Skip to main content
NCBI home page
Biology of Reproduction logoLink to Biology of Reproduction
. 2013 Oct 2;89(6):148. doi: 10.1095/biolreprod.113.107649

Investigating the Role of Tbx4 in the Female Germline in Mice1

Nataki C Douglas 3,2,, Ripla Arora 4, Cayla Yiyu Chen 3, Mark V Sauer 3, Virginia E Papaioannou 4
PMCID: PMC4076358  PMID: 24089201

ABSTRACT

Normal development of germ cells is essential for fertility and mammalian reproduction. Although abnormal development of oocytes or follicles may lead to primary ovarian insufficiency (POI), a disorder that causes infertility in 1% of women less than 40 yr of age, the genes and signaling pathways activated in POI are not as yet fully elucidated. Tbx4, a member of the T-box family of transcription factors, is expressed in embryonic germ cells and postnatal oocytes at all stages of folliculogenesis. To investigate the requirement for Tbx4 in the germline, we analyzed germ cell development in the absence of Tbx4. We show that primordial germ cells (PGCs) are reduced in Tbx4 homozygous null (Tbx4−/−) embryos at Embryonic Day (E) 10.0. Tbx4−/− embryos die by E10.5; to study later time points in vitro, a tamoxifen-inducible estrogen receptor Cre recombinase was used to delete Tbx4 conditional mutant alleles. In addition, Gdf9cre and Zp3cre, two oocyte-specific Cre recombinases, were used to delete Tbx4 from postnatal primordial and primary follicles, respectively. We show that in vitro differentiation of the gonad into morphologically distinct testes and ovaries occurs normally starting at E11.5 when Tbx4 is deleted. In Gdf9cre; Tbx4fl/− and Zp3cre; Tbx4fl/− adult females, primordial, primary, secondary, and antral follicles form, ovulation occurs, corpus luteum formation is normal, and the mice are fertile without any evidence of diminished ovarian reserve. Although postnatal deletion of Tbx4 in oocytes does not obviously impair fertility, it is possible that the reduction in PGCs observed in Tbx4 homozygous null mutant embryos could affect long-term fertility in adults.

Keywords: fertility, oocytes, primordial germ cells, T-box, Tbx4

Postnatal oocyte-specific deletion of Tbx4 does not obviously impair fertility but reduces primordial germ cell number.

INTRODUCTION

The development of the mammalian female reproductive system starts during gestation and is completed postnatally with the onset of puberty. Sexually dimorphic gene expression, activation of multiple signaling pathways, and complex interactions between somatic and germ cells are required for normal development and function of ovarian follicles, the functional units of the ovary. In humans, abnormal development of oocytes or follicles may lead to premature ovarian failure, recently termed primary ovarian insufficiency (POI), which affects 1% of women less than 40 yr of age [1]. POI causes a premature reduction in endogenous estrogen production and leads to significant morbidities in young women, including increased risks of cardiovascular and neurodegenerative diseases, osteoporosis, and infertility. In order to evaluate the causes of POI, it is important to understand the molecular framework that underlies oocyte development and function.

Bipotential gonads, precursors to the testes and ovaries, reside within the urogenital (UG) ridges and contain both germ cells that will mature into oocytes and somatic cells. In mice, primordial germ cells (PGCs) arise in the epiblast between Embryonic Day (E) 7 and E7.5, migrate to the UG ridges, and enter the gonads by E11.5. Upon entry into the UG ridges, PGCs are termed gonocytes or germ cells [2]. Germ cell fate determination depends on interactions with differentiated somatic cells in the gonads, and by E13.5, male and female germ cells are distinguishable from one another [3–6]. Germ cells in testes reside within testis cords and undergo mitotic arrest. In contrast, germ cells in ovaries enter meiosis, undergo arrest in prophase I, and remain loosely connected in germ cell cysts from E13.5 until birth. Primordial follicle formation occurs in early postnatal life [7]. As follicle formation is initiated, a single occyte is surrounded by pre-granulosa cells. Ovarian folliculogenesis involves oocyte maturation, somatic cell differentiation, and progression through distinct primordial, primary, secondary, preantral, antral, and preovulatory follicle stages [8, 9].

We recently described the expression of the TBX2 subfamily of T-box transcription factor genes, which includes Tbx4, during development of the mammalian reproductive system [10]. The T-box family of transcription factors, defined by a conserved DNA-binding domain, the T-box, plays a critical role in the determination of cell fate decisions during organogenesis and in the differentiation of many organ systems [11–13]. Mutations in Tbx4 are associated with developmental defects in both mice and humans. In mice, homozygous null Tbx4 mutants lack chorio-allantoic fusion, which prevents formation of the umbilical vessels and results in death by E10.5. In addition, Tbx4-null embryos have abnormal hind limb development [14]. Heterozygous mutations in the human TBX4 gene cause small patella syndrome, an autosomal, dominant, skeletal dyplasia characterized by patellar, pelvic bone, and foot anomalies [15]. The similarities between mice and human phenotypes underlie the importance of T-box genes in mammalian development and strongly suggest conservation of function among different species. Tbx4 is the only member of the TBX2 subfamily expressed in germ cells [10]. Despite this expression, the function of Tbx4 in germ cell development in mice or humans has not as yet been described. In other systems, such as the respiratory system, Tbx4 interacts with members of the FGF and WNT families, genes that play important roles in germ cell differentiation and gonad development [16–19]. Thus, Tbx4 may have a role in the signaling pathways activated during normal reproductive-system development. In this report, we utilize Tbx4-null and Tbx4 conditional mutant alleles to investigate the requirement for Tbx4 during embryonic germ cell development and postnatal folliculogenesis.

MATERIALS AND METHODS

Mice

Mice carrying the following alleles were genotyped as previously described: a Tbx4-null allele, Tbx4tm1.1Pa [14], hereafter referred to as Tbx4−; a Tbx4 conditional “floxed” allele, Tbx4tm1.2Pa [20], hereafter referred to as Tbx4fl; ROSA26cre-ERT2, a ubiquitously expressed tamoxifen-inducible cre transgene [21], hereafter referred to as creER; and two oocyte-specific cre transgenes (both from Jackson Laboratory), Gdf9-icre [22], hereafter referred to as Gdf9cre, and Tg (Zp3-Cre)93Knw [23], hereafter referred to as Zp3cre. Ovaries from Gdf9cre; Tbx4fl/fl, Gdf9cre; Tbx4fl/−, Zp3cre; Tbx4fl/fl, and Zp3cre; Tbx4fl/− females were analyzed at postnatal time points.

Gdf9cre; Tbx4fl/fl males bred to Tbx4fl/fl females generated both Gdf9cre; Tbx4fl/fl and, unexpectedly, Gdf9cre; Tbx4fl/− progeny, indicating Cre activity in male germ cells. Excision of Tbx4 in the male germ cells was observed in three different Gdf9cre; Tbx4fl/fl males in our colony (Supplemental Table S1; all Supplemental Data are available online at www.biolreprod.org). To confirm activity of Gdf9cre in both male and female germ cells, Gdf9cre males were mated to ROSA26 LacZ reporter (Rosa-lox-stop-lox-LacZ; R26R) homozygous females [24]. β-Galactosidase staining of 10-μm frozen sections demonstrated sporadic Gdf9cre activity in the adult testis and Gdf9cre activity in oocytes, starting at the primordial follicle stage (Supplemental Fig. S1).

Wild-type adult ICR mice (Taconic) were bred to generate pups at Postnatal Day (P) 8 and P16. All lines of mice were kept on mixed genetic backgrounds. Noon (1200 h) on the day a mating plug was observed was designated E0.5. Embryos and UG ridges were dissected from timed matings. Somites were counted. Yolk sacs were removed for PCR genotyping and determination of sex with primers specific for the sex-determining gene on the Y chromosome (Sry) (forward: TTCCAGGAGGCACAGAGATT and reverse: GTCCCACTGCAGAAGGTTGT). The Institutional Animal Care and Use Committee of Columbia University Medical Center approved all animal protocols.

In Situ Hybridization, Immunofluorescence, and Histology

For section in situ hybridization (ISH), colorimetric immunohistochemistry (IHC), and histology, ovaries from postnatal mice were isolated and fixed in 4% paraformaldehyde. Following fixation, samples were infiltrated with sucrose, embedded in Tissue-Tek O.C.T. Compound (Sakura Fine Technical Co., Ltd.), snap frozen on dry ice in ethanol, and stored at −80°C. Frozen sections of 10 μm were made. Each ISH and IHC was performed at least three times, and five different ovaries were analyzed. For histology, sectioned ovaries from three to five mice of each genotype were stained with Harris modified hematoxylin (Fisher Scientific) and eosin (Sigma-Aldrich) (H&E).

Whole-mount and section ISH and colorimetric IHC on frozen sections were performed using standard protocols [25–27]. Antisense probes for Tbx2, Tbx4, Tbx5, and Pou5f1 were used as previously described [10], and sections were counterstained with Nuclear Fast Red (Sigma) unless otherwise indicated. ISH with sense RNA controls showed no background staining. For IHC, anti-PECAM antibody (#553370; 1:350; BD Biosciences), biotin goat anti-rat IgG (#559286; 1:750; BD Biosciences), the avidin/biotin blocking kit (Vector Labs), the Vectastain ABC kit, and the DAB substrate kit (Vector Labs) were used. Sections were counterstained with hematoxylin.

Two whole-mount immunofluorescence (IF) protocols were utilized. First, for single-fluorochrome IF, embryos at the 30- to 33-somite stage (E10.0) were dissected in cold PBS with 0.1% bovine serum albumin (BSA; Sigma-Aldrich), fixed in 4:1 methanol and dimethylsulfoxide (DMSO) at −20°C overnight, and stored in 100% methanol. Embryos were rehydrated and then incubated overnight at 4°C with anti-SSEA1 primary antibody (1:200; Developmental Studies Hybridoma Bank) in PBS/2% nonfat dry milk/0.5% Tween (PBSMT), followed by washes in PBSMT and an overnight incubation with goat anti mouse-IgG Alexa-Fluor 555 (1:250; Invitrogen). Embryos were dehydrated and cleared in methyl salicylate for viewing.

Second, for two-fluorochrome IF, embryos were dissected in PBS with 0.1% BSA, fixed in 0.5% paraformaldehyde at 4°C overnight, washed in cold PBS/0.1% Triton X-100 (PBT; Fisher Scientific), and stored in PBT/0.3% sodium azide (Sigma-Aldrich). Embryos were incubated with blocking solution (1% BSA/10% heat-inactivated serum/1% Triton X-100 in PBS) for 5 h at 4°C, followed by an overnight incubation at 4°C with the primary antibody, a wash with PBT, and an overnight incubation at 4°C with the secondary antibody. All antibodies were diluted in 1% BSA/2% heat-inactivated serum/1% Triton X-100 in PBS. Primary antibodies used included: anti-Stella (ab19878; 1:200; Abcam), anti-SSEA1 (1:200; Developmental Studies Hybridoma Bank), anti-phospho-Histone H3 (anti-phH3; H9908; 1:100; Sigma-Aldrich), and anti-cleaved caspase-3 (#9661; 1:200; Cell Signaling). Secondary antibodies, all diluted 1:250, included: goat anti-mouse-IgG Alexa-Fluor 488 (Invitrogen), donkey anti-rat-IgG Alexa-Fluor 594 (Invitrogen), donkey anti-rat-IgG Alexa-Fluor 488 (Invitrogen), and donkey anti-rabbit-IgG Alexa-Fluor 594 (Invitrogen). Samples were stained with 4′,6-diamidino-2-phenylindole (DAPI; Sigma-Aldrich) for nuclear visualization. Embryos at the four- to seven-somite stage (E8.0) were cleared and mounted with Vectashield (Vector Labs) for viewing. Embryos at the 23- to 25-somite (E9.5) and 29- to 33-somite (E10.0) stages were dehydrated and cleared in methyl salicylate for viewing. To confirm specificity of primary antibodies, stage-matched control embryos were processed as described above but were incubated with secondary antibodies only.

Depletion of Embryonic Germ Cells

Administration (i.p.) of busulfan (Sigma-Aldrich) was utilized to deplete embryonic gonads of germ cells. Pregnant female mice were injected with 150 μl of busulfan solution (20 mg/ml in 50% DMSO) or 50% DMSO in PBS (control) at E11.5. Gonads were removed from the embryos at E13.5 and processed for ISH with probes for Tbx4 and Pou5f1 as described above.

UG Ridge Culture

UG ridges were dissected from embryos at the 48- to 50-somite stage (E11–E11.5) in cold PBS with 0.1% BSA and cultured on 3-μm filters (Millipore) in media containing Dulbecco modified Eagle medium (Invitrogen), 10% fetal bovine serum (Hyclone), 1% penicillin/streptomycin (Invitrogen), and 2 μM 4-hydroxy (OH) tamoxifen (Sigma-Aldrich). After 3 days of culture at 37°C in 5% CO2, UG ridges were washed in cold PBS, fixed in 4% paraformaldehyde, dehydrated in a methanol series, and stored at −20°C until use.

Ovarian Reserve Measures

Ovaries were removed from mice at approximately 3 wk of age (P21–P23), fixed in Bouin solution (Sigma-Aldrich), embedded in paraffin, sectioned at 6 μm, and stained with H&E. Ovarian volume was determined by using the total number of 6-μm sections per ovary, assuming a spherical shape. In every fifth section, follicles containing the nucleolus of the oocyte were classified and counted. The total count was multiplied by five, to account for the fraction of the ovary sampled [28]. The following classification was used: a primordial follicle contains an oocyte with a single layer of flattened granulosa cells; a primary follicle has a single layer of cuboidal granulosa cells; pre-antral follicles have two or more layers of cuboidal granulosa cells without an antral cavity; and antral follicles have a fluid-filled cavity, the antrum [8, 9, 29]. Follicles were counted from four ovaries from each of the following genotypes: Gdf9cre; Tbx4fl/fl, Gdf9cre; Tbx4fl/−, Gdf9cre; Tbx4fl/+, Zp3cre; Tbx4fl/fl, and Zp3cre; Tbx4fl/+, representing three mice of each genotype.

Fertility Studies

Female mice 6 wk of age of the following genotypes were housed continuously with wild-type ICR males of proven fertility: Gdf9cre; Tbx4fl/fl, Gdf9cre; Tbx4fl/−, Gdf9cre; Tbx4fl/+, Zp3cre; Tbx4fl/−, and Zp3cre; Tbx4fl/+. The number of litters and numbers of pups per litter were recorded over a 6-mo period.

Imaging

Images of whole-mount samples were taken under bright field on a Nikon SMZ1500 microscope (Nikon). Section ISH was examined with a Nikon MICROPHOT-FXA microscope (Nikon), and images were captured using NIS-Elements D3.10 software. Images of whole-mount, single-flurorochrome IF samples were taken with the Nikon SMZ1500 microscope and the 25× objective (APO LWD 25x/1.10W) of a Nikon A1R MP multiphoton confocal microscope. Each embryo was imaged sagittally with an estimated optical section thickness of 600 nm and a 3-μm step size, selected to ensure that all cells would be counted. Overlapping images, two to four per embryo, were captured and stitched together digitally. The number of PGCs per embryo was determined by manually counting cells in the confocal stacks with NIS Elements version 4.000.07 software (Build 787; Nikon). For each image, counts were performed twice, with the investigator blind to the genotypes, and averaged. There was a <5% difference in the two counts for each image.

Images of whole-mount, two-fluorochrome IF samples were taken with the Nikon SMZ1500 microscope and the 25× objective (APO LWD 25x/1.10W) of a Nikon A1R MP confocal microscope over a depth of 200–300 μm, with an estimated optical section thickness of 1.6 μm and a 5-μm step size, selected to ensure that all cells would be counted. At the four- to seven-somite stage, the posterior end of each embryo was imaged along the dorsal-ventral axis. At the 23- to 25-somite and 29- to 33-somite stages, each embryo was imaged sagittally to sample the migratory and gonadal PGCs between the forelimb and hind limb buds. The number of PGCs and the number of proliferating (phH3) or apoptotic (cleaved caspase 3) PGCs were determined by manually counting cells in the confocal stacks with NIS elements version 4.000.07 (Build 787) software. Counts were performed with the investigator blind to the genotypes.

Statistics

Medians were compared using the Mann-Whitney U-test or the Kruskal-Wallis test for ANOVA. For normally distributed data, means ± SD, compared with the Student t-test, are reported. Statistical analyses were performed using Prism Version 5.0d (GraphPad).

RESULTS

Expression of Tbx4 in Germ Cells

Tbx4 expression in the bipotential embryonic gonad is first detected at E11.5 [10]. To determine if Tbx4 is expressed in embryonic germ cells or in the surrounding tissue, we analyzed Tbx4 and Pou5f1 expression following busulfan treatment of pregnant females. Busulfan is an alkylating agent that depletes germ cells. After busulfan administration, a reduction in Pou5f1 expression, a marker of germ cells, in gonads would be expected [10, 30]. We found that both Tbx4 and Pou5f1 expression were greatly reduced in embryonic ovaries after germ cell depletion (Fig. 1, A–D). These data are consistent with Tbx4 expression in female germ cells at E13.5. As shown in Figure 1, Tbx4 was expressed in oocytes at all stages of folliculogenesis: primordial follicles at P8 (Fig. 1E, yellow arrow); primary, secondary, and antral follicles at P16 (Fig. 1, F and G); and preovulatory follicles in 6-wk-old adult mice (Fig. 1H).

FIG. 1.

FIG. 1

Open in a new tab

Tbx4 is expressed in embryonic female germ cells and postnatal oocytes. A–D) Whole-mount ISH for Tbx4 and Pou5f1 on E13.5 ovaries after exposure to vehicle (control) or busulfan. Tbx4 (A) and Pou5f1 (B) were expressed throughout embryonic ovaries. After administration of busulfan to deplete germ cells, expression of both Tbx4 (C) and Pou5f1 (D) was reduced. E–H) ISH for Tbx4 on frozen sections of prepubertal (E–G) and adult (H) ovaries. Tbx4 was expressed in oocytes of primordial (yellow arrows, E), primary (F), secondary, and antral follicles (G) and in preovulatory, antral follicles (asterisk, H). a, antral; p, primary; s, secondary. Nuclear fast red counterstain was not used in E. Original magnification ×500 (A–D); bars = 10 μm (E), 50 μm (F and G), and 100 μm (H).

PGC Depletion in Embryos Lacking Tbx4

At E8.0, PGCs are present at the base of the allantois. Endogenous alkaline phosphatase staining was used to show that PGCs are in similar locations and in comparable numbers in wild-type and Tbx4-null embryos at E8.0 [14]. Between E8.5 and E11.0, PGCs migrate along the wall of the hindgut and dorsal mesentery into the gonads within the UG ridges. PGCs migrating through the dorsal mesentery and within the UG ridge at E10.0 (30–33 somites) were visualized with SSEA1, an antibody that reacts with a carbohydrate antigen on PGCs (Fig. 2) [31, 32], to determine if Tbx4 is required for germ cell development between E8.0 and E10.5, the stage beyond which Tbx4-null embryos die [14]. There was no increase in midline extragonadal PGCs in Tbx4−/− as compared to that of Tbx4+/− embryos at E10.0 (Fig. 2, A and B). Confocal imaging and 3D analysis were used to quantify the total number of PGCs in the UG ridges (Fig. 2, C and D). The mean number of PGCs in the UG ridges of Tbx4−/− embryos is significantly lower than that in stage-matched Tbx4+/− littermate controls (256 ± 78 vs. 418 ± 102; P = 0.002; Fig. 2E).

FIG. 2.

FIG. 2

Open in a new tab

PGCs are reduced in Tbx4-null embryos. Tbx4+/− and Tbx4−/− embryos at the 30- to 33-somite stage were isolated and stained with anti-SSEA1 antibody to identify PGCs. Right lateral views of the trunk region with anterior to the left are shown. PGCs are identified as white punctate staining, and the dotted yellow line (A) indicates the position of the gonad. There was no increase in midline extragonadal PGCs in Tbx4−/− embryos (B) as compared to Tbx4+/− (A) littermate controls. A representation of the stitched images (C, D) generated via extended depth of focus processing of confocal stacks is presented. E) The number of PGCs in individual embryos is shown. There were significantly fewer PGCs in Tbx4−/− embryos as compared to Tbx4+/− embryos. Data were compared with the Student t-test. Mean and SD are indicated. **P < 0.01. Original magnification ×700 (A and B); bars = 100 μm (C and D).

Neither Decreased Proliferation nor Increased Apoptosis Fully Explain PGC Reduction in Embryos Lacking Tbx4

To investigate the cellular mechanism underlying PGC depletion in Tbx4−/− embryos at E10.0, proliferation and apoptosis of PGCs were assessed. At the four- to seven-somite stage (E8.0), proliferating PGCs were detected as cells expressing both phH3 and Stella, a marker of PGCs expressed as early as E7.0 (Fig. 3A) [33]. The median percentage of proliferating PGCs is similar in Tbx4−/− embryos (5.4%; interquartile range [IQR] 0, 8) and stage-matched Tbx4+/− controls (2.9% [IQR 0.7, 6.3]) at the four- to seven-somite stage (Fig. 3E). At the 23- to 25-somite (E9.5) and the 29- to 33-somite (E10.0) stages, proliferating PGCs were detected as cells expressing both phH3 and SSEA1 (Fig. 3, B and C). The median percentage of proliferating PGCs is similar in Tbx4−/− embryos and stage-matched Tbx4+/− controls at both the 23- to 25-somite stage (4.4% [IQR 3.6, 8.7] vs. 7.1% [IQR 4.1, 8.3]) and the 29- to 33-somite stage (2.9% [IQR 0.7, 3.4] vs. 4.0% [IQR 1.9, 6.4]; Fig. 3, F and G). Apoptotic PGCs at the 29- to 33-somite stage were identified as cells expressing both cleaved caspase 3 and SSEA1 (Fig. 3D). The median percentage of apoptotic PGCs is slightly higher in Tbx4−/− embryos (2.0% [IQR 1.6, 3.4]) than in stage-matched Tbx4+/− controls (0.9% [IQR 0.3, 1.9]), but the difference does not reach statistical significance (Fig. 3H). Taken together, these results suggest that neither reduced proliferation nor increased apoptosis is responsible for the reduction in PGCs observed at E10.0 in embryos lacking Tbx4.

FIG. 3.

FIG. 3

Open in a new tab

PGC proliferation and apoptosis are not significantly altered in Tbx4-null embryos. Double-staining IF of whole-mount embryos was performed to detect proliferating and apoptotic PGCs. A–D) Representative IF images from single optical sections of the confocal stacks are shown, and insets highlight double positive cells. A) PGCs are labeled with Stella antibody (red), and proliferating cells are labeled with phH3 (green). B, C) PGCs are labeled with SSEA1 antibody (green), and proliferating cells are labeled with phH3 (red). D) PGCs are labeled with SSEA1 antibody (green), apoptotic cells are labeled with cleaved caspase 3 (red), and DAPI identifies all nuclei. E–G) The percentages of phH3+ PGCs were similar in Tbx4+/− and Tbx4−/− embryos at the four- to seven-somite (E8.0), 23- to 25-somite (E9.5), and 29- to 33-somite (E10.0) stages. H) The percentage of cleaved caspase 3+ PGCs at the 29- to 33-somite stage was slightly higher in Tbx4−/− embryos. For A–D, dorsal is down in all images except C, where dorsal is to the right. Bars = 100 μm (A–D).

Loss of Tbx4 Does Not Prevent Sexual Differentiation of the Gonad In Vitro

To investigate the ability of bipotential germ cells lacking Tbx4 to differentiate into sexually dimorphic male and female germ cells, isolated UG ridges from Tbx4fl/fl embryos at the 48- to 50-somite stage with and without the creER transgene were cultured in the presence of 4-OH tamoxifen for 3 days (Fig. 4). Similar to prior studies with tamoxifen administration in vivo, Tbx4fl/fl was roughly 75%–95% excised after 24 h in vitro as determined by PCR for Tbx4fl and excised alleles (data not shown) [20]. Thus, complete excision is anticipated by 3 days. Whole-mount ISH with Pou5f1 was used to visualize germ cells. The wild-type pattern of Pou5f1 expression in cultured UG ridges from female embryos differs from that of males. In females, Pou5f1 expression is largely homogeneous throughout the gonad (Fig. 4A), whereas in males, Pou5f1-positive germ cells are clustered to give a testis cord-like pattern (Fig. 4C). The pattern of Pou5f1 expression in UG ridge cultures from Tbx4fl/fl female (Fig. 4B) and male (Fig. 4D) embryos with the creER transgene is similar to those of controls. This finding indicates that following the migration of germ cells into the UG ridges, germ cell sex determination and sexual differentiation of the gonad occurs in the absence of Tbx4, in vitro.

FIG. 4.

FIG. 4

Open in a new tab

Loss of Tbx4 does not prevent sexual differentiation of the gonad in vitro. UG ridges isolated at the 48- to 50-somite stage were cultured for 3 days in the presence of 4-OH tamoxifen. Pou5f1 expression was analyzed using ISH on cultured UG ridges. Tbx4 conditional alleles in cultured UG ridges were excised from female (B) and male (D) embryos using 4-OH tamoxifen with creER. In vitro differentiation of male gonads with Pou5f1-stained germ cells arranged in a testis cord-like pattern occurred in control (C) and creER-containing Tbx4 mutant (D) UG ridge cultures. A, anterior; P, posterior; g, gonad; k, kidney. Original magnification ×400 (A–D).

Normal Ovarian Histology in Mice with Oocyte-Specific Deletion of Tbx4

To deactivate Tbx4 exclusively in oocytes, we used two oocyte-specific Cre recombinases, Gdf9cre and Zp3cre, that express Cre recombinase in oocytes of primordial follicles and primary follicles, respectively [22, 23, 34]. Ovarian histology was analyzed by H&E staining, and vasculature was analyzed by PECAM IHC in ovaries from 6-wk-old mice (Fig. 5). Ovaries from mice with oocyte-specific deletion of Tbx4 were compared to Gdf9cre; Tbx4fl/+ and Zp3cre; Tbx4fl/+ littermate controls. Ovarian histology and PECAM expression in all genotypes are similar (Fig. 5, A–D, data are not shown for Gdf9cre; Tbx4fl/fl and Zp3cre; Tbx4fl/fl). Primordial (Fig. 5, E, J, and O, arrows), primary (Fig. 5, F, K, and P), secondary (Fig. 5, G, L, and Q), and antral (Fig. 5, H, M, and R) follicles from control and Tbx4 mutant ovaries are shown. PECAM expression was used to examine the vasculature of the corpora lutea that form after ovulation of oocytes (Fig. 5, I, N, and S). The pattern of PECAM expression is similar in corpora lutea from Gdf9cre; Tbx4fl/−, Zp3cre; Tbx4fl/−, and control ovaries. In order to rule out compensation of Tbx4 function by other T-box genes, Tbx2 and Tbx5 expression was assessed. Neither Tbx2 nor Tbx5 is expressed in ovaries from Gdf9cre; Tbx4fl/− mice (Supplemental Fig. S2). The similar ovarian histology and pattern of PECAM expression in all genotypes suggests that oocyte-specific deletion of Tbx4 does not impair folliculogenesis, ovulation, or corpus luteum formation.

FIG. 5.

FIG. 5

Open in a new tab

Normal follicular development and corpus luteum formation in adult ovaries with oocyte-specific deletion of Tbx4. Morphological analysis of ovaries from 6-wk-old control (A and E–H), Gdf9cre; Tbx4fl/− (C and J–M), and Zp3cre; Tbx4fl/− (O–R) mice was performed. Primordial (E, J, and O, yellow arrows), primary (F, K, and P), secondary (G, L, and Q), and antral (H, M, and R) follicles from control and Tbx4 mutant ovaries are shown. Ovaries were stained with anti-PECAM antibody to visualize vasculature (brown stain) of corpora lutea in control (B and I), Gdf9cre; Tbx4fl/− (D and N), and Zp3cre; Tbx4fl/− (S) mice. The vascular pattern in corpora lutea of control (I), Gdf9cre; Tbx4fl/− (N), and Zp3cre; Tbx4f/− mutant (S) ovaries is similar. Bar in D = 100 μm for A–D; bars in O and P = 10 μm for E, F, J, K, O and P; and bars in Q, R, and S = 100 μm for G–I, L–N, and Q–S. a, antral; cl, corpus luteum.

Ovarian Reserve Is Not Diminished in Mice with Oocyte-Specific Deletion of Tbx4

The presence of all types of ovarian follicles in adult mice does not reflect baseline ovarian reserve, the capacity of the ovary to provide fertilizable oocytes. Ovarian reserve can be assessed by measuring ovarian volume and/or quantifying the number and types of follicles present in an ovary. To assess ovarian reserve in mice with oocyte-specific deletion of Tbx4, follicles were classified and counted at approximately 3 wk of age (P21–P23). Median ovarian volumes (Table 1) and the median number of follicles are similar in Gdf9cre mutants (Gdf9cre; Tbx4fl/fl and Gdf9cre; Tbx4fl/−) compared to those in Gdf9cre controls (Fig. 6A and Table 1) and in Zp3cre; Tbx4fl/fl mutants compared to those in Zp3cre controls (Fig. 6B and Table 1). These data suggest that postnatal, oocyte-specific deletion of Tbx4 does not deplete ovarian reserve.

TABLE 1.

Measures of ovarian reserve in mice with oocyte-specific deletion of Tbx4 compared to controls.*

graphic file with name i0006-3363-89-6-148-t01.jpg

Open in a new tab
* 

No statistically significant differences were noted when median ovarian volumes or median follicle numbers were compared.

FIG. 6.

FIG. 6

Open in a new tab

Oocyte-specific deletion of Tbx4 does not diminish ovarian reserve. Histomorphometric analysis of ovaries from 3-wk-old mice (P21–P23) was performed to assess ovarian reserve. A) The median number of primordial, primary, preantral, and antral follicles per ovary was similar in Gdf9cre control, Gdf9cre; Tbx4fl/fl, and Gdf9cre; Tbx4fl/− mice. B) The median number of primordial, primary, preantral, and antral follicles per ovary was similar in Zp3cre control and Zp3cre; Tbx4fl/fl mice. Box plots with median, minimum, and maximum values are shown. Data were compared with the Kruskal-Wallis and Mann-Whitney U-tests. No significant differences were observed.

Normal Fertility in Mice with Oocyte-Specific Deletion of Tbx4

To determine if Tbx4 excision was complete in oocytes that yielded offspring upon fertilization, pups from matings of Gdf9cre; Tbx4fl/fl or Zp3cre; Tbx4fl/− female mice with wild-type males were tested by PCR analysis of genomic DNA to assess the Tbx4 genotype. Wild-type and excised Tbx4 DNA, but no Tbx4fl DNA, was amplified in all of the offspring (Fig. 7A), indicating that the Tbx4fl allele was completely excised by Gdf9cre and Zp3cre recombinases in the oocytes that resulted in live offspring (data not shown for Zp3cre).

FIG. 7.

FIG. 7

Open in a new tab

Oocyte-specific deletion of Tbx4 does not impair fertility. Gdf9cre; Tbx4fl/fl and Zp3cre; Tbx4fl/fl female mice were mated with wild-type males to assess excision of Tbx4 in oocytes yielding offspring. A) PCR analysis of genomic DNA shows excision of Tbx4 in all offspring. B) Litter sizes from Gdf9cre mutant females (Gdf9cre; Tbx4fl/fl and Gdf9cre; Tbx4fl/−) and Gdf9 controls (Gdf9cre; Tbx4fl/+) were compared. Litter sizes from Gdf9cre mutant females (n = 38 litters) were similar to those of Gdf9 control females (n = 23 litters), with P = 0.322. C) Litter sizes from Zp3cre mutant females (Zp3cre; Tbx4fl/−, n = 14 litters) were similar to Zp3cre controls (Zp3cre; Tbx4fl/+, n = 6 litters), with P = 0.122. Data were compared with the Mann-Whitney U-test. Median litter sizes and interquartile ranges are indicated. cond, conditional; wt, wild type.

To test the fertility of mice with deletion of Tbx4 in oocytes, 6-wk-old Tbx4fl/−, Tbx4fl/fl, and control Tbx4fl/+ female mice carrying one or the other oocyte-specific Cre transgene were bred to wild-type stud males, and litters were recorded for 6 mo. Comparing months 1–3 to months 4–6, the median litter size in each genotype remained the same (i.e., Gdf9cre; Tbx4fl/fl: 11 vs. 11 pups; Gdf9cre; Tbx4fl/−: 9.5 vs. 11 pups; Zp3cre; Tbx4fl/−: 8 vs. 9 pups), suggesting that there was no age-related decline in fertility. As the median litter size from Gdf9cre; Tbx4fl/fl and Gdf9cre; Tbx4fl/− mice are similar, litters from these genotypes were combined for further analyses. The median litter size from Gdf9cre mutants (Gdf9cre; Tbx4fl/fl and Gdf9cre; Tbx4fl/−) and Gdf9cre controls (10 vs. 11 pups; Fig. 7B) and the median litter size from Zp3cre mutants and Zp3cre controls (10 vs. 8.5 pups; Fig. 7C) are similar. Taken together, these data suggest that expression of Tbx4 in oocytes of primordial and primary follicles is not required for normal fertility.

DISCUSSION

The establishment and maintenance of female fertility is a complex process that includes development of bipotential germ cells capable of committing to the female lineage, formation of ovarian follicles, bidirectional communication between oocytes and somatic cells within follicles to promote oocyte maturation, and extrusion of mature oocytes from follicles to achieve pregnancy. Normal oocyte development is absolutely required for ovarian follicle formation; if germ cells fail to enter the UG ridges, somatic cells do not differentiate and the ovaries undergo atresia [35]. Knowledge of the molecular framework underlying the development and differentiation of ooctyes has lagged behind in comparison to what is known about the development of sperm. We sought to contribute to the understanding of the molecular mechanisms underlying development of the female reproductive system by investigating the requirement for the transcription factor Tbx4 in the female germ line. Our previous work established the expression of Tbx4 in embryonic germ cells and postnatal, prepubertal oocytes [10].

Between E8.0 and E11.5, migration and cellular proliferation are the key processes involved in PGC development. Prior to E9.0 and entry into the hindgut, PGC migration is thought to be passive as PGCs move with the invaginating endoderm. In contrast, PGCs in the hindgut are motile [2, 36, 37]. Active PGC migration from the hindgut to the UG ridges involves cellular contact with other PGCs, communication with other cells and molecules that promote PGC survival, and interactions with the extracellular matrix (ECM) via glycoproteins such as laminin and integrins [2, 37, 38]. At these developmental stages, cellular proliferation occurs with a PGC doubling time of approximately 16–17 h, and there is minimal apoptosis of PGCs [37, 39].

Our results show that embryos lacking Tbx4 have significantly fewer PGCs in the UG ridges at E10.0, with no reduction in cellular proliferation, as measured by the percentage of cells in mitosis. However, it is important to consider that even a slight increase in PGC doubling time, starting at E8.0, could result in fewer PGCs at later developmental stages. The slight increase in PGC apoptosis, as measured by the percentage of cells expressing cleaved caspase 3, does not support a statistically significant increase in PGC apoptosis in embryos lacking Tbx4. As expression of cleaved caspase 3 does not reflect cumulative PGC apoptosis, the slight increase in PGC apoptosis we observe could result in fewer PGCs over time. Although we did not observe an increase in extragonadal midline PGCs at E10.0, which would indicate slower or impaired migration, PGC migration may be impaired earlier, as Tbx4 mutant embryos are known to lack some ECM components [40]. These processes have not as yet been investigated but may contribute to the reduction in gonadal PGCs in embryos lacking Tbx4.

Using tissue-nonspecific alkaline phosphatase Cre recombinase (TNAPCre), a PGC-specific Cre recombinase, it has been shown that a reduction in PGCs in the gonads by E11.5 does not always result in a reduced number of postnatal oocytes or infertility. Loss of either Pou5f1 [41] or REST, a transcriptional regulator that promotes PGC survival [42], leads to apoptosis of PGCs by E10. Whereas adult female mice with PGC-specific Pou5f1 deletion are depleted of oocytes [41, 42], mice with PGC-specific deletion of REST have normal fertility.

In order to determine if a reduced number of PGCs at E10.0 due to early deletion of Tbx4 could affect ovarian reserve and fertility, we considered Cre recombinases specific to PGCs but found them to have limited utility for excising Tbx4. TNAPCre is expressed in PGCs with a recombination efficiency of approximately 60%, and is expressed in the placenta after E10 [43]. Given the requirement for Tbx4 in the developing allantois [14, 44], we were unable to use TNAPCre to delete Tbx4 in PGCs. Prdm1-cre is expressed in 55%–76% of PGCs by E7.5 [45]. We initiated studies with Prdm1-cre. However, Prdm1-cre induced early embryonic lethality when combined with Tbx4fl/fl or R26Rfl/fl reporter mice (data not shown) and, therefore, could not be used for analysis of Tbx4 function. Thus, with existing PGC-specific Cre transgenes, we cannot determine if deletion of Tbx4 in PGCs will affect female fertility.

In vitro development of the bipotential gonads into morphologically distinguishable testes and ovaries has been well described [46, 47]. In vivo, Sertoli cells, differentiated somatic cells in the male gonad, aggregate around PGCs to form testis cords, and by E13.5 male and female gonads are morphologically distinct. Germ cells differentiate according to their somatic cell environment, e.g., E11.5 female germ cells cultured with UG ridges from male embryos will develop into prospermatogonia or immature sperm [3, 6]. Based on the pattern of Pou5f1 expression, our data indicate that in vitro differentiation of the gonad into morphologically distinct testes and ovaries occurs when Tbx4 is deleted starting at E11.5. This finding indicates that Tbx4-null germ cells are competent to respond to paracrine signals and suggests that bipotential germ cells lacking Tbx4 can differentiate into male and female germ cells in vivo.

Gdf9cre and Zp3cre are two well-described oocyte-specific cre recombinase transgenes that have slightly different patterns of cre expression and activity; Gdf9cre activity begins in primordial follicles by P3, and Zp3cre activity begins in primary follicles by P5 [22]. Expression of both Gdf9cre and Zp3cre continues through later stages of follicle development and into adulthood. Stage-specific deletion of genes in oocytes may produce different phenotypes. Whereas deletion of Pten in oocytes of primordial follicles with Gdf9cre causes premature activation of primordial follicles and POI in mice [48], deletion of Pten in primary follicles using Zp3cre does not affect ovarian follicular development or fertility [49].

Tbx4 is expressed in oocytes of primordial and primary follicles. Because Gdf9cre and Zp3cre are expressed at different stages of oocyte development [22], we utilized both Cre recombinases to investigate the requirement for Tbx4 in oocytes during folliculogenesis and to determine if loss of Tbx4 in oocytes impairs fertility. In Gdf9cre; Tbx4fl/− and Zp3cre; Tbx4fl/− adult females, primordial, primary, secondary, and antral follicles form, ovulation occurs, and corpus luteum formation is not impaired. We determined that ovarian reserve, as measured by ovarian volume and total number of follicles, is not diminished in Gdf9cre; Tbx4fl/− or Zp3cre; Tbx4fl/fl mice at 3 wk of age, just prior to the onset of puberty and ovulatory estrous cycles. We also demonstrated that oocyte-specific deletion of Tbx4 in primordial and primary follicles does not obviously impair fertility. To address the concern that fertility could have been conferred by the oocytes that escaped Cre-mediated deletion of Tbx4, we showed that all oocytes that gave rise to offspring lacked Tbx4 and that litter sizes resulting were normal. One limitation of our postnatal analyses worthy of consideration is that we demonstrated Tbx4 RNA but not protein expression in postnatal oocytes. If Tbx4 mRNA is not translated into Tbx4 protein in oocytes, we would not expect a phenotype after oocyte-specific deletion of Tbx4.

Although postnatal, oocyte-specific deletion of Tbx4 does not affect fertility, the reduced number of PGCs resulting from loss of Tbx4 during embryonic development could affect long-term fertility in adults. Because of embryonic lethality and the limited usefulness of Cre recombinases that excise Tbx4 in PGCs, these fertility studies cannot be readily performed in mice. However, our data set the stage for performing TBX4 mutation analyses in women with POI. Nonsyndromic POI affects 1:100 women <40 yr of age and 1:1000 women <30 yr of age, and genetic alterations have been found in 20%–25% of cases that were initially considered idiopathic [50]. If TBX4 mutations are found to be associated with POI, murine studies to determine the mechanisms whereby PGCs are decreased in Tbx4 mutant embryos would contribute to our understanding of the molecular framework underlying POI.

Supplementary Material

Supplemental Data
supp_89_6_148__index.html (2KB, html)

ACKNOWLEDGMENT

We would like to thank members of the Papaiaonnou laboratory for critical reading of the manuscript. The SSEA1 monoclonal antibody developed by David Solter and Barbara Knowles was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the Eunice Kennedy Shriver National Institute of Child Health and Human Development and maintained by The University of Iowa, Department of Biology, Iowa City, IA 52242.

Footnotes

1

Supported by the National Institutes of Health/NICHD (to N.C.D. and V.E.P.) and the Robert Wood Johnson Foundation (to N.C.D.).

REFERENCES

  1. Simpson JL. Genetic and phenotypic heterogeneity in ovarian failure: overview of selected candidate genes. Ann N Y Acad Sci 2008; 1135: 146–154. [DOI] [PubMed] [Google Scholar]
  2. McLaren A. Primordial germ cells in the mouse. Dev Biol 2003; 262: 1–15. [DOI] [PubMed] [Google Scholar]
  3. Adams IR, McLaren A. Sexually dimorphic development of mouse primordial germ cells: switching from oogenesis to spermatogenesis. Development 2002; 129: 1155–1164. [DOI] [PubMed] [Google Scholar]
  4. McLaren A. Germ and somatic cell lineages in the developing gonad. Mol Cell Endocrinol 2000; 163: 3–9. [DOI] [PubMed] [Google Scholar]
  5. Bowles J, Koopman P. Retinoic acid, meiosis and germ cell fate in mammals. Development 2007; 134: 3401–3411. [DOI] [PubMed] [Google Scholar]
  6. McLaren A. Germ cells and germ cell sex. Philos Trans R Soc Lond B Biol Sci 1995; 350: 229–233. [DOI] [PubMed] [Google Scholar]
  7. Pepling ME, Spradling AC. Mouse ovarian germ cell cysts undergo programmed breakdown to form primordial follicles. Dev Biol 2001; 234: 339–351. [DOI] [PubMed] [Google Scholar]
  8. Pedersen T, Peters H. Proposal for a classification of oocytes and follicles in the mouse ovary. J Reprod Fertil 1968; 17: 555–557. [DOI] [PubMed] [Google Scholar]
  9. Nicholas CR, Chavez SL, Baker VL. Reijo Pera RA. Instructing an embryonic stem cell-derived oocyte fate: lessons from endogenous oogenesis. Endocr Rev 2009; 30: 264–283. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Douglas NC, Heng K, Sauer MV, Papaioannou VE. Dynamic expression of Tbx2 subfamily genes in development of the mouse reproductive system. Dev Dyn 2012; 241: 365–375. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Naiche LA, Harrelson Z, Kelly RG, Papaioannou VE. T-box genes in vertebrate development. Annu Rev Genet 2005; 39: 219–239. [DOI] [PubMed] [Google Scholar]
  12. Wardle FC, Papaioannou VE. Teasing out T-box targets in early mesoderm. Curr Opin Genet Dev 2008; 18: 418–425. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Conlon FL, Yutzey KE. T-box factors. : Rosenthal N, Harvey RP. (eds.), Heart Development and Regeneration. Burlington, MA: Academic Press; 2010: 651–671. [Google Scholar]
  14. Naiche LA, Papaioannou VE. Loss of Tbx4 blocks hindlimb development and affects vascularization and fusion of the allantois. Development 2003; 130: 2681–2693. [DOI] [PubMed] [Google Scholar]
  15. Bongers EM, Duijf PH, van Beersum SE, Schoots J, Van Kampen A, Burckhardt A, Hamel BC, Losan F, Hoefsloot LH, Yntema HG, Knoers NV, van Bokhoven H. Mutations in the human TBX4 gene cause small patella syndrome. Am J Hum Genet 2004; 74: 1239–1248. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Tevosian SG, Manuylov NL. To β or not to β: canonical β-catenin signaling pathway and ovarian development. Dev Dyn 2008; 237: 3672–3680. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Arora R, Metzger RJ, Papaioannou VE. Multiple roles and interactions of Tbx4 and Tbx5 in development of the respiratory system. PLoS Genet 2012; 8: e1002866. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Wilhelm D, Palmer S, Koopman P. Sex determination and gonadal development in mammals. Physiol Rev 2007; 87: 1–28. [DOI] [PubMed] [Google Scholar]
  19. Kim Y, Kobayashi A, Sekido R, DiNapoli L, Brennan J, Chaboissier MC, Poulat F, Behringer RR, Lovell-Badge R, Capel B. Fgf9 and Wnt4 act as antagonistic signals to regulate mammalian sex determination. PLoS Biol 2006; 4: e187. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Naiche LA, Papaioannou VE. Tbx4 is not required for hindlimb identity or post-bud hindlimb outgrowth. Development 2007; 134: 93–103. [DOI] [PubMed] [Google Scholar]
  21. de Luca C, Kowalski TJ, Zhang Y, Elmquist JK, Lee C, Kilimann MW, Ludwig T, Liu SM, Chua SC Jr. Complete rescue of obesity, diabetes, and infertility in db/db mice by neuron-specific LEPR-B transgenes. J Clin Invest 2005; 115: 3484–3493. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Lan ZJ, Xu X, Cooney AJ. Differential oocyte-specific expression of Cre recombinase activity in GDF-9-iCre, Zp3cre, and Msx2Cre transgenic mice. Biol Reprod 2004; 71: 1469–1474. [DOI] [PubMed] [Google Scholar]
  23. de Vries WN, Binns LT, Fancher KS, Dean J, Moore R, Kemler R, Knowles BB. Expression of Cre recombinase in mouse oocytes: a means to study maternal effect genes. Genesis 2000; 26: 110–112. [PubMed] [Google Scholar]
  24. Soriano P. Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat Genet 1999; 21: 70–71. [DOI] [PubMed] [Google Scholar]
  25. Grieshammer U, Le M, Plump AS, Wang F, Tessier-Lavigne M, Martin GR. SLIT2-mediated ROBO2 signaling restricts kidney induction to a single site. Dev Cell 2004; 6: 709–717. [DOI] [PubMed] [Google Scholar]
  26. Wilkinson DG, Nieto MA. Detection of messenger RNA by in situ hybridization to tissue sections and whole mounts. Methods Enzymol 1993; 225: 361–373. [DOI] [PubMed] [Google Scholar]
  27. Douglas NC, Tang H, Gomez R, Pytowski B, Hicklin DJ, Sauer CM, Kitajewski J, Sauer MV, Zimmermann RC. Vascular endothelial growth factor receptor 2 (VEGFR-2) functions to promote uterine decidual angiogenesis during early pregnancy in the mouse. Endocrinology 2009; 150: 3845–3854. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Tilly JL. Ovarian follicle counts—not as simple as 1, 2, 3. Reprod Biol Endocrinol 2003; 1: 11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Balla A, Danilovich N, Yang Y, Sairam MR. Dynamics of ovarian development in the FORKO immature mouse: structural and functional implications for ovarian reserve. Biol Reprod 2003; 69: 1281–1293. [DOI] [PubMed] [Google Scholar]
  30. Ross A, Munger S, Capel B. Bmp7 regulates germ cell proliferation in mouse fetal gonads. Sex Dev 2007; 1: 127–137. [DOI] [PubMed] [Google Scholar]
  31. Marani E, van Oers JW, Tetteroo PA, Poelmann RE, van der Veeken J, Deenen MG. Stage specific embryonic carbohydrate surface antigens of primordial germ cells in mouse embryos: FAL (S.S.E.A.-1) and globoside (S.S.E.A.-3). Acta Morphol Neerl Scand 1986; 24: 103–110. [PubMed] [Google Scholar]
  32. Laird DJ, Altshuler-Keylin S, Kissner MD, Zhou X, Anderson KV. Ror2 enhances polarity and directional migration of primordial germ cells. PLoS Genet 2011; 7: e1002428. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Saitou M, Barton SC, Surani MA. A molecular programme for the specification of germ cell fate in mice. Nature 2002; 418: 293–300. [DOI] [PubMed] [Google Scholar]
  34. Sun QY, Liu K, Kikuchi K. Oocyte-specific knockout: a novel in vivo approach for studying gene functions during folliculogenesis, oocyte maturation, fertilization, and embryogenesis. Biol Reprod 2008; 79: 1014–1020. [DOI] [PubMed] [Google Scholar]
  35. McLaren A. Development of the mammalian gonad: the fate of the supporting cell lineage. Bioessays 1991; 13: 151–156. [DOI] [PubMed] [Google Scholar]
  36. Molyneaux KA, Stallock J, Schaible K, Wylie C. Time-lapse analysis of living mouse germ cell migration. Dev Biol 2001; 240: 488–498. [DOI] [PubMed] [Google Scholar]
  37. Bendel-Stenzel M, Anderson R, Heasman J, Wylie C. The origin and migration of primordial germ cells in the mouse. Semin Cell Dev Biol 1998; 9: 393–400. [DOI] [PubMed] [Google Scholar]
  38. Gomperts M, Garcia-Castro M, Wylie C, Heasman J. Interactions between primordial germ cells play a role in their migration in mouse embryos. Development 1994; 120: 135–141. [DOI] [PubMed] [Google Scholar]
  39. Tam PP, Snow MH. Proliferation and migration of primordial germ cells during compensatory growth in mouse embryos. J Embryol Exp Morphol 1981; 64: 133–147. [PubMed] [Google Scholar]
  40. Arora R, Papaioannou VE. The murine allantois: a model system for the study of blood vessel formation. Blood 2012; 120: 2562–2572. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Kehler J, Tolkunova E, Koschorz B, Pesce M, Gentile L, Boiani M, Lomeli H, Nagy A, McLaughlin KJ, Scholer HR, Tomilin A. Oct4 is required for primordial germ cell survival. EMBO Rep 2004; 5: 1078–1083. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Okamura D, Mochizuki K, Taniguchi H, Tokitake Y, Ikeda M, Yamada Y, Tournier C, Yamaguchi S, Tada T, Scholer HR, Matsui Y. REST. and its downstream molecule Mek5 regulate survival of primordial germ cells. Dev Biol 2012; 372: 190–202. [DOI] [PubMed] [Google Scholar]
  43. Hammond SS, Matin A. Tools for the genetic analysis of germ cells. Genesis 2009; 47: 617–627. [DOI] [PubMed] [Google Scholar]
  44. Arora R, del Alcazar CM, Morrisey EE, Naiche LA, Papaioannou VE. Candidate gene approach identifies multiple genes and signaling pathways downstream of Tbx4 in the developing allantois. PLoS One 2012; 7: e43581. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Ohinata Y, Payer B, O'Carroll D, Ancelin K, Ono Y, Sano M, Barton SC, Obukhanych T, Nussenzweig M, Tarakhovsky A, Saitou M, Surani MA. Blimp1 is a critical determinant of the germ cell lineage in mice. Nature 2005; 436: 207–213. [DOI] [PubMed] [Google Scholar]
  46. Tilmann C, Capel B. Mesonephric cell migration induces testis cord formation and Sertoli cell differentiation in the mammalian gonad. Development 1999; 126: 2883–2890. [DOI] [PubMed] [Google Scholar]
  47. Martineau J, Nordqvist K, Tilmann C, Lovell-Badge R, Capel B. Male-specific cell migration into the developing gonad. Curr Biol 1997; 7: 958–968. [DOI] [PubMed] [Google Scholar]
  48. Reddy P, Liu L, Adhikari D, Jagarlamudi K, Rajareddy S, Shen Y, Du C, Tang W, Hamalainen T, Peng SL, Lan ZJ, Cooney AJ, et al. Oocyte-specific deletion of Pten causes premature activation of the primordial follicle pool. Science 2008; 319: 611–613. [DOI] [PubMed] [Google Scholar]
  49. Jagarlamudi K, Liu L, Adhikari D, Reddy P, Idahl A, Ottander U, Lundin E, Liu K. Oocyte-specific deletion of Pten in mice reveals a stage-specific function of PTEN/PI3K signaling in oocytes in controlling follicular activation. PLoS One 2009; 4: e6186. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Persani L, Rossetti R, Cacciatore C. Genes involved in human premature ovarian failure. J Mol Endocrinol 2010; 45: 257–279. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental Data
supp_89_6_148__index.html (2KB, html)
supp_113.107649_107649SupDataV2.pdf (3.3MB, pdf)

Articles from Biology of Reproduction are provided here courtesy of Oxford University Press

RESOURCES