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 Table of Contents  
Year : 2021  |  Volume : 5  |  Issue : 4  |  Page : 220-236

Molecular mechanisms underlying cell-fate specification and cellular diversity of the trophoblast lineage during placental morphogenesis in mice

1 Department of Obstetrics and Gynecology, The First Affiliated Hospital of Xiamen University, School of Medicine, Xiamen University, Xiamen, Fujian 361003, China
2 Fujian Provincial Key Laboratory of Reproductive Health Research, School of Medicine, Xiamen University, Xiamen, Fujian 361102, China
3 Department of Obstetrics and Gynecology, The First Affiliated Hospital of Xiamen University, School of Medicine, Xiamen University, Xiamen, Fujian 361003, China; Fujian Provincial Key Laboratory of Reproductive Health Research, School of Medicine, Xiamen University, Xiamen, Fujian 361102, China

Date of Submission30-Nov-2021
Date of Decision08-Dec-2021
Date of Acceptance19-Dec-2021
Date of Web Publication30-Dec-2021

Correspondence Address:
Hai-Bin Wang
Chengyi building, School of Medicine, Xiamen University, Xiang'an South Road, Xiang'an District, Xiamen, Fujian 361003
Jin-Hua Lu
Chengyi building, School of Medicine, Xiamen University, Xiang'an South Road, Xiang'an District, Xiamen, Fujian 361102
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Source of Support: None, Conflict of Interest: None

DOI: 10.4103/2096-2924.334381

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Placental morphogenesis is a highly dynamic process involving mutual recognition and interlacing between the trophoblast–uterus and ultimately the initiation of the maternal–fetal circulatory system. During placental morphogenesis in mice, the trophoblast lineage, which integrates maternal and fetal signaling, undergoes stage-specific changes in gene regulatory programs directing cellular proliferation and fate specification, generating diverse trophoblast subtypes. While accumulating evidence from studies on genetically engineered and mutant mice has revealed the involvement of cell-specific core transcription factors in certain key events during placental morphogenesis, the precise molecular mechanisms by which multipotent trophoblasts gradually differentiate into different subtypes are still largely unknown. In this review, we primarily focus on mutant mouse models with placental phenotypes to provide a comprehensive understanding of the molecular mechanisms underlying cell-fate specification and cellular diversity of the trophoblast lineage during the placental morphogenesis.

Keywords: Differentiation; Morphogenesis; Multipotency; Placenta; Trophoblast

How to cite this article:
Wang JQ, Mu C, Sun Y, Lu JH, Wang HB. Molecular mechanisms underlying cell-fate specification and cellular diversity of the trophoblast lineage during placental morphogenesis in mice. Reprod Dev Med 2021;5:220-36

How to cite this URL:
Wang JQ, Mu C, Sun Y, Lu JH, Wang HB. Molecular mechanisms underlying cell-fate specification and cellular diversity of the trophoblast lineage during placental morphogenesis in mice. Reprod Dev Med [serial online] 2021 [cited 2022 May 21];5:220-36. Available from: https://www.repdevmed.org/text.asp?2021/5/4/220/334381

  Introduction Top

The placenta, a comparatively recently evolved organ that serves as the interface between the maternal and fetal compartments, is responsible for the exchange of gas and nutrients, production of hormones to maintain the pregnancy,[1],[2] and maternal tolerance of the fetal allograft. In addition, it acts as a selective barrier protecting the fetus from mostly maternal perils, such as glucocorticoids, xenobiotics, pathogens, and parasites, thus creating a protective milieu for normal fetal development.[3] The placenta functions as the lungs, gut, kidneys, and liver of the fetus during pregnancy.[4] Disturbances to placental development and functions often lead to pregnancy-related diseases or fetal death.[5],[6]

Throughout mammalian evolution, the placenta has evolved into diverse shapes, structures, and cell types, even between closely related species. These differences are manifested in four categories, including different types of placental interface (epitheliochorial, synepitheliochorial, endotheliochorial, and hemochorial), placental shape (diffuse, cotyledonary, zonary, and discoidal), fetomaternal interdigitation (folded, lamellar, villous, trabecular, and labyrinthine), and cell types (polyploid trophoblast giant cells [TGCs], diploid glycogen trophoblasts [GlyTs], spongiotrophoblasts, and syncytiotrophoblasts [STs] in mice) [Figure 1].[7],[8] The mice possess discoid hemochorial placenta with labyrinth interdigitation formed during the placental morphogenesis [Figure 1]. Placental morphogenesis is a highly dynamic process involving several key events, including trophectoderm (TE) specification, development of the extraembryonic ectoderm (ExE) and ectoplacental cone (EPC) by multipotent trophoblast proliferation and differentiation, chorioallantoic attachment and branching morphogenesis, and maternal vascular remodeling [Figure 2], [Figure 3], [Figure 4], [Figure 5]. The occurrence of these events during trophoblast lineage development is synchronously regulated by maternal and fetal signaling and needs to undergo stage-specific changes in gene regulatory programs and cell morphology. In this review, we discuss the mechanisms underlying the occurrence of these key events during placental morphogenesis from genetic insights.
Figure 1: Placental variation among mammals. (a) Gross placental shapes in different animals. Based on the gross shape of the placenta and area of contact between fetal and maternal tissues, there are four major types of placenta: diffuse (e.g., pigs and horses), cotyledonary (e.g., ruminants), discoid (e.g., primates and rodents), and zonary (e.g., dogs and cats). This image was modified.[130] (b) Degrees of fetal trophoblast invasiveness in different types of placenta. Based on the maternal tissue in contact with the chorionic epithelium of the fetus, there are three types of placenta: epitheliochorial (fetal chorionic epithelium is in contact with the maternal endometrial epithelium), endotheliochorial (fetal chorionic epithelium is in contact with the maternal endothelium), and hemochorial (fetal chorionic epithelium is bathed in maternal blood). The red arrows and words indicate the maternal tissue directly in contact with the chorionic epithelium of the fetus in different types of placenta. MV: Maternal vessel; MMe: maternal mesothelium; FMe: Fetal mesothelium; FV: Fetal vessel; Tr: Trophoblast; Epi: Maternal epithelium; Fen: Fetal endothelium; FB: Fetal blood; Men: Maternal endothelium; MB: Maternal blood; Cyto-Tr: Cytotrophoblast; Syncytio-Tr: Syncytiotrophoblast; Syntr-I: Syncytiotrophoblast I; Syntr-II: Syncytiotrophoblasts II; STGC: Sinusoidal trophoblast giant cell.

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Figure 2: Specification of TE fate in mice. (a) Preimplantation embryo development in mice. After fertilization, the oosperm divides to form two blastomeres. The blastomeres constantly proliferate and then undergo cell polarization and compaction at the late 8-cell stage to form morula. The 8–16-cell division stage gives rise to inner (yellow) and outer (blue) cells that contribute, respectively, to the ICM and TE of the blastocyst. The primitive endoderm cell (green) lineage emerges from the ICM at the early blastula stage. At the late blastocyst stage, the mouse embryo gives rise to three tissues: TE, ICM, and primitive endoderm. (b) Key molecular pathways to bias ICM fate or TE fate in early mouse embryo. Between the late 2-cell and 4-cell stages, the high level of Carm1 biases the fate of ICM. Baf155 promotes TE specification and is negatively regulated by Carm1. At the late 8-cell stage, keratins function as asymmetrically inherited factors that specify the first TE cells of the embryo. At the 8–16-cell stage, Yap1 nuclear localization and interaction with the trophoblast -specific factor, Tead4, is the main signal for the specification of the TE. (c) Yap1 translocation to the nuclei determines TE specification at the 8–16-cell stage. The inhibition of the Hippo pathway promotes Yap1/Taz translocation to the nuclei, where they interact with Tead4 to activate the expression of target genes. ZGA: Zygotic gene activation; TE: Trophectoderm; ICM: Inner cell mass; PrE: Primitive endoderm; Epi: Epiblast.

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Figure 3: Maintenance of the TSC niche in mice. (a) Multipotency and proliferation of trophoblast is regulated by trophoblast–epiblast interactions. Fgf4 and Fgfr2 are specifically expressed in the ICM/epiblast and trophoblasts, respectively. Due to Fgf4/Fgfr signaling, the polar trophectoderm cells overlying the ICM continue to proliferate and retain the expression of Cdx2. Bmp4 can induce Wnt3 for amplifying Nodal expression in the epiblast. Nodal in turn induces Fgf4 to regulate Bmp4 expression in the ExE. Thus, the four factors form a signaling circuitry at the ExE–Epi interface. (b) Fgf4-Erk pathway to regulate trophoblast multipotency. Fgf4/Fgfr signal promotes Erk phosphorylation, which activates the transcription of Esrrb and Sox2, thereby enhancing transcriptional networks of TSC self-renewal. (c) Occupancies of multipotency factors on the promoter region of target genes for transcription. For example, Tfap2c, Smarca4, Eomes, Elf5, and Cdx2 are enriched in Elf5. TK: Tyrosine kinase; ICM: Inner cell mass; TSC: Trophoblast stem cell; ExE: Extraembryonic ectoderm; Epi: Epiblast.

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Figure 4: Placental morphogenesis in mice. Placental morphogenesis in mice undergoes several key events, including trophectoderm specification (preimplantation); development of the ExE (E5.5–E6.5), chorion (E7.0–E8.5), and EPC (E5.5–E9.5); chorioallantoic attachment (E8.5); and branching morphogenesis (E9.5–E10.5). EPC: Ectoplacental cone; TGC, Trophoblast giant cell.

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Figure 5: Development of the trophoblast lineage in mice. (a) Structural diagram of maternal blood flow through the intervillous space in the mature mouse placenta. Five subtypes of TGCs are present, including parietal TGCs, spiral artery-associated TGCs, cannel TGCs, sinusoidal-TGCs, and channel TGCs. The maternal blood is led to the placenta along the spiral artery in which the vascular endothelial cells are replaced by Spa-TGCs, and then enters the spongiotrophoblast layer where cannel TGCs converge blood to form the main stream. Next, the blood disperses into the labyrinthine layer, where the maternal sinus is lined by S-TGCs. Finally, the maternal blood inversely flows into the vein lined by channel TGCs. Furthermore, glycogen trophoblasts in spongiotrophoblast layer and maternal decidua also have migratory capacity. (b) Molecular markers show the differentiation of trophoblast lineage in mice. The image refers to single nucleus RNA-seq of the mouse placental labyrinth.[131] TGCs: Trophoblast giant cells; S-TGCs: Sinus-TGCs.

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  Specification of Trophectoderm Fate: The First Lineage Segregation in Mammals Top

The first two cell types that differentiate during mammalian development are the TE, the extraembryonic tissue giving rise to the trophoblast lineages of the placenta, and the inner cell mass (ICM), which separates into the primitive endoderm and epiblast, developing to form the yolk sac tissue and embryo proper, respectively. The cleavage and proliferation of blastomeres during the 2–4-cell stage lead to distinct bidirectional cellular contexts, which is the prelude to the first lineage separation in mammals [Figure 2]a.[9],[10],[11] Studies have attempted to identify the molecules that are expressed differentially between different blastomeres and are essential for cell-fate commitment during the first lineage segregation. The critical molecules are as follows.

Coactivator-associated arginine methyltransferase 1

Coactivator-associated arginine methyltransferase 1 (Carm1) is a type I arginine methyltransferase that methylates the arginine residues of histones and nonhistones and important for first lineage segregation during embryo development [Figure 2]b. It has been shown that Carm1 is a maternally-inherited mRNA.[9] Carm1 expression is low and equal in each blastomere at the early 2-cell stage and increases to varying degrees in each blastomere at the late 2-cell stage, while the catalytic product of Carm1 H3R26me2 shows no difference. At the 4-cell stage, the unequal distribution of Carm1 expression is maintained, and different intensities of H3R26me2 are also observed between blastomeres. However, Carm1 expression becomes weaker and diffuses in the nucleoplasm by the late 8-cell stage.[10],[11] Torres-Padilla et al. confirmed that the higher Carm1 and H3R26me2 levels between blastomeres at the 4-cell stage could bias the subsequent cell fate toward ICM, as Carm1 overexpression in one blastomere at the 2-cell stage by injecting synthetic Carm1 mRNA promoted the future progeny to an ICM fate.[9] Moreover, recent evidence has shown that Carm1 drives ICM fate commitment through the Carm1/Prdm14/H3R26me2-Oct4/Sox2-Sox21 axis.[12],[13]

Carm1 is mainly assembled into the paraspeckles that are within interchromatin granule clusters and enriched in characteristic RNA-binding proteins, including Pspc1, Nono, and Sfpq, around the scaffolds of a specific long-noncoding RNA, Neat1. In addition, almost all paraspeckles in the mouse embryo at the 4-cell stage contain Carm1. On the one hand, elevated Carm1 expression by exogenous mRNA injection in the zygote results in disruption of paraspeckles and relocalization of their components. On the other hand, the knockdown of Nono or Neat1 RNA reduces the number of Carm1 speckles and the intensity of H3R26me2.[11] These results demonstrate that the functions of Carm1 are highly dependent on the paraspeckles. The depletion of Nono or Neat1 results in the developmental arrest at the 16–32-cell stage while elevating the expression of the trophoblast-specific transcription factor (TF), Cdx2, indicating that the cells are stepping for the extraembryonic fate. Furthermore, maternal and zygotic deletion of Carm1 might not affect the first lineage separation, as evidenced by the formation of blastocysts. However, the development of Carm1-deleted embryos is delayed and blastocysts with reduced numbers of cells are formed, followed by the ectopic expression of Nanog in the TE and the coexpression of Cdx2 and Nanog in the ICM.[11] These results demonstrate that Carm1 in the paraspeckles plays an essential role in regulating cell-fate determination during mouse embryo development. It has also been reported that Carm1 physically binds to an endogenous retrovirus-associated long-noncoding RNA, LincGET.[10] Consistent with Carm1, LincGET is transiently and asymmetrically expressed in the nucleus of mouse embryos during the 2–4-cell stage. The knockdown of the LincGET expression by injecting the exogenous interference RNA into one blastomere of the 2-cell embryo prevents the blastomere from undergoing an ICM fate, demonstrating that the asymmetrical expression of LincGET is essential for cell-fate determination during embryonic development. Further examination showed an interdependence of LincGET and Carm1 in guiding the ICM fate. LincGET–Carm1 interaction promotes H3R26me2 modification and activates gene expression in ICM, including Sox2, Sox21, and Nanog, but not Pou5f1. Moreover, the LincGET/Carm1 complex increases global chromatin accessibility and promotes the transcriptional activation of transposon elements, such as GLN, ERVL, ERVK, and LINE-1. However, it should be noted that several truncated LincGET mutants remained bound to Carm1 but failed to upregulate the expression of Sox2 and Sox21.[10] It was proposed that the functional domains of LincGET probably act as “anchors” to guide the LincGET–Carm1 complex to the correct location in the nucleus, without which CARM1 cannot function effectively even if the LincGET–Carm1 complex is formed.


Although the cellular heterogeneity in 2-4-cell blastomeres bias the future cell fate, the first lineage separation actually occurs at 8-16-cell stage and completes at the 16-cell stage [Figure 2]a. At the 8-cell stage, the blastomeres undergo the first morphogenetic event (compaction) to form a small compacted cell mass morula. E-cadherin is a cell adhesion molecule that plays the most important role in the process of compaction and the formation of cell–cell adhesion during embryo development. Before compaction, E-cadherin is present throughout the plasma membrane of all 8-cell stage blastomeres. During compaction, E-cadherin is localized to the basolateral cell-to-cell contact sites to form adherens junctions.[14] E-cadherin is a maternally-inherited protein. On the removal of both maternal and zygotic Cdh1 genes (encoding E-cadherin), the embryos fail to compact utterly.[15] Moreover, the disruption of E-cadherin-mediated cell adhesion via blocking antibody ECCD1 in compacted 8-cell embryos results in decompaction of small-sized embryos or those with undetectable ICM.[16] During the initiation of compaction, E-cadherin is phosphorylated, and Ca2+-phospholipid-dependent protein kinase C (PKC) is activated.[17] In addition, there is evidence that the cellular actomyosin cortex contributing to increased surface tension within contactless apical domains is also involved in embryo compaction.[18]

Polarity proteins

When the blastomeres of the embryo undergo compaction, they also undergo a process of intracellular polarization. Before compaction at the 8-cell stage, the individual blastomeres are not polarized with a round morphology and microvilli across their entire cell surface. With the compaction in progress, the blastomeres polarize along their apical-basolateral axes, and the microvilli disappear at the cell-to-cell contact regions but persist on the contact-free apical membrane.[19] The subcellular localization of ezrin is directly in accordance with the dynamics of microvilli distribution.[20] With the onset of polarization, ezrin protein is phosphorylated by atypical PKC, which is regulated by Rho GTPases and MEK/ERK.[21] The members of apical polarity protein/partitioning defective complex composed of Pard3, Pard6, and Mark2 and the basolateral polarity members, including Scrib and Llgl1, are all expressed in the preimplantation mouse embryo. It has been reported that the expression of Pard6b is regulated by Tfap2c, since the failure of the polarization, tight junction, and TE cell lineage specification in the Tfap2c-downregulated embryos are attributed to the dysregulated Pard6b expression. Moreover, the apical polarity proteins have been shown to be responsible for the formation of tight junctions and blastocyst cavitation.[17]


It has been reported that Yap1 activity is associated with compaction and the formation of intercellular polarization during early embryo development.[16] During cleavage, nuclear Yap1 is first detected in embryos at the 4-cell stage, but the signal is weak and variable among different blastomeres. By the early 8-cell stage, nuclear Yap1 is detected in all blastomeres. After the 8-cell stage, Yap1 increasingly accumulates in the nucleus of outside cells and is restricted to the TE cells at the mid/late blastocyst stage, while nuclear Yap1 accumulation decreases in inside cells and appears to be excluded from the nuclei in the ICM [Figure 2]c. It has been shown that Yap1 localization is influenced by compaction as Yap1 is not strictly excluded from the nuclei of inside cells in the embryos treated with ECCD1, the blocking antibody of E-cadherin, at 18–22-cell stages. Cell position is the secondary predicted clue that influences Yap1 localization and cell fate. Nishioka et al. aggregated individual blastomeres from three different embryos into one large chimera with some blastomeres occupying a position internal to others. In these reaggregated embryos, outside cells reestablished polarity, and nuclear Yap1 and Cdx2 were detected, while neither nuclear Yap1 nor Cdx2 was detected in inside cells.[16] Moreover, in the ICM, the Hippo pathway promotes the phosphorylation of Lats1/2 kinases, which further phosphorylates the Yap1 and results in Yap1 degradation [Figure 2]c. In the TE, mechanical stress, probably due to cell polarity or apical extracellular matrix (ECM) signaling, activates Rho GTPase to stabilize the actin cytoskeleton, which in turn inhibits the Hippo pathway core kinases. Inhibition of the Hippo pathway promotes Yap1/Taz translocation to the nucleus, where it interacts with Tead4 to activate the expression of target genes. Cdx2 is a major regulator of Tead4-dependent changes in trophoblast gene expression. However, Tead4 promotes trophoblast fate through both Cdx2-dependent and -independent pathways.[16]


At the 16-cell stage, Cdx2 is exclusively expressed in the outer layer of cells of the morula.[22] The overexpression of single Cdx2 in embryonic stem cells (ESCs) is sufficient for promoting the fate programming toward trophoblast stem cells (TSCs).[23] However, Cdx2 mutants could form the blastula, but the TE shows increased expression of the Nanog gene, demonstrating that Cdx2 is essential for inhibiting the pluripotent gene network but is dispensable for TE differentiation. During cell-fate conversion from ESCs to TSCs via the overexpression of Cdx2, Cdx2 first represses a preexisting ES cell-associated gene expression program, followed by the activation of TS cell-specific genes.[23] In preimplantation embryos, Cdx2 is initially co-expressed with Pou5f1, and they form a complex for the reciprocal repression of their target genes in ESCs.[24] Regarding the regulation of Cdx2 expression, Tead4 is the critical factor for Cdx2 expression as the total loss of Cdx2 in Tead4 null mutant embryos that fail to develop into blastulas.[25] Moreover, the expression of Tead4 is directly regulated by Sox2, a maternally-inherited factor that is essential for both cell pluripotency and extraembryonic derivatives. In embryos with Sox2 knockdown, the expressions of Tead4 and Cdx2 were downregulated.[26] Other factors, such as Tfap2c and Gata3, are also involved in the regulation of Cdx2 expression.[27],[28],[29] However, they are not the critical factors for the first lineage separation as their null mutants went through blastulas.

  Maintenance of the Trophoblast Stem Cell Niche: Epiblast–Trophoblast Interactions via Fgf4/Fgfr Signaling Top

The mature blastocyst hatches out of the pellucid zone and initiates implantation response after the second lineage segregation at approximately E4.0 in the mouse. During the process of blastocyst implantation, the mural TE that is not in contact with the ICM differentiates into primary TGCs with reduced expression of Cdx2. The polar TE overlying the ICM continues to proliferate, mediated by Fgf4/Fgfr signaling, and retain the expression of Cdx2 [Figure 3]a and [Figure 3]b.[22],[30] It has been shown that Fgf4 and Fgfr2 are specifically expressed in ICM and trophoblasts, respectively.[30] While embryos homozygous for the Fgf4 null allele develop to the blastocyst stage, complete implantation, and induce uterine decidualization, they fail to develop substantially thereafter due to severely impaired cell proliferation either in the ICM or in the TE.[30],[31] Moreover, Fgfr2 null mice showed peri-implantation lethal phenotypes, similar to those observed in the Fgf4−/− mice.[32] Therefore, it is not surprise that permanent mouse TSCs could be derived and cultured under Fgf4 conditions. The derived TSC lines contribute to the trophoblast lineage in vivo in chimeras and differentiate into other trophoblast subtypes in vitro in the absence of Fgf4.

In addition to FGF4, the derivation of mouse TSCs requires the presence of embryonic fibroblast-conditioned media (EFCM).[30] Further research found that the major active protein components of the EFCM are Tgfβ and activin as the long-term continuous proliferation of TSCs is maintained by either Tgfβ or activin but requires the presence of Fgf4.[33] Fgf4, through its downstream effector, ERK, directly upregulates the expression of trophoblast core TFs, Cdx2, Eomes, and Sox2.[34],[35] Sox2 is likely a critical downstream target of Fgf4 signaling in the TSCs as the Fgf4-independent self-renewal of TSCs can be maintained by the sustained expression of Sox2 and Esrrb [Figure 3]b.[34] Tgfβ/activin promotes the formation of the activated Smad2/3/4 heterocomplex that then accumulates in the nucleus to regulate the expression of target genes.[36] Activin can either maintain the self-renewal of TSCs or promote ST differentiation by cellular-context-specific TFs or transcription cofactors, depending on the availability of Fgf4.[37] It has been shown that Bcor, the X chromosome-linked gene, is specifically downregulated by activin A in a dose-dependent manner in TSCs but immediately upregulated upon TSC differentiation. Moreover, Bcor represses the expression of Eomes and Cebpa by binding to their promoter regions.[38]

Kubaczka et al. and Benchetrit et al. reported that induced TSC (iTSC) lines were obtained by TF-induced transdifferentiation directly from mouse fibroblasts. Both laboratories identified that Gata3/Eomes/Ets2 coexpression was involved in cellular reprogramming and was sufficient to drive TSC fate. The iTSCs showed similar transcriptome, methylation, and H3K27ac of specific loci and faithful H2A.X deposition to the TSCs derived from the embryos. The iTSCs also displayed competent trophoblast differentiation potential as they formed hemorrhagic lesions and formed chimerized E13.5 placenta.[39],[40]

There is evidence that Eomes-null embryos develop to form blastocysts, but they are arrested soon after implantation and fail to form organized embryonic or extraembryonic structures.[41] When the Eomes−/− blastocysts were cultured in the presence of Fgf4, they failed to grow out and retained the blastocyst-like structure, indicating that Eomes is required, cell autonomously, for the transition from TE to trophoblast.[41] Contrastingly, some Ets2-null embryos, which developed to form trophoblast-lineage structures with a generally complete EPC and ExE at E6.5, died before E8.5, due to poor connection to the maternal circulation and possible abnormal basement membrane metabolism.[42] Another study showed that Ets2 is essential for TSC self-renewal as the inactivation of Ets2 in TSCs resulted in slower proliferation and intensive differentiation.[43] Moreover, Gata2 and Gata3 are specifically coexpressed in the TE of mature blastocysts. While the knockdown of Gata3 transcript partially impairs blastocyst maturation, the deletion of Gata2 has no effect on maturation of blastocyst. However, the double loss of Gata2 and Gata3 leads to failed blastocyst development in most embryos and no embryo implantation, indicating that the functions of Gata2 and Gata3 overlap to ensure the progression of peri-implantation development.[44]

The precise mechanisms underlying transcriptional and epigenetic reprogramming events in TSCs remain unknown. Kidder and Palmer reported the transcriptional regulator network of Tfap2c, Eomes, Ets2, Gata3, and a chromatin remodeling factor, Smarca4, which plays an important role in maintaining mouse TSC self-renewal.[45] The majority of regions bound by the five factors are the distal promoter regions (>300 bp upstream or downstream of the transcription start site (TSS) of target genes) in TSCs. Tfap2c was found to bind predominantly to the palindrome motif (GCCNNNGGC) and occupy many more promoters (2,672, about 10% of 28,000 promoters) than Eomes, Ets2, Gata3, and Smarca4 (1,747, 215, 835, and 1, 118, respectively). The five TFs were found to bind to a total of 4,319 genes. Of note, the occupancy of multiple factors occurs in many target genes [Figure 3]c. Among these occupancies, Tfap2c and Eomes shared the highest number of target genes. Significantly, Tfap2c, Smarca4, and Eomes co-occupied 73% of the total genes (432 of 591) bound by any three of the five TFs. The three TF bindings were found in genes that are important for TSC self-renewal and differentiation, such as Elf5, Tead4, Hand1, Notch1–4, Tgf-β receptors (Tgfβr1/3), and Wnt signaling genes (Wnt2b/3a/5a/7a/7b/9a, and Smo). The three TFs also bound additional genes enriched in ESCs and were important for ESC self-renewal and pluripotency, as well as genes of epigenetic regulators, including the de novo DNA methyltransferase Dnmt3b, Dnmt3l, and histone deacetylases (Hdac6/9/10). Compared with genes occupied by single factors, genes occupied by multiple factors were found to be more highly expressed in TSCs than differentiated TSCs or ESCs cells. However, Tfap2c, Smarca4, Eomes, and Gata3 were also enriched in a subset of target genes expressed in differentiated TSCs, suggesting that TF occupancy might inactivate some genes in the TSCs. The regions bound by Tfap2c, Smarca4, and Eomes are also more closely related to H3ac marks than those bound by Ets2 and Gata3, in accordance with the known roles of Smarca4 through its bromodomain in association with acetylated lysine residues on histone H3 and H4 tails for the activation of gene transcripts, suggesting that the mechanism of TSC self-renewal mediated by Eomes may be different from that of Gata3 and Ets2. Eomes bind the promoter of Tfap2c, and the self-promoter binding of Tfap2c and Eomes, show self-regulatory feedback loops between Tfap2c and Eomes in the TSCs. Tfap2c and Eomes further bind to the promoters of Tead4, Elf5, Fgfr, and Gata3. Furthermore, Eomes and Gata3 are also downstream target genes of Tead4 as their expression is lost in Tead4 null mutant embryos.[25],[46] Moreover, Ets2 is a downstream target of Eomes. Thus, Tead4 could be considered as the trophoblast specification factor to initiate the transcriptional regulatory circuitry via Yap1 signaling, whereas Eomes may be in the core of interaction networks to maintain TSC self-renewal via Fgf4/Fgfr signaling.

Gata3 mainly binds to the GATA motif,[45] including Elf5, Esrrb, and Bmp4.[44] Esrrb, the major downstream target gene of Gata3, is also involved in transcriptional regulatory circuitry in the TSCs.[45] Esrrb homozygous mutant embryos died at 10.5 days postcoitum (dpc). The Esrrb mutant placenta showed abnormal chorion and TGC development as early as E7.5. By E9.5, the chorion size was not significantly reduced, and the population of TGCs was extended throughout the EPC.[47] Essentially, Esrrb is an early target of Fgf/Erk signaling in the TSCs as the expression of Esrrb was more rapidly reduced than that of the other TFs (Sox2, Eomes, Elf5, and Cdx2) upon Mek/Erk inhibition by the inhibitor PD0325901.[48] Moreover, Esrrb peaks are highly enriched in the canonical Esrrβ/Esrrα-binding motifs and additionally enriched in a secondary motif that is a Cdx2 binding site. Cdx2 was found to interact with Tead4, Eomes, and Tfap2c, but not with Esrrb, in the TSCs. Indeed, only a small subset of Esrrb peaks (4.1%) are co-bound by Cdx2, suggesting that the core TFs for TSC self-renewal are fitted to diverse motifs to firm up the transcriptional regulatory circuitry. Globally, 60% of Esrrb peaks are co-occupied by Lsd1, a histone demethylase that selectively removes monomethyl and dimethyl groups from either H3K4 or H3K9, which was found to have the ability to maintain the state of the TSC by preventing early onset of differentiation. In TSCs, Lsd1 binds to the core set of Esrrb targets including Elf5, Eomes, Bmp4, and Sox2, showing that the interactors of Esrrb are involved in the epigenetic regulation of transcription. Moreover, Esrrb also interacts with the TFs and transcription cofactor complexes that directly interact with RNAPII, suggesting Esrrb-mediated RNAPII recruitment and activation at its target. While the depletion of the Esrrb gene results in a rapid differentiation of TSCs due to the downregulation of certain key TSC-specific TFs, and these Esrrb-deficient TSCs lost the ability to form hemorrhagic lesions in vivo, ectopically expressed Esrrb can partially block the rapid differentiation of TSCs in the absence of Fgf4. Furthermore, Esrrb can substitute for Eomes to generate iTSCs.[49]

After hatching from the pellucid zone, the proliferation and differentiation of TE cells lead to the development of ExE in proximity to the extraembryonic endoderm and epiblast and the formation of EPC near the maternal decidua. The ExE, especially its distal region near the epiblast, is still in the Fgf4/Fgfr signaling environment. This may be why TSC lines could also be obtained from explant cultures of ExE at E6.5.[30]

Although Elf5 homozygous mutant embryos develop to form EPC, no ExE is formed as early as E5.5.[50] Thus, special attention should be paid to the role of Elf5 during ExE development. Furthermore, Pou2f1 null mutant embryos show no ExE development as the cells inside the extraembryonic endoderm all expressed Pou5f1 but not Cdx2.[51] Both Elf5 and Ets2 are members of the ETS superfamily. Homozygous mutants of either Elf5 or Ets2 result in various degrees of ExE loss, whereas heterozygotes have no phenotype. However, the compound heterozygous mutants (Elf5+/−/Ets2+/−) show a phenotype intermediate to that of the more severe Elf5−/− and milder Ets2−/− mutants.[52] These findings indicate that Elf5 and Ets2 maintain the mouse ExE in a dose-dependent and synergistic manner.

At E6.5, the expression of Eomes, Cdx2, and Esrrb is predominantly localized to the distal ExE near the epiblast, hardly in the EPC.[22],[41],[47] However, the gradient of Tfap2c mRNA in the ExE at E6.5 is observed, with a higher level in proximal ExE than in distal ExE, and with expression throughout the EPC. Tfap2c null mutants lost the capacity to establish a normal maternal–embryonic interface with malformed extraembryonic tissues since E7.5.[27] The expression of Gata3 in the EPC is clearly higher than that in the entire ExE.[53] Elf5 mRNA is mainly limited to the ExE or chorion before E7.5, but its protein is present throughout the extraembryonic trophoblasts, including ExE cells with self-renewal potential and differentiated EPCs.[48],[52] From the pattern of expression of the Elf5, it can be deduced that the EPC are derived from the ExE.

Elf5 and Ets2 play important roles in the maintenance of stemness and differentiation of trophoblasts.[48] In TSCs, the interaction of Elf5 with Eomes recruits Tfap2c to triply occupy the sites of TSC-specific genes and drive their expression, at which state the amount of Tfap2c is comparatively low. However, on differentiation, Elf5 and Tfap2c levels increase, and their interactions no longer predominantly contain other TSC-specific TFs. Consistently, short-term and long-term overexpression of Elf5 and Tfap2c, either alone or in combination, in the TSCs results in their differentiation, mostly toward TGCs and similar to the early stage of differentiation induced by the withdrawal of Fgf4 and EFCM.[48] It has been reported that Ets2 is also required to promote the differentiation of TGC and junctional zone (Jz) trophoblasts. TGC differentiation involves the expression of Ets2-dependent Hand1, a gene required for the differentiation of all TGC types.[54]

The development of ExE is closely associated with the progression of gastrulation primordial germ cell specification in the developing mouse embryo; thus, it is also involved in the functions of Nodal, Bmp4, and Wnt signaling in the two events. Nodal is a member of the TGF superfamily. While immature Nodal protein is secreted from the epiblast and visceral endoderm cells, its activation is mediated by the convertases, Furin and Pace4/Pcsk6, which are expressed in the ExE. The mature Nodal ligand, in turn, promotes the expression of Fgf4,[55] demonstrating that Nodal plays a role in maintaining trophoblast stemness. Similarly, the overexpression of Nodal in trophoblasts significantly inhibits TGC differentiation[56] and an insertional null mutation in Nodal resulted in the hyperplasia of both embryonic ectoderm and ExE as well as failed mesoderm formation in the egg cylinder-stage embryo.[57] With the progression of gastrulation, the most distal ends of the ExE join together, which is mediated by Bmp2 signaling secreted from the extraembryonic mesoderm, as the Bmp2 null mutant mice have defective closed proamniotic canals.[58] The ExE gradually departs from the embryonic proper and develops into the chorion. Similar to Eomes and Cdx2 transcripts, Bmp4 mRNA is specifically expressed in the distal ExE at E6.5. At E7.5, Bmp4 mRNA is extensively expressed in the chorion, with an expression level higher than that in the EPC.[59] However, the Bmp4 null mutant mice appeared to form a complete chorion but have defective allantois, thus lacking the chorion plate at E8.5.[60] Moreover, Bmp4 could induce the generation of trophoblasts from mouse ESCs under defined culture conditions on laminin, while serum inhibits this transformation.[61] Before the derivation of permanent human TSC lines directly from villous cytotrophoblasts, human TSCs were obtained from human embryonic stem cells (hESCs) by the induction of BMP4,[62] suggesting that Bmp4 plays an important role in trophoblast fate determination. However, this conclusion was subsequently challenged by Bernardo et al., who proposed that BMP4 induces human and mouse pluripotent stem cells primarily to form mesoderm, rather than trophoblast, through Brachyury and Cdx2.[63] In addition, at the prestreak stage, Wnt3 transcripts are detected in the proximal epiblast, with higher expression detected posteriorly. Compared with the wild-type embryos consisting of three germ layers and developing the posterior amniotic fold at 7.0 dpc, Wnt3−/− embryos retain the bi-layered structures with one cavity resembling a pregastrula and with a thin layer of ExE.[64] It has been shown that Bmp4 induces Wnt3 to amplify Nodal expression in the epiblast.[65] In turn, Nodal induces Fgf4 to regulate Bmp4 expression in the ExE;[55] thus, the four factors form a signaling circuitry at the ExE–Epi interface.

  Placental Morphogenesis: Orderly Differentiation of Trophoblast Lineage Top

The embryo initiates placental morphogenesis after implantation, during which different trophoblast subtypes are specified and differentiated [Figure 4], finally forming a placenta with mature structure and functions.

The formation of labyrinth layer

The development of the placental labyrinth begins with the critical event chorioallantoic attachment, during which the chorion develops from the EXE attached to the allantois derived from the extraembryonic mesoderm at E8.5. Chorioallantoic attachment has been considered to be mainly mediated by the binding of the adhesion receptor α4-integrin to its ligand Vcam1, which are expressed specifically on the basal surface of the chorion, derived from the mesepithelium of extraembryonic mesoderm, and at the tip of the outgrowing allantois, respectively.[66] While the majority of α4-integrin or Vcam1 null mutant embryos show deficiency in chorioallantoic attachment, the allantois and chorion appear to develop normally.[66],[67] Wnt7b null mutant mice show failure in chorioallantoic attachment due to defective α4-integrin expression, but Vcam1 is normally expressed in allantois,[68] again demonstrating that the interactions between α4-integrin and Vcam1 are fundamental for chorioallantoic attachment. In addition, homozygous Mrj mutants died at mid-gestation due to failed chorioallantoic attachment at E8.5, but the expression of α4-integrin and Vcam1 is normal, indicating that Vcam1/α4-integrin interaction alone is not sufficient for the successful union of chorion and allantois and other molecules might be involved in chorioallantoic attachment.[69]

On initiating chorioallantoic attachment at approximately E8.5, the chorionic mesothelium tends to disappear gradually, and the chorionic epithelium undergoes branching morphogenesis. The chorioallantoic branching morphogenesis, which forms placental villi, is a key milestone during mouse placenta development, guiding allantoic fetal blood vessels toward the maternal blood sinuses. Gene-targeting studies in mice have identified dozens of genes that play essential roles in the morphogenesis of chorioallantoic branching morphogenesis.[5] However, the key molecular interactions between cells in relation to driving branching morphogenesis are still largely unknown. It should be noted that Gcm1 is considered to be the initial TF for driving branching morphogenesis as Gcm1 is specifically expressed in the initial branching site[70] and Gcm1 null mutant mice result in a complete block of branching.[71]

As early as E7.5, Gcm1 mRNA is expressed in a few cells in the chorion near the EPC. At E8.5, the expression of Gcm1 mRNA is limited to trophoblast cells at the tips of the chorion, where trophoblasts directly connect with the allantois. By contrast, most trophoblast cells in the unfolded regions of the chorionic plate are negative for Gcm1, and those expressing Rhox4b are highly proliferative due to coexpression with TSC markers, Cdx2 and Esrrb, and are thought to comprise a reserve of less restricted labyrinth progenitors.[72] Gcm1-positive cell clusters are not overrated with phospho-histone H3-positive cells, indicating that Gcm1-positive cells exit the mitotic cell cycle.[73] A similar phenomenon was also observed in Gcm1-overexpressed TSCs, whose cell cycle is rapidly arrested.[74] The target genes of Gcm1 during placental development in mice remain largely unknown. It has been shown that Gcm1 directly binds to the promoters of Itga4 and Rb1 genes to upregulate their expression.[75] The expression of Gcm1 marks one type of progenitor cell for placental labyrinth development. Moreover, Epcamhigh trophoblasts are multipotent progenitors of the labyrinth in the mouse placenta. HGF/c-Met signaling is found to sustain the proliferation of Epcamhigh trophoblast progenitors and is involved in trophoblast differentiation in the labyrinth.[76],[77] In addition, stem cell antigen (Sca)-1, also called Ly6a, is expressed in trophoblast subtypes with multipotent potential in the chorion and labyrinth layers and persists in the mid to late gestation mouse placenta, marking a subpopulation of trophoblasts that can differentiate to form labyrinth trophoblasts.[37] Recently, Hand1 was reported to be localized in labyrinth trophoblast progenitor cells. Hand1 deletion mediated by Nkx2-5-Cre leads to defective labyrinth development.[78] While different progenitor cells have been identified by distinct markers in the placental labyrinth, the difference or relationship among these progenitor cells needs further investigation.

Although the mechanistic process model of molecular and cellular behavior contributing to branching morphogenesis is still unknown, clues from the changes in cell morphology during branching morphogenesis may provide insight into this process. At the flat chorion stage, Gcm1-positive cells are cuboidal; upon the onset of branching, they become elongated.[73] In Mrj null mutant embryos, Mrj−/− trophoblasts in the chorion plate were more disorganized and rounded, with the collapse of the actin cytoskeleton, failure to establish a membrane-associated deposition of E-cadherin and β-catenin, and disorganization of the ECM.[79],[80] The collapse of the actin cytoskeleton in Mrj−/− trophoblasts resulted from the formation of large keratin aggregates as inclusion bodies, which disrupted cell–cell adhesion and organization. Mrj is required for proteasome degradation of keratin filaments in cultured TGCs, and chorioallantoic attachment is rescued in Mrj−/−/(keratin 18) K18+− and Mrj−/−/K18−/− conceptuses, demonstrating that the cytotoxic effect of keratin aggregates on chorionic trophoblast organization and function is the major cause of disturbance to the chorioallantoic attachment.[80] Furthermore, failure in the deposition of E-cadherin and the disorganization of the ECM are also associated with defective cell behavior.[79] The majority of Cdh1 siRNA-treated trophoblasts were small and rounded upon Fgf4 withdrawal, similar to that of Mrj deficiency. Moreover, Mrj−/− trophoblasts on exogenous laminin-511 substrate regained their morphology in a manner similar to that of wild-type TS and formed attachments to neighboring cells due to more stable adherens junctions. The tetraploid aggregation technique showed that Mrj−/ embryos fail to undergo chorioallantoic attachment because of defects within the chorionic trophoblast layer, rather than from fetal allantois or chorionic mesothelium.[80] Chimeric placentas proceed normally up to E9.5 and show the initiation of branching morphogenesis. Vcam1 and α4-integrin are expressed normally in the allantois and chorion, respectively. In addition, Mrj−/− chorions at E8.5 tend to develop normally but have expanded regions with Rhox4b expression but no Gcm1 expression.[79] These results demonstrate that the functional and structural abnormalities of chorion cells directly and adversely affect chorioallantoic attachment.

Signaling from the fetal blood–vascular system is also required for branching morphogenesis. It has been reported that Rbpj null mutation in mice resulted in failed placental branching morphogenesis, even in the mutants in which chorioallantoic attachment occurred normally. The conditional knockout of Rbpj gene in trophoblast lineage by Cyp19-driving Cre generated a normal labyrinth layer, demonstrating that trophoblastic Rbpj was dispensable for branching morphogenesis. Tetraploid aggregation in combination with specific deletion of Rbpj gene in the epiblast cells (and thus in the allanotic vessels) driven by Sox2-Cre resulted in defective branching morphogenesis, demonstrating that failed branching morphogenesis in Rbpj−/− embryos resulted from the defects within the extraembryonic mesoderm.[81] Under culture conditions, allantois promoted observable morphological changes in TSCs. However, this effect is restricted to trophoblasts directly contacting the allantois, indicating that the signal occurs in a cell–cell paired manner, not in a highly diffusible manner.[73] Both Fzd5 and Wnt2 null mutants have poorly developed labyrinth layers due to failure in branching morphogenesis. Fzd5, a receptor for Wnt proteins, is mainly expressed in chorionic trophoblasts at the branching points, whereas Wnt2 is expressed in the allantois. Wnt2 interacts with Fzd5, especially at the branch point, and upregulates local Gcm1 levels, revealing a mechanism regulating the expression of Gcm1 and a dialog between chorionic trophoblasts and allantois. Fzd5-mediated activation of downstream signaling pathways induces the dissociation of tight junctions at branch points by downregulating tight junction components, such as Tjp1, Cldn4, and Cldn7. Furthermore, Fzd5-mediated signaling pathways are also involved in the expression of Vegfa in the chorionic trophoblast.[82]

During the chorionic branching elongation process, the dialog between chorionic trophoblasts and blood vessels also plays an important role in the expansion of the chorioallantoic interface. Conditional deletion of the Vhl gene in the endothelial cells by Tie2-driving Cre results in small labyrinth layers and defective embryonic vascularization.[83] Esx1 is a maternally-inherited X-chromosome imprinted gene that is specifically expressed in extraembryonic tissues. Heterozygous females who inherited the Esx1 mutation from their mother resulted in both defective labyrinthine morphogenesis and vascularization at the maternal–fetal interface.[84] Apela, a circulating hormone secreted by trophoblasts of the labyrinth layer, interacts with its receptor, Aplnr, expressed specifically in fetal endothelial cells and plays a critical role in preventing preeclampsia.[85]

Several hours after the chorioallantoic attachment, at approximately E9.2, chorionic trophoblasts undergo multinucleation and form STs of the labyrinth layer.[86] There are two layers of STs, ST-I and ST-II, with the latter opposed to fetal vessel endothelial cells. Of note, although ST layers I and II are all syncytial cells, they are very divergent with readily distinguishable intercellular territory and intracellular ultrastructure, as observed by electron micrographs.[70] The molecular properties are even more different between the two layers, such as cell surface markers and the allocation of transport protein.[3],[70],[87] It has been shown that Gcm1−/− mutants result in the complete loss of both ST layers,[71],[88] demonstrating that the formation of the ST layer as a cascade event of branching morphogenesis is controlled directly by Gcm1. A syncytium is formed by cell–cell fusion mediated by SynA and SynB, two proteins that are essential for the generation of multinucleated STs in the placental labyrinth of mice.[89],[90] SynA and its receptor, Ly6e, are localized to the ST-I.[89],[91] Homozygous SynA null mouse embryos die between embryonic days 11.5 and 13.5 (E11.5 and E13.5) with fusion deficiency in the interhemal syncytial layer.[89] Ly6e−/− embryos are morphologically normal at ~ E13.5 and die by E15.5 with a phenotype similar to that in SynA−/− embryos.[92] SynB is expressed in ST-II. While the SynB null mouse embryos exhibited disrupted placenta architecture with impaired formation of ST-II, unfused apposed cells, and enlarged maternal lacunae, were viable except for the limited late-onset growth retardation and reduced neonate number.[90] Therefore, the two ST layers appear to be individually formed as the deletion of either SynA or SynB has no effect on the formation of ST-II or ST-I, respectively. In addition, the expression of Gcm1 is restricted to the basal chorion, where Synb expression is probably directly activated by Gcm1.[71],[75] Recently, TMEM16F was reported to mediate trophoblast fusion and placental development. The deletion of Tmem16f led to defective trophoblast syncytialization in ST-II and subsequent deficiency in maternofetal exchange.[93]

The development of spongiotrophoblast layer

After implantation, the polar ExE continues to proliferate and gives rise to cellular growth protruding into the uterine lumen in the mesometrial region, the EPC. It is generally agreed that the EPC is derived from the proliferation and differentiation of ExE cells and further develop and differentiate to form spongiotrophoblasts.[94] Trophoblasts at the apical part of the EPC near the decidua express Tpbpa, the perfective marker of spongiotrophoblast layer cells.[95]

Ascl2 is a maternally inherited allele expressed in the EPCs and is specifically related to the cellular differentiation of EPCs into spongiotrophoblast layer cells. At E7.5, Ascl2 mRNA is expressed in the proximal chorion cells and mostly in the EPCs.[96] However, at E10.5, the proportion of Ascl2-positive cells is lower than that of Tpbpa-positive cells based on their transcripts.[97] Heterozygous females who inherited the Ascl2 mutation from their mother exhibited embryonic lethality with the loss of spongiotrophoblasts and their precursors and an expanded TGC layer at 10 dpc.[96] However, the morphological structure of the EPC in mutant embryos appeared normal at E7.5. Moreover, Ascl2−/− chimeras with tetraploid wild-type embryos developed normally, and adult Ascl2−/− mice were viable, demonstrating that Ascl2 is essential for placental development.[98] Further studies showed that chimeras had no Ascl2−/− cells in the spongiotrophoblast layer at E10.5 and E12.5, but high contributions of Ascl2−/− cells were observed in the labyrinthine layer, demonstrating that Ascl2 is required for spongiotrophoblasts.[99] Overall, these results show that Ascl2 functions as a bridge for the EPC to form a spongiotrophoblast layer rather than directly becoming secondary TGCs. In addition, Ascl2 and Hand1, another bHLH TFs, are reported to exhibit a reciprocal exclusionary expression pattern as their expression is generally distributed in nonoverlapping regions. The inverse relationship between Ascl2 and Hand1 plays an important role in maintaining the balance between TGCs and spongiotrophoblasts and thus may explain why the TGC layer is expanded in Ascl2−/− mutants.[100]

Ascl2, as an imprinted pathway, controls the spongiotrophoblast layer size in a dose-dependent manner. A Del7AI allele, generated by deleting the interval region between two clusters of imprinted genes (IC1–IC2) on distal mouse chromosome 7, resulted in locally reduced Ascl2 expression by one-half in maternal Del7AI/+ heterozygous pups.[101] Such mutant embryos were viable, although with growth retardation, and the three main layers of the mutant placenta displayed many developmental abnormalities, including reduced placental weight and spongiotrophoblast population, no GlyTs, and an expanded TGC layer. While the mutant placenta showed increased production of the trilaminar labyrinth trophoblast types, the labyrinthine vasculature was disorganized. Moreover, in the mutant placenta, the expression of Phlda2, another maternally expressed imprinted gene in this gene locus, was elevated twice as much as that in the wild type (WT), similar to that in Ascl2−/− mutants, suggesting that it also plays an important role in this phenotype.[101] In addition, the Phlda2 null mutant mice showed an overgrowth of the placenta with a 27% increase in placental weights at E16.5. The hypertrophies of both the labyrinthine and spongiotrophoblast layers, and an increase in the number of GlyTs,[102] demonstrated that Phlda2 acts as an antagonist of placental growth. A transgenic mouse model with elevated Ascl2 expression also exhibited placental and fetal growth restriction. The placentae showed a 40% reduction in the number of parietal TGCs (P-TGCs), a marked loss of spongiotrophoblasts, and a substantial mislocalization of GlyTs into the labyrinth.[103]

Therefore, any genetic, epigenetic, cellular, or maternal factor affecting Ascl2 expression may change the placental structure and composition, as well as pregnancy outcomes. Mst1/2 double-knockout mice exhibited an extremely similar phenotype to Ascl2−/− mutants and conformably had no Ascl2 expression in either EPC or chorion. Furthermore, the failure to reduce the number of trophoblasts did not appear to result from the proliferation abnormality or apoptosis of TGCs and spongiotrophoblasts.[104] While Hand1 null mutant embryos died mostly due to the reduced number of Pl1-positive TGCs, they also showed very small EPC sizes.[100],[105]

Tpbpa-positive trophoblasts in the spongiotrophoblast layer are the progenitors of spiral artery TGCs (SpA-TGCs), canal TGCs, and partial P-TGCs. Hu and Cross eliminated Tpbpa-positive trophoblasts using a Cre recombinase transgene driven by the Tpbpa promoter to irreversibly activate a diphtheria toxin A transgene. Such conceptus showed a reduced number of TGC subtypes associated with maternal blood space remodeling, including SpA-TGCs and canal TGCs, resulting in insufficient spiral artery remodeling with smaller diameters and shallow invasion at E10.5, and death at approximately E11.5, suggesting the essential role of these cells in modulating the maternal vasculature. In contrast, there was an increase in the population of Ascl2-positive/Tpbpa-negative cells, while there was no significant reduction in the population of the P-TGCs population, probably due to the compensation from the Ascl2-positive trophoblasts.[106]

Prdm1-positive trophoblasts, another lineage-restricted progenitor population residing in the spongiotrophoblast layer, not only contribute to the expansion of the spongiotrophoblast layer but also secondarily cause the expansion of the underlying labyrinth layer.[107],[108] Prdm1 is particularly important for SpA-TGC specification since Prdm1 null mutant embryos lose SpA-TGCs.[107] Furthermore, at E9.5 and E11.5, Nodal expression in the placenta is limited to the spongiotrophoblast layer. A hypomorphic mutation in Nodal results in an expansion of both the TGCs and spongiotrophoblast layer and a decrease in labyrinthine development.[56] It is likely that the Nodal secreted from the spongiotrophoblast layer at the mid-gestation stage contributes to the expansion of the labyrinth and maintains the cellular fate of the trophoblasts residing in the spongiotrophoblast layer. These results demonstrate that the deficiency or compromise of the spongiotrophoblast layer disables placental blood circulation.

There was a reduction in the junction zone of HtrA1−/− placenta and fewer SpA-TGCs in the decidua; however, there was more mislocalization of GlyTs in the labyrinth. A significant downregulation of total HtrA1 expression has also been observed in preeclampsia placenta with intrauterine growth restriction.[109] Flt1 is expressed in the spongiotrophoblast layer.[110] The elevation of its soluble form, sFlt1, an antagonist of Vegfa, is inseparably linked with preeclampsia.[111] The relative abundance of sFlt1 or Flt1 mRNAs changed markedly as gestation progressed. At E10.5, sFlt1 mRNA was undetectable, and Flt1 was readily apparent. However, sFlt1 mRNA was abundant, but Flt1 was barely detectable at E16.5.[112] Flt1 appears to be the downstream gene of Ascl2, due to its negatively correlated expression pattern during TSC differentiation.[113] Moreover, Pigf is another receptor of sFlt1, and it appears that Pigf can reduce maternal serum concentration of sFlt1 based on the evidence that Pigf null mutant embryos had increased the levels of sFlt1. Conversely, Pigf-null mutation did not lead to any symptoms of preeclampsia but unexpectedly ameliorated maternal hypertension and preeclampsia in a disease-model mouse with a pregnant catechol-O-methyltransferase deficiency.[114] One possible explanation is that the placenta structure with increased Jz and simultaneously decreased labyrinth area in Pigf-null embryos automatically prevents the incidence of preeclampsia.

Tfap2c is a TF essential for the development of both EPC and ExE as the Tfap2c null mutation results in the disorganization or underdevelopment of extraembryonic structures.[27] The deletion of Tpbpa-Cre-mediated Tfap2c resulted in the initial growth arrest of the Jz in the placenta after E10.5. Tfap2c acts on the proliferation of trophoblasts by repressing Cdkn1a and activating the MAPK pathway, further supporting the specification of GlyTs by activating the Akt pathway.[115]

The initial specification of GlyTs may occur as early as the EPC stage, as glycogen-containing cells identified by PAS staining have been present at E7.5.[116] Starting at approximately E14.5, GlyTs invade the mesometrial decidua. When the spongiotrophoblast volume doubles, as estimated from E12.5 to E14.5, the total number of GlyTs increases by approximately 80 times. The number of GlyTs increases steadily between E14.5 and E16.5, followed by a significant decrease by more than two times in both the spongiotrophoblast layer and the decidua by E18.5.[117] Consistent with this, placental glycogen levels fell by nearly 50% from E15 to E18. Ascl2 may be one of the most important specification factors of GlyTs, as the partial loss of Ascl2 function resulted in the absence of GlyTs.[101] Igf2, a paternally expressed imprinted gene, is extensively expressed in the spongiotrophoblast layer and is then restricted to GlyTs, the tight clusters of cells with abundant glycogen, at E9.5–12.5.[118] However, it appears that Igf2 is not very critical for the entire process of individual development, as the mutant animals are apparently normal and fertile. However, they are growth restricted during gestation, with few GlyTs (only ~20% of wild-type at E15–18) and decreased total placental glycogen content.[119] The biallelic expression of Igf2 can be activated via deletion of the H19 DMR, which results in a greater number of GlyTs and increased total placental glycogen content.[120] Moreover, the conditional loss of Igf2 in the Jzs decreased the volumes of spongiotrophoblasts and GlyTs in the placentae of female but not male fetuses.[121]

Trophoblast giant cell differentiation

TSCs, characterized by an extremely large cytoplasm and polyploid nuclei, are formed at different stages during placental development. Until recently, there were approximately five TGC subtypes present/identified during placental development, including P-TGCs, spiral artery-associated TGCs (Spa-TGCs), cannel TGCs (C-TGCs), sinusoidal TGCs (S-TGCs), and channel TGCs (Ch-TGCs). These TGCs are located in different compartments and play important roles in the placental development [Figure 5]a.[1]

The mural TE of the blastocyst mediates the first interaction events between the implanting blastocyst and the maternal environment in mice. After implantation, the mural TE begins to differentiate and forms primary TGCs. Several secondary TGC subtypes are gradually differentiated during placental development, as described above.

It has been shown that the polyploid nuclei of TGCs are formed via endoreduplication. Unlike the proliferating diploid cells, TGCs undergo polycyclic DNA replication without cell division and have the ability to amass more than 1,000 copies of genomic DNA. As reviewed by Hu and Cross, the endoreduplication of TGCs actually changes the dynamic cyclin profiles at the following three cell-cycle points: G2 decision point, at which the cyclin B/Cdk1 complex is inactivated, allowing cells to escape the M phase and directly transit to the G1 phase; G1 to S checkpoint, at which cyclin D1 substitutes for cyclin D3, and expressions of p53 and Rb decline, allowing cells to go through repeated S phases without intervening mitoses; S phase, during which geminin is degraded and the expression of p57Kip2 resets the periodic S phase during endoreduplication, and the expression of both cyclin A and E maintains phase cyclin/Cdk activities.[1]

While endoreduplication is essential for the development of TGC, Hannibal and Baker[122] reported that endoreduplication in the TGCs did not cover the entire genome but selectively amplified the genome surrounding some key placental genes. High-resolution quantification of DNA content showed that relatively under- and over-replicated loci were within small regions of the genome, with 5% of the genome in 47 loci and 0.2% of the genome in 5 loci, respectively. The underrepresented regions are 28%–54% less than the overamplified regions and 22%–37% more regions than the rest of the genomes have an average of 30–900 copies. However, the underrepresented regions still maintain a greater-than-diploid karyotype. Over-replicated parts of the genome occur in five regions containing gene families of prolactin, serpins, cathepsins, and the natural killer (NK)/CLEC, which are expressed in the placenta and thought to be important for normal pregnancy.[122]

Considering the important roles of TGCs during embryo development, one of the most important physiological functions of TGCs is to remodel the maternal spiral artery via its invasive behavior. The maternal blood is led to the placenta along the spiral artery in which the vascular endothelial cells are replaced by Spa-TGCs and then enters the spongiotrophoblast layer where the C-TGCs converge blood to form the main stream, following which it is dispersed into the labyrinthine layer where the maternal sinus is lined by S-TGCs. Finally, maternal blood inversely flows into the vein lined by Ch-TGCs [Figure 5]a. There is evidence that many over-replicated genes or gene families are responsible for trophoblast invasion and vascular remodeling. The cathepsin family is composed of proteases and cathepsin pro-peptides (Tpbpa, Tpbpb, Ctla2a, and Ctlab). The canonical function of cathepsins is to promote placental invasion by degrading ECM proteins and digesting phagocytosed maternal cells and matrix materials.[123] Other proteinases also have the same function, such as matrix metalloproteinases (Mmp2, 3, 9, 13), inhibitors of metalloproteinases (Timp1, 2, 3, 4), and urokinase plasminogen activators.[1] During human pregnancy, uterine NK cells expressing NK immunoglobulin-like receptors interact with extravillous trophoblasts expressing human leukocyte antigen ligands, two members of the NK/CLEC complex mentioned above, to control both the depth of trophoblast invasion and extent of vascular remodeling.[124]

In addition to the invasive characteristics of TGCs, they also exhibit endocrine behavior. As reviewed by Hu and Cross, Pl1 and Pl2 act on the brain to participate in maternal adaptive behavior. They also act on the mammary gland to promote the proliferation and differentiation of mammary epithelial cells and on the ovaries to stimulate corpus luteum maintenance and progesterone production, as well as on the pancreas, to promote beta-cell proliferation and insulin synthesis and secretion. In addition, other TGC-derived hormones are involved in hematopoiesis, the systemic immune system, and the adaptive function of the liver. Furthermore, hormones and other factors secreted from TGCs are also involved in embryo implantation, uterine decidualization, vasculature remodeling, and the uterine immune system during pregnancy, in a paracrine manner.[1]

With respect to the control of TGC development, Hand1 is considered the most important TF determining the fate of TGC, as Hand1 is expressed in all TGC subtypes. The TF, Hand1, is a member of the helix–loop–helix DNA-binding domain superfamily. The maternal Hand1 mRNA probably contributes to the formation of primary TGCs, due to its presence in the oocyte.[125] At E6.5, Hand1 mRNA is extensively expressed in the EPCs.[126] The homozygous deletion of Hand1 resulted in embryo lethality at approximately E8.5, with defective TGC differentiation.[105] In cultured TSCs, the expression of Hand1 is rapidly elevated after 1-day of differentiation via the withdrawal of Fgf4/EFCM (differentiation), and trophoblasts express TGC markers. Moreover, Hand1 overexpression promotes TGC differentiation even in the presence of Fgf4/EFCM.[74] Accumulated evidence indicates that Hand1 function is critically regulated. Hand1 was found to regulate TGC development as a homodimer whose expression dosage is critical for survival.[100] Sox15 is expressed at a higher level in TGCs than in embryos or various adult tissues. It has been shown that Sox15 interacted with Hand1 through its HMG domain to enhance the Hand1-driven transcription.[127] In addition, it has been shown that the nucleolar release of Hand1 is a crucial step for carrying out its biological activity. In rat choriocarcinoma (Rcho-1) cells, HICp40 interacts with Hand1 to negatively regulate its transcriptional activity through nucleolar sequestration.[128] The phosphorylation of threonine T107 and serine S109 residues of Hand1 mediated by the noncanonical polo-like kinase Plk4 (Sak) is a key step during trophoblast differentiation as it determines the nucleolar–nuclear translocation and dimerization of Hand1.[128],[129]

  Concluding Remarks and Prospects Top

The placenta is a relatively more recently evolved organ. Although the placental architecture varies considerably between species, the gross anatomy of the placenta is not very complicated. Roughly speaking, the placenta is the primary site where the fetus exchanges material and gases with the mother via maternal blood circulation [Figure 1]. However, the trophoblast lineage in the mouse placenta uses a variety of strategies to address placental morphogenesis, including gene duplication and gene amplification, cyto-architectonic changes, such as polyploidization and sensitization, and cellular behavior adaption, such as invasion and angioarchitecture. Why does the placenta, rather than other organs, use these strategies? What changes in the genetic and epigenetic regulatory programs effectively promote the diversity of cell fate? How many trophoblast subtypes does the placenta need and what are their functions? New insights from single-cell differences in the transcriptome and epigenome may help us answer these questions [Figure 5b. In addition, when a single-cell strategy is employed, special attention should be paid to the following points: (1) the true developmental trajectory of different trophoblast subtypes based on the lineage tracing Cre mouse model is needed; (2) the interactions between different cell types in the placenta during placental development could be analyzed based on the scRNA-Seq data, but further experimental evidence is needed.

In this review, we introduce different types of postimplantation defects in placental morphogenesis, resulting from genetically engineered and mutant mice [Supplementary Table 1]. Special attention should be paid to some of them, including: (a) the trophoblasts stop proliferation shortly after implantation, as seen in Eomes and Fgf4 null mutants; (b) the trophoblasts lost multipotency and failed to develop to extraectoderm, which was observed in Elf5 or Pou2f1 null mutant embryos; (c) the EPC degenerated and disappeared after gastrulation, as seen in Tfap2c and Ezh2 homozygous mutant embryos; (d) the specification of TGC fate is seriously compromised, such as in Hand1 null mutant embryos, (e) chorioallantoic attachment completely failed, as seen in Mrj−/− conceptuses, (f) branching morphogenesis failed to occur, as seen in Gcm1 and Fzd5 null mutant embryos, (g) spongiotrophoblast layer is completely lost, as seen in Ascl2 mutant allele from the mother, (h) Spa-TGCs are lost and maternal vascular remodeling is defective, as seen in Prdm1-null embryos.

In this review, we focused on the mouse model to provide a comprehensive understanding of the molecular mechanisms underlying cell-fate specification and the cellular diversity of trophoblast lineage during placental morphogenesis. This is to gain a better understanding of the important organ–placenta during pregnancy and provide theoretical evidence for the prevention and treatment of placenta-related diseases in humans.[130],[131]

Supplementary information is linked to the online version of the paper on the Reproductive and Developmental Medicine website.

Financial support and sponsorship

This work was supported in parts by the National Key R&D Program of China (2018YFC1004102 to J.L.) and the National Natural Science Foundation of China (81830045 to H.W., 31971071 and 82171660 to J.L., and 31701016 to J. W.).

Conflicts of interest

There are no conflicts of interest.

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  [Figure 1], [Figure 2], [Figure 3], [Figure 4], [Figure 5]


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