Molecular Human Reproduction, Vol.20, No.7 pp. 599–618, 2014
Advanced Access publication on April 3, 2014 doi:10.1093/molehr/gau027
REVIEW
Totipotency and lineage segregation
in the human embryo
C. De Paepe 1,†, M. Krivega1,†, G. Cauffman 1,2, M. Geens 1,
and H. Van de Velde 1,2,*
1
2
Research Group Reproduction and Genetics (REGE), Vrije Universiteit Brussel (VUB), Laarbeeklaan 103, B-1090 Brussel, Belgium
Centre for Reproductive Medicine (CRG), Universitair Ziekenhuis Brussel (UZ Brussel), Laarbeeklaan 101, B-1090 Brussel, Belgium
*Correspondence address. E-mail: [email protected]
Submitted on October 18, 2013; resubmitted on March 7, 2014; accepted on March 14, 2014
abstract: During human preimplantation development the totipotent zygote divides and undergoes a number of changes that lead to the
first lineage differentiation in the blastocyst displaying trophectoderm (TE) and inner cell mass (ICM) on Day 5. The TE is a differentiated epithelium
needed for implantation and the ICM forms the embryo proper and serves as a source for pluripotent embryonic stem cells (ESCs). The blastocyst
implants around Day 7. The second lineage differentiation occurs in the ICM after implantation resulting in specification of primitive endoderm and
epiblast. Knowledge on human preimplantation development is limited due to ethical and legal restrictions on embryo research and scarcity of
materials. Studies in the human are mainly descriptive and lack functional evidence. Most information on embryo development is obtained from
animal models and ESC cultures and should be extrapolated with caution. This paper reviews totipotency and the molecular determinants and
pathways involved in lineage segregation in the human embryo, as well as the role of embryonic genome activation, cell cycle features and epigenetic modifications.
Key words: totipotency / human preimplantation embryo / human embryonic stem cells / lineage segregation / epigenetic modifications
Introduction
Human preimplantation development starts with the fusion of two
highly differentiated cells—oocyte and spermatozoon—resulting in a
totipotent zygote. During the first 5 days of embryogenesis, the zygote
divides, changes morphologically and forms a blastocyst. The development from a single totipotent cell into a multicellular organism encompasses intermingling of the maternal and paternal chromosomes,
cleavage divisions of the cells (blastomeres), embryonic genome activation (EGA), particular cell cycle characteristics and epigenetic reprogramming. The embryo undergoes compaction on Day 4 which is
characterized by increased intercellular adhesion and flattening of the
blastomeres. Subsequent cell divisions and cavitation on Day 5 lead to
the formation of a blastocyst (first lineage segregation). It is comprised
of a fluid-filled blastocoel cavity with a compact inner cell mass (ICM)
surrounded by trophectoderm (TE) cells that form a cohesive onelayer epithelium. The blastocyst further expands and hatches out of
the zona pellucida. Just after blastocyst implantation into the endometrium, the ICM diverges into primitive endoderm (PE, also referred
to as hypoblast) and primitive ectoderm (also referred to as epiblast,
EPI; second lineage segregation). Lineage studies in mice indicate
that TE cells contribute to the placenta, PE cells to the yolk sac and
EPI cells to the fetus and extra-embryonic mesoderm (Gardner and
†
Johnson, 1973; Papaioannou et al., 1975; Gardner and Rossant,
1979; Gardner, 1985).
Even though human and mouse embryos seem to be morphologically
similar during preimplantation development, data cannot be extrapolated without caution because important differences exist. First, blastocyst formation corresponds to 3 –3.5 days post-coitus (dpc) in mice,
in contrast to Day 5 after insemination in humans (Hertig et al., 1959;
Brinster, 1963; Steptoe et al., 1971). Secondly, mouse blastocysts
implant at 4–4.5 dpc, whereas human embryos undergo at least one
additional round of cell division before implantation occurs between
Day 7 and Day 9 after insemination (Hertig et al., 1959; Finn and
McLaren, 1967; Norwitz et al., 2001; Cockburn and Rossant, 2010).
Thirdly, human embryos invade into the endometrium (interstitial implantation) whereas mouse embryos attach to the endometrium and are
encapsulated (secondary interstitial implantation; James et al., 2012a, b).
At this moment, our understanding of human preimplantation development and the underlying regulatory mechanisms of totipotency and
differentiation are limited. This is due to the scarcity of human research
materials and the ethical and legal restrictions regarding the use of human
embryos for research purposes in many countries. Early human embryogenesis can only be studied in vitro. Moreover, functional studies are
lacking and data have been extrapolated from embryonic stem cell
(ESC) lines and animal models.
These authors contributed equally to the work.
& The Author 2014. Published by Oxford University Press on behalf of the European Society of Human Reproduction and Embryology. All rights reserved.
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This paper aims to review the current knowledge on totipotency and
differentiation in the human embryo. The molecular determinants and
pathways involved in lineage segregation are discussed as well as the contribution of EGA, cell cycle features and epigenetic modifications. We
also discuss some of the data reported on human embryonic stem cell
(hESC) lines. Finally, we discuss data found in human embryos and
hESC that have been created by somatic cell nuclear transfer (SCNT).
We refer to animal studies, mainly in the mouse, to emphasize similarities
and differences between the species.
Totipotency
At present, there are two definitions of totipotency. According to the
strict definition, totipotency refers to the ability of a single cell to
develop into an adult organism and generate offspring (Edwards and
Beard, 1997), of course, the cell can only demonstrate this potency
after transfer into a uterus. The zygote is the ultimate totipotent cell
because it is able to develop all by itself into the embryo proper with
all embryonic and extra-embryonic lineages (including trophoblast supporting implantation). During preimplantation development, the embryonic cells (blastomeres) progressively lose totipotency, but it is not
known when and how this occurs. Totipotency is lost because the cell
is either committed or too small. Cell commitment or fate refers to an
irreversible developmental restriction (i.e. differentiation) of a cell.
However, the blastomeres become smaller during the early cleavage
divisions. Their size, which is inversely correlated with time, may restrict
their potency to develop into an organism. This limitation may be evaded
using a second less stringent definition of totipotency referring to the
ability of a cell to contribute to all lineages in an organism (Ishiuchi and
Torres-Padilla, 2013). However, this definition may be interpreted differently by scientists going down the ‘slippery slope’ (Box 1). Plasticity has
been used to define an intermediate state between totipotency and differentiation. Plasticity allows the cell to have a developmental preference
toward a certain cell lineage, but this preference is still reversible and thus
the cell is not yet committed.
In animal models, totipotency according to the strict definition has
been investigated by embryo splitting. Totipotency of both blastomeres
at the 2-cell stage has been demonstrated in sheep (Willadsen, 1979),
but not in the mouse. Some cleavage stage blastomeres are proved to
be totipotent at the 2-cell stage in mice (Tarkowski, 1959), the 4-cell
stage in rhesus monkey (Chan et al., 2000) and the 8-cell stage in pigs
(Saito and Niemann, 1991). Bovine is the only model where it has
been demonstrated that the four blastomeres of a 4-cell stage embryo
can develop into four genetically identical calves, proving that the sister
blastomeres are equal and totipotent (Johnson et al., 1995). Several
studies in mice demonstrated that from the 4-cell stage onwards single
blastomeres need carrier cells to develop further into an organism and
thus they are no longer totipotent according the strict definition
(Suwińska et al., 2005; Tarkowski et al., 2010). Each blastomere of the
4-cell mouse embryo was shown by tracing experiments to give rise to
trophoblast and ICM (Hillman et al., 1972). Moreover, some 4- and
8-cell stage blastomeres contribute to all lineages in chimeric mice
(Rossant, 1976; Kelly, 1977; Bałakier and Pedersen, 1982) and thus are
totipotent according to the less stringent definition. Similarly, aggregated
inner and outer blastomeres of mouse compacted embryos (16-cell
stage) are able to develop into live offspring (Suwińska et al., 2008).
Finally, aggregated inner cells of early mouse blastocysts (32-cell stage)
De Paepe et al.
Box 1: The ‘slippery slope’ of totipotency.
1. Strict definition: One cell develops on its own into a fertile organism
a. Zygote
b. Some single early cleavage stage blastomeres
2. Less stringent definition: One cella contributes§ to all lineagesb in an
organismc
a. One cell
- One blastomere
- More (aggregated) blastomeres
- One (or more) pluripotent stem cell(s)8 in vivo and/or in vitro
W Naive
W Primed
b. All lineages
- Cells in all tissues and organs in vivo (embryonic and extra-embryonic
layers including placenta)
- Cells representing all embryonic and extra-embryonic layers in vitro
based on the expression of specific markers and/or functional assays
c. Organism
- Live birth
- Blastocyst in vitro
W Post-implantation with three lineages (EPI, TE and PE)
W Preimplantation with two lineages (ICM and TE)
§
(1) In vivo chimera assay (the cell(s) is (are) injected into a carrier embryo
supporting its growth and development and its descendants are found in all
organs and tissues including the placenta; or (2) in vivo teratoma formation to
test pluripotency (obtained after injection of undifferentiated ESC into
immunocompromised mice and confirming the presence of embryonic
ectoderm, mesoderm and endoderm); or (3) in vitro embryoid bodies
formation to test pluripotency (by formation of 3D multicellular structures
formed by non-adherent cultures of differentiating ES cells and confirming the
presence of embryonic ectoderm, mesoderm and endoderm); or (4) in vitro
by specific lineage differentiation.
8The definition of pluripotency, in particular the capacity to differentiate into
the three embryonic germ layers, becomes problematic. Depending of their
origin in the embryo, primed ESC may be pluripotent (derived from the
pluripotent post-implantation EPI which develops into the three embryonic
germ layers in the embryo) but naı̈ve ESC may be more than pluripotent
(derived from the preimplantation ICM which develops into embryonic and
extra-embryonic lineages in the embryo but not into trophoblast cells).
However, since they display more potency, they may be totipotent according
the less stringent criteria (Supplementary Information, File 1). Paradoxically,
primed ESC lines are more potent in vitro than naı̈ve ESC lines.
develop into blastocysts with ICM and TE cells but do not implant
anymore, however, according to the least stringent definition they are
still totipotent. At this stage, aggregated outer cells only develop into
trophoblast vesicles and thus lost totipotency.
In humans, only the zygote is proved to be totipotent according to the
strict definition (Fig. 1a). One may argue that the phenomenon of bichorionic biamniotic monozygotic twinning provides evidence for totipotency of the 2-cell stage blastomeres. However, it is not known how
this rare event occurs and, despite decades of in vitro culture of human
preimplantation embryos in IVF laboratories, the observation of two
morulas within one zona pellucida has never been reported. Moreover,
according to the strict definition of totipotency, the 2-cell stage blastomeres should behave as two distinct zygotes (Herranz, 2013) thus
their descendant cells should not intermingle during division and compaction (Fig. 1b). A case report describing the birth of a child after the
transfer of a Day 2 cryopreserved embryo, of which only one out of
four cells had survived the procedure, provided evidence that at least
one of the blastomeres at the 4-cell stage is totipotent (Veiga et al.,
Totipotency and lineage segregation in the human embryo
601
Figure 1 Totipotency in the human according to the strict definition: (a) the zygote is totipotent because it can develop into a fertile human being after
implantation into the uterus; (b) the 2-cell stage blastomeres are totipotent if they develop each individually into a human being, the descendants of the
blastomeres do not intermingle during development. Manipulated human embryos cannot be transferred into a uterus to test their potency, but (c) the
sister blastomeres of a 4-cell stage human embryo can develop individually into blastocysts with ICM and TE cells; (d) the descendent cells of one 4-cell
stage blastomere injected with a dye contribute to both ICM and TE lineages.
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1987; Van de Velde et al., 2008; Veiga, personal communication). Indirect support for totipotency at the 4-cell stage in the human was given by
splitting Day 2 embryos into four sister blastomeres that all developed
in vitro into blastocysts with a compact ICM and a cohesive TE monolayer
(Van de Velde et al., 2008; Fig. 1c). Finally, it has been shown that if one
4-cell stage blastomere is injected with a dye, the descendent cells contribute to both ICM and TE (Mottla et al., 1995). Obviously, (some) 4-cell
stage blastomeres can contribute to both lineages and thus are not committed (Fig. 1d). Recently, it has been shown that TE as well as ICM cells
from a full blastocyst can develop into ICM and TE cells, indicating that
they are not yet committed (De Paepe et al., 2013), display plasticity
and are totipotent according to the least stringent definition. The TE
cells lose this potency from expansion onwards. For legal and ethical
reasons manipulated human embryos are not transferred into a uterus
to test their potency and thus totipotency according to the strict definition will never be proved in the human (Alikani and Willadsen, 2002;
Van de Velde et al., 2008; De Paepe et al., 2013).
In summary, the potency of the human cleavage stage blastomeres
remains largely unknown, but, at least, one of the 4-cell stage blastomeres is totipotent according to the strict definition. Human blastomeres
are uncommitted until the full blastocyst stage and, according to the least
stringent definition, full blastocysts’ ICM and TE cells are totipotent.
Human embryonic stem cells
ESC are pluripotent cell lines usually derived from the ICM of the blastocyst and considered as a model to study embryogenesis. They can be
propagated indefinitely in culture in an undifferentiated state (which is
obviously not a characteristic of the zygote or blastomeres). Pluripotency
refers to the capacity of a cell to develop into cells from the three germ
layers in vitro and in vivo. The transcription factors POU5F1 (formerly
called OCT4), SOX2 and NANOG play a major role in sustaining the
undifferentiated state (Boyer et al., 2005, 2006).
In mice, ESC, trophoblast stem cells (TS) and extra-embryonic endoderm stem cells (XEN) have been derived from the blastocyst
(Yamanaka et al., 2006). MESC and mTS lines have also been derived
from single 8-cell stage blastomeres (Chung et al., 2006). The stem cell
lines exclusively contribute to their progenitor lineage in chimeric
animals (Yamanaka et al., 2006). ESC lines have been studied primarily
in the mouse. It is clear now that there are at least two ground states
of mESC (Supplementary data, File 1 and 2): (i) naı̈ve mESC which
correspond to ICM cells from preimplantation blastocysts and depend
upon LIF and BMP4; and (ii) primed mEpiSC which correspond to
post-implantation EPI cells and depend upon FGF and Activin A.
In the human, pluripotent hESC lines (Thomson et al., 1998; Reubinoff
et al., 2000) have been derived from preimplantation blastocysts and
characterized by cell surface markers, differentiation capacity, transcriptomics and (epi)-genomics. The cultures show a high degree of heterogeneity that is partly due to variations in derivation and culture
conditions (Enver et al., 2005; Osafune et al., 2008; Hough et al., 2009;
Pera and Tam, 2010; Nguyen et al., 2013) and/or genetic background
(Chen et al., 2009). HESC are able to differentiate in vitro into extraembryonic endoderm cells (Thomson et al., 1998; Lee et al., 2013)
and trophoblast cells (Thomson et al., 1998; Xu et al., 2002; GeramiNaini et al., 2004; Harun et al., 2006). HESC lines differentiating into
trophoblast cells express specific transcriptions factors (e.g. CDX2 and
GATA3), genes associated with the cytoskeleton (e.g. KRT7 and KRT8)
De Paepe et al.
and the extracellular matrix (e.g. COLA4), genes involved in invasion
(e.g. IGF2, CDH1 or E-cadherin) and hormones (e.g. b-hCG) (Marchand
et al., 2011).
Initially, hESC were derived from ICM cells (Thomson et al., 1998;
Reubinoff et al., 2000) with the highest derivation rate from Day 6 blastocysts (Chen et al., 2009). HESC have also been derived from single 4and 8-cell stage blastomeres (Klimanskaya et al., 2006, 2007; Feki et al.,
2008; Geens et al., 2009; Ilic et al., 2009), indicating that these early blastomeres are at least pluripotent. ICM-derived and blastomere-derived
hESC have similar transcriptional profiles suggesting that during in vitro
culture and derivation the cells that give rise to hESC have a similar precursor cell in the embryo (Giritharan et al., 2011; Galan et al., 2013).
The origin of hESC in the human embryo is unclear. Although they are
generally derived from the ICM, hESC are not the counterpart of ICM
cells because they do not have the same transcriptional profile (Reijo
Pera et al., 2009). HESC and ICM of the full blastocyst both express
surface membrane HLA-G molecules (Verloes et al., 2011), but it is
not the case for all hESC lines (Drukker et al., 2006). HLA-G expression
can be induced in vitro by specific culture conditions such as low
oxygen (Das et al., 2007). Recently, it was shown that isolated and
plated human ICM cells first develop in vitro further toward a post-ICM
intermediate stage and subsequently grow out into a hESC colony
(O’Leary et al., 2012). Based on the expression of the early germ cell
markers DAZL and STELLAR it was suggested that hESC are the equivalent
of early germ cells (Zwaka and Thomson, 2005). However, this hypothesis is doubtful because hESC do not easily differentiate into germ
cells (Geijsen et al., 2004; Nayernia et al., 2006; Aflatoonian et al., 2009).
HESC differ morphologically and functionally from mESC (Ginis et al.,
2004; Schnerch et al., 2010). HESC grow as flat colonies, their undifferentiated state is maintained by adding FGF2 and/or Activin A to the
culture medium. These are similar culture conditions as required for
the derivation and propagation of mTS lines (Yamanaka et al., 2006)
and mEpiSC lines (Brons et al., 2007; Tesar et al., 2007). The derivation
of stable human XEN and TS lines has not yet been reported either after
single blastomere plating or after blastocyst plating (Douglas et al., 2009).
The latter may be correlated with the fast differentiation of trophoblast
into syncytiotrophoblast cells (Rossant, 2008). TS lines have been
derived from rhesus monkey blastocysts but they tend to differentiate
into syncytial-like cells during long-term culture (Vandevoort et al.,
2007). It is now generally accepted that hESC and induced pluripotent
stem cells (iPSCs) obtained after somatic cell reprogramming (genetically
modified by introducing transcription factors Pou5F1, Sox2, Klf4 and
c-Myc; Yamanaka et al., 2006) resemble more mEpiSC than mESC.
Therefore, it has been suggested that hESC may originate from progenitor EPI cells and thus represent primed hESC. This may also explain their
heterogeneity (Osafune et al., 2008; Pera and Tam, 2010; Nguyen et al.,
2013) which has also been described in mEpiSC derived from the
heterogeneous EPI (Brons et al., 2007; Tesar et al., 2007). Moreover,
upon exposure to BMP4 hESC and mEpiSC both differentiate into PE
and trophoblast cells (Xu et al., 2002; Brons et al., 2007). It is a
mystery why hESC and mEpiSC have more in vitro differentiation capacity
than mESC.
For ethical reasons, it is not possible to derive ESC lines from human
post-implantation embryos. Culturing hESC lines with LIF and 2i turns
them into a more naı̈ve state but these naı̈ve hESC cannot be propagated
in the long-term and differentiate (Hanna et al., 2010). Very recently,
however, hESC have been stably converted into a naı̈ve state without
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Totipotency and lineage segregation in the human embryo
genetic modification by using medium supplemented with a cocktail of
chemical inhibitors (Gafni et al., 2013). This supplemented medium
has also been successfully used to derive stable naı̈ve pluripotent stem
cell lines from human preimplantation blastocysts.
In summary, hESC represent a model to study embryogenesis in vitro.
However, undifferentiated embryonic cells are only transiently present in
the embryo. Moreover, hESC lines have been adapted to long-term in
vitro culture conditions and may even be an in vitro artifact. Finally,
hESC rather resemble mEpiSC than mESC, it has been suggested that
hESC represent a primed stem cell state derived from post-implantation
EPI cells that arise in culture after explanting the preimplantation blastocyst. Recently, a more naı̈ve state of hESC has been obtained using specific culture conditions indicating that, similar to the mouse, there are
distinct states of hESC. Therefore, data from ESC cultures should be
extrapolated to the human embryo with caution.
Lineage segregation
Lineage segregation into TE, PE and EPI is mainly controlled by transcription factors. In mouse embryos, the first segregation is the consequence
of reciprocal inhibition of POU5F1 and CDX2 in ICM and TE (Niwa et al.,
2005; Ralston and Rossant, 2005; Strumpf et al., 2005). The second segregation resulting in PE and EPI is the result of mutual interaction between
NANOG and GATA6 (Chazaud et al., 2006). Several models may
explain the segregation of the lineages in the mouse embryo (Box 2):
sorting (Chazaud et al., 2006; Dietrich and Hiiragi, 2007); position
(Tarkowski and Wróblewska, 1967); polarization (Johnson and McConnell, 2004); waves of division (Bruce and Zernicka-Goetz, 2010). In mice,
some of the key regulatory pathways involved in lineage segregation have
been identified: Hippo signaling (Nishioka et al., 2008) and BMP4 (Home
et al., 2012) in the first differentiation (Box 3) and FGF/Grb2 (Chazaud
et al., 2006) in the second differentiation (Box 4). The interaction
between POU5F1, NANOG and GATA6 has also been thoroughly
investigated in mESC (Niwa et al., 2000; Boyer et al., 2006; Nishiyama
et al., 2009) and hESC (Hay et al., 2004; Chew et al., 2005; Hyslop
et al., 2005; Zaehres et al., 2005; Darr et al., 2006; Fong et al., 2008).
Interestingly, during reprogramming of differentiated murine somatic
cells into iPSC (another type a pluripotent cells) by Pou5f1, Sox2, Klf4
and c-Myc (Takahashi et al., 2007), Pou5f1 can be replaced by E-cadherin
suggesting that Pou5f1 expression is regulated by cell– cell contact via
E-cadherin. Interestingly, E-cadherin is linked to the WNT signaling
pathway by b-catenin (Redmer et al., 2011).
The first differentiation
In the human, morphological differences between cells (polarization
and/or position) have not been reported before compaction (Nikas
et al., 1996) which establishes onset of the first differentiation. E-cadherin
molecules act as homotypic receptors and contribute to adhesion of
compacting cells concentrating in the areas of blastomere–blastomere
contact (Alikani, 2005). At the blastocyst stage, TE cells show a strong
membrane localization of E-cadherin. Gap junctions (marked by connexin CX43) between the blastomeres are already detected at the
4-cell stage, but they become more dense during development and
more apparent in the TE layer when compared with ICM cells (Hardy
et al., 1996). Tight junctions (marked by ZO1) and desmosomes are
exclusively established between the outer cells at compaction and
Box 2: Lineage segregation models in
mouse embryos.
(1) A stochastic model was proposed in the mouse to explain the first
differentiation at the compaction stage illustrating inter-blastomere variation
in the amount of master proteins NANOG, POU5F1 and CDX2 followed by
a phase of positional change (sorting) depending on the global differences in
gene expression (Dietrich and Hiiragi, 2007). Using time lapse video, it was
shown that the blastomeres move extensively at each cleavage stage
(Kurotaki et al., 2007), supporting the model of cell sorting and consistent
with the highly regulative capacity of the embryo. The second differentiation
occurs in a similar way. Initially, the EPI- and PE-specific transcription factors
NANOG and GATA6, respectively, are expressed in a random
“salt-and-pepper” pattern in the ICM, followed by segregation into the
appropriate cell lineages (Chazaud et al., 2006).
(2) The “inside– outside model” proposes that lineage segregation is directed
by the position of the cell (Tarkowski and Wróblewska, 1967): outside cells
develop into TE and inside cells develop into ICM. According to this model,
cells on the inside and on the outside are exposed to distinct environments
and different amounts of cell contact resulting into distinct fates.
(3) The “cell polarity model” proposes that polarization is associated with
differences in transcription factor expression. At the 8-cell stage, blastomeres
undergo an increase in intercellular contact (compaction) and polarize along
their apical– basal axis (Johnson and McConnell, 2004). Polarization is
characterized by the apical localization of members of the Par complex (Par3,
Par6 and aPKC); (Plusa et al., 2005; Alarcon, 2010). Also Cdx2 mRNA
becomes polarized at the apical cortex of polarized cells (Jedrusik et al., 2008).
During the two subsequent divisions (8– 16 cells and 16– 32 cells), the
inheritance of the polarized state is influenced by the orientation of the
cleavage plane in the blastomere: symmetric (conservative) divisions generate
polarized outer cells, whereas asymmetric (differentiative) divisions generate
polar outer cells and apolar inner cells. At the 32-cell stage, the polar cells
become TE whereas the apolar cells form the ICM and differentiate into EPI
and PE. The two models—position and polarization—may work in concert to
direct cell lineage segregation. Individual blastomeres separated from 2- to
32-cell stage embryos do not show a lineage-specific pattern but rather
develop a unique pattern that is similar to TE (Lorthongpanich et al., 2012). It
seems that the correct patterning of lineage-specific gene expression requires
positional signals and cell– cell interaction.
(4) Another model was proposed suggesting that the first wave of asymmetric
divisions would generate most of the EPI lineage whereas the second wave
would generate most of the PE lineage (Bruce and Zernicka-Goetz, 2010).
Cells that are not appropriately positioned change their position, gene
expression profile or die by apoptosis.
become clearly apparent at blastocyst expansion to support the integrity
of the TE cells (Hardy et al., 1996). KRT18 expression is present in the
cytoskeleton of some outer cells in the compacting embryo and
further on it is found in TE at all blastocyst stages and in ICM cells
facing the cavity (Cauffman et al., 2009). HLA-G, another marker for
TE lineage differentiation, is present in the membrane of inner and
outer cells at compaction (Yao et al., 2005). It is also transiently
present in the membrane of early ICM cells (Verloes et al., 2011), but
it becomes restricted to the TE cells and ICM cells facing the cavity at
the moment of hatching (Verloes et al., 2011). These observations indicate that the outer cells become polarized at compaction and already
obtain epithelial features induced by the environment although they
are not yet committed to the TE lineage (De Paepe et al., 2013). The
TE-defining transcription factor CDX2 is only detectable in the nuclei
of the outer layer from blastocyst expansion onwards (Niakan and
Eggan, 2013). During a short period, at the onset of expansion,
POU5F1 and CDX2 are co-localized in the nuclei of TE cells. The
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De Paepe et al.
Box 3: The first differentiation in mouse
embryos.
Box 4: The second differentiation in mouse
embryos.
The establishment of ICM and TE lineages in mice begins with the
up-regulation of Cdx2 in outside cells followed by down-regulation of Pou5f1,
Sox2 and Nanog in the same cells. Initially, Cdx2 and Pou5f1 are co-expressed
in the morula; the reciprocal repression occurs in blastocysts and in mESC
(Niwa et al., 2005). Depletion of maternal and zygotic Cdx2 mRNA results in
delayed embryo development with increased cell cycle length and problems
to initiate compaction (Jedrusik et al. 2010). Inconsistent with this observation
are two studies in which it was demonstrated that Cdx2 null embryos reach
the blastocyst stage and collapse around the time of implantation (Wu et al.,
2010; Blij et al., 2012). The embryos still cavitate and form a distinct ICM after
elimination of maternal and zygotic Pou5f1 expression (Wu et al., 2013). In
maternal/zygotic knockout embryos, CDX2 is not found in ICM cells and
NANOG is found in cells that are scattered apart in the ICM. Later on,
NANOG and CDX2 are co-localized in some EPI nuclei. Thus, the reciprocal
POU5F1/CDX2 interaction does not result into the first lineage
differentiation but rather maintains the ICM fate. Pou5f1 null embryos form
ICM but plating these ICM does not lead to the derivation of mESC lines and
the outgrowth containing a lot of Cdx2 expressing trophoblast cells. Thus,
although POU5F1 is the major regulator of pluripotency in mESC (Boyer et al.,
2006), maternal POU5F1 it is not a major regulator of pluripotency in oocytes
(Wu et al., 2013).
Recently the Hippo signaling pathway, which plays a role in cell contact in
cultured cells (Zhao et al., 2007a), has been described in the reciprocal Cdx2/
Pou5f1 repression in the embryo (Nishioka et al., 2009). The Hippo pathway
involves the transcription factor TEAD4 and its co-activator YAP. TEAD4 acts
upstream of CDX2 and is present in the nuclei of all the cells (Nishioka et al.,
2008). Cell contact and/or position may activate the Hippo signaling, resulting
in YAP phosphorylation and subsequent nuclear exclusion in inside cells.
Without the presence of YAP in the nucleus, TEAD4 is inactive and Cdx2
expression is silenced (Cockburn and Rossant, 2010). In outside cells, Yap is
not phosphorylated and localized in the nuclei where it can, in cooperation
with Tead4, activate Cdx2 expression. Tead2/2 embryos fail to cavitate (Yagi
et al., 2007; Nishioka et al., 2008); Cdx22/2 embryos cavitate but fail to
maintain TE (Strumpf et al., 2005; Jedrusik et al., 2010) and Pou5f12/2
embryos display a defective ICM (Nichols et al., 1998).
The Hippo pathway may not be the only pathway involved in the first
lineage segregation in the mouse embryo. Culturing mouse embryos with
BMP4 blocks their development at the compaction stage and results in the
unusual co-localization of CDX2 and TEAD4 in the nuclei of inner cells
(Nishioka et al., 2009; Home et al., 2012).
Signaling through the fibroblast growth factor (FGF)/mitogen-activated
protein kinase (MAPK) pathway is the earliest event known influencing
differentiation of the mouse ICM into the EPI and PE. This pathway leads to the
expression of the GATA transcription factors, GATA4 and GATA6, which
become restricted to the PE (Feldman et al., 1995; Arman et al., 1998; Cheng
et al., 1998; Chazaud et al., 2006) and the EPI marker NANOG (Mitsui et al.,
2003). The transcription factor NANOG is initially present in all cells from the
morula stage onwards but it becomes down-regulated in the outer cells at the
blastocyst stage. Within the ICM cells, the progenitor EPI cells express Nanog
and produce FGF4 whereas the progenitor PE cells express Gata6 and the
Fgf2r receptor. Laminin expression seems to play a role in this lineage
segregation and remains restricted to PE cells. Initially, the progenitor EPI and
PE cells are distributed in a random salt-and-pepper way, the distinct lineages
segregate after sorting. Grb22/2 embryos only display EPI cells in the ICM
(Chazaud et al., 2006). Nanog2/2 blastocysts have ICM cells but they fail to
generate EPI (Mitsui et al., 2003). Heterozygous Nanog+/2 blastocysts have
similar numbers of ICM cells in the early blastocysts when compared with
Nanog+/+ blastocysts but they have fewer ICM cells in the EPI. In Nanog+/2
blastocysts fewer ICM cells are found displaying NANOG and PE formation,
which depends upon functional EPI, is delayed.
Another transcription factor, SOX17, has also been described in the
specification of PE cell from EPI cells within the mouse ICM (Morris et al.,
2010; Niakan et al., 2010; Artus et al., 2011).
phenomenon of co-expression has also been observed in mouse, bovine
and rhesus monkey embryos (Degrelle et al., 2005; Berg et al., 2011) suggesting that the segregation of the TE and ICM markers is initiated just
prior to implantation. By Day 8, POU5F1 becomes restricted to a
small population in the EPI indicating that distinct populations arise
within this lineage (Chen et al., 2009). At this time, CDX2 appears to
be down-regulated in TE. This coincides with the time when the trophoblast cells adhere and invade into the endometrium. However, whether
these data represent the true in vivo situation or result from in vitro culture
conditions, in particular absence of implantation into endometrial cells, is
not known.
Finally, whereas transcription factor binding sites for TCFAP2 that
mediate CDX2-independent repression of the pluripotency marker
POU5F1 are present in the mouse, they were not found in humans
and cattle, suggesting alternative mechanisms for lineage commitment
in different species (Berg et al., 2011; James et al., 2012a, b). So far,
the molecular mechanisms that mediate the first lineage segregation in
the human remain largely unknown. The Hippo signaling pathway
might be conserved between species, but information about this
pathway in the human is currently not available.
The other lineage, ICM cells, has also been investigated in the human.
The ICM markers POU5F1, SOX2 and NANOG were already well
described in hESC (Boyer et al., 2005; Hyslop et al., 2005; Zaehres
et al., 2005). They bind to the promoters of their own genes forming
an interconnected auto-regulatory loop controlling pluripotency and
self-renewal (Boyer et al., 2005). NANOG is only present in the nuclei
of some ICM cells in the full/expanding blastocyst (Hyslop et al., 2005;
Cauffman et al., 2009; Niakan and Eggan, 2013). POU5F1 is found
earlier in the nuclei of inner and outer cells at compaction and in ICM
and TE cells at the full blastocyst stage (Cauffman et al., 2005b; Niakan
and Eggan, 2013). It is down-regulated in the outer cells in the expanded
blastocyst. Interestingly, NANOG is restricted to ICM cells earlier than
POU5F1 (Niakan and Eggan, 2013). SOX-2 expression starts from
the 8-cell stage onwards; but its nuclear expression is not restricted to
the inner cells at compaction nor to ICM cells at the full blastocyst
stage. SOX2 is only down-regulated in the TE cells after expansion of
the blastocyst. Another transcription factor associated with the undifferentiated state, SALL4, is expressed in the nuclei of inner and outer cells at
all stages from compaction till blastocyst expansion when it becomes
restricted to the ICM cells. Thus, none of the markers for the undifferentiated state can be used to identify cells allocated to the ICM until
expansion (Cauffman et al., 2009).
The co-localization of lineage markers such as POU5F1, SOX2,
SALL4, KRT18, HLA-G and the absence of CDX2 in human TE cells displaying plasticity at the full blastocyst stage explain the ability of isolated
and reaggregated TE cells to reconstitute a blastocyst with a compact
ICM comprising NANOG expressing cells and a cohesive TE layer
(De Paepe et al., 2013). Additionally, full blastocyst TE cells can change
lineage direction when they are placed in an inner position. These data
suggest that full human blastocyst TE cells are not yet committed
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Totipotency and lineage segregation in the human embryo
toward the TE lineage and may thus be a potential source of hESC. This
potency is lost during expansion since isolated and reaggregated TE cells
at this stage do not recompact anymore. This coincides with the onset of
CDX2 expression (Niakan and Eggan, 2013) and the up-regulation of
ZO-1 (Hardy et al., 1996) supporting the establishment of an integer
and functional TE monolayer. Commitment occurs at the early blastocyst
stage in mice (Suwińska et al., 2008) and in the expanded blastocyst stage
in cattle (Berg et al., 2011) corresponding with the reciprocal localization
of CDX2 and POU5F1.
Mouse early blastocyst inner cells (Suwińska et al., 2008) and human
full blastocyst ICM cells (De Paepe et al., 2013) have been shown to be
capable of generating TE cells. The ability of a mouse ICM cells to differentiate into TE is controversial. Recently, it has been described that isolated mouse ICM is not able to differentiate into trophectoderm
(Szczepanska et al., 2011); however, in these experiments, more
advanced blastocysts were used in combination with distinct experimental procedures. Using hESC as a model to study embryogenesis in vitro, it
has been found that they have the ability to spontaneously differentiate
into trophoblast cells (Thomson et al., 1998; Gerami-Naini et al.,
2004; Harun et al., 2006). Long-term culture of hESC lines in an undifferentiated state depends upon WNT, FGF and TGFb pathways (Box 5).
The role of WNT is unclear, most likely it enhances proliferation
(Dravid et al., 2005) but it has also been correlated with differentiation
(Sokol, 2011). FGF and Activin A are required for the self-renewal of
hESC (Amit et al., 2000; James et al., 2005; Lu et al., 2006; Xiao et al.,
2006). Activin A is a member of the TGFb superfamily. BMP4, another
member of the TGFb superfamily, antagonizes with Activin A and
induces differentiation toward the TE lineage (Xu et al., 2002, 2008;
Wu et al., 2008). Activin A and BMP4 have distinct receptors. Receptor
binding transduces signals through R-SMAD proteins SMAD2/3 and
SMAD1/5/8, respectively. The phosphorylated R-SMADs bind to the
common SMAD4 and together they form a complex. This complex
enters the nucleus where it directly activates transcription of distinct
target genes. It has been suggested that Activin A and BMP4 may antagonize and play a role in the balance between pluripotency and differentiation by competing for SMAD4 (Xu et al., 2008). Recently, it has
been found that SMAD2 plays a major role in sustaining the self-renewal
of both hESC and mEpiSC by binding directly to the NANOG proximal
promoter leading to its up-regulation (Sakaki-Yumoto et al., 2013b).
Down-regulation of SMAD2 in hESC results in BMP4 signaling activation.
This leads to differentiation toward endoderm (SOX17) and trophoblast
(CDX2) lineages. The reciprocal interaction between CDX2 and
POU5F1 further supports differentiation. This result not only supports
the hypothesis that hESC and mEpiSC are similar, but it also demonstrates that pluripotency-versus-lineage segregation is controlled by antagonistic Activin A-versus-BMP4 interaction (Fig. 2). Finally, in mESC
NANOG binds to SMAD1 limiting BMP signaling that promotes differentiation into mesoderm (Suzuki et al., 2006). It would be interesting to
know whether, in the human, NANOG also binds to SMAD1/5/8 inhibiting trophoblast differentiation. FGF2 supports Activin A-dependent
self-renewal of hESC via NANOG expression, but the exact mechanism
is not understood (Vallier and Pedersen, 2005; Greber et al., 2008;
Greber et al., 2010).
The second differentiation
GATA6, GATA4 and SOX17 proteins have been identified in progenitor
PE cells in human expanded blastocysts (Kuijk et al., 2012; Roode et al.,
2012). At this stage, some inner cells exhibit high levels of NANOG and
low levels of GATA6, whereas in other cells both markers are expressed
at about the same level. This pattern is similar to the salt-and-pepper distribution described in mouse blastocyst ICMs (Chazaud et al., 2006).
After hatching GATA6 and NANOG are expressed in a mutually exclusive manner indicating segregation of PE and EPI, respectively (Kuijk et al.,
Box 5: Signaling pathways associated with
pluripotency and differentiation in human
embryos and hESC.
(1) FGF: Binding of FGF to FGF receptor homodimers leads to MAPK signaling
which activates transcription factors in the nucleus (Stephenson et al., 2012).
(2) TGFb superfamily: Binding of homodimers of BMP4, Activin A/Nodal or
TGFb to heterodimers of the Type I and Type II TGFb receptors leads to
phosphorylation of cytoplasmic SMADS. The phosphorylated R-SMADs bind
to the common SMAD (coSMAD4) forming a complex that acts as a
transcription factor for distinct target genes. Next to SMAD signaling, other
non-SMAD pathways can be initiated by TGFb receptor activation, including
MAPK. For example, TGFbII can phosphorylate PAR6 resulting in the
dissemblance of tight junctions and epithelial to mesenchymal transition
(Moustakas and Heldin, 2009).
(3) WNT: Upon activation of the canonical WNT pathway, the b-catenin
regulatory complex (Axin, APC and GSK3) is degraded. b-catenin, an
E-cadherin adaptor protein which is normally degraded through
phosphorylation by its regulatory complex, is accumulated in the cytoplasm
and translocated into the nucleus where it will act as a transcriptional
co-activator (Sokol, 2011).
There is no doubt that these pathways interact with each other, e.g. GSK3
plays a key role in WNT signaling but also interferes with SMAD signaling;
MAPK which plays a role in FGF signaling and the non-SMAD TGFb signaling
pathways also has an effect on the SMAD TGFb pathway (Sakaki-Yumoto
et al., 2013a).
Figure 2 TGFb signaling in hESC. Activin A and BMP4 antagonize in
sustaining pluripotency in hESC: SMAD2 induces NANOG expression,
down-regulation of SMAD2 results in CDX2 expression via BMP4 signaling (Sakaki-Yumoto et al. 2013b). The interaction between
NANOG and SMAD1/5/8 has not yet been demonstrated in hESC
(dashed line).
606
2012; Roode et al., 2012). SOX17 is initially detectable in early blastocysts (Niakan and Eggan, 2013). At the expanded blastocyst stage,
SOX17 is highly expressed in the nuclei of all ICM cells, whereas in
hatched blastocysts SOX17 expression is restricted to the putative PE
within the ICM. Limited SOX17 expression has also been described in
hatched blastocysts coinciding with GATA4 expression (Roode et al.,
2012). GATA6 expression is detectable in the majority of SOX17
expressing cells, except for a few SOX17-positive cells within the ICM
(Niakan and Eggan, 2013). In contrast to the mouse, human blastocysts
express laminin in TE cells and not in PE cells, suggesting that PE lineage
specification may be distinct between these two species. Finally, the epithelium markers HLA-G (Verloes et al., 2011) and KRT-18 (Cauffman
et al., 2009) are present in the ICM cells facing the cavity of full and
expanded blastocysts. This may be induced by the environment (blastocoel fluid) but it does not fit into the model of salt-and-pepper distribution of progenitor PE and EPI cells followed by sorting into the two
distinct lineage layers.
FGF/MAPK signaling plays a major role in the second lineage differentiation in mice and bovine. Bovine embryos cultured with FGF4 and
heparin develop into blastocysts with an ICM that is entirely composed
of PE cells (Kuijk et al., 2012). However, MAPK signaling inhibitors do
not fully ablate the PE progenitor cells in bovine embryos implying
other signaling pathways for second lineage segregation. In the mouse,
however, pharmacological inhibition of MAPK signaling or FGF receptor
inhibition in mouse embryos blocks the appearance of PE cells (Nichols
et al., 2009; Yamanaka and Ralston, 2010). Mouse embryos cultured in 2i
conditions exclusively give rise to the EPI lineage in the ICM. On the other
hand, co-culture with FGF4 solely induces the PE lineage in the ICM
(Nichols et al., 2009). Studies on human blastocysts have demonstrated
that FGF/MAPK signaling is not an evolutionary conserved mechanism
for the specification of EPI and PE lineages. The 2i conditions have no
effect on the EPI cells in the human embryo (Roode et al., 2012). This
has been confirmed by inhibiting MAPK signaling (Kuijk et al., 2012;
Roode et al., 2012), indicating that FGF/MAPK signaling is not imperative
for this lineage segregation in the human. The mechanism of PE lineage
specification in the human remains unknown.
In summary, reports on lineage differentiation in the human embryo
mostly describe the expression of specific markers known from animal
models and hESC. The models described in lineage segregation in mice
have not been validated in human embryos. Very few functional
studies have been reported. A number of studies in hESC point out distinct signaling pathways which play a role in sustaining the pluripotent state,
but these pathways have not been investigated in the human embryo. Although the data in the human are limited, they are of great value because
they indicate differences between distinct species and provide new insights
into lineage segregation during early human embryogenesis.
Embryonic genome activation
Embryonic gene expression does not start immediately after fertilization.
First, during early mitotic divisions, maternal mRNA and proteins are
degraded. Secondly, the newly formed embryo has to activate its
genome, i.e. to start up a transcriptional and translational machinery to
support its own growth and development. It is likely that the mRNAs
and proteins required for oocyte maturation and fertilization, which
are dispensable for further embryo development, are degraded fast.
Those that are preserved until this point are necessary to sustain the
De Paepe et al.
first mitotic divisions and to activate the embryonic genome and, therefore, they are only degraded after EGA. EGA is one of the most important
events in embryogenesis, but how exactly it is triggered is not yet completely understood. The cytoplasmic content changes dramatically
during the first cleavage divisions by maternal mRNA degradation and
EGA. These changes may have an effect on the totipotency of the cleavage stage blastomeres. The timing of EGA onset differs between distinct
species. In cats EGA starts at the 2-cell stage (Waurich et al., 2010), in
sheep and cattle it starts at the 8-cell stage (Crosby et al., 1988; Kues
et al., 2008) and in rabbits it starts around the first differentiation in the
blastocyst (Léandri et al., 2009).
In mice, clearance of RNA starts shortly after fertilization. The major
part of the maternal transcripts is degraded in response to deadenylation
while elimination of the other part requires participation of the products
of the embryonic genome (Tadros and Lipshitz, 2009). EGA starts
between the 1- and 2-cell stages followed by a peak activity between
the 2- and 4-cell stages (Aoki et al., 1997; Hamatani et al., 2004; Wang
et al., 2004). A relationship between the undifferentiated state and
EGA has been shown for POU5F1 that appears to be critical for the expression of regulatory genes involved in transcription, translation, RNA
polyadenylation and RNA degradation and thus can act as an upstream
regulator of EGA in mice (Foygel et al., 2008).
In the human embryo, the majority of maternal mRNA is degraded
between the 2- and 4-cell stage (Dobson et al., 2004; Wong et al.,
2010; Vassena et al., 2011), followed by a gradual disappearance of the
remaining transcripts over time (Vassena et al., 2011). Three successive
waves of transcription have been found during the cleavage stages. EGA
starts at the 2-cell stage with a minor wave of transcription that correspond with factors involved in transcription, protein synthesis and metabolism (Vassena et al., 2011). The second minor wave follows at the 4-cell
stage. The third and major wave of transcription occurs at the 8-cell stage
(Braude et al., 1988; Vassena et al., 2011) and coincides with the expression of genes involved in mRNA and protein metabolism, development
and differentiation. Later on, at the blastocyst stage, another major wave
of specific gene expression starts involving genes that regulate further
embryo development and organogenesis, implantation and placentation
(Zhang et al., 2009a; Wong et al., 2010; Vassena et al., 2011). A subset of
transcripts also consists of genes that are stably maintained throughout
preimplantation development, e.g. housekeeping genes.
Multiple waves of EGA may affect the balance between totipotency
and differentiation. Several groups analyzed the temporal and spatial localization of the lineage-defining transcription factors during human preimplantation development (Cauffman et al., 2005b, 2009; Niakan and
Eggan, 2013). It is clear that the totipotent human zygote does not
display any nuclear expression of the three key transcription factors
(SOX2, POU5F1 and NANOG) sustaining the undifferentiated state in
hESC (Cauffman et al., 2009). The low cytoplasmic staining of SOX2
and POU5F1 in early cleavage stages can most likely be attributed to proteins present from the maternal stock. However, it cannot be excluded
that the assays were not sensitive enough to detect nuclear localization of
the proteins. Another explanation could be that the antibodies (or
primers in case of mRNA) are directed against a specific isoform. For instance, in the case of POU5F1_iA and POU5F1F_iB (isoforms formerly
called OCT4A and OCT4B), only POU5F1_iA is associated with the undifferentiated state in human embryos and hESC (Cauffman et al., 2006).
And finally, one should keep in mind that mRNA expression precedes
protein synthesis and transport. Thus, not the presence of the
Totipotency and lineage segregation in the human embryo
transcription factors’ mRNA, but the proteins in the nuclei should be
correlated with differentiation.
The findings in human early cleavage stage embryos are supported by
data in mice. The blastomeres of early mouse embryos (before and immediately after EGA) do not express these factors (Dietrich and Hiiragi,
2007). Pou5f1_ia (formerly Oct4a isoform) null female mice are fertile,
strongly indicating that maternal Pou5f1 mRNA, which is present in the
oocyte, is not a major regulator of totipotency (Wu et al., 2013).
Embryos lacking Sall4, another marker for the undifferentiated state, are
also totipotent (Elling et al., 2006). Thus, the founder lineages can arise
in the blastocyst without the transcription factors POU5F1 and SALL4.
Based on data currently available, the transcription factors are only
produced and become active in the embryonic nuclei after EGA. The
major wave of transcription at the 8-cell stage in human preimplantation
embryos is particularly interesting for lineage segregation. In the human
embryo, POU5F1 transcripts are already present at the 4-cell stage followed by SOX2 and NANOG at the 6-cell stage (Vassena et al., 2011). Initially, a mutually exclusive expression of b-HCG/b-LH and POU5F1 was
found in cleavage stage blastomeres from Day 3 human embryos suggesting early commitment to TE and ICM lineages (Hansis et al., 2004; Hansis,
2006). Contradictory data, however, showed that single 5- to 8-cell stage
blastomeres display a common gene expression pattern corresponding
with both the undifferentiated (NANOG, POU5F1 and SOX2) and trophoblast (CDX2 and EOMES) phenotypes (however, we cannot confirm the
expression of CDX2 and EOMES at these early stages). This illustrates
that the blastomeres are not immediately committed after EGA.
Variability in RNA expression between embryos of the same stage and
between individual blastomeres of the same embryo on Day 3 should be
interpreted with caution. First, embryo quality may affect the outcome of
gene expression analysis. Embryos arrested at the early cleavage stages
undergo EGA (Dobson et al., 2004). However, they show severe repression of genes involved in cytokinesis, micro-RNA biogenesis and poly(A)
tail length modulation, whereas mRNA levels of housekeeping genes,
hormone receptors and maternal factors are not significantly changed
(Wong et al., 2010). Secondly, the sister blastomeres display a high
grade of heterogeneity, partially because of the different timing of
major wave of EGA onset in each of them, and autonomous development (Hansis et al., 2004; Cauffman et al., 2005a; Edwards and Hansis,
2005; Wong et al., 2010).
In summary, the kinetics of mRNA degradation and EGA correlate
with a number of intriguing observations in the human. First, maternal
transcripts are largely degraded before the major onset of EGA at the
8-cell stage. Maternal mRNA degradation and EGA alter the cytoplasmic
content of the blastomeres during the first cleavage divisions and this may
have an effect on the totipotency of the blastomeres. Secondly, before
the 8-cell stage the embryo does not display any of the nuclear transcription factors that have been associated with the undifferentiated state.
Finally, lineage segregation does not start immediately after the first
major wave of EGA.
Cell cycle characteristics
Besides multiple signaling cascades, also the cell cycle (in general 24 h)
plays a role in the undifferentiated and differentiated states. The cell
cycle (G1, S, G2 and M phases) is regulated by cyclins and cyclindependent kinases (CDKs; Fig. 3). Cyclins are synthesized at specific
stages of the cell cycle whereas CDKs are constitutively expressed.
607
Various cyclin/CDK complexes determine progression through the distinct phases.
Interestingly, a short cell cycle (12 h) has been observed in all the
pluripotent cell types, including ESC and early stages of embryo development (Chisholm, 1988; Savatier et al., 1994; Becker et al., 2010; Wong
et al., 2010). In fact, a higher capacity of obtaining the pluripotent state
from somatic cells has been associated with an ultrafast cell cycle (8 h
versus normal 24 h; Guo et al., 2014).
During the first cell cycles (cleavage divisions), the timing of the G1
phase is significantly reduced in mouse embryos (Smith and Johnson,
1986; Chisholm, 1988; Moore et al., 1996). At the third cell cycle, the
G1 phase takes only 1 h compared with 11 h of the normal cell cycle
(Smith and Johnson, 1986). The heterogeneity in the cell cycle duration
starts from the third and increases already by the fourth cell cycle (Smith
and Johnson, 1986). Even during the fifth cell cycle of the cleaving mouse
embryo the length of the G1 and G2 phases is about four to five times less
than in normal somatic cells (Chisholm, 1988). The cell cycle duration of
the rodent pluripotent EPI cells is significantly reduced up to 3–9 h also
due to dramatic reduction of G1 and G2 phases, and even in some cases
the S phase as well (Mac Auley et al., 1993; Stead et al., 2002). Based on
relative RNA expression analysis in mouse oocytes and 1- to 2-cell stage
embryos, it has been shown that G1 phase-specific cyclins D1, E, CDK2
and p21 (or CDK inhibitor 1) are significantly up-regulated during the
second mitotic cell cycle which correlates with the onset of EGA
(Moore et al., 1996). The length of the first mitotic division (1- to
2-cell stage) in mouse embryos, which is controlled by maternal
factors, is associated with the capacity of the embryo to develop into
blastocyst (Balbach et al., 2012). EGA occurs at the late 2-cell stage
and coincides with a longer second cell cycle (Schultz, 2002). Maternal
genes are overrepresented in 8-cell stage mouse embryos cleaving
slowly to the 3-cell stage (first and second cycle); indicating that the degradation of maternal transcripts is delayed or their stability is increased.
On the other hand, embryonic genes are overrepresented in fast cleaving
embryos pointing out that they more efficiently proceed through EGA.
Similarly, human embryos display short cell cycles during the early
cleavage divisions (Wong et al., 2010). A number of cell cycle drivers, including G1 phase-specific factors, are intensively activated after EGA but
the check point proteins are lacking (Kiessling et al., 2010). Owing to the
absence of appropriate DNA quality control at early cleavage stages,
human embryos do not clear of the cells with aneuploidy, chromosome
breakage or segmental aberrations. Consequently, they take part further
in development and this may cause the genetic mosaicism that is
observed until Day 4 of human preimplantation development (Vanneste
et al., 2009; Mertzanidou et al., 2013a, b).
In the human, proper cell cycle progression during the early cleavage
stages determines the success of blastocyst formation as well. The combination of time lapse and gene expression studies on human IVF
embryos showed a correlation between the character of the cell divisions
and the transcription profile at the early cleavage stage (Wong et al.,
2010). Three cell cycle parameters have been found to predict blastocyst
development: (i) the duration of the first cytokinesis, (ii) the time interval
between the 2- and 3-cell stage and (iii) the time interval between the
second and the third mitoses (i.e. synchronicity third and fourth blastomeres). Precise cell cycle progression to the 4-cell stage allows predicting
embryo quality already on Day 2 before the major wave of EGA. Moreover, the data indicate that embryo quality is determined before EGA.
Interestingly, embryos arrested at the 2-cell stage expose a decrease in
608
De Paepe et al.
Figure 3 Cell cycle in somatic and embryonic (stem) cells. The cell cycle of a classical proliferating cell takes 24 h. It consists of G1 phase (11 h), S phase
(8 h), G2 phase (4 h) and mitosis (M; 1 h). The cell cycle of embryonic cells is shorter (15 – 16 h) due to the truncated G1 phase. This unique characteristic of
embryonic cells, including stem cells, allows them to proliferate rapidly and avoid easy onset to differentiation (red cross). The well-known critical regulators
of the cell cycle are CDKs and their binding partners cyclin proteins. Cyclin/CDK complex formation induces activation of CDK and consequently cell cycle
progression. Inhibitors of CDKs (CKIs) prevent formation of this complex, which prevents transition from one phase to another and, therefore, inhibits the
cell cycle. There are several different Cyclin and CDK proteins, each of them specific for the transition of the certain cell cycle phases. Complete lack of most
of the Cyclins D has been proposed to be necessary for the truncated G1 phase and, therefore, shortened cell cycle in hESC (dashed red cross).
expression of a number of cytokine genes, while those arrested at the
4-cell stage have down-regulated only a few of them. This observation
supports the role of the cell cycle and the importance of properly organized cleavages before the 4-cell stage in successful blastocyst formation.
Finally, the precise progression to the 4-cell stage, in particular the
second and the third parameter, is associated with the absence of
mitotic errors (aneuploidy) at this stage (Chavez et al., 2012). Interestingly, fragments which arise during cell division may contain micronuclei with
chromosomes. These fragments may be reabsorbed by the originating
cells or by neighboring cells explaining the high grade of aneuploidy in
fragmented embryos. Scoring cell cycle parameters in combination
with fragmentation may become the revolutionary approach needed
to select the best embryo for transfer in IVF clinics.
Short cell cycles have also been observed in mESC (Savatier et al.,
1994). This is correlated with the absence of certain critical regulators
of cytokinesis such as cyclin D and retinoblastoma proteins (Rb, prevents
progression from G1 into S; Savatier et al., 1994, 1996). Moreover,
CDK2, cyclin A and E complexes are very active in mESC and lack cell
cycle-dependent periodicity (Stead et al., 2002). Inhibiting CDK2 activity
with a specific inhibitor causes the cell cycle to slow down but this is not
associated with any specific cell cycle phase.
In their turn, hESC expose a shortened cell cycle as well (15– 16 h)
because of a truncated G1 phase (3–4 h) and the lack of a G0 phase
(Becker et al., 2006, 2010). The majority of hESC are kept in the
S phase. Their cell cycle is changed to normal once hESC are committed
to differentiate (Neganova et al., 2009; Becker et al., 2010; Calder et al.,
2013). Therefore, the unique cell cycle characteristics of hESC are connected to their pluripotent capacity. A lot of data exist on the expression
and function of key regulators of the cell cycle in hESC. However, most of
them are contradictory and this variation is most likely caused by culture
Totipotency and lineage segregation in the human embryo
conditions and experimental design (Barta et al., 2013). All groups agree
on the presence of high levels of CDKs (CDK2, CDK4 and CDK6) and
low or almost undetectable levels of CDK inhibitors (CKIs: p16, p18,
p19, p20, p21, p27 and p57; Egozi et al., 2007; Neganova et al., 2009;
Sengupta et al., 2009; Zhang et al., 2009b; Bárta et al., 2010; Dolezalova
et al., 2012). CDKs are necessary for intensive proliferation and CKIs are
normally up-regulated in differentiating stem cells assisting the exit from
the cell cycle (Dolezalova et al., 2012). Cyclins A and B are highly
expressed in hESC. High levels of cyclin D2 and its partner CDK4 have
been observed suggesting that those are important for G1 phase shortening, while others cyclins D1, D3 and E1 are lacking or almost undetectable in hESC (Becker et al., 2006, 2010). Nevertheless, the complete lack
of cyclin D proteins in combination with the constitutive expression of
cyclin E1 has also been reported (Filipczyk et al., 2007). These data
recall the situation in mESC where truncation of the G1 phase is achieved
by constitutively high levels of CDK2/cyclin E1 allowing direct transition
from M to late G1 phases (Becker et al., 2010). This explains the complete lack of most of the cyclins D necessary for the truncated G1
phase (Burdon et al., 2002).
There is an association between genes sustaining the undifferentiated
state and cell cycle components in hESC. NANOG directly activates
transcription of CDK6 and CDC25A and therefore stimulates the beginning of the S-phase (Zhang et al., 2009b). Down-regulation of POU5F1
results in a decrease of cyclins and increase in CKI p21 in human mesenchymal stem cells and hESC, while up-regulation of POU5F1 correlates
with an increase in specific CDK4 and CDC25A (Greco et al., 2007;
Lee et al., 2010). Increased cell cycle duration, in particular the G1
phase, has been associated with the onset of neural differentiation in
mESC and hESC (Lange et al., 2009; Borghese et al., 2010), e.g. CKI
p21 directly inhibits SOX2 expression in neural stem cells (MarquésTorrejón et al., 2013). POU5F1 and SOX2 together regulate the G1
phase-specific low expression of cyclin D1 (Card et al., 2008). Similar
findings have been reported in mESC (White and Dalton, 2005).
In summary, proper cell cycle progression in early cleavage stages
determines the success of human blastocyst formation. Undifferentiated
cells display short cell cycles. There is a remarkable relationship between
genes required for the undifferentiated state and cell cycle components in
hESC. It would be interesting to know whether human blastomeres, ICM
cells and hESC lines possess similar cell cycle properties.
Epigenetic modifications
The identity of a cell is determined by its epigenetic state. Epigenetic
changes such as DNA methylation and histone modifications influence
the expression of specific genes without altering the DNA sequence
(Supplementary data, File 2). The zygote, which results from the
fusion of two highly differentiated gametes, is characterized by a
unique epigenetic state. The parental genomes have to be reset into a
totipotent state after fertilization. During preimplantation development
extensive epigenetic remodeling results in lineage segregation.
Drastic epigenetic remodeling occurs during both gametogenesis and
embryogenesis. For instance, during gametogenesis all DNA methylation
marks are erased and re-established and imprinted genes are set in a sexspecific way (pre-zygotic reprogramming). After fertilization, there is a
second global wave of DNA demethylation, except for imprinted
genes (post-zygotic reprogramming; Jaenisch, 2004; Perera and Herbstman, 2011). The paternal genome, which is packaged by protamines
609
instead of histones, is actively demethylated by factors in the cytoplasm
of the oocyte. The maternal genome, on the other hand, is more resistant
to the demethylating activity because it has a distinct chromatin configuration and is passively demethylated during cleavage divisions. Preimplantation stages are characterized by DNA hypomethylation. A global wave
of DNA remethylation is observed at the time of implantation.
Epigenetic events have been thoroughly investigated in mice (Supplementary data, File 3). Very few data on epigenetic remodeling have been
reported in the human embryo. The global level of DNA methylation has
been investigated in human preimplantation embryos obtained from in
vitro matured oocytes. Some zygotes have pronuclei with a similar methylation state, whereas in other zygotes a hypomethylated paternal pronucleus is observed (Fulka et al., 2004). The nature of this asymmetric
pattern has not been investigated yet. Human embryos progressively
obtain a weaker DNA methylation pattern during the cleavage stages
which might be associated with the potency of the blastomeres. A
global wave of remethylation occurs around the time of implantation
where the TE cells shows higher DNA methylation levels than the ICM
cells (Fulka et al., 2004). These data suggest that there is an epigenetic
asymmetry between the two lineages, ICM and TE, in the human.
DNA methylation is orchestrated by DNA methyltransferases that
methylate cytosine residues of CpG dinucleotides. The maintenance
DNA methyltransferase (DNMT1) and the de novo DNA methyltransferases (DNMT3a and DNMT3b) are expressed at almost all stages of
human preimplantation development (Huntriss et al., 2004). Transcripts
of CpG-binding protein (CGBP) protecting unmethylated CpG sites
from methylation were detected in ICM but not in TE cells of human blastocysts (Huntriss et al., 2004), which is consistent with the asymmetric
DNA methylation pattern (Fulka et al., 2004).
X chromosome inactivation (XCI) may represent another example of
asymmetry between ICM and TE cells. XCI occurs in female cells to
obtain a similar dosage of X-linked gene expression levels as in male
cells and is initiated by the transcription of the long non-coding XIST
RNA from the X chromosome that will subsequently be inactivated. It
has been reported that female human 8-cell stage embryos display pinpoints of XIST RNA on one of the two X chromosomes (van den Berg
et al., 2009). Further in development XIST RNA is expressed in TE and
PE cells from one of the X chromosomes and accumulates in a cloud
on another X chromosome indicating XCI initiation. Unfortunately,
the status of the ICM cells was not reported in this study, while asymmetric XCI was described in mouse embryos (Okamoto and Heard, 2006).
Contradictory results have been reported showing that in human blastocysts XIST is expressed on both X chromosomes that remain transcriptionally active, despite XIST expression, until the time of implantation
(Okamoto et al., 2011). These opposing results may be due to technical
complications and/or distinct embryo culture conditions.
The XCI state of hESC is highly variable. HESC are believed to display
two active X chromosomes upon derivation, but often initiate XCI
due to suboptimal culture conditions (Silva et al., 2008; Lengner
et al., 2010). Female mESC lines display two active X chromosomes
at the undifferentiated state, but randomly inactivate one of the X chromosomes upon differentiation (van den Berg et al., 2011). HESC,
however, maintain their pluripotent phenotype even in the presence
of completed XCI (Bruck and Benvenisty, 2011). This supports the hypothesis that hESC resemble mEpiSCs that also display XCI and represent a later developmental stage than naı̈ve mESC. Whereas in
female human somatic cells a random XCI pattern is usually observed
610
(Moreira de Mello et al., 2010), the XCI pattern in TE derivatives is under
discussion. A study using hESC differentiating toward the trophoblast
lineage suggested that XCI in human trophoblast cells, as in mouse, is
under imprinting control (Dhara and Benvenisty, 2004). Studies analyzing
extra-embryonic tissues and term placenta, however, reported that
random XCI was common in these tissues, which may indicate that, in
contrast to the mouse, XCI may not be under imprinting control in
normal human placentation (Zeng and Yankowitz, 2003; Moreira de
Mello et al., 2010).
Although global DNA methylation is important for the regulation of
gene expression, other mechanisms are known to control gene expression as well, e.g. histone modifications. Histone acetylation is generally
associated with transcriptional activity, whereas histone lysine methylation can be either activating (e.g. H3K4me3) or repressive (H3K27me3).
Genes associated with H3K4 and H3K27 trimethylation have been investigated in hESC (Zhao et al., 2007b). Three distinct categories of genes
are identified: genes containing only H3K4me3 that are mainly involved
in self-renewal; bivalent genes containing both H3K4me3 (active mark)
and H3K27me3 (repressive mark) involved in early development; and
genes containing neither modification involved, e.g. in immunological
events. The so-called bivalent histone methylation poises transcription
and enables cells to flexibly modulate developmental gene expression
in response to different environmental factors.
The sequence of the promoter can contribute to the epigenetic mechanism that affects its regulation. In general, H3K27me3 and DNA methylation are both inversely correlated with gene expression. How they
regulate pluripotency and differentiation is not fully understood. In
hESC, the promoters for genes expressed late in embryo development
(e.g. mesenchymal stem cells) are often CG poor and they mainly
employ DNA methylation upon silencing, suggesting that DNA methylation may be required for gene repression in terminally differentiated cells
(Xie et al., 2013). On the other hand, the promoters of genes active in
early embryo development (ESC, trophoblast and mesendoderm) are
CG rich but they mostly engage H3K27me3 upon silencing (e.g. SOX2,
Eomes, SOX17). Exceptionally, the promoter of POU5F1 and
NANOG employ DNA methylation upon silencing in differentiated
cells. The early embryo developmental genes remain largely unmethylated upon differentiation. In fact, early embryo developmental genes
(transcription factors, signaling pathways, homeobox) are often
located in large genomic domains. These domains are devoid of DNA
methylation and, therefore, called DNA methylation valleys (DMVs).
Many DMV genes show a bivalent state of H3K4me3 and H3K27me3,
but they mostly become monovalent in differentiated cells. The flexibility
fits well with the nature of early embryonic cells and stem cells keeping
developmental genes poised for activation during early differentiation.
These events have not been investigated during lineage segregation in
the human embryo.
Interestingly, although the contribution of the spermatozoon to
embryo development is thought to be limited since histones are replaced
by protamines, a small fraction of histone-bound DNA has been found in
human sperm (Hammoud et al., 2009). In particular, H3K4me3 is
enriched at promoters of, e.g. paternally expressed imprinted loci and
certain transcription and signaling factors involved in development. On
the other hand, H3K27me3 is enriched at developmental promoters
that are repressed in early embryos, including many bivalent
H3K4me3/H3K27me3 promoters described in hESC. These data indicate that in sperm certain loci are poised for development depending
De Paepe et al.
on their histone packaging. In general, developmental promoters are
DNA hypomethylated in sperm but they acquire methylation during
differentiation.
There are points to take into consideration when studying epigenetic
mechanisms:
(i) It is important to distinguish between patterns crucial for lineage
segregation (i.e. the preimplantation period) and the defects that can
occur during preimplantation without implications for the initial differentiation of the embryonic cells but with consequences for the period after
implantation and eventually later during life. Epigenetic imprinting disorders such as Beckwith–Wiedemann disease (Maher et al., 2003) and
Angelman syndrome (Cox et al., 2002) have been associated with
ART, but the origin of these diseases is unknown.
(ii) Epigenetic changes may be induced by the study procedure itself,
e.g. by manipulation (Torres-Padilla, 2008). Extrinsic factors such as
serum (Thompson et al., 1995) and light (Schultz, 2007) have been
shown to have an effect on development through epigenetic changes.
Consequently, developmental abnormalities may be introduced by
assisted reproductive technologies such as hormonal stimulation, manipulation, cryopreservation, cloning (Li et al., 2005; Gebert et al.,
2009; Thaler et al., 2012) and associated cell culture conditions (Dumoulin et al., 2010; Nelissen et al., 2012; McEwen et al., 2013). For example,
vitrification has been shown to have an effect on the promoter methylation of Pou5f1, Nanog and Cdx2 in mouse blastocysts (Zhao et al., 2012).
This also applies to hESC, which undergo epigenetic changes upon differentiation toward different cell types, and additionally display epigenetic
variation that may be inherent to the genotype or obtained after
culture adaptation (Bibikova et al., 2006; Allegrucci et al., 2007; Doi
et al., 2009; Lund et al., 2012; McEwen et al., 2013; Nguyen et al.,
2013), the latter being stably passed on to the differentiated cells
(Nazor et al., 2012).
In summary, there are very few data reported on epigenetic modifications observed during human preimplantation development. Human
preimplantation embryos are characterized by a global DNA hypomethylation state which might be associated with the potency of the
cells. There seems to be an epigenetic asymmetry between ICM and
TE lineages, but the nature of this event is largely unknown. In the
human embryo there are currently no reports on histone modifications
and DNA methylation patterns playing a role in lineage segregation as
described in mice. However, hESC have been studied thoroughly and
the nature of the cells has been associated with bivalent histone methylation marks enabling the cells to modulate developmental gene expression in response to the environment.
Somatic cell nuclear transfer
Oocyte quality plays a major role in reproduction, it is a prerequisite for live
birth and thus for totipotency. It is reflected by genetic constitution on the
one hand and cytoplasmic content supporting EGA on the other hand. The
cloning efficiency is largely dependent upon the reprogramming capacity of
the cytoplasm. Both the oocyte and the zygote can be used for SCNT
(Lorthongpanich et al., 2010). The zygote is totipotent, but the ooplasm
of a mature oocyte contains all the factors necessary to reprogram the
DNA (two pronuclei after normal fertilization or somatic cell DNA
after SCNT) into a totipotent state and support embryo development
(post-zygotic reprogramming). Differences between the oocyte and the
zygote may play a role in SCNT experiments.
Totipotency and lineage segregation in the human embryo
In the mouse model, therapeutic cloning (SCNT-ESC derivation for
transplantation purposes) has a much higher efficiency than reproductive
cloning (live birth; Supplementary data, File 4). In humans, only therapeutic cloning is allowed. Data are limited because of ethical and legal
restrictions and the scarcity of human oocytes/zygotes. To reprogram
adult human cells by SCNT for patient-specific hESC derivation mostly
enucleated mature oocytes are used. These enucleated oocytes are
artificially activated (e.g. by electropulse and/or ionomycin; Tachibana
et al., 2009, 2013), which in normal fertilization is done by the sperm
cell completing meiosis and initiating mitosis. Transcriptome comparison
between human mature and fertilized oocytes revealed that their mRNA
content is similar (Dobson et al., 2004). There is already some up- and
down-regulation of transcripts in the zygote, but the implications of
these changes for SCNT experiments in the human are unknown. In
IVF/ICSI embryos, the major wave of EGA occurs at the 8-cell stage
(Braude et al., 1988; Vassena et al., 2011). Human SCNT embryos
often arrest at the 4- to 8-cell stage, suggesting that the somatic donor
cell nucleus cannot activate embryonic genes crucial for further development (Noggle et al., 2011). However, this arrest is not observed using
611
caffeine during the procedure (Tachibana et al., 2013). Caffeine, a phosphatase inhibitor, is used to avoid premature activation of the cytoplasm
which may be induced by the micromanipulation and results in poor
embryo development. Moreover, oocytes were stained with Hoechst
and subjected to UV irradiation to remove the DNA which can additionally damage oocyte quality (in particular, the mitochondria) and resolve
into the early developmental arrest of the cloned embryos. The developmental arrest has also been observed in a study using human zygotes for
SCNT (Egli et al., 2011). In this study, the mitotic spindle was removed
after nuclear envelope breakdown in the first mitosis; however, the
arrest may also have been due to other technical differences.
In rhesus monkey SCNT-ESC lines have been obtained (Byrne et al.,
2007), but the live birth of cloned animals has not yet been reported.
Compared with normal embryos, the ICM cells of cloned embryos maintain a high level of DNA methylation and this may disturb normal embryo
development after SCNT (Yang et al., 2007).
IPSC represent another model to study the reprogramming of somatic
cells. Interestingly, a privileged somatic cell state for obtaining pluripotent
capacity has been associated with an ultrafast cell cycle (Guo et al., 2014).
Figure 4 General overview of the gradual loss in potency and lineage segregation in the human embryo. From experiments with isolated and reaggregated
blastomeres it is clear that the early blastomeres are not yet committed toward ICM or TE. The cells that descend from the totipotent zygote gradually lose
totipotency and finally develop into one of the three lineages (TE, EPI or PE) in the blastocyst. The segregation of cell lineages does not occur immediately
after EGA. When and how the cells get committed is linked to cell cycle features and epigenetic modifications that generate distinct transcriptional programs.
612
However, we do not discuss iPSC in this paper because they are pluripotent. The data on iPSC have been reviewed recently (Liang and Zhang,
2013).
In summary, mature and fertilized human oocytes have a similar
mRNA content. However, it is not clear whether their cytoplasm have
similar capacities to reprogram the DNA of a human somatic cell.
Cloned human embryos and hESC are useful to study EGA, lineage
differentiation, cell cycle progression and epigenetic phenomena.
De Paepe et al.
Funding
Our research is supported by grants from the Scientific Research Foundation—Flanders (FWO-Vlaanderen) and the Research Council (OZR)
of the VUB.
Conflict of interest
None declared.
Conclusion
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Information on totipotency and cell fate during embryogenesis is mainly
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