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First published online 3 August 2006
doi: 10.1242/dev.02506

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Primordial germ cell deficiency in the connexin 43 knockout mouse arises from apoptosis associated with abnormal p53 activation

Laboratory of Developmental Biology, National Heart Lung and Blood Institute, National Institutes of Health, Building 50/Room 4537, 9000 Rockville Pike, Bethesda, MD 20892, USA.

^* Author for correspondence (e-mail: loc{at}nhlbi.nih.gov)

Accepted 21 June 2006

	SUMMARY

TOP SUMMARY INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Connexin 43 knockout (Cx431KO) mice exhibit germ cell deficiency, but the underlying cause for the germ cell defect was unknown. Using an Oct4-GFP reporter transgene, we tracked the distribution and migration of primordial germ cells (PGCs) in the Cx431KO mouse embryo. Analysis with dye injections showed PGCs are gap-junction-communication competent, with dye coupling being markedly reduced in Cx431-deficient PGCs. Time-lapse videomicroscopy and motion analysis showed that the directionality and speed of cell motility were reduced in the Cx431KO PGCs. This was observed both in E8.5 and E11.5 embryos. By contrast, PGC abundance did not differ between wild-type and heterozygous/homozygous Cx431KO embryos until E11.5, when a marked reduction in PGC abundance was detected in the homozygous Cx431KO embryos. This was accompanied by increased PGC apoptosis and increased expression of activated p53. Injection of -pifithrin, a p53 antagonist, inhibited PGC apoptosis and prevented the loss of PGC. Analysis using a cell adhesion assay indicated a reduction in ß1-integrin function in the Cx431KO PGCs. Together with the abnormal activation of p53, these findings suggest the possibility of anoikis-mediated apoptosis. Overall, these findings show Cx431 is essential for PGC survival, with abnormal p53 activation playing a crucial role in the apoptotic loss of PGCs in the Cx431KO mouse embryos.

Key words: PGC, Cx43, Migration, p53, ß1 integrin, Apoptosis, Mouse

	INTRODUCTION

TOP SUMMARY INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The connexin multigene family encode proteins that oligomerize to form gap junction membrane channels that span the extracellular space. They allow the direct transfer of ions, metabolites and other small second messengers between neighboring cells (Bruzzone et al., 1996; White and Paul, 1999). The connexin gene Gja1 encodes a 43 kd protein known as connexin 43 or Cx43 (Goodenough et al., 1996; Sohl and Willecke, 2004), also known as ₁ connexin (Kumar and Gilula, 1996), and is referred to here as Cx431. Studies of knockout mouse models have revealed an essential role for a number of the connexins in development and disease (White and Paul, 1999). Cx431 was shown to be essential in heart development, as the Cx431 knockout mice Gja1^tm1Kdr (Cx431KO) die at birth due to conotruncal heart malformations and pulmonary outflow obstruction (Reaume et al., 1995). Cx431KO mice also displayed gonadal defects, with newborn pups exhibiting hypoplastic ovaries and testes deficient in primordial germ cells (PGCs) (Juneja et al., 1999; Roscoe et al., 2001).

In mouse embryos, cytochemical detection of alkaline-phosphatase activity indicated that PGCs first appear at embryonic day (E) 7.0, in the extra-embryonic mesoderm at the base of the allantois (Bendel-Stenzel et al., 1998). However, PGCs may be linage specified earlier, as PGCs emerge when epiblasts from E5.5 embryos are co-cultured with extra-embryonic ectoderm (Yoshimizu et al., 2001). From the allantois, PGCs migrate into the hindgut endoderm (E8.0), and travel along the dorsal mesentery (E9.5), finally arriving at and populating the genital ridges (E10.5-E11.5) (Bendel-Stenzel et al., 1998; McLaren, 2003). From a founding population of about ten cells at E7.0 (Ginsburg et al., 1990), PGCs rapidly expand to approximately 25,000 cells at E13.5, when gonadal differentiation begins in conjunction with sex determination (Tam and Snow, 1981). Recent studies have provided extensive evidence of an essential role for Cx431 in gonadogenesis. Cx431 is seen at all stages of ovary development (Perez-Armendariz et al., 2003) and is required for the growth and expansion of granulosa cells. Ovaries in the Cx431KO mice are PGC deficient (Juneja et al., 1999). In the absence of Cx431, granulosa cells are not functionally coupled and stop growing at an early pre-antral stage (Gittens et al., 2003). Lack of Cx431 restricted to the granulosa cells is sufficient to prevent the normal development of both oocyte and follicle, but chimera studies showed that Cx431 expression in oocytes is not required for oocytes to reach maturity (Gittens and Kidder, 2005). These findings indicate an essential role for Cx431 in folliculogenesis. Cx431 is also found at all stages of testicular development, and provides gap junctions that link germ cells with Sertoli cells (Perez-Armendariz et al., 2001). Testes isolated from Cx431-deficient fetuses display a `Sertoli cell only' phenotype characterized by an almost complete lack of germ cells (Roscoe et al., 2001).

Cx431KO embryos are known to be PGC deficient from E11.5, the time when genital ridges are first formed (Juneja et al., 1999). This would suggest an early requirement for Cx431 in germ cell development. Actively migrating PGCs are known to form extensive cell-cell contacts, such that more than 90% of the migrating PGCs are linked via cell processes (Gomperts et al., 1994). This suggests the possibility that cell-cell interactions mediated by Cx431 may play a role in regulating the migration and targeting of PGCs to the genital ridge. We previously showed Cx431 modulates the migratory behavior of two extracardiac cell populations, the cardiac neural crest (Huang et al., 1998; Sullivan et al., 1998; Xu et al., 2001) and pro-epicardial cells (Li et al., 2002). Thus it seemed plausible that the PGC deficiency of the Cx431KO mouse may involve a germ cell migration defect due to the loss of Cx431 gap junction contacts. To investigate this, we utilized an Oct4 (Pou5f1 - Mouse Genome Informatics) promoter-driven GFP transgene as a marker for tracking PGCs in the mouse embryo (Yoshimizu et al., 1999). Oct4 encodes a transcription factor belonging to the POU family of proteins. Its promoter has been used to drive PGC-specific gene expression (Pesce and Scholer, 2001). An Oct4-GFP transgene has been shown to be an ideal marker for PGCs spanning E8.5 to E11.5 (Yoshimizu et al., 1999). The transgene is initially expressed ubiquitously from the morula stage, but at E8.0 its expression becomes restricted to PGCs at the base of the allantois, remaining PGC restricted until E14.5, when expression in female gonads is significantly downregulated (Yoshimizu et al., 1999). This pattern of transgene expression closely matches the expression pattern of endogenous Oct4 transcripts observed by whole-mount in situ hybridization analysis (Kawase et al., 2004). Using the Oct4-GFP transgene, we tracked PGC migration in the E8.5 to E11.5 mouse embryos. With microelectrode impalements into GFP-expressing PGCs, we showed that migrating PGCs make functional gap junction contacts with surrounding cells. We found no difference in the distribution or abundance of PGCs in wild type versus homozygous/heterozygous Cx431KO mouse embryos from E8.5 to E10.5. However, at E11.5, there was a marked loss of PGC in the homozygous Cx431KO mouse embryos. Although Cx431-deficient PGCs showed decreased cell motility, PGCs from the heterozygous and homozygous Cx431KO mouse embryos showed the same change in cell motile behavior. By contrast, studies using TUNEL labeling indicated an abnormal elevation of apoptosis only at E11.5, the time when PGC deficiency is first detected in the Cx431KO embryos. This was associated with an abnormal elevation of activated p53, and injections with a p53 antagonist, -pifitrhin, into pregnant mice prevented the loss of PGCs in the Cx431KO embryo. p53 is known to mediate apoptosis associated with integrin-mediated anoikis (Wang et al., 2002; Zhang et al., 2004), and using a cell adhesion assay, we further showed ß1-integrin-mediated adhesion was reduced in Cx431-deficient PGCs. Together these findings indicate that Cx431 is required for germ cell survival, with Cx431-deficient PGCs undergoing apoptosis in a p53-dependent manner.

	MATERIALS AND METHODS

TOP SUMMARY INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Mouse breeding and genotyping
All experiments were conducted in accordance with an approved animal protocol of the National Heart Lung Blood Institute (Protocol Number H-0086). All experiments were carried out using embryos obtained from the mating of heterozygous Cx431KO mice that were homozygous for the Oct4-GFP transgene (Yoshimizu et al., 1999). Embryonic day 0.5 (E0.5) refers to the day the vaginal plug was found. Cx431 genotyping was conducted as previously described (Xu et al., 2001).

Analysis of PGC distribution and abundance in whole-mount specimens
Embryos were collected at E8.5 to E11.5 and fixed overnight at 4°C in 10% formalin. For E9.5 to E11.5 embryos, the left body wall was dissected free before fixation to facilitate PGC visualization. The following day embryos were washed 3x in PBS, and embedded in 2% agarose in 35 mm Petri dishes. E8.5 embryos were positioned primitive streak side down to allow easy visualization of PGCs at the base of the allantois and in the prospective hindgut. Embryos at E9.5 to E11.5 were placed on their right side to facilitate visualization of PGCs in the hindgut, hindgut/developing genital ridge, and genital ridge, respectively. Embryos were viewed under a Leica DMLFSA microscope with a water immersion lens and a 100-watt mercury lamp using GFP filters (425/60 nm excitation and 480 nm emission). For quantitation of PGC abundance, whole-mount fluorescence images were captured for embryos of the same developmental stage using a uniform imaging protocol. This involved capturing z-stacks consisting of an identical number of optical slices (see below) under the same magnification, and using identical camera settings and exposure time. The highest magnification was used at each developmental stage to allow visualization of all of the PGC-containing tissue in the embryo - i.e. the allantois for E8.5, hindgut for E9.5-E10.5 and genital ridge for E11.5. Under these conditions, the relative abundance of PGC in embryos at the same developmental stage can be directly compared via the mean intensity of GFP fluorescence per unit area (i.e. average brightness/pixel). For these quantitative assessments, 500 µm image stacks were obtained via 75 optical slices using an ORCA-ER digital camera (Hamamatsu). The image stacks were deconvolved using Openlab volume deconvolution (Openlab 3.1.7, Improvision), then merged to generate a single 2D image, from which the mean fluorescence intensity was determined using Openlab. To assess PGC distribution along the hindgut, E9.5 stage embryos were collected and fixed overnight in 10% formalin at 4°C. The following day brightfield and GFP fluorescent images were taken using a Leica MZFLIII stereomicroscope. Somites were used as morphological landmarks to assess PGC abundance along the hindgut. Maximal PGC migration distance was also determined by measuring the distance from the base of the allantois to the PGC migration front situated rostrally along the anterior hindgut.

Time-lapse videomicroscopy of live tissue explants
For time-lapse imaging of E8.5 embryos, the caudal end of the embryo spanning the base of the allantois was embedded in a thin layer of agarose ventral side up in a 35 mm Petri dish using low melt agarose. For E11.5 embryos, tissue explants containing the genital ridge were obtained by dissecting free a short abdominal section of the embryo spanning the developing fore and hindlimb buds. After removing one side of the body wall, the tissue was embedded in low melt agarose with the exposed body wall facing up, allowing easy viewing of the PGCs in the genital ridge. To embed the dissected tissue in low melt agarose, the Petri dish was chilled on ice for 45-60 seconds to set the agarose, after which 4-5 ml of pre-warmed (37°C) L-15 medium was added. The explants were imaged as above using darkfield epifluorescence illumination under a Leica DMLFS microscope. A small amount of mineral oil (Sigma) was used to cover the medium to prevent evaporation. Temperature was maintained at 37.0±0.5°C using a Peltier heated stage (Omega, CT). To further ensure temperature stability, an air curtain was maintained around the microscope using two heating fans (Nevtek, VA). Time-lapse images were captured under darkfield illumination. Z-image stacks were obtained comprising 11-13 µm slices spanning 165-220 µm thickness captured every 6 minutes over 6-8 hours. To minimize phototoxicity, epifluorescent illumination was reduced by using a neutral density filter and a UV cutoff filter to block transmission below 400 nm. Exposure time was minimized, with each embryo having a cumulative exposure of 140 seconds over the course of 8 hours. Image stacks were processed by volume deconvolution and subsequently merged into a single 2D image for fluorescence quantitation. These serial 2D images were used to calculate the speed and directionality of cell motility, assessed using the measurements module of Volocity (Volocity 3.5, Improvision Ltd). Directionality is derived from the net migration distance achieved divided by the total real distance traveled. It measures the degree to which the migratory path strayed from a straight line, with a maximal directionality of 1 corresponding to a cell moving in a straight line, while a cell meandering extensively from a straight path would have a directionality much less than 1.

Apoptosis and cell proliferation analyses
PGC apoptosis was first examined using a sulforhodamine poly-caspase-binding kit (ATCC, Manassas, VA). This entailed culturing embryo explants with a cell permeable and noncytotoxic sulforhodamine derivative of valylalanylaspartic acid fluoromethyl ketone that binds and fluorescently labels caspases 1, 3, 4, 5, 6, 7, 8 and 9. Further analysis was carried out using TUNEL labeling. For these studies, E11.5 genital ridges were fixed in 2% paraformaldehyde (20 minutes), followed by three washes in PBS and permeabilization overnight with PBT (PBS with 10% FBS and 0.2% Titron X-100) (12+ hours at 4°C). The following day, the genital ridges were processed using an APO-BrdU TUNEL Assay Kit from Invitrogen (Carlsbad, CA) using the manufacturer's specifications.

For cell proliferation analysis, pregnant mice at E11.5 gestation were injected intraperitoneally with 3 mg/ml BrdU (1 ml/100 g body weight; Zymed Laboratories, San Francisco, CA). After 45 minutes, the embryos were harvested and the genital ridges retrieved, fixed in 2% paraformaldehyde (20 minutes), washed in PBS (x3), followed by PBS containing 10% FBS, then permeabilized with PBT overnight. The following day the genital ridges were incubated for 30 minutes with the denaturation solution supplied in the Zymed BrdU staining kit (Zymed Laboratories Inc., San Francisco, CA). BrdU detection for both TUNEL and cell proliferation analyses was carried out using an anti-bromodeoxyuridine monoclonal antibody conjugated with Alexa Fluor 546 (Invitrogen, Carlsbad, CA), and visualized using a Cy3 fluorescence filter (535/50 nm excitation and 610/75 nm emission).

PGC dye-coupling analysis
To quantitate dye coupling in PGCs, hindgut explants from E9.5 embryos were isolated and cultured on coverslips overnight in DMEM containing 10% FBS, 100 U/ml penicillin/streptomycin, and supplemented with growth factors (2 ng/ml TNF- and 50 ng/ml soluble SCF from RD Systems Minneapolis, MN; and 10 ng/ml LIF from Chemicon International, Temecula, CA). For dye injection, coverslips were secured to the bottom of 35 mm culture dishes using a small amount of silicon grease, and L-15 medium containing 10% FBS added to the dish. The dish was then placed on a heated stage of a Leica DMLFSA microscope for microelectrode impalements. Micropipettes were pulled from borosilicate capillary glass (with filament O.D. 1 mm, I.D. 0.58 mm) using a model P-97 Flaming/Brown micropipette puller (Sutter Instruments, Novato, CA) and back-filled with sulforhodamine 101 dye (12.5 mg/ml, Invitrogen, Carlsbad, CA). Oct4-GFP-expressing PGCs were iontophoretically injected with sulforhodamine 101 dye for 2 minutes using 0.5 nA current pulses at 1 Hz. Dye spread was observed for a further 3 minutes following cessation of iontophoresis. Dye spread was quantified by measuring total dye spread area.

ß1-integrin-function-blocking antibody treatment on PGC migration
Tissue from the base of the allantois was isolated from E8.5 embryos containing the Oct4-GFP transgene, and cultured overnight in 8-chamber glass slides (BD Biosciences, San Jose, CA) with DMEM containing 10% FBS and supplemented with growth factors as described above. The following day the culture medium was replaced with L-15 medium with the same growth factors and covered with a thin layer of paraffin oil to prevent evaporation. Rat monoclonal anti-mouse ß1-integrin-function-blocking antibody (1:200 dilution; BD Biosciences, San Jose, CA) was added and the explant was then examined by time-lapse imaging. Image stacks comprised 15-20 optical slices (11-13 µm z-steps) were captured every 8 minutes over 6-8 hours. Image stacks were deconvolved, merged into a single 2D image, and the 2D images spanning the time-lapse interval were used to analyze the speed and directionality of PGC cell migration using Volocity (Volocity 3.5, Improvision, Ltd, UK).

Assessment of ß1-integrin-mediated adhesion
ß1-integrin-mediated adhesion was assessed using a Chemicon cell adhesion assay (Temecula, CA). Genital ridges were isolated from E11.5 embryos, incubated for 20 minutes in PBS containing 0.5 mg/ml collagenase/dispase (Sigma, St Louis, MO), then washed in PBS and transferred into 200 µl of DMEM with 10% FBS (per genital ridge). This was followed by gentle aspiration with a 1 ml pipette, and the disaggregated cells were then placed in wells pre-coated with a ß1-integrin antibody (Chemicon Kit ECM522). After 2 hours' incubation (5% CO₂, 37°C), five random areas from each well were imaged using a 10x objective under darkfield illumination to quantitate the number of bound PGCs. The wells were then washed with 1 ml of PBS, after which another five random areas from each well were imaged. The number of PGCs remaining was counted, and the percent of bound PGCs after washing was calculated to compare the relative level of ß1-integrin adhesion in wild-type, heterozygous, and homozygous Cx431 KO PGCs.

Treatment with -pifithrin
Pregnant female mice at E10.5 gestation were injected intraperitoneally with either -pifithrin (2.2 mg/kg, Calbiochem, La Jolla, CA) or vehicle (DMSO/0.9% NaCl) at midday and again at midnight before harvesting at E11.5. The genital ridges were retrieved and GFP fluorescence, TUNEL labeling and PGC proliferation were assayed as described above. To examine the expression of activated p53, E11.5 genital ridges were fixed in 4% paraformaldehyde for 20 minutes, then washed and incubated with antibody to phosphoSer¹⁵-p53 (Cell Signaling Technology, Inc., Danvers, MA) at 4°C overnight, followed by incubation with a Cy3-conjugated secondary antibody. Epiflourescence imaging was carried out on a Leica DMRE microscope with a 40x objective.

Statistical analyses
Our results were expressed as mean ± standard error of the mean, with the statistical analyses carried out using the InStat software package (3.0b, GraphPad Software, Inc.). Differences between genotypes were compared using unpaired analysis of variance (ANOVA), while the means of all three genotypes were compared using the Bonferroni test. P<0.05 was considered statistically significant.

View larger version (45K): [in this window] [in a new window]   Fig. 1. Dye coupling of PGC cells with their neighbours. (A,B) Representative images obtained following intracellular microinjection of fluorescent dye sulforhodamine 101 into PGCs contained in hindgut explants from both Cx431 wild-type (+/+) and homozygous Cx431KO (-/-) embryos (E9.5). Both A and B comprise three images superimposed: a DIC image, green Oct4-GFP fluorescence image (PGC), and a red sulforhodamine 101 fluorescence image. (C) Dye spread was quantified by calculating total area of dye spread after 5 minutes (2 minutes' active microinjection plus 3 minutes' passive spread). PGCs in Cx431KO (-/-) explants displayed a significant reduction in their ability to pass dye into their neighbors. For this analysis, 8, 4 and 7 microelectrode impalements were carried out, respectively in four Cx431 +/+, four Cx431 +/- and five Cx431 -/- embryo explants.

	RESULTS

TOP SUMMARY INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Distribution and abundance of PGCs
To determine if the distribution and abundance of PGCs may be perturbed by the loss of Cx431 function, Cx431KO mouse embryos from E8.5 to E11.5 were examined by epifluorescence imaging to visualize the migrating PGCs. At E8.5, the GFP-expressing PGCs can be seen in the caudal aspect of the embryo, at E9.5/E10.5 in the region spanning the hindgut, and at E11.5, the genital ridge (Fig. 2). This same pattern was observed in wild-type and heterozygous/homozygous Cx431KO mouse embryos. To determine whether there may be changes in PGC abundance with Cx431 deficiency, we quantitatively assessed the GFP fluorescence in wild-type, heterozygous and homozygous Cx431KO mouse embryos from E8.5 to E11.5. No difference was detected between wild-type, heterozygous, and homozygous KO embryos at E8.5, E9.5 or E10.5 (Table 1). However, at E11.5, GFP fluorescence in the homozygous Cx431KO mouse embryos was markedly reduced compared with wild-type and heterozygous KO embryos (Table 1). This difference was evident even by direct visual observation of the genital ridges (Fig. 2). The decrease in GFP fluorescence is not due to changes in intrinsic GFP fluorescence in the Cx431-deficient PGCs, as GFP fluorescence intensity in individual PGCs, whether wild type or Cx431 deficient, was indistinguishable (data not shown).

View this table: [in this window] [in a new window]   Table 1. Changes in PGC abundance in Cx431 KO mouse embryos

View larger version (80K): [in this window] [in a new window]   Fig. 2. PGC distribution in the Cx431 KO mouse embryo. The distribution of PGCs in Cx431 wild-type (+/+) and homozygous Cx431KO (-/-) mouse embryos from E8.5 to E11.5 was observed via GFP expression mediated by an Oct4-GFP transgene. No difference was seen in the overall distribution of GFP-expressing cells in the wild-type versus KO mouse embryos. However, at E11.5 there was a noticeable reduction in GFP fluorescence in the genital ridges of the KO mouse embryos. Scale bars: 100 µm.

Analysis of PGC proliferation and apoptosis
While examining the time-lapse movies, we observed PGCs undergoing mitotic cell division (see Movie S2 in the supplementary material). In addition, some PGCs were observed to fragment and disintegrate (see Movie S3 in the supplementary material). Shown in Fig. 5 are images from a time-lapse movie showing the progressive fragmentation of a PGC (Fig. 5A). Immunostaining using a poly-caspase marker showed that these cell fragments were caspase-positive, suggesting that they may represent apoptotic cells (Fig. 5B). To compare the rate of PGC proliferation or apoptotic cell death in wild-type versus Cx431KO embryos, we counted the number of mitotic and fragmenting cells observed in the time-lapse movies. An apoptotic index generated by normalizing the number of apoptotic cells by the number of mitotic cells showed more than twofold increase in the relative incidence of apoptosis in the homozygous KO PGCs (Fig. 5C).

View larger version (58K): [in this window] [in a new window]   Fig. 3. Analysis of PGC migration in Cx431KO mouse embryos. (A,B) An example of how GFP-expressing PGCs were visualized and tracked in an E8.5 mouse embryo by time-lapse imaging. The migratory paths of individual PGCs are superimposed on the image of the embryo seen at the beginning of the experiment (A), and for clarity are plotted with a single origin in B. (C,D) The migratory paths of PGCs in E8.5 and E11.5 embryos were used to determine the speed (C) and directionality (D) of cell locomotion. PGCs from both homozygous and heterozygous Cx431KO embryos demonstrated significant decreases in both speed and directionality. This data entailed the analysis of a total of 72 Cx431 +/+, 74 Cx431 +/- and 66 Cx431 -/- independent cell migration tracings obtained from five embryo explants for each of the three Cx431 genotypes. Scale bar: 100 µm.

View larger version (72K): [in this window] [in a new window]   Fig. 4. PGC migration along the hindgut of Cx431KO embryos.Phase contrast (A) and GFP fluorescence (B) images of an E9.5 wild-type mouse embryo demonstrate the distribution of PGCs along the hindgut. We used somites as morphological landmarks (outlined in yellow) to assess the abundance of PGCs along the hindgut. We also determined the maximal PGC migration distance by measuring the distance from the base of the allantois (arrowhead), where PGCs originate, to the migration front situated rostrally along the anterior hindgut (arrow). Scale bars: 1 mm.

View larger version (52K): [in this window] [in a new window]   Fig. 5. PGC fragmentation seen by time-lapse imaging. (A) Images from a time-lapse series, showing the progressive fragmentation of a PGC seen in an E11.5 genital ridge explant from a homozygous Cx431KO mouse embryo. The images 1 through 5 are 24 minutes apart. (B) After time-lapse imaging, the genital ridges were stained with a sulforhodamine derivative of valylalanylaspartic acid fluoromethyl ketone that binds caspases 1, 3, 4, 5, 6, 7, 8 and 9 (see Cy3 panel). In the merged image, regions of co-localization of the polycaspase staining (Cy3 panel) with the PGC cell fragments (GFP panel) can be seen. (C) From analysis of the time-lapse movies, the number of apoptotic and mitotic cells was quantitated to generate an apoptotic index comprising the number of apoptotic cells normalized by the number of mitotic cells. This showed a significant increase in apoptosis in the homozygous Cx431KO genital ridges. The number of embryos used in this analysis included seven Cx431 +/+, four Cx431 +/- and five Cx431 -/- embryos. Scale bars: 25 µm.

View larger version (114K): [in this window] [in a new window]   Fig. 6. PGC apoptosis and proliferation in E11.5 genital ridges. (A-C) BrdU labeling was used to assess the cell proliferation rate of PGCs in E11.5 wild-type (A) heterozygous and homozygous Cx431KO (B) embryos. BrdU immunodetection using an Alexa Fluor 546 conjugated anti-BrdU antibody (red) showed some regions of co-localization with GFP-expressing PGCs (seen as yellow). No statistical difference was seen in the proliferation rate of PGCs in wild-type versus heterozygous/homozygous Cx431KO mouse embryos (C). Note that the data are normalized to the frequency of BrdU-positive PGCs in wild-type embryos. This analysis included eight Cx431 +/+, 25 Cx431 +/- and six Cx431 -/- genital ridges. (D-F) TUNEL labeling in E11.5 genital ridges from wild-type (D) and Cx431KO (E) mouse embryos. TUNEL labeling is visualized by BrdU incorporation, detected using Alexa Fluor 546 conjugated anti-BrdU antibody (red). In the genital ridges from the KO mouse embryo, we noted many regions containing TUNEL-positive GFP-expressing PGCs (see white arrows in E). Quantitative assessments showed a significant increase in TUNEL-positive PGCs in the homozygous Cx431KO embryos (F). Note that the data are normalized to the frequency of TUNEL-positive PGCs in wild-type embryos. This analysis included 12 Cx431 +/+, 16 Cx431 +/- and 10 Cx431 -/- genital ridges. Scale bars: 100 µm.

View larger version (19K): [in this window] [in a new window]   Fig. 7. Analysis of ß1-integrin function in PGC motility. (A) Treatment of wild-type PGCs with a ß1-integrin-function-blocking antibody caused a significant reduction in both the speed and directionality of PGC migration in E11.5 genital ridge explants. The migration data was obtained from the analysis of 48 cells derived from five embryo explants for both sham and function-blocking antibody treatments. (B) PGC attachment to ß1-integrin-antibody-coated dishes was used to assess ß1-integrin-mediated cell adhesion. Homozygous Cx431KO PGCs showed a marked reduction in ß1-integrin-mediated adhesion, while control assays using plates coated with anti-mouse IgG antibodies showed no difference in adhesion between the different genotypes. It should be noted that the heterozygous KO PGCs also showed some reduction in adhesion to ß1-integrin-antibody-coated dishes, but less than that observed in the homozygous KO PGCs. For assessments of ß1-integrin-mediated adhesion, assays were carried out using PGCs derived from 14 Cx431 +/+, 20 Cx431 +/- and nine Cx431 -/- embryos, while control assays using anti-mouse IgG were carried out using PCGs from three Cx431 +/+, 14 Cx431 +/- and eight Cx431 -/- embryos.

	DISCUSSION

TOP SUMMARY INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

View larger version (58K): [in this window] [in a new window]   Fig. 8. Abnormal p53 activation in Cx431KO PGCs. (A-C) Genital ridges from E11.5 wild-type and Cx431KO mouse embryos were whole-mount stained using antibody against activated p53 (phospho-serine 15). PGCs (green) expressing activated p53 (red) are denoted by arrowheads. Quantitative analysis (C) revealed a marked increase in the expression of activated p53 in homozygous Cx431KO PGCs. This analysis was carried out using nine Cx431 +/+, 16 Cx431 +/- and 11 Cx431 -/- explants. (D) Assessment of PGC abundance in E11.5 genital ridges obtained from embryos of mice injected with -pifithrin or with vehicle alone (DMSO). With vehicle injection, homozygous Cx431KO mouse embryos showed a significant reduction in PGC abundance. By contrast, following -pifithrin treatment there was no difference in PGC abundance in the wild-type versus heterozygous or homozygous KO embryos. This analysis included 22 Cx431 +/+, 28 Cx431 +/- and ten Cx431 -/- -pifithrin-treated embryos, and 15 Cx431 +/+, 18 Cx431 +/- and ten Cx431 -/- vehicle-treated embryos. (E) TUNEL labeling showed that following -pifithrin treatment the rate of apoptosis was similar in homozygous KO, heterozygous KO or wild-type PGCs. Note that the data are normalized to the frequency of TUNEL-positive PGCs detected in wild-type embryos. This analysis included five Cx431 +/+, 37 Cx431 +/- and 16 Cx431 -/- -pifithrin-treated genital ridges. Scale bars: 25 µm.

We note that a number of mouse mutations have been identified that affect germ cell survival, the best characterized being those involving mutations in c-kit and the c-kit ligand, stem cell factor (SCF) (Bendel-Stenzel et al., 1998). These mutants exhibit a triad of anomalies that include spotting phenotype related to melanocyte defect, germ cell deficiency and hematopoietic defects. Although Cx431KO mice are not known to have pigmentation defects, we have observed ectopic pigment cells in the heart of Cx431KO mice (C.W.L., unpublished). In addition, recent studies suggest that Cx431 also may be important in hematopoiesis, perhaps through mediating stromal-dependent interactions (Cancelas et al., 2000; Durig et al., 2000; Ploemacher et al., 2000). Together with the germ cell deficiency, this would suggest the intriguing possibility that Cx431 regulation of PGC survival may entail downstream signaling involving c-kit. Given that PGCs are gap junction communication competent, this could be mediated via direct transfer of second messengers, metabolites, and even peptides between cells (Bruzzone et al., 1996; Neijssen et al., 2005), and could involve direct coupling of PGCs with PGCs, or PGCs with somatic cells. Although our dye-coupling analysis did not detect dye transfer between PGCs, PGCs are extensively networked via direct cell-cell contacts, and gap junctional communication between PGCs cannot be ruled out (Gomperts et al., 1994).

Given the important role of integrins in modulating cell motility and cell survival, we also examined integrin function in migrating PGCs. We note that a number of previous studies have suggested that gap junctions and integrins may be coordinately regulated. For example, ß1-integrin loss following RGD peptide treatment was associated with dysregulated expression of Cx431 (Czyz et al., 2005). Another study showed that Cx431 gap junctions were lost when cells were treated with antibodies against fibronectin or integrin (Guo et al., 2002). Using ß1-integrin-function-blocking antibody, we showed that ß1-integrin is required for PGC motility. In Cx431-deficient PGCs, we found a reduction in ß1-integrin-mediated adhesion. These observations suggested the possibility that the loss of PGCs may involve anoikis - cell death elicited by perturbation of integrin-mediated adhesion. Cx431-deficient PGCs were also found to have elevated expression of activated p53, which is known to play an important role in anoikis (Grossmann, 2002). Moreover, pifithrin, a p53 antagonist, prevented the apoptotic loss of PGCs in the Cx431KO embryos. Together these findings suggest that Cx431 regulation of PGC cell survival involves downstream regulation of p53 cell signaling. Recent studies suggest that p53 translocation into the mitochondria may be an important initial step in p53-mediated induction of apoptosis (Zhao et al., 2005). Interestingly, Cx431 also has been shown to translocate into the mitochondria (Schulz and Heusch, 2004), and this has been suggested to protect against ischemic cell death (Boengler et al., 2005). Whether Cx431 and p53 are translocated into the mitochondria in PGCs undergoing apoptosis is not known.

The unique property of gap junctions in providing a conduit for the direct coupling of metabolic pools between cells makes a compelling argument for gap-junction-mediated cell-cell communication playing a role in apoptosis. Treatment of well-coupled GFSHR-17 granulosa cells with gap junction uncouplers has been reported to promote apoptosis (Ngezahayo et al., 2005). However, in primary granulosa cells, the induction of apoptosis was actually associated with increased cell coupling, and upon gap junction blockade, apoptosis was inhibited (Krysko et al., 2004). We note that a specific role for Cx431 in apoptosis has been suggested in a number of other studies (Albright et al., 2001; Furlan et al., 2001; Nakase et al., 2004; Yasui et al., 2000). Thus Cx431 knockdown using antisense was shown to elicit apoptosis in cultured rat cardiomyocytes (Yasui et al., 2000), while another study showed cell surface expression of Cx431 inhibited apoptosis induced by choline deficiency (Albright et al., 2001). Elevated apoptosis was also found in conjunction with focal ischemic brain injury in mice with astrocyte-targeted deletion of Cx431 (Nakase et al., 2004). In contrast to these findings, several studies showed that ectopic Cx431 expression is positively correlated with apoptosis (Huang et al., 2001; Hur et al., 2003; Kalvelyte et al., 2003). Thus the precise role of gap-junction-mediated coupling in modulating apoptosis remains unclear, and potentially could be cell and tissue context-dependent.

Overall, our studies indicate that apoptosis underlies the germ cell deficiency of the Cx431KO mouse. Our findings suggest that this may involve anoikis and the abnormal activation of p53 elicited by altered integrin function in the Cx431-deficient PGCs. How integrin function may be modulated by Cx431 and whether this involves signaling mediated by gap-junctional coupling and/or Cx431-mediated protein-protein interactions are challenging questions that will need to be addressed in future studies. It is interesting to note that recent reports have suggested that gap-junctional coupling may not be essential to the modulation of cell motility and cell proliferation by Cx431 (Huang et al., 1998; Moorby and Patel, 2001; Qin et al., 2002; Xu et al., 2001). Whether these findings have any relevance to apoptosis will need to be investigated in future studies.

Supplementary material
Supplementary material for this article is available at http://dev.biologists.org/cgi/content/full/133/17/3451/DC1

	ACKNOWLEDGMENTS

 
We thank Dr H. R. Scholer for providing the Oct4-GFP transgenic mouse line. This work was supported by funding from the Division of Intramural Research of National Heart Lung Blood Institute/National Institutes of Health.

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