Histone chaperone APLF level dictates the implantation of mouse embryos Free

Infertility has been one of the major challenges in reproductive medicine. Unrecognized pregnancy losses in the early stage of pregnancy could be due to failure in embryo implantation. The migration and invasion of cells of extra-embryonic lineage is critical for implantation. Identification of the molecular signatures essential for the crosstalk between the embryo and uterus is of utmost importance. Following fertilization, the zygote undergoes a well-orchestrated cleavage forming blastomeres that polarize as the symmetry divaricate to form the blastocyst after subsequent divisions (Yamanaka et al., 2006). During this relatively short-lived stage, blastomeres bifurcate into undifferentiated inner cell mass (ICM) and outer differentiated primary epithelia called trophectoderm (TE). TE is the first tissue type to be specified in mammalian development. During the late blastocyst stage, trophoblast cells differentiate and invade the maternal endometrium to eventually allow the establishment of the maternal-fetal interface (Soncin et al., 2015). This juncture of maternal-fetal union is the placenta, which encloses the embryo during implantation. Synchronized development of the embryo within the placenta and uterine receptivity determines the ‘window of implantation’. The preparatory phase of implantation is marked by the dramatic alteration of the cytoarchitecture of both the embryo and the maternal endometrium. These alterations are brought about by the cellular transition cycles – the mesenchymal to epithelial transition (MET) and the epithelial to mesenchymal transition (EMT). EMT is marked by the loss in the epithelial character of polarity and cell-cell adhesion while gaining the invasive and migratory character of mesenchymal cells (Kalluri and Weinberg, 2009). An intricate network of transcription factors coordinates this tightly regulated process. However, epigenetic control over the cellular transitions during blastocyst implantation has yet to be explored.

We demonstrated that downregulation of histone chaperone Aprataxin and PNK-like factor (APLF) engendered MET associated with cellular reprogramming of mouse embryonic fibroblasts (MEFs) (Syed et al., 2016). APLF is a DNA damage responsive factor involved in non-homologous end joining (NHEJ) repair (Grundy et al., 2013). Additionally, APLF demonstrates histone chaperone activity by its acidic domain (Mehrotra et al., 2011). It acts as an H2A-H2B histone chaperone by binding through its DNA interaction surface (Corbeski et al., 2018). APLF interacts with the core histones and recruits the macroH2A variant to the damaged DNA sites and within the promoters of genes implicated in EMT (Mehrotra et al., 2011; Syed et al., 2016). Recently, we established the presence of an enhanced level of APLF in breast cancer and how it could regulate EMT associated with breast cancer metastasis (Majumder et al., 2018). In our first study, we found that APLF levels were significantly less or negligible in mouse embryonic stem cells (ESCs), as well as in induced pluripotent stem cells (iPSCs) (Syed et al., 2016). As ESCs are derived from the ICM of the blastocyst and the first cellular transition in development happens to be EMT during implantation, we were intrigued to study APLF in development.

Here, we show that APLF is required during mouse pre- and post-implantation development involving EMT. We demonstrated that the absence of APLF induced abnormal hatching during development in vitro and loss in implantation during development in vivo. Gene expression analyses indicated inhibition in EMT of TE or trophoblast cells due to induction of cadherin 1 (CDH1, also known as E-cadherin). Loss in implantation could be attributed to the significantly elevated level of Cdx2, Cdh1 and soluble Fms-like tyrosine kinase 1 (sFlt1, also known as Flt1) during TE or trophoblast stem cells (TSCs) differentiation and invasion. Hence, our study suggests a significant role for APLF in implantation and the post-implantation stages of mouse embryonic development.

The pre-implantation development of an embryo starts with the formation of the 1-cell zygote. We determined the level of APLF at different stages of mouse pre-implantation development along with the cell adhesion molecule CDH1. Epithelial marker CDH1 is influenced by APLF level, as mentioned in our earlier studies (Syed et al., 2016; Majumder et al., 2018). Immunofluorescence analysis demonstrated the presence of APLF in the 4-cell stage within the nucleus, as well as in the cytosol (Fig. 1A; Fig. S1A). Next, we observed an enhanced APLF expression in 16-cell to early morula and late morula stage embryos (Fig. 1B,C; Fig. S1B). However, when the embryo underwent the first differentiation to form ICM and TE within a blastocyst (32-cell stage), the APLF level was significantly downregulated (Fig. 1C; Fig. S1C). Quantitative RT-PCR (qRT-PCR) analysis of the expression of Aplf in different stages of embryo development demonstrated the same trend as observed in immunofluorescence analysis (Fig. 1D). A significant increase in Aplf was detected in the compaction stage of morula compared to other stages (Fig. 1D). Statistical significance demonstrated the differential expression pattern in Aplf expression throughout the different stages of development (Fig. 1D). However, a striking picture was observed with nuclear expression of APLF in 4-cell stage embryo (Fig. 1A; Fig. S1A) versus cytoplasmic expression of APLF in higher stages of pre-implantation development of the embryo.

Interestingly, exploration of APLF expression in mouse blastocyst demonstrated that a significant level of APLF was enhanced within the TE (Fig. 1C). Also, TEAD4-APLF and OCT3/4-APLF co-immunostaining further confirmed the fact that the APLF expression was almost negligible within the epiblast region of the blastocyst (32-64 cell stage) (Fig. 1E; Fig. S1D). Intensity measurement showed significant enrichment of APLF in the TE versus the ICM in embryonic day (E)3.5 mouse blastocyst (Fig. 2A-C).

Upon investigation of APLF expression in later stages of post-implantation development at E7.5, we observed the presence of APLF within the trophoblast giant cells, TE, ectoplacental cone and allantois but almost negligible presence in other lineages responsible for the formation of the embryo proper (Fig. 2D-I). Surface intensity profile further validates the immunofluorescence analysis for the enhanced expression of APLF in cells of TE lineage (Fig. S1E). It could be noted here that the presence of APLF in the hypoblast was evidenced in the co-immunostained APLF-TEAD4 embryo (Fig. 1E).

Owing to the manifestation of differential expression patterns at different stages, we anticipated that APLF might regulate the pre- or post-implantation development of mouse embryos. So, we introduced empty vector pLKO.1 and Aplf short hairpin (sh)RNA at the 1-cell stage of mouse embryo by micromanipulation (Fig. 3A). The Aplf shRNA used in this study has been validated in our earlier report describing the role of APLF in the induction of pluripotency (Syed et al., 2016). APLF knockdown in MEFs, adapted from Syed et al. (2016), showed ∼70% downregulation of APLF at the protein level (Fig. S2A). In silico analysis for checking Aplf shRNA specificity using SpliceCenter (Ryan et al., 2008), a web-based bioinformatics tool, could not detect any off-targets for this Aplf shRNA (Fig. S2B). The knockdown of APLF in embryos was confirmed by western blot analysis within a sample size of n=35 and n=38 for vector control and Aplf shRNA, respectively (Fig. 3B). Further validation of shRNA is discussed in the rescue experiments mentioned later in the Results section. This experiment to understand the role of APLF in in vitro development was performed three times independently. Uninjected, vector injected and Aplf shRNA-injected mouse embryos developed normally until the morula stage (Fig. S2C). Interestingly, a few hatched embryos started appearing earlier in the Aplf-shRNA set than in the control set (Fig. S2C, bottom right panel, red circle). But, at E3.75, the number of hatched embryos injected with Aplf shRNA were significantly higher in comparison to control and empty vector (Fig. 3C,D; Fig. S2D). Therefore, we were intrigued to understand what caused the abnormal increase in the hatching of embryos upon downregulation of APLF.

Transcription factor CDX2 marks the TE (Strumpf et al., 2005). Owing to the enhanced expression of APLF within the TE (Figs 1, 2), we investigated the level of CDX2 in the manipulated embryos. Hatching of an embryo is succeeded by its implantation, and it is known that implantation involves EMT (Thiery et al., 2009). Therefore, we analyzed the expression of CDH1, the epithelial marker, along with other lineage-specific markers in the manipulated embryos. Immunofluorescence analysis demonstrated a significant increase in CDH1 and CDX2 levels in Aplf-sh embryos compared to vector control embryos (Fig. 3E). Three-dimensional projections further proved the induced expression of CDX2 and CDH1 in Aplf-sh embryos at the early blastocyst stage of 32-64 cells (Fig. S3A). Interestingly, it was the increase in the number of cells expressing CDX2 in Aplf-sh embryos compared to the vector control that contributed to the induced expression of CDX2 (Fig. S3B). A qRT-PCR analysis of embryo samples also demonstrated a significant increase in Cdh1 and Cdx2 levels in Aplf shRNA-injected embryos (Fig. 3F). Aplf knockdown in embryos failed to significantly affect the level of expression of other lineage markers, including Nestin, Sox17, α-Sma (also known as Acta2) or Eomes (Fig. S3C). Next, we segregated the hatched embryos of the vector control and Aplf shRNA sets and determined the expression of Cdx2 and Cdh1 (Fig. S3D). Loss of Aplf was associated with a significant increase in Cdh1 and Cdx2 within hatched embryos generated from Aplf knockdown compared to hatched embryos in uninjected and vector control sets (Fig. S3D). Next, it was interesting to determine the Cdx2 and Cdh1 expression pattern in the hatched and unhatched embryos from vector control and Aplf-knockdown sets. Vector-injected embryos demonstrated no significant change in Cdx2 and Cdh1 expression in hatched to unhatched embryos (Fig. S3E). A significant increase in the expression of both Cdx2 and Cdh1 was detected in hatched Aplf-shRNA embryos versus unhatched counterparts (Fig. S3F). We wondered whether an induced level of Cdh1and Cdx2, and an abnormal hatching pattern of Aplf shRNA-manipulated embryos, would hamper the implantation or the development of the embryo in vivo.

Micromanipulated two-cell embryos were transferred into the oviduct of 0.5 days post coitum (dpc) pseudopregnant females and dissected at E8.5. Uninjected and vector-injected embryos developed to E8.5 (Fig. 4A, left and middle panel; Fig. S4A, left and middle panels). Interestingly, Aplf shRNA-injected embryos either failed to implant or had significantly fewer implantation sites and hence could not develop to E8.5 (Fig. 4A, right panel; Fig. S4A, right panel). These injections were performed in triplicate and, among these, a similar fate was observed for the Aplf shRNA embryos on two occasions. However, in the third set of experiments, embryos were formed but we could not detect any downregulation of Aplf in those embryos (Fig. S4A, right panel bottom). The proportion of successful implantation to failed implantation of Aplf shRNA-injected embryos was significantly less compared to uninjected and vector sets (Fig. 4B). Therefore, downregulation of APLF indeed resulted in a defect in implantation and thereby impaired the development of the embryo.

Rescue experiments with the injection of Aplf-cDNA along with Aplf-shRNA (Fig. S4B,C) demonstrated normal development of mouse embryos (E3.75-E4 mid blastocyst stages) both in vitro and in vivo, thereby confirming the specificity of the Aplf shRNA used in this entire study, and hence the role of APLF in embryonic development (Fig. 4C,D; Fig. S4D,E). The in vitro development experiments were repeated three times independently; however, the in vivo experiments could only be successfully performed two times. Technical glitches resulted in the loss of pregnancy in one set (vector/pUltra) of the experiment. Gene expression analysis with these embryos showed no significant difference in the levels of Aplf, Cdx2 or Cdh1 (Fig. S4F).

Next, we asked whether the predominant expression of APLF in TE is responsible for this outcome in vitro or in vivo. Therefore, to understand the involvement of TE towards abnormal hatching or loss in implantation in the absence of APLF, we exploited the mouse trophoblast stem cell (TSC) model.

TSCs were derived from E3.5 mouse blastocyst cultured on a mitotically inactivated MEF feeder layer in the presence of FGF4 and heparin. The TSC-like clump (Fig. S5A) was dissected manually and cultured for multiple passages on MEF feeder layer to derive mouse TSCs. To ascertain the character of TSCs, the cells were checked for the presence of CDX2 and the absence of OCT4 (also known as Pou5f1) by immunofluorescence analysis in the first passages (Fig. S5B). The micrograph in Fig. S5C shows TSC colonies after passage 8. Western blot analysis for the expression of CDX2 in these cells further validated the character of TSCs (Fig. S5D). Aplf shRNA and empty vector pLKO.1 was introduced individually into TSCs and puromycin-resistant TSCs were screened for further analysis. The knockdown was validated by western blot analysis (Fig. 4E). Differentiation of TSCs followed by EMT is involved during implantation. Hence, we differentiated the vector control and Aplf shRNA-expressing TSCs for 8 days and determined the expression of genes by qRT-PCR analysis. Interestingly, differentiated Aplf shRNA TSCs showed a significant increase in expression of Cdh1 but a loss in Snai2 expression and an insignificant increase in Snai1 level compared to differentiated vector control TSCs (Fig. 4F). Differential expression patterns of genes involved in EMT/MET further consolidated the role of APLF in the cellular transition. Tead4, indispensable for embryo implantation and a promoter of EMT (Home et al., 2012; Zhang et al., 2018), was significantly downregulated in Aplf-shRNA TSCs compared to vector control differentiated TSCs (Fig. 4F). However, we failed to observe any significant change in the level of Cdx2 (Fig. 4F). It has been shown that induced expression of placental sFlt1 along with elevated sFlt1 in maternal serum contribute to loss of pregnancy and symptoms associated with pre-eclampsia in mothers (Fan et al., 2014). Our studies on the development of Aplf shRNA embryos resulted in a significant loss in implantation/pregnancy (Fig. 4A). Therefore, we determined the sFlt1 level in differentiating TSCs. The qRT-PCR analysis showed a significant increase of sFlt1 level in Aplf-shRNA TSCs compared to vector control TSCs or undifferentiated (proliferating) TSCs (Fig. 4G), whereas we failed to observe any significant change in total Flt1 level (Fig. S5E). Loss in Gcm1 transcription factor, a trophoblast differentiation marker, has been associated with an increase in sFLT1 level in pre-eclampsia (Bainbridge et al., 2012) and, interestingly, Gcm1 level was significantly downregulated in Aplf-shRNA TSCs compared to the vector control or undifferentiated (proliferating) TSCs (Fig. 4G).

Thus, we demonstrated here that histone chaperone APLF is differentially expressed at pre- and post-implantation stages of development of mouse embryos (Fig. 5). Loss of APLF resulted in abnormal hatching in micromanipulated embryos in vitro, whereas a significant loss of implantation sites was observed in vivo. Mechanistic analysis revealed that induced expression of the epithelial marker Cdh1 and loss of Snai2 and Tead4 expression could inhibit EMT associated with trophoblast invasion and implantation. Additionally, a significant increase in Cdx2 and sFlt1 might be responsible for the observed phenotype of abnormal hatching and loss in implantation, respectively. We predict that APLF could be exploited as a parameter to predict the chances of developing complications during pregnancy.

Quite a significant number of studies have been reported regarding the role of APLF as a DNA repair factor in the context of NHEJ repair. But, for the first time, our group reported its functional role as a barrier to reprogramming (Syed et al., 2016). We demonstrated how downregulation of APLF could induce MET during induction of pluripotency (Syed et al., 2016). Later, we also showed how it could regulate EMT associated with breast cancer metastasis (Majumder et al., 2018). APLF level was significantly lower in mouse ESCs, as well as in iPSCs (Syed et al., 2016). The first differentiation in development marks the distinct formation of ICM and TE in the blastocyst, which then undergoes EMT during implantation and gastrulation. Therefore, we investigated the relevance of APLF in this context.

In this study, we observed a dynamic expression of APLF in mouse pre-implantation development with the least expression in the blastocyst stage (Fig. 1). But APLF expression was significantly induced in the peri-implantation state (late blastocyst to E7.5) that involves EMT. It is to be noted here that, at the 4-cell stage, APLF expression was higher in the nucleus compared to the complete absence of nuclear APLF in the later stages of development (Fig. 1); whereas, in the E7.5 embryo, trophoblast giant cells very prominently expressed nuclear APLF only (Fig. 2B). This pattern of expression was checked by two different antibodies, which had been validated in earlier studies (Syed et al., 2016; Mehrotra et al., 2011; Majumder et al., 2018). It might be worth noting that our earlier study on breast cancer showed both nuclear and cytosolic expression of APLF (Majumder et al., 2018), which was further supported by human breast cancer atlas data. Also, lung carcinoma cells demonstrated a significant level of cytosolic APLF (Rulten et al., 2008). Therefore, we anticipate that either concentration or context might be the guiding force that dictates the localization of APLF or vice versa. However, a major highlight is that, in spite of being mostly present in the cytosol, APLF can regulate Cdx2 expressed in the nucleus (Fig. 3E). We would stress that whether the regulation of Cdx2, Cdh1, Snai2, Tead4 or sFlt1 is a direct effect of APLF level or not is still uncertain. This might be an indirect effect such that when there is abnormal hatching the level of Cdx2 or Cdh1 is increased. Or in TSCs, the loss in differentiation might result in the reduced expression of Tead4 or enhanced expression of sFlt1. Thus, the entire mechanistic machinery involving APLF in pre-implantation development is still to be precisely uncovered. However, upon determination of the interacting partners of APLF by mass spectrometry analysis (data not shown), we observed that APLF has both nuclear and cytoplasmic partners, which further proves the fact that its expression is really context specific. Among them, the interaction between cytoskeleton proteins stands out predominantly. Adhesion molecules, including cadherin proteins, play a significant role in blastocyst apposition and during embryo implantation. Interestingly, Transgelin-2 (TAGLN2), an actin-binding protein, interacts with APLF and has been implicated in embryo implantation (Liang et al., 2019). Loss of TAGLN2 leads to the reduction in migration and the invasive character of TSCs (Liang et al., 2019). Furthermore, in some instances, TAGLN2 has been reported to be a negative regulator of NFκβ (Zhou et al., 2019). Upon activation, NFκβ binds to the Cdx2 promoter, resulting in the induction of Cdx2 (Asano et al., 2016). We anticipate that the interaction of APLF with different cytoplasmic proteins during the blastocyst stage thereby might contribute towards the regulation of the level of Cdx2.

Microinjection-mediated knockdown studies using Aplf shRNA showed abnormal hatching of embryos around E3.75. Hatching is a process that requires protease thinning of zona pellucida, along with proper biophysical characteristics of epithelial TE cells, regulated by the E-cadherin-Amotl2 complex (Hildebrand et al., 2017). We observed a significant increase in CDH1 and CDX2 level during the hatching of Aplf shRNA embryos (Fig. 3C; Fig. S3F). CDX2 is required to reduce the epithelial character in the embryo to contribute to EMT required for implantation (Kurowski et al., 2019). CDX2, which is highly expressed in TE of E3.5 embryos, should be downregulated at the peri-implantation period (Kurowski et al., 2019). A balanced CDX2 level is required to retain the apicobasal polarity mediated by CDH1 (Sutherland, 2003). Hence, an unwarranted sustained level of Cdx2 and Cdh1 could contribute to the abnormal hatching of Aplf shRNA embryos.

However, that is not the complete picture. Upon further investigation of TSCs for examining the function of APLF, we observed that there is an inhibition in the EMT, as revealed by the gene expression analyses (Fig. 4F). TSCs, when maintained in the presence of FGF4 and heparin, are in the undifferentiated state, whereas removal of these factors induces the EMT of TSCs (Mobley et al., 2017). Ideally, these cells then lose the epithelial marker, Cdh1, and a subset of them acquires the mesenchymal or invasive property to invade the maternal tissue for implantation. Implantation is one of the pioneer developmental stages in which the EMT comes into action (Thiery et al., 2009). The TSCs differentiate to invasive trophoblast giant cells and this results in placentation (Tanaka et al., 1998). In higher primates, implantation is mediated by synctiotrophoblasts in the earlier stages and then by both cytotrophoblasts and synctiotrophoblasts. Cytotrophoblast cells undergo the EMT, forming multilayered cell columns infiltrating the maternal decidua and blood vessels. The penetrative nature of placentation resembles the invasiveness of tumors, but during development it is a well-regulated process (Vićovac and Aplin, 1996). On the contrary, we observed that upon differentiation, Aplf-shRNA TSCs demonstrated a significant increase in the epithelial marker (Cdh1) but a reduction in the mesenchymal marker (Snai2). Although the P-value of 0.0562 for Snai2 expression between the groups was on the borderline of statistical significance, that does not mean the values are ‘not different’ or the ‘same’. It is just that the value did not satisfy the arbitrary cutoff for statistical significance. Additionally, we performed an array of EMTs using the same samples and we observed that the expression of Snai2 and other EMT genes was downregulated in Aplf-shRNA TSCs (data not shown). Our earlier studies also reported a significant increase in the panel of MET markers, including BMP7, Ocln and Smad2, and a reduction in EMT markers, including MMPs, Vimentin, N-cadherin (also known as CDH2), Zeb1 and Twist1 upon downregulation of APLF in MEFs (Syed et al., 2016). Therefore, it is very clear from the expression pattern of genes, that EMT is indeed inhibited upon APLF knockdown during development. However, another EMT marker, Snai1, was upregulated in Aplf-shRNA TSCs, although not significantly (Fig. 4E). Interestingly, overexpression of Snai1 has been associated with the inhibition of trophoblast giant cell differentiation involved in implantation (Nakayama et al., 1998). In addition to its role in TE specification and implantation, Tead4 overexpression has been shown to promote EMT in head neck squamous cell carcinoma (Zhang et al., 2018). This perfectly supports our conclusion that APLF downregulation indeed inhibits EMT. However, we did not observe any significant change in the level of Cdx2 in spite of Tead4, its upstream regulator, being significantly downregulated. We can put forward two theories in this regard: (1) the change is only associated with EMT-related phenomena; or (2), in TSCs, the Tead4-Gata3 axis is working independently in the context of APLF downregulation. These theories remain to be further explored.

The next question that arises is how APLF is regulating Cdh1 expression? A previous paper from our lab showed that downregulation of APLF in MEF results in a loss of recruitment of repressive histone variant macroH2A.1 from the Cdh1 promoter, thereby inducing CDH1 expression (Syed et al., 2016). A similar mechanism could be operational, as macroH2A.1 is highly enriched in TE compared to ICM (Pasque et al., 2012), analogous to the localization of APLF reported here (Fig. 1C,E).

Another significant finding from the Aplf-shRNA TSC differentiation studies was the presence of a higher level of sFlt1, which is devoid of transmembrane and cytoplasmic domains. It acts antagonistically to vascular endothelial growth factor (VEGF; also known as VEGFA) and placental growth factor (PIGF) (Kendall and Thomas, 1993). The elevated presence of sFlt1 in the placenta and maternal serum gives rise to pre-eclampsia, causing loss in pregnancy as well (Fan et al., 2014). Coordinated vascular development from both sides of the maternal-fetal interface is crucial for successful implantation. sFlt1 is antagonistic to VEGF and a balance between positive and negative regulators of angiogenesis is important (Clark and Charnock-Jones, 1998). An imbalanced production of endometrial VEGF induced placental sFlt1 (Fan et al., 2014). As per this study, we suggest that an optimal level of APLF in the placenta should be there to regulate the level of sFlt1 to avoid any complications during pregnancy. Consequently, we could infer that a higher level of sFlt1 in embryos devoid of APLF would have disrupted the balance of angiogenesis crucial for implantation, and hence contributed to the loss of implantation.

We have convincingly shown how a loss in APLF level contributes to failure in implantation and the development of the mouse embryo (Fig. 4). However, two other studies in the context of DNA repair activity of APLF have generated Aplf knockout mice (Rulten et al., 2011; Tong et al., 2016). The possibility of such contradicting results in knockdown and knockout studies could be either due to off-targets effects of shRNA or genetic compensation. We have already negated the first option by checking the off targets of the Aplf shRNA (Fig. S2B). The next possibility is the phenomenon of genetic compensation, in which a mutation can induce the expression of related genes, which in turn would assume the function of the mutated gene (El-Brolosy and Stainier, 2017). This transcriptional adaptation operates via the mRNA surveillance machinery, triggered by the defective mRNA formed by the knockout gene (El-Brolosy et al., 2019). In both studies, there is a formation of defective mRNA because one is knocked out by frameshift mutation (Tong et al., 2016), whereas the other would result in a defective protein lacking the two domains (PBZ and AD) at the 3′ end of APLF protein (Rulten et al., 2011). Moreover, no proof for the complete loss in APLF expression at the protein level was provided in these two reports. The possibility of transcriptional adaptation can be refuted only if the knockout is produced either by deleting the promoter or the entire gene locus. It would be interesting to examine gene compensation in the context of Aplf but this is presently outside the scope of this study.

Recent reports showed that APLF is underexpressed in human infertile male cohort studies, along with other DNA repair genes CYB5R4, ERCC4 and TNRFSF21 (El-Brolosy et al., 2019; Cheung et al., 2019). These data further support our observation regarding APLF in regulating the fate of the fertilized zygote. The presence of APLF in the placenta and endometrium has already been documented in the Human Protein Atlas database.

Our study for the first time investigated the role of the histone chaperone APLF, in early mammalian development and gives an insight into its expression pattern and localization. It also demonstrates the significance of APLF in development in virtue of the phenotypes shown by the knockdown studies in vitro and in vivo. However, there is an immense scope to investigate the actual mechanism resulting in these remarkable phenotypes. We are looking further into the clinical correlation with human placental samples in order to broaden the scope of the study. Our findings have the potential to be explored further and can be implicated to rectify implantation failure and improve pregnancy-related issues. In conclusion, we provided novel evidence for the significantly enriched expression of histone chaperone APLF in TE of mouse pre-implantation embryo and among TE-derived lineages in post-implantation embryo, which could regulate development-associated EMT involved in the implantation of the mouse embryo.

HEK293T and MEFs were cultured in Dulbecco's modified Eagle's Medium (DMEM), 10% fetal bovine serum (FBS; Invitrogen, 1600044), 1% penicillin and streptomycin (Invitrogen, 10378016), and 1% antimycotic-antibiotic (Invitrogen, 10378016) (Syed et al., 2016). HEK293T cell line was a gift from Dr Soumen Paul (University of Kansas Medical Center, USA; validated by Majumder et al., 2019). W9.5 ESCs were cultured in feeder-free conditions in ESC medium comprising Iscove's modified Dulbecco's medium (Invitrogen, 12440053), 15% ESC qualified serum (Invitrogen, 10439024), 0.0124% mono-thioglycerol (Sigma-Aldrich, M6145), 1% penicillin/streptomycin, 1% antimycotic-antibiotic (Majumder et al., 2015), and were a gift from Prof. Peter J. Scambler (University College London, UK; validated by Majumder et al., 2019).

All animal-related experiments were performed at the host institute (Rajiv Gandhi Centre for Biotechnology) in accordance with the Committee for the Purpose of Control and Supervision of Experiments on Animals guidelines and the institutional protocol approved by the Institutional Animal Ethics Committee (IAEC/160/DSD/2012, IAEC/265/DSD/2014 and IAEC/678/DSD/2018). Micromanipulation and post-implantation related studies were performed at the National Centre for Biological Sciences (NCBS) in accordance with the institutional ethics clearance [AJ-1/2015(R1-E)]. The strain type, age, sex, purpose and ethical clearance number has been summarized in Table S1.

Pregnant C57Bl/6J-Rgcb mice at E13.5 post coitum were euthanized. Uterine horns were removed and embryos collected, finely minced and incubated with 0.125% trypsin for 30 min to complete the digestion. Adding MEF medium inactivated trypsin and the suspension was centrifuged at low speed. The supernatant was collected and cultured in MEF medium (Syed et al., 2016). Mitotically inactivated MEFs were used to culture ESCs or TSCs and to generate conditioned medium. Approximately 90% of confluent MEFs were treated with 10 µg/ml mitomycin C (Sigma-Aldrich, M4287) for 2 h followed by multiple washes with 1× PBS.

ShRNA in pLKO.1 vector targeting mouse APLF was purchased from Sigma-Aldrich (SHCLNG-NM_024251; TRCN0000250398). Stable transfection using calcium chloride was performed in HEK293T cells to produce lentiviral particles harboring shRNA against Aplf as mentioned in earlier reports (Syed et al., 2016). Another set of MEFs was transduced with lentiviral particles having empty pLKO.1 vector and was treated as the control. This Aplf shRNA has been used in earlier studies reported by this group (Syed et al., 2016).

For TSCs, total RNA was isolated from ∼500,000 cells using a Qiagen RNeasy Kit (Applied Biosystems, 74106) according to the manufacturer's protocol. cDNA was prepared and SYBR Green Master Mix was used for qRT-PCR analysis (Syed et al., 2016). Primers used are listed in Table S2.

From pre-implantation embryos, total RNA was isolated from embryos at different pre-implantation stages using the Arcturus Pico Pure RNA Isolation Kit (Applied Biosystems, Thermo Fisher, KIT0204) following the manufacturer's protocol. The number of embryos used for RNA isolation is listed in Table S3. cDNA was prepared and SYBR Green Master Mix was used for qRT-PCR analysis (Syed et al., 2016). Primers used are listed in Table S2.

Cell pellets were suspended and lysed in radio immunoprecipitation assay buffer and the protein concentrations were determined by the Bradford Reagent (Bio-Rad, 500-0006) (Majumder et al., 2018) and resolved by 10% SDS-PAGE. Antibodies used are listed in Table S4.

At 48 h before mating, donor female superovulation was induced and synchronized via intraperitoneal injections of gonadotropins. The timing and dosage of hormones depended on several factors such as the light/dark cycle maintained in the facility, as well as the strain and age of donor animals used (Luo et al., 2011; Masters and Wheeler, 1996). All animal housing and breeding rooms had 14 h of light (from 06.00 to 20.00 h) followed by 10 h of dark (from 20.00 h to 06.00 h). With this light/dark cycle regime, a pool of 8 to 10 B6D2 F1 females of 3 to 4 weeks of age (9-12 g) are superovulated via intraperitoneal injections of 5 IU of pregnant mare serum gonadotropin (PMSG; Prospec, HOR-272) at ∼17.00 h on day 1, followed by human chorionic gonadotropin (hCG; Choluron Intervet International) at 5 IU/female at 16.00 h on day 3 (hCG therefore being injected between 47-48 h after PMSG). Females were then immediately individually placed with B6D2 F1 stud males for mating. PMSG mimics follicle-stimulating hormone and hCG mimics luteinizing hormone, and together result in superovulation and relatively well-synchronized ovulation of oocytes of multiple females at a time. Ovulation occurs at ∼8-12 h post hCG, and oocytes retain a fertilization potential for up to 12 to 15 h post hCG. This superovulation regime, therefore, allowed us to first maximize the number of embryos collected per female (up to 20-30 zygotes per collected embryo per female), as well as to synchronize the ovulation and fertilization of multiple females. Mating was confirmed by observation of a vaginal plug the next morning and zygotes were then harvested in M2 medium from the oviducts of the superovulated donor females on day 4 at ∼10.00 h, ∼18 h post hCG. The harvested zygotes from 8 to 10 superovulated females were pooled together and some were processed for zygote microinjection of plasmid DNA of various test and control constructs, whereas other control zygotes were not injected with any constructs, and all the embyros were further processed as mentioned below. All sets of embryos were then incubated at 37°C in kalium simplex optimized medium (KSOM) for in vitro culture until they matured to hatching blastocysts. Embryos were imaged at regular times during in vitro developmental progression. Pictures were taken daily at the same time five times a day for all embryo sets at ∼08.30 h, ∼10.30 h, ∼01.00 h-02.00 h, ∼16.00 h-17.00 h and ∼20.00 h-21.00 h to compare developmental progression differences between each set of embryos. Table S5 summarizes the time points.

B6D2F1/Ncbs and C57Bl6/J-Rgcb female mice were euthanized at different embryonic day points post-observation of plug. B6D2F1/Ncbs were superovulated, whereas C57Bl6/J-Rgcb female mice were not superovulated. Uterine horn along with the ovaries was dissected out intact and kept in a dish containing 1× PBS. Ovaries were removed and two horns were cut near the cervix. A 1 ml syringe containing DMEM with FBS was inserted onto the uterine horn by holding it with a pair of forceps and then flushed out into a four-well dish to collect the embryos. They were further manipulated for other processes using a mouth pipette.

The entire set of experiments was performed at the NCBS using an approved animal protocol [#AJ-1/2015(R1-E)]. To knockdown APLF, Aplf shRNA (Sigma-Aldrich) cloned into the pLKO.1 lentiviral vector (Sigma-Aldrich) was used (as mentioned above), along with the vector control. Circular non-linearized vectors were suspended in Mi buffer at a concentration of 10 ng/µl. One-cell-embryos obtained by superovulating B6D2F1/Ncbs mice were subjected to random cytoplasmic injections and were grown in M16 medium (Sigma-Aldrich, M7292) until the two-cell stage and then in KSOM medium plus amino acids (Sigma-Aldrich, MR-121-D) until late blastocyst stage at 37°C in a Cook premixed tri-gas chamber (5% CO₂ and 5% O₂, balanced with nitrogen)/MINC bench top incubator. Embryos were visualized and photographed daily for 4 days to visualize the in vitro development. For the in vivo development, two-cell embryos injected with Aplf shRNA, pLKO.1 empty vector and uninjected ones were transferred to a 0.5 dpc pseudopregnant CD1/CrlNcbsor B6NCBAF1/Ncbs female mouse, and were maintained until E8.5 in vivo. On E8.5, they were euthanized and uterine horns were dissected intact and photographed for the development of implantation sites and the embryos. This experiment was repeated three times independently.

Pregnant female mice were sacrificed and the abdominal cavity was opened. Using fine forceps and scissors uterine horn was cut intact and placed in a dish containing 1× PBS. Individual embryos were cut apart; the muscle layer around the embryo was removed with forceps. Decidua were pierced with forceps and opened to tear out the two sides separately. Embryos were retained in one half, shelled out and kept in DMEM containing 10% FBS. Embryos were processed accordingly for whole-mount immunofluorescence.

Mouse Aplf full-length cDNA (accession number NM_001170489) was amplified from MEFs by performing PCR using a superscript reverse transcriptase kit (Invitrogen, 18080044). The amplified Aplf cDNA was cloned into XbaI and BamHI sites in the pUltra lentiviral vector (a gift from Malcolm Moore, Memorial Sloan-Kettering Cancer Center, New York City, NY, USA; Addgene, 24129; Lou et al., 2012) generating pUltra/AplfcDNA construct. The expression of pUltra/Aplf was validated by transfecting MEFs. Lentiviral particles were generated as described above. MEFs were transfected with these particles and grown for 72 h to analyze the expression of APLF by GFP. After validation, for the rescue of APLF expression, mouse embryos were co-injected with either empty pLKO.1 and pUltra vectors (5 ng/μl of each plasmid) or Aplf-shRNA and pUltra/AplfcDNA (5 ng/μl of each plasmid). One set was allowed to develop in vitro until E4.5, whereas another set was injected into pseudopregnant mice and dissected at E8.5. This experiment was repeated three times independently.

Isolated four-cell embryos were fixed with 4% paraformaldehyde (Sigma-Aldrich, P6148) for 10 min at room temperature, whereas the rest of the embryos used in this study were fixed for 30 min at room temperature, followed by washing three times with 1× PBS. Samples were permeabilized by submergence in 0.25% Triton X-100 (Sigma-Aldrich, T9284) for 1 h at room temperature, followed by two washes with 1× PBS, and blocking with 10% goat serum (Jackson ImmunoResearch, 005-00-121) and 0.1% Triton X-100 in 1× PBS overnight at 4°C. Embryos were incubated with primary antibody (1:100) dilution in blocking solution overnight at 4°C and washed three times with 1× PBS. Embryos were then incubated with fluorescence-conjugated secondary antibody (anti-rabbit IgG Alexa-Fluor 488, Invitrogen, A11008; anti-mouse IgG Alexa-Fluor 568; Invitrogen, A11004) at room temperature for 1 h, followed by three washes with 1× PBS. Nuclei were stained with Hoechst 33258 dye (Sigma-Aldrich, B1155) at a 1:10,000 dilution, incubated at room temperature for 10 min and washed once with 1× PBS wash. Embryos were then mounted on a slide using anti-fade reagent (Thermo Fisher Scientific ProLong Gold antifade reagent, P36934), and images were captured using confocal microscopy (NIKON A1R si) (Home et al., 2009).

Trophoblast stem cells were derived from C57Bl/6J-Rgcb mouse embryo at the E3.5 blastocyst stage. Blastocysts were plated on inactivated MEFs and cultured in RPMI medium (HyClone, SH30027.01) with 20% FBS, 1 mM sodium pyruvate (Thermo Fisher Scientific, 11360070) and 50 μm β-mercaptoethanol (Sigma-Aldrich, M3148) supplemented with 25 ng/ml FGF4 (Sigma-Aldrich, F8424) and 1 μg/ml heparin (V-parin, VHB Medi Sciences). After 72 h, TSC-like clumps were further manually picked up, trypsinized and replated on MEF feeder layers in the TSC medium (Ray et al., 2009). Cells were analyzed for the expression of CDX2, a TE marker, and OCT4, an ESC-specific marker. CDX2⁺ TSCs were used for experiments mentioned in this study. TSCs were maintained on an inactivated MEF feeder layer. For experiments, mouse TSCs were expanded in a proliferative state without MEF feeders by culturing in the presence of 70% MEF-conditioned medium, 30% TS cell medium containing 20% FBS, 25 ng/ml FGF4 and 1 μg/ml heparin. Conditioned medium was produced by the addition of 10.5 ml of TSC medium to 100 mm culture plates with ∼90% confluent mitotically inactivated MEFs. For differentiation, TSCs were cultured under feeder-free conditions in TSC medium devoid of FGF4/heparin/conditioned medium for 8 days.

Data were collected and represented as three biological replicates of the experiments, and statistical analyses were performed using GraphPad Prism 8 with specific tests indicated in the figure legends.

W9.5 ESCs were a kind gift from Prof. Peter J. Scambler. The APLF antibody was a kind gift from Prof. Ivan Ahel (Oxford University, UK).

The peer review history is available at https://jcs.biologists.org/lookup/doi/10.1242/jcs.246900.reviewer-comments.pdf