscle-like” structure is further confirmed by immunofluorescence for smooth muscle action alpha (SMAα, green; DAPI blue), demonstrating that this rudimentary tissue is likely the outer muscle of the presumptive uterine tissue in Lhx1 cKO females. Scale bars represent 500 μm in A & B and I–L; 1 cm in C & D; 250 μm in E & F and 100 μm in G & H (50 μm in insets). O, ovary; Ovd, oviduct; Ut, uterus.

Hematoxylin and eosin staining showed the well-organized luminal epithelium (Fig. 2G) surrounded by condensed mesenchymal cells in the control uterus at P0. In Lhx1 cKO neonates, there were no epithelial cells in the membranous tissue and the surrounding mesenchymal cells were loose and showed histological indications of cell death (Fig. 2H). In the uteri of adult control mice, the two muscle layers and the entire endometrium including the luminal epithelium (LE), glandular epithelium (GE) and stroma (Str) developed as expected (Fig. 2I). Interestingly, in Lhx1 cKO females though mesenchymal cells did not appear normal at P0, they do develop into tissue that resembles the outer longitudinal muscle (Fig. 2J). Immunofluorescence for smooth muscle actin alpha (SMAα) further supported the notion that this membrane-like structure in Lhx1 cKO adults was likely the outer longitudinal myometrial layer (Fig. 2K and L). These data suggest that Lhx1 is cell-autonomously required for the development of epithelial lineages derived from the Müllerian duct and supports the idea that there are extensive interactions between the epithelium and adjacent mesenchyme postnatally to induce the differentiation of the stroma and inner circular myometrial layer of the uterus. Furthermore, differentiation of the outer longitudinal myometrial layer appears to be independent of the Müllerian duct.

To determine the onset of the reproductive organ phenotypes in Lhx1 cKO mice, embryos at different developmental time points were analyzed. Urogenital organs were isolated at various developmental stages and processed for X-gal staining to track Lhx1lacZ expression and monitor tissue morphology. At E12.5, the Müllerian ducts in the control embryos were marked by β-gal and had extended caudally along the Wolffian ducts (Fig. 3A). Although the Müllerian ducts were able to invaginate from the mesonephric epithelium, they failed to elongate farther in the Lhx1 cKO embryos (Fig. 3B). At E13.5, the Müllerian ducts in control embryos were completely formed and connected to the urogenital sinus (Fig. 3C), while only the very rostral part of the Müllerian ducts were found in Lhx1 cKO embryos. In mutant embryos, the caudal ends of Müllerian ducts never connected to the urogenital sinus (Fig. 3D). At E14.5, the Müllerian ducts further developed and expressed strong Lhx1lacZ signals in controls (Fig. 3E), whereas only the rostral part of the Müllerian ducts formed in the mutants (Fig. 3F). At E16.5, Lhx1lacZ expression was reduced in the caudal Müllerian ducts of control embryos (Fig. 3G). In contrast, there was no caudal Müllerian duct in Lhx1 cKO embryos (Fig. 3H). The Wolffian ducts and kidneys appeared to be normal in both controls and mutants. There were also no differences with regard to Müllerian duct morphology between sexes from E12.5 to E14.5 (Eusterschulte et al., 1992 and data not shown). These results indicate that Lhx1 acts cell-autonomously in the Müllerian duct epithelium during female reproductive tract development for ductal elongation.

Fig. 3. Loss of Müllerian duct elongation in the Lhx1 cKO.

Fig. 3

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X-gal stained urogenital tracts. Red arrows mark the tip of the Müllerian duct (MD). A, B, At E12.5, in control embryos, the Müllerian ducts have elongated about two thirds of the entire length of Wolffian ducts (WD). In Lhx1 cKO embryos, the Müllerian ducts have only elongated less than 25% of the entire length of the Wolffian ducts. C, D, At E13.5, in control embryos, the Müllerian ducts complete their formation by connecting to the urogenital sinus (blue arrow head). In Lhx1cKO mice, the Müllerian ducts are still truncated in comparison to controls. E, F, At E14.5, in control embryos, the Müllerian ducts, Wolffian ducts and kidneys are marked by Lhx1lacZ expression. In Lhx1cKO embryos, only the rostral Müllerian ducts have formed and their Wolffian ducts and kidneys are not affected. G, H, At E16.5, Lhx1lacZ weakly labels the Müllerian duct cells and reveals Müllerian fusion at the sinus in control embryos. In Lhx1 cKOs, Müllerian duct elongation stops immediately caudal to the gonads. Scale bars represent 200 μm.

Increased cell death and decreased cell proliferation in the Müllerian duct epithelium of Lhx1 cKO embryos

The failure of the elongation of the Müllerian duct epithelium led us to hypothesize that Lhx1 promotes cell survival and/or proliferation in the Müllerian duct epithelium. The terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay was used to assess cell death in the Müllerian duct epithelium. At E12.5 there were very few TUNEL-positive cells in the control Müllerian duct epithelium (Fig. 4A and C, mean and SEM: 0.7 ± 0.3% of total 881 MDE cells counted, n=5). In contrast, the Müllerian duct epithelial cells of Lhx1 cKO at E12.5 showed a significant increase (T-test: p < 0.01) of TUNEL-positive signals (Fig. 4B and C, mean and SEM: 16.2 ± 3.7% of total 838 MDE cells counted, n=4). Immunofluorescence against activated-Caspase3 also showed that while very few Müllerian duct epithelial cells were undergoing apoptosis in controls (Fig. 4D and F, mean and SEM: 1.1 ± 0.3%, 371 MDE cells counted, n=3), there were significantly more apoptotic cells (T-test: p < 0.01) in the Müllerian duct epithelium of Lhx1 cKO embryos (Fig. 4E and F, mean and SEM: 12.1 ± 2.3%, 333 MDE cells counted, n=3).

Fig. 4. Ablation of Lhx1 leads to increased cell death in the Müllerian duct epithelium.

Fig. 4

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A,B, TUNEL assay demonstrates there is almost no cell death in the Müllerian duct epithelium of control embryos, whereas cell death (green signal) is found in the Müllerian duct epithelium of the Lhx1 cKO embryos at E12.5. C, Statistical analysis shows the difference in cell death between control (0.7 ± 0.3%, mean ± SEM) and Lhx1 cKO (16.2 ± 3.7%) embryos is significant (T-test: p < 0.01). Error bars represent SEM. D, E, Immunofluorescence for activated-caspase 3 (red) demonstrates there is almost no apoptosis in the Müllerian duct epithelium of control embryos, whereas there is more apoptosis in the Müllerian duct epithelium of Lhx1 cKO embryos at E12.5. Green, immunofluorescent labeling of pan-cytokeratin. Blue, DAPI. F, Statistical analysis shows the difference in apoptotic cell death between control (1.1 ± 0.3%) and Lhx1 cKO (12.1 ± 2.3%) embryos is significant (T-test: p < 0.01). Error bars represent SEM. Scale bars represent 50 μm in A, B, D and E and 5 μm in insets of D and E.

PAX2 is a paired-box homeodomain transcription factor expressed in both the Wolffian and Müllerian ducts. Pax2 is critical for Müllerian duct development and for cell survival in a polycystic kidney disease model (Torres et al., 1995; Ostrom et al., 2000). Pax2 has also been shown to regulate Bcl2, an anti-apoptotic gene which is expressed in the Müllerian duct (Park et al., 2006; Lee et al., 1998). Whether LHX1 regulates Pax2 expression during the elongation of Müllerian ducts and inhibits cell death/apoptosis is unknown. Whole mount in situ hybridization was performed to visualize Lhx1 and Pax2 expression in E13.5 control and Lhx1 cKO embryos. Lhx1 is down-regulated in the partially formed Müllerian ducts, but not in the Wolffian ducts of Lhx1 cKO embryos (Fig. 5A and B), whereas Pax2 expression levels did not seem to vary between the control and Lhx1 cKO Müllerian ducts (Fig. 5C and D). These results indicate that Pax2 acts upstream of Lhx1 or that Lhx1 and Pax2 act in parallel to support the survival of Müllerian duct epithelial cells.

Fig. 5. Lhx1 andPax2 expression in Lhx1 cKO embryos.

Fig. 5

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A, B, At E13.5, Lhx1 mRNA signals are relatively strong in the Müllerian ducts compared to the Wolffian ducts of control embryos. Lhx1 RNA signals in the Wolffian ducts of Lhx1 cKO embryos were similar to controls. Very weak signals were found only in the rostral Müllerian duct. C, D, At the same stage, Pax2 mRNA levels in the Müllerian and Wolffian ducts of the control and Lhx1 cKO embryos are comparable. MD, Müllerian duct; WD, Wolffian duct. Scale bar represents 200 μm.

Cell proliferation in the E12.5 Müllerian duct epithelium was assessed by immunofluorescence for phospho-Histone H3 (pH3), a marker of mitosis. Trunk and tip regions of the developing Müllerian ducts were compared between control and Lhx1 cKO embryos. In control embryos, approximately one-third of the total Müllerian duct epithelial cells in the trunk region were proliferating (Fig. 6A and C, mean and SEM: 36.4 ± 2.4%, 322 MDE cells counted, n=3), whereas there was a significant decrease (T-test: p < 0.01) in cell proliferation of Lhx1 cKO trunk Müllerian duct epithelium (Fig. 6B and C, mean and SEM: 15.5 ± 0.5%, 335 MDE cells counted, n=3). In control embryos, tip cells of the Müllerian duct showed similar proliferative activity as in the truck region (Fig. 6D and F, mean and SEM: 36.0 ± 4.2%, 371 MDE cells counted, n=3). Interestingly in Lhx1 cKO embryos, a significantly decreased proliferation rate (T-test: p < 0.05) was also observed in the tip cell population (Fig. 6E and F, mean and SEM: 22.7 ± 2.1%, 252 MDE cells counted, n=3). The above data are consistent with the notion that Lhx1 promotes epithelial cell survival and proliferation during Müllerian duct elongation.

Fig. 6. Ablation of Lhx1 leads to decreased cell proliferation in the Müllerian duct epithelium.

Fig. 6

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A, B, Immunofluorescence for phospho-histone H3 (pH3, red) at E12.5 in the trunk region demonstrates that there is a large amount of cell proliferation in the control Müllerian duct epithelium, whereas in Lhx1 cKO embryos there is much less proliferation in the Müllerian duct epithelium. C, Statistical analysis shows a significant difference in mitotic rate of the Müllerian duct between control (36.4 ± 2.4%, mean ± SEM) and Lhx1 cKO (15.5 ± 0.5%) embryos. T-test: p ≤ 0.01. Error bars represent SEM. D, E, Immunofluorescence for pH3 (red) at E12.5 shows a large amount of cell proliferation in the control Müllerian duct epithelium but much less proliferation is detected in the Müllerian duct epithelium tip region of the Lhx1 cKO embryos. Green, immunofluorescent labeling of pan-cytokeratin; blue, DAPI. F, Statistical analysis shows the difference in mitotic rate in the Müllerian tip between control (36.0 ± 4.2%) and Lhx1 cKO (22.7 ± 2.1%) embryos is also significant. T-test: p < 0.05. Error bars SEM. Scale bars represent 50μm in A, B, D and E and 5 μm in insets in A and B.

Visualization of cell behaviors during Müllerian duct elongation in control and Lhx1 cKO embryos

To visualize the behaviors of Müllerian duct cells during elongation, we crossed Wnt7a-Cre transgenic mice with R26R-YFP fluorescent reporter mice and generated Wnt7a-Cre; R26R-YFP embryos at E12.0–E12.5. The embryonic urogenital system was isolated and cultured in vitro. Fluorescence time-lapse imaging was used to follow the cell behaviors in the elongating Müllerian duct, revealing several interesting features not previously reported (Fig. 7, Supplementary Movie 1). The leading tip cells of the elongating Müllerian duct were very dynamic, forming and retracting numerous protrusion-like structures (Fig. 7). The growth rate of the developing Müllerian duct was about 25 to 35 μm per hour under our culture conditions, which was comparable to results suggested from previous in vivo studies (Orvis and Behringer, 2007).

Fig. 7. Time-lapse images of Müllerian duct elongation.

Fig. 7

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Mesonephros/ovary explant from E12.5 Wnt7a-Cre; R26R-YFP embryo was cultured in vitro and imaged for 28 hours. The developing Müllerian ducts were fluorescently labeled with YFP, including the migrating tips (yellow arrows). Caudal is to the right in each panel. Hours indicate time in culture. The Müllerian duct elongates along the Wolffian duct (not labeled) beneath the mesonephric epithelium (ME) in a rostral to caudal direction. The tips of the Müllerian duct exhibit multiple extensions (green arrows) that appear to have an exploratory behavior. The trailing Müllerian duct caudal to the tip becomes narrower in width. Scale bar represents 100 μm (30 μm in insets). MDE, Müllerian duct epithelium; ME, mesonephric epithelium; O, ovary.

To visualize the cell behaviors during Müllerian duct elongation in Lhx1 cKO embryos, we established a colony of compound homozygous mice carrying the Lhx1fx and R26R-YFP reporter alleles (Lhx1fx/fx; R26R-YFP). Lhx1lacZ/+; Wnt7a Tg/+ males were bred with Lhx1fx/fx; R26R-YFP females to generate YFP-labeled embryos at E12.0 to E12.5 (TS22–TS28). The gonad and mesonephros, that includes the forming Müllerian duct, were dissected from embryos and cultured in vitro for time-lapse imaging. In control embryos, the YFP-labeled Müllerian duct extended caudally as described above. As expected, tip cells in the control Müllerian duct produced numerous protrusions and showed dynamic morphological changes during Müllerian duct elongation (Fig. 8A–E and Supplementary Movie 2). In contrast, in Lhx1 cKO embryos the Müllerian duct elongated slowly and the leading tip showed few protrusions (Fig. 8F–J). Mutant tip cells also exhibited compromised migration and at later time points during the imaging process appeared to become static. This created a bulbous-like structure at the tip of the duct (Fig. 3F, 8G–H and Supplementary Movie 3).

Fig. 8. Time-lapse images of Müllerian duct cell behaviors in control and Lhx1cKO embryos.

Fig. 8

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Control and Lhx1 cKO Müllerian ducts were labeled with YFP using the Wnt7a-Cre and R26R-YFP alleles. A–E, Control Müllerian ducts; F–J, Lhx1 cKO Müllerian ducts. Hours in culture are indicated. Caudal is to the right in each panel. A and F, At the initial time of culture, the tips of the developing Müllerian ducts in the control and Lhx1 cKO embryos appeared similar. B and G, After two hours in culture, the control Müllerian duct tip elaborate multiple, thin cytoplasmic extensions and elongate caudally along the Wolffian duct. In Lhx1 cKO embryos, the tip cells seem to be more static and elongation appears slower than the control. C and H, After four hours in culture, the control Müllerian duct tips continued to form cytoplasmic extensions and seem to explore the extracellular space in the direction of elongation. In Lhx1 cKO embryos, the tip cells were less motile and do not migrate forward. D, E, I, J, After 6 hours and 8 hours in culture, the control Müllerian duct tips continued to move caudally and continued to generate multiple, thin cytoplasmic extensions. In the Lhx1cKO embryos, the cells appeared to pile up at the tip, which is essentially immobile. Scale bar represents 50 μm.

Discussion

Lhx1 acts cell autonomously in Müllerian duct epithelium for female reproductive tract development

The Müllerian duct-specific knockout of Lhx1 results in viable female mice that are infertile because they lack a uterus. We found that the Müllerian duct is specified, invaginates from the mesonephic epithelium, and elongates sufficiently to induce the differentiation of the rostral oviduct. However, the Müllerian duct fails to elongate more caudally and does not connect to the urogenital sinus. This results in shortened oviducts, uterine aplasia, and infertility. These findings along with previous chimera studies are consistent with the idea that Lhx1 acts cell autonomously in the Müllerian duct for female reproductive tract elongation (Kobayashi et al., 2004).

We previously reported that no Müllerian duct derivatives, including oviduct, uterus, cervix and upper vagina were found in Lhx1 null newborn mice (Kobayashi et al., 2004). The interpretation of this phenotype is complicated by the fact that Lhx1 is expressed in both the Müllerian and Wolffian ducts (Kobayashi and Behringer, 2003; Orvis and Behringer, 2007). The elongation of the Müllerian duct requires the Wolffian duct because it produces WNT9B (Gruenwald, 1941; Carroll et al., 2005; Orvis and Behringer, 2007). Lhx1 null newborn male mice lack Wolffian duct derivatives and a Wolffian duct-specific knockout of Lhx1 leads to the loss of the Wolffian duct and non cell-autonomous loss of Müllerian duct tissue (Kobayashi et al., 2005). Thus, the absence of Müllerian duct derivatives in Lhx1 null mice could be secondary to Wolffian duct defects. However, our Müllerian duct-specific knockout suggests that Lhx1 does have a cell autonomous role in Müllerian duct elongation.

Lhx1 encodes a LIM homeodomain transcription factor that regulates downstream target genes. The identity of these target genes for Müllerian duct development is currently unknown. Our findings identify the first transcription factor that is required in Müllerian duct epithelial cells for ductal development, providing a molecular entry point to identify other genes that regulate the development of this genital duct, a precursor tissue for the female reproductive tract. Interestingly, there are two other transcription factors, EMX2 and PAX2, which are also required for Müllerian duct development (Torres et al., 1995; Miyamoto et al., 1997). However, like Lhx1, they are also expressed in the Müllerian and Wolffian ducts. The roles of Emx2 and Pax2 in the Müllerian duct epithelium have not yet been determined. The persistence of Pax2 expression in the partially formed Müllerian duct in the Lhx1 cKO mutants suggests that Pax2 may act upstream of or in parallel with Lhx1. The Müllerian aplasia phenotype of Lhx1 cKO mice is very similar to the defects found in Dach1 +/−; Dach2 −/− mutant female mice (Davis et al., 2008). In these mutant females, the uterus lacked an epithelium or stroma but smooth muscle was present. Dach1 and Dach2 encode transcriptional cofactors that are expressed in the Müllerian and Wolffian ducts. In Dach1 −/−; Dach2/Y males, the Müllerian duct apparently does not form. These findings suggest that Dach genes act redundantly for Müllerian duct development. The transcriptional factors that act with the DACHSHUND cofactors are currently unknown although LHX1, EMX2, and PAX2 are candidates.

We found that the very rostral part of Müllerian duct formed in Lhx1 cKO embryos and probably differentiated into the shortened oviducts found in adult mutant females. Interestingly, at E16.5, the Müllerian duct present in the Lhx1 cKO mutants had elongated to the approximate position of the flexura medialis that may be the future oviduct-uterine boundary (Stewart and Behringer, 2012). In contrast, we showed in a study of female chimeras composed of Lhx1 wild-type and null cells that the null cells did not contribute to the epithelial compartment of the reproductive tract at birth (Kobayashi et al., 2004). One explanation for this difference in phenotypes between the conditional knockout and chimera studies could be the timing of the onset of Wnt7a-Cre activity, Lhx1 locus deletion, and stability of Lhx1 mRNA and LHX1 protein in the Müllerian duct epithelium of the cKO mice. Indeed, some residual Lhx1 mRNA was detected in the truncated Müllerian duct by whole mount in situ hybridization in E13.5 Lhx1 cKO embryos. Thus, there may be sufficient levels of Lhx1 in the rostral Müllerian duct for its formation, persistence and subsequent differentiation into oviductal tissue. Alternatively, the complete absence of Lhx1 null epithelial cells in the reproductive tract of newborn female chimeras may be because null cells are outcompeted earlier in development by wild-type cells. It is also possible there is redundant Lhx family expression in the rostral Müllerian duct (Zhao et al., 2007).

Lhx1 regulates cell survival and proliferation for Müllerian duct elongation

Previously, we showed that the tip of the elongating Müllerian duct contains proliferating progenitor cells that contribute to duct elongation (Orvis and Behringer, 2007). Homogeneous cell proliferation throughout the entire Müllerian duct epithelium has been reported that also contributes to duct elongation (Guioli et al., 2007; Fujino et al., 2009). In the present study, we demonstrate that Müllerian duct epithelial cells in the trunk and tip regions undergo significant cell death and reduced proliferation in the absence of Lhx1. This likely contributes to the block in ductal elongation found in the Lhx1 cKO females. This observation suggests that Lhx1 is critical for the maintenance and/or proliferation of developing Müllerian duct epithelial cells. Lhx1 has also been shown to regulate cell survival and differentiation of the developing kidney and cerebellum (Carroll and Vize, 1999; Kobayashi et al., 2005; Zhao et al., 2007).

Cell behaviors during Müllerian duct elongation

To our knowledge, we provide the first visual time-lapse documentation of cell behaviors of the developing Müllerian duct. Data acquired by combining ex vivo organ culture and time-lapse imaging of live cells shows that the cells at the tip of the elongating Müllerian duct generate dynamic cytoplasmic processes that appear to be exploratory in nature. The exploratory behavior of tip cells in the developing Müllerian duct appears to be an important ability for Müllerian duct elongation and this ability is severely compromised upon loss of Lhx1. These cellular characteristics may correlate with their “mesoepithelial” phenotype during initial Müllerian duct elongation and contribute to the rapid generation of the duct within a relatively short developmental window (Jacob et al., 1999; Orvis and Behringer, 2007). The forming Wolffian duct has also been to extend cytoplasmic processes and is regulated by a Gata3-Raldh2-Ret network (Chia et al., 2011). In contrast to our observations of Müllerian duct elongation, the caudal Wolffian duct elicits numerous lateral cytoplasmic extensions. In addition, the tip of the elongating Wolffian duct appears to be composed of numerous streams of cells, whereas the Müllerian duct tip appears more cohesive primarily extending cytoplasmic processes. Perhaps this is because Müllerian duct elongation uses the Wolffian duct as a guide, whereas the Wolffian duct extends caudally through the mesonephric mesenchyme perhaps attracted by a caudal signal. Ret and Gata3 expression is not detected in the Müllerian duct epithelium (Pachnis et al., 1993; Grote et al., 2006). Therefore, the cytoplasmic processes observed at the tip of the elongating Müllerian duct are likely regulated by factors that are different from those of the Wolffian duct.

Interestingly, the cell behaviors observed in the developing Müllerian duct are highly reminiscent of collective cell migration (Fujino et al., 2009; Rørth, 2012; Theveneau and Mayor, 2013). N-Cadherin has been implicated to be involved in both Müllerian duct development and collective cell migration (Friedl and Gilmour, 2009; Smith et al., 2010; Theveneau et al., 2010). In addition, the more rostral region of the forming Müllerian duct became thinner, perhaps through convergent extension mechanisms (Wallingford et al., 2002). WNT family members WNT4, WNT7A and WNT9B participate in the development and differentiation of female reproductive tract (Vainio et al., 1999; Miller and Sassoon, 1998; Carroll et al., 2005). Although these WNT proteins are generally involved in the canonical WNT pathway, it has been shown that they can also mediate non-canonical WNT-PCP signaling to promote convergent extension (Le Grand et al., 2009; Karner et al., 2009). Consistent with this idea, one of the non-canonical WNT receptor genes, Frizzled10, is expressed in the developing Müllerian duct (Nunnally and Parr, 2004) and could potentially convey WNT-PCP signals to facilitate convergent extension in the Müllerian duct.

Our live imaging results and studies by others suggest that elongation of Müllerian duct may be accomplished by collective cell migration, convergent extension and/or cell shape changes, and loss of Lhx1 can impede these important cellular processes (Fujino et al., 2009). Whether and how the LHX1 transcriptional program intersects with the WNT-PCP signaling pathway in female reproductive tract development is unclear. Nevertheless, Lhx1 has been shown to regulate nephric duct extension and ureteric bud morphogenesis, a process that depends on WNT-PCP signaling. (Kobayashi et al., 2005; Pedersen et al., 2005; Karner et al., 2009; Lienkamp et al., 2012). Thus, Lhx1 may have a broader role in ductal extension beyond the Müllerian system.

In addition to the aforementioned factors, Fujino et al. (2009) demonstrated that PI3K/AKT regulates Müllerian duct migration and elongation by performing incision of the mesonephros and Müllerian duct and chemical inhibitor exposure of cultured urogenital ridges (Fujino et al., 2009). PI3K/AKT signaling has previously been shown to regulate actin organization and cell migration (Jiménez et al., 2000; Enomoto et al., 2005) and thus may provide another molecular control of tip cell motility and polarity in the developing Müllerian duct. Further defining the molecular cascades of Müllerian duct development should shed light on the etiology of diseases in the female reproductive tract, including Müllerian duct agenesis, hypoplasia and perhaps gynecological cancers.

Epithelial-mesenchymal interactions are required for stromal and myometrial differentiation

Reciprocal interactions between epithelium and mesenchyme are critical for the development of the female reproductive tract and other organs, including the mammary glands and the prostate (Cunha et al., 2004). Mesenchymal or stromal cells in the female reproductive tract secrete paracrine signals to instruct the morphological and functional differentiation of the epithelial cells. Mesenchymal cells also mediate physiological actions of estradiol and progesterone and regulate epithelial cell proliferation and apoptosis in adult females (Lubahn et al., 1993; Cooke et al., 1997).

Histological analysis of Lhx1 cKO females revealed that there are no epithelial cells in the reproductive tract at birth except in the shortened oviduct. Consequently, the uterine endometrium does not differentiate. The Müllerian duct epithelium likely gives rise to the luminal and glandular epithelium of the endometrium (Lin et al., 2009). Therefore, the Müllerian duct not only gives rise to the epithelial compartment of the endometrium but is also required to induce the differentiation of the adjacent mesenchyme into the uterine stroma. Surprisingly some smooth muscle differentiates that appears structurally like the outer myometrium. This suggests that the epithelial compartment of the developing uterus is required for differentiation of the inner circular muscle layer. This may be direct or indirect through the epithelial-directed differentiation of the stroma. Another explanation for the development of the outer longitudinal muscle structure is that the uterine serosa provides instructive cues for uterine mesenchymal cell differentiation (Brauer et al., 1989). Thus, uterine tissue layer specification, survival and differentiation are highly dependent on the presence of epithelial cells. This observation also underscores the importance of proper epithelial-mesenchymal interaction in female reproductive tract development and diseases.

Methods

Mice

Lhx1tm1Tmj (Lhx1lacZ; Kania et al., 2000), Lhx1tm2.1Bhr (Lhx1fx; Kwan and Behringer, 2002), Gt(ROSA)26Sortm1Sor (R26R-lacZ; Soriano, 1999) and Gt(ROSA)26Sortm1(EYFP)Cos (R26R-YFP; Srinivas et al., 2001) mice were maintained on a C57BL/6J, 129S6/SvEvTac mixed genetic background. Tg(Wnt7a-EGFP/cre)1Bhr (Wnt7a-Cre Tg/+; Winuthayanaon et al., 2010) mice were maintained on a C57BL/6J, SJL/J, 129S6/SvEvTac mixed genetic background. All animal procedures were approved by the Institutional Animal Care and Use Committee of the University of Texas M.D. Anderson Cancer Center.

PCR genotyping

Lhx1lacZ, Lhx1fx and R26R-YFP alleles were genotyped as previously described (Kania et al., 2000; Kwan and Behringer, 2002; Soriano, 1999). Wnt7a-Cre Tg/+ mice were genotyped by PCR using the following primers: CreFw: 5′-GGACATGTTCAGGGATCGCCAGGC-3′ and CreRv: 5′-CGACGATGAAGCATGTTTAGCTG-3′. Wnt7a-Cre PCR was performed on a DNA Thermal Cycler 480 (Perkin-Elmer Corp., Wellesley, MA) with 35 cycles at 95°C for 30s, 56°C for 30s, 72°C for 45s. The expected size of the PCR product is 219 bp.

X-gal staining of embryos

X-gal staining for β-gal activity was performed as described (Nagy et al., 2003) with minor modifications. Embryos were collected at different stages. Lower body trunks with urogenital tissues were isolated and fixed briefly in 4% paraformaldehyde (PFA) in PBS at 4°C (10 minutes for E11.5–E12.5; 15 minutes for E14.6–E16.5). Tissues were then washed in X-gal rinse solution (PBS containing 0.02% NP-40 and 0.01% deoxycholate) 3 times for 15 minutes each. The wash solution was removed and tissues were stained with X-gal: 5 mM K3Fe(CN)6, 5 mM K4Fe(CN)6, 0.02% NP-40, 0.01% deoxycholate, 2 mM MgCl2, 5 mM EGTA and 2 mg/ml X-gal solution dissolved in DMSO in PBS. Tissues were incubated at 37°C for 12 to 18 hours until the intense blue signal was observed. Tissues were washed 3 times in PBS and then postfixed in 2% PFA/PBS for 20 minutes. Fixed tissues were cleared in glycerol/PBS (1:1 ratio) overnight at 4°C and photographed.

Tissue processing

Embryos and reproductive tissues from neonates or adults were dissected and fixed in 4% PFA at 4°C overnight. Fixed tissues were further processed for cryo- or paraffin sectioning. For cryosectioning, tissues were washed 3 times in PBS and immersed in 30% sucrose/PBS solution overnight at 4°C. Tissues were then incubated in 30% sucrose/O.C.T. (Tissue-Tek) at room temperature for 1 hour and frozen blocks were made with the same sucrose/O.C.T. mixture. Frozen sections were cut at 10 μm and stored at −80°C for further histological analysis and immunofluorescence.

For paraffin sectioning, fixed tissues were rinsed 3 times in PBS and stored in 70% ethanol. Serial dehydration and paraffin infiltration of fixed tissues were performed using a tissue processor, Leica TP1020 (Leica Microsystems NusslochGmbh, Heidelberger). Paraffin blocks were made using a Leica EG1160 embedding station. Paraffin sections were cut at 4 μm and store at 4°C for histological analysis and TUNEL assay.

Immunofluorescence and quantification

Immunofluorescence was performed on 10 μm frozen sections. In brief, sections were dried at room temperature for 20 minutes and then washed 3 times in 1x PBS. Sections were incubated in blocking solution containing 5% normal goat serum and 3% bovine serum albumin in PBS for 60 minutes at room temperature. Rabbit anti-activated caspase 3 antibody (1:100, Cell Signaling Technologies, Inc.) or rabbit antibody against phospho histone H3 (1:300, Millipore) and rat antibody against pan-cytokeratin (1:300, Developmental Studies Hybridoma Bank) were applied to samples and incubated overnight at 4 °C. Sections were next washed in PBS three times and incubated with fluorophore-conjugated goat anti-rabbit (1:600, for activated caspase 3) or donkey anti-rat (1:800, for pan-cytokeratin) IgG (Invitrogen, Inc.) for 45 minutes at room temperature. Samples were then washed in PBS 5 times and covered with DAPI-containing Vectasheld mounting medium (Vector Laboratories, Inc.). Immunofluorescent signal for pan-cytokeratin marked the Wolffian duct and weakly labeled the Müllerian duct at E12.5 (Orvis and Behringer, 2007). Cytokeratin immunofluorescence and DAPI signal were both used to distinguish Müllerian duct from nearby structures and for MDE cell counting.

TUNEL assay

TUNEL was performed to detect cell death. Paraffin sections were deparaffinized and incubated in proteinase K solution (20 μg/ ml in 10 mM Tris-HCl, pH 7.4–8) for 20 minutes at 37°C. After washing three times in PBS, sections were permeabilized with 0.1% Triton X-100 in 0.1% sodium citrate for 2 minutes on ice, and then in 50 μl TUNEL reaction mixture (Roche Applied Science) for 1 hour at 37°C in the dark. Samples were then washed in PBS for three times and covered with DAPI-containing Vectasheld mounting medium.

Whole mount in situ hybridization

Preparation of Digoxigenin-11-UTP-labeled antisense cRNA probes was performed as previously reported (Lufkin, 2007). Lhx1 and Pax2 antisense cRNA probes were generated from plasmids described previously (Dressler et al., 1990; Shawlot and Behringer, 1995). Whole-mount in situ hybridization of E13.5 embryos was performed as described (Wilkinson and Nieto, 1993) with minor modifications. In brief, embryos were stained in glass scintillation vials using BM purple (Roche Applied Science) in alkaline phosphatase buffer at room temperature in the dark. The color reaction was slowed by moving samples to 4°C and subsequently by washing three times in MABT buffer (0.1M Maleic acid, 0.15M NaCl and 0.1% Tween-20, pH 7.5). Stained tissues were cleared in glycerol/PBS mix (1:1 ratio) for photographs.

Organ culture and time-lapse imaging

E12.0–E12.5 urogenital ridges were dissected in cold PBS and then cultured in organ culture medium consisting of Dulbecco’s modified Eagle’s medium (with 4.5 g/ L D-glucose, without sodium pyruvate and phenol red), 10% fetal bovine serum, 2 mM glutamine and 1 mM sodium pyruvate (Invitrogen Inc.). A culture system for fetal gonads and urogenital tissues was described previously (Nel-Themaat et al., 2009). Briefly, urogenital ridges were placed on a Millicell tissue culture plate insert (Millipore Corporation) and cultured at the air-medium interface in a 35 mm glass bottom culture dish (MatTek Corporation). Time-lapse images were acquired on a PerkinElmer spinning disc laser confocal microscope at 37°C and 5% CO2 in a humidified chamber. A 514 nm laser was used to acquire YFP images. Images were taken at 100x or 200x magnifications (with 10x and 20x long working-distance objective lens) using 50% laser power and 300 millisecond exposure with 2 × 2 binning. The Z-plane depth was 40–60 μm. Images were processed by Ultra VIEW ERS ImageSuite Software (PerkinElmer Life and Analytical Sciences, Inc.) and Adobe Photoshop.

Supplementary Material

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Acknowledgments

We thank Drs. Francesco DeMayo, Mary Dickinson, Richard Kelley and Ming-Jer Tsai for their insightful suggestions. We are grateful to Hank Adams and Ying Wang for their technical assistance. Supported by National Institutes of Health (NIH) grant HD030284, the Ben F. Love Endowment, and the Kleberg Foundation to R.R.B. G.D.O. was supported by National Cancer Institute T32 grant CA09299. Veterinary resources were supported by NIH grant CA16672.

Footnotes

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